We are IntechOpen, the world's leading publisher of Open Access books Built by scientists, for scientists

4,200+

Open access books available

116,000+

International authors and editors

125M+

Downloads

151 Countries delivered to Our authors are among the

Top 1%

most cited scientists

12.2% Contributors from top 500 universities

Selection of our books indexed in the Book Citation Index in Web of Science™ Core Collection (BKCI)

## Interested in publishing with us? Contact book.department@intechopen.com

Numbers displayed above are based on latest data collected. For more information visit www.intechopen.com

## **Meet the editors**

Rolando Chamy is a Biochemical Engineer and Ph.D. in Chemical Engineering from the University of Santiago de Compostela, Spain. He holds a professor chair at the Pontificia Universidad Católica de Valparaíso, and is the Director of the Biotechnology Center of Curauma, a center of applied research, which belongs to the same university. Currently, he also leads the research line

in Renewable Bioresource at Fraunhofer Chile Research. His main lines of research are environmental biotechnology, renewable energy, climate change, and innovation management.

Additionally, he has developed an extensive body of work in the field of applied research, with more than 100 as project director, principal investigator or co-investigator, supported with more than 100 ISI publications and 19 contributions to book chapters.

Francisca Rosenkranz, Biochemical Civil Engineer (Pontificia Universidad Católica de Valparaíso) and PhD in Biotechnology (Pontificia Universidad Católica de Valparaíso and Universidad Técnica Federico Santa María) with the co-supervision of the University of Santiago de Compostela, Spain. Her specialization areas include biogas production, anaerobic treatment of hard biodegradable compounds and studies of microbial communities in anaerobic systems. Her actual positions are at Fraunhofer Chile Research, as research project coordinator in the bioenergy area, and at Biotechnology Center of Curauma, as Chief of Operations.

Contents

**Preface VII**

Nina Djapic

**by Tsunami 39**

**Coconut Fiber 105** Yasko Kodama

**Degradation 129**

Wojcieszyńska

Chapter 2 **Chlorophyll Biodegradation 17**

Chapter 3 **Bioremediation of Agricultural Land Damaged**

M. Azizul Moqsud and K. Omine

Yu-Huei Peng and Yang-hsin Shih

Anita Rani Santal and Nater Pal Singh

Chapter 1 **Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil 1**

Chapter 4 **Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features 51**

Chapter 5 **Biodegradation of Melanoidin from Distillery Effluent: Role of**

Chapter 6 **Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with**

Chapter 7 **Intradiol Dioxygenases — The Key Enzymes in Xenobiotics**

Urszula Guzik, Katarzyna Hupert-Kocurek and Danuta

**Microbes and Their Potential Enzymes 71**

L. B. Yin, L. Z. Zhao, Y. Liu, D. Y. Zhang, S. B. Zhang and K. Xiao

### Contents

#### **Preface XI**


Wojcieszyńska

Chapter 8 **Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils 155** Ildefonso Díaz-Ramírez, Erika Escalante-Espinosa, Randy Adams Schroeder , Reyna Fócil-Monterrubio and Hugo Ramírez-Saad

Preface

damaged soils.

This book contains a collection of different research activities that include the biodegrada‐ tion compounds with contaminant characteristics and special products of different interests

The chapters consider the degradation of contaminant compounds generated by industrial activities, i.e., oil industry by-product compounds and halogen compounds or compound generated by natural phenomena such as tsunamis, which require interventions to recover

In addition, the book contains chapters that involve special product degradation processes such as chlorophyll, which corresponds to a biological process indicator as photosynthesis.

**Rolando Chamy and Francisca Rosenkranz**

Pontificia Universidad Católica de > Valparaíso, Chilr

Pontificia Universidad Católica de > Valparaíso, Chile

Biotechnology Center of Curauma

School of Biochemical Engineering

as an added value product or that allows following up various biological processes.

### Preface

Chapter 8 **Hydrocarbon Biodegradation Potential of Native and**

**VI** Contents

**Exogenous Microbial Inocula in Mexican Tropical Soils 155** Ildefonso Díaz-Ramírez, Erika Escalante-Espinosa, Randy Adams Schroeder , Reyna Fócil-Monterrubio and Hugo Ramírez-Saad

> This book contains a collection of different research activities that include the biodegrada‐ tion compounds with contaminant characteristics and special products of different interests as an added value product or that allows following up various biological processes.

> The chapters consider the degradation of contaminant compounds generated by industrial activities, i.e., oil industry by-product compounds and halogen compounds or compound generated by natural phenomena such as tsunamis, which require interventions to recover damaged soils.

> In addition, the book contains chapters that involve special product degradation processes such as chlorophyll, which corresponds to a biological process indicator as photosynthesis.

> > **Rolando Chamy and Francisca Rosenkranz** Biotechnology Center of Curauma Pontificia Universidad Católica de > Valparaíso, Chilr

School of Biochemical Engineering Pontificia Universidad Católica de > Valparaíso, Chile

**Chapter 1**

**Isolation and Characterization**

L. B. Yin, L. Z. Zhao, Y. Liu, D. Y. Zhang,

Additional information is available at the end of the chapter

agricultural and urban settings around the world [5].

sensitive to the toxic effects of SPs [12-15].

A current environmental concern is the contamination of aquatic ecosystem due to pesticide discharges from manufacturing plant, agricultural runoff, leaching, accidental spills and other sources [1, 2]. Synthetic pyrethroid insecticides were introduced into widespread use for the control of insect pests and disease vectors more than three decades ago. In addition to their value in controlling agricultural pests, pyrethroids are at the forefront of efforts to combat malaria and other mosquito-borne diseases [3] and are also common ingredients of household insecticide and companion animal ectoparasite control products [4]. Cypermethrin is a type of synthetic pyrethroids (SPs), a class of pesticides widely used for insect control in both

The use of SPs in China has increased sharply since many organophosphate products, such as methamidophos and parathion, are being phased out for agricultural use. With such extensive application, many adverse effects, such as pest resistance, residues in foods, and environmen‐ tal contamination are public safety concerns [6, 7]. Although SPs are widely considered safe for humans, numerous studies have shown that exposure to very high concentrations of SPs might cause human health problems [8]. Such effects include bioaccumulation toxicity; immune suppression, endocrine disruption; modify electrical activity in various parts of the nervous system, neurotoxicity, lymph node and splenic damage, and carcinogenesis [9-11]. In addition, bees, fish, crabs, tadpoles, arthropods, and other non-target organisms are extremely

> © 2013 Yin et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

S. B. Zhang and K. Xiao

http://dx.doi.org/10.5772/56339

**1. Introduction**

**of Cypermethrin Degrading Bacteria**

**Screened from Contaminated Soil**

**Chapter 1**

## **Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil**

L. B. Yin, L. Z. Zhao, Y. Liu, D. Y. Zhang,

S. B. Zhang and K. Xiao

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56339

#### **1. Introduction**

A current environmental concern is the contamination of aquatic ecosystem due to pesticide discharges from manufacturing plant, agricultural runoff, leaching, accidental spills and other sources [1, 2]. Synthetic pyrethroid insecticides were introduced into widespread use for the control of insect pests and disease vectors more than three decades ago. In addition to their value in controlling agricultural pests, pyrethroids are at the forefront of efforts to combat malaria and other mosquito-borne diseases [3] and are also common ingredients of household insecticide and companion animal ectoparasite control products [4]. Cypermethrin is a type of synthetic pyrethroids (SPs), a class of pesticides widely used for insect control in both agricultural and urban settings around the world [5].

The use of SPs in China has increased sharply since many organophosphate products, such as methamidophos and parathion, are being phased out for agricultural use. With such extensive application, many adverse effects, such as pest resistance, residues in foods, and environmen‐ tal contamination are public safety concerns [6, 7]. Although SPs are widely considered safe for humans, numerous studies have shown that exposure to very high concentrations of SPs might cause human health problems [8]. Such effects include bioaccumulation toxicity; immune suppression, endocrine disruption; modify electrical activity in various parts of the nervous system, neurotoxicity, lymph node and splenic damage, and carcinogenesis [9-11]. In addition, bees, fish, crabs, tadpoles, arthropods, and other non-target organisms are extremely sensitive to the toxic effects of SPs [12-15].

© 2013 Yin et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Cypermethrin is more effective against pests including moth pests of cotton, fruits and vegetable crops. Extensive and improper use of this kind chemicals leads to greater health risk to plants, animals and human population which had been reviewed time to time by several researchers [16]. One of the major problems asides from toxicity and carcinogenicity of pesticides is their long persistence in nature that amplifies the toxicity and health risk problems in the area of contamination [17].

**2. Materials and methods**

Standard analytical grade sample of 100 μg/mL cypermethrin (99.8% purity) was purchased from the Agro-Environmental Protection Institute, Ministry of Agriculture (Tianjin, China). Acetonitrile, methanol and hexane were of chromatographic grade while other chemicals were of analytical grade. Cypermethrin dissolved in acetone solution was added to desirable concentration in medium as the sole carbon source. Mineral Salts Medium (MSM) (g/L): 1.0 NH4NO3, 1.0 NaCl, 1.5 K2HPO4, 0.5 KH2PO4, 0.2 MgSO4 7H2O, pH 7.0. For solid plate, 1.5% (w/ v) agar was added. Medium were sterilized by autoclaving at 121℃ for 30 min before use.

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

3

An activated sludge sample was collected from the wastewater treatment pool of a pesticide plant located in Changsha (Hunan, China), which had produced cypermetrin over 5 years. Wastewater sludge enrichment was performed by placing 10 g activated sludge in a 250 mL-Erlenmeyer flask containing 100 mL sterilized MSM media with an initial cypermethrin concentration at 20 mg/L, and incubated in a light incubator (PRX-450D, China) at 37℃ and 7500 lux; the flasks were shaken 3–5 times per day. After 10 days or so, the medium turned red-brown, a 5 mL aliquot of the culture was inoculated into 100 mL of fresh MSM medium containing 50 mg/L cypermetrin, and the new mixture was incubated for another 10 days under the same conditions. The medium was gradually acclimated to increasing concentrations of cypermetrin ranging from 50 to 200 mg/L at intervals of a week. After about 10 transfers, a mixed microbial population was diluted in series, and then streaked on MSM agar medium plate containing 100 mg/L cypermethrin. The dilution series was repeated at least 5 times, until single colony was achieved. The abilities of isolates to degrade cypermethrin were determined by gas chromatography (GC) according to Yin et al and Chen et al [23, 27]. The relatively higher degradation ability colonies were selected for further degrading studies. These organisms were stored long-term on porous beads in a cryopreservative fluid at -20℃ and short-term on agar

**2.3. Characterization and identification of the cypermethrin degrading isolates**

(JEM-6360, JEOL) and/or scanning electron microscope (JSM-6360LV, JEOL).

A cypermethrin degrading isolate designated as S10-1 showed the highest degradation rate was selected for further study. The purified S10-1 was identified on the basis of its morphological characteristics and results of biochemical tests and 16S rRNA gene sequence analysis. The isolate S10-1 was grown on MSM agar plates containing 50 mg/L cypermethrin at 37℃ and 7500 lux for 7 days, its cell morphology, method of reproduction, and the structure of its inner photosynthesis membrane and flagella were observed by transmission electron microscope

The isolate S10-1 was further confirmed by 16S rRNA gene sequence. The DNA was extracted and purified using the Qiagen genomic DNA buffer set. PCR amplification was performed as described by Mirnejad et al [28]. The 16S rRNA sequencing was performed by Beijing Liuhe

**2.2. Enrichment, isolation and screening of bacterial strains**

**2.1. Chemicals and media**

plates at 4℃

Therefore, it is necessary to develop a rapid and efficient disposal process to eliminate or minimize the concentrations of SPs in the environment. A variety of physical and chemical methods are available to treat the soils contaminated with hazardous materials but many of these physical and chemical treatments do not actually destroy the hazardous compounds but are bound in a modified matrix or transferred from one phase to another [18, 19], hence biological transforming is essential. The biological treatment of chemically contaminated soil involves the transformation of complex or simple chemical compounds into non-hazardous forms [20]. For biodegradation, ideally the target pesticide will be able to serve as the sole carbon source and energy for microorganisms, including the synthesis of appropriate enzymes if need able. The specificity of enzymes active against xenobiotic compounds differs from one microorganism to another.

In the light of this fact, biodegradation, especially microbial degrading, has proven to be a suitable method for insecticide elimination. Previous studies indicated that microbes play important roles in degrading and detoxifying SPs residues in the environment. Thus far, many reports have described the biodegradation of cypermethrin by various bacteria, including *Ochrobactrum lupini*, *Pseudomonas* aeruginosa, *Streptomyces aureus*, and *Serratia* spp.[21-23], but there is few research describing biodegradation of pesticides by *Rhodobacter sphaeroides*. Among the different genera of pesticide-degrading bacteria, the photosynthetic (PSB) genus *Rhodo‐ bacter* has a special status in the ecosystem, since its metabolic functions are extraordinarily versatile, including degradation of various organic compounds, nitrogen fixation, hydrogen production [24], as a biofertilizer for promoting plant growth and increasing grain yield [25], and 5-aminolevulinic acid production which has multiple functions including a relatively strong herbicidal effect in clover [26]; therefore, microbes belonging to this genus are ideal choice for degrading pesticide residues.

The research aim was to identify the potential microbial strain able to utilize cypermethrin from the contaminated soil. In this study, the pesticide degrading potential of a bacterial culture is examined with the hope of isolation and characterization of cypermethrin degrading potentials in the contaminated soil. In addition, the optimum dose and the suitable conditions for cypermethrin degradation using laboratory scale were also evaluated. The results of the present study suggest that the use of potential microorganisms in the treatment system can successfully overcome many of the disadvantages associated with the conventional method used for the degradation of inhibitory compound.

### **2. Materials and methods**

#### **2.1. Chemicals and media**

Cypermethrin is more effective against pests including moth pests of cotton, fruits and vegetable crops. Extensive and improper use of this kind chemicals leads to greater health risk to plants, animals and human population which had been reviewed time to time by several researchers [16]. One of the major problems asides from toxicity and carcinogenicity of pesticides is their long persistence in nature that amplifies the toxicity and health risk problems

Therefore, it is necessary to develop a rapid and efficient disposal process to eliminate or minimize the concentrations of SPs in the environment. A variety of physical and chemical methods are available to treat the soils contaminated with hazardous materials but many of these physical and chemical treatments do not actually destroy the hazardous compounds but are bound in a modified matrix or transferred from one phase to another [18, 19], hence biological transforming is essential. The biological treatment of chemically contaminated soil involves the transformation of complex or simple chemical compounds into non-hazardous forms [20]. For biodegradation, ideally the target pesticide will be able to serve as the sole carbon source and energy for microorganisms, including the synthesis of appropriate enzymes if need able. The specificity of enzymes active against xenobiotic compounds differs from one

In the light of this fact, biodegradation, especially microbial degrading, has proven to be a suitable method for insecticide elimination. Previous studies indicated that microbes play important roles in degrading and detoxifying SPs residues in the environment. Thus far, many reports have described the biodegradation of cypermethrin by various bacteria, including *Ochrobactrum lupini*, *Pseudomonas* aeruginosa, *Streptomyces aureus*, and *Serratia* spp.[21-23], but there is few research describing biodegradation of pesticides by *Rhodobacter sphaeroides*. Among the different genera of pesticide-degrading bacteria, the photosynthetic (PSB) genus *Rhodo‐ bacter* has a special status in the ecosystem, since its metabolic functions are extraordinarily versatile, including degradation of various organic compounds, nitrogen fixation, hydrogen production [24], as a biofertilizer for promoting plant growth and increasing grain yield [25], and 5-aminolevulinic acid production which has multiple functions including a relatively strong herbicidal effect in clover [26]; therefore, microbes belonging to this genus are ideal

The research aim was to identify the potential microbial strain able to utilize cypermethrin from the contaminated soil. In this study, the pesticide degrading potential of a bacterial culture is examined with the hope of isolation and characterization of cypermethrin degrading potentials in the contaminated soil. In addition, the optimum dose and the suitable conditions for cypermethrin degradation using laboratory scale were also evaluated. The results of the present study suggest that the use of potential microorganisms in the treatment system can successfully overcome many of the disadvantages associated with the conventional method

in the area of contamination [17].

2 Biodegradation of Hazardous and Special Products

microorganism to another.

choice for degrading pesticide residues.

used for the degradation of inhibitory compound.

Standard analytical grade sample of 100 μg/mL cypermethrin (99.8% purity) was purchased from the Agro-Environmental Protection Institute, Ministry of Agriculture (Tianjin, China). Acetonitrile, methanol and hexane were of chromatographic grade while other chemicals were of analytical grade. Cypermethrin dissolved in acetone solution was added to desirable concentration in medium as the sole carbon source. Mineral Salts Medium (MSM) (g/L): 1.0 NH4NO3, 1.0 NaCl, 1.5 K2HPO4, 0.5 KH2PO4, 0.2 MgSO4 7H2O, pH 7.0. For solid plate, 1.5% (w/ v) agar was added. Medium were sterilized by autoclaving at 121℃ for 30 min before use.

#### **2.2. Enrichment, isolation and screening of bacterial strains**

An activated sludge sample was collected from the wastewater treatment pool of a pesticide plant located in Changsha (Hunan, China), which had produced cypermetrin over 5 years. Wastewater sludge enrichment was performed by placing 10 g activated sludge in a 250 mL-Erlenmeyer flask containing 100 mL sterilized MSM media with an initial cypermethrin concentration at 20 mg/L, and incubated in a light incubator (PRX-450D, China) at 37℃ and 7500 lux; the flasks were shaken 3–5 times per day. After 10 days or so, the medium turned red-brown, a 5 mL aliquot of the culture was inoculated into 100 mL of fresh MSM medium containing 50 mg/L cypermetrin, and the new mixture was incubated for another 10 days under the same conditions. The medium was gradually acclimated to increasing concentrations of cypermetrin ranging from 50 to 200 mg/L at intervals of a week. After about 10 transfers, a mixed microbial population was diluted in series, and then streaked on MSM agar medium plate containing 100 mg/L cypermethrin. The dilution series was repeated at least 5 times, until single colony was achieved. The abilities of isolates to degrade cypermethrin were determined by gas chromatography (GC) according to Yin et al and Chen et al [23, 27]. The relatively higher degradation ability colonies were selected for further degrading studies. These organisms were stored long-term on porous beads in a cryopreservative fluid at -20℃ and short-term on agar plates at 4℃

#### **2.3. Characterization and identification of the cypermethrin degrading isolates**

A cypermethrin degrading isolate designated as S10-1 showed the highest degradation rate was selected for further study. The purified S10-1 was identified on the basis of its morphological characteristics and results of biochemical tests and 16S rRNA gene sequence analysis. The isolate S10-1 was grown on MSM agar plates containing 50 mg/L cypermethrin at 37℃ and 7500 lux for 7 days, its cell morphology, method of reproduction, and the structure of its inner photosynthesis membrane and flagella were observed by transmission electron microscope (JEM-6360, JEOL) and/or scanning electron microscope (JSM-6360LV, JEOL).

The isolate S10-1 was further confirmed by 16S rRNA gene sequence. The DNA was extracted and purified using the Qiagen genomic DNA buffer set. PCR amplification was performed as described by Mirnejad et al [28]. The 16S rRNA sequencing was performed by Beijing Liuhe Huada Genomic Company (Beijing, China). The sequences with the highest 16S rDNA partial sequence similarity were selected and compared by CLUSTAL W. Phylogenetic and molecular evolutionary analyses were conducted by MEGA 4.0 software with the Kimura 2-paremeter model and the neighbor joining algorithm [29]. Confidence estimates of branching order were determined by bootstrap resampling analysis with 1000 replicates.

HP-5 5% phenyl methyl siloxane capillary column (30 m × 320 μm × 0.25 μm; Agilent Tech‐ nologies, USA) was used for separation, with helium as the carrier gas (flow rate, 1 mL/min). Other GC parameters included an inlet temperature of 250℃ and a detector temperature of 300℃; initially, the oven temperature was 150℃ for 2.0 min, was ramped to 280℃ at 15℃/min, and then maintained at 280℃ for 5.0 min. The injection volume was 1.0 μL. Samples were introduced in split-less mode. Concentrations were determined by analyzing peak area with

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

5

Metabolites were isolated from the culture filtrates of the organism grown in cypermethrin (100 mg/L) by extraction with acetonitrile, before and after acidification to pH 2 with 2 M HCl, and the residue obtained was dissolved in hexane [22]. The metabolites were identified and analyzed using the GC/MS system (Agilent 7890A/5975, Agilent Technologies, USA) equipped with electron ionization (EI). EI (70 eV) was performed with a trap current of 100 mA and a source temperature of 200℃. Full scan spectra were acquired at m/z 45–500 at 2 sec per scan. The metabolites were confirmed by standard MS, data collection and processing were performed using Agilent MSD ChemStation software containing the Agilent chemical library.

After repeated enrichment and purification processes, we obtained approximately 20 strains of organisms with different colony morphologies from the activated sludge samples. But the degradation experiments showed the isolate S10-1 possessed the relatively higher degradation, capacity of degrading cypermethrin (100 mg/L) by 90.4% after incubating 7 days at pH 7.0 and temperature 35℃ (Fig. 3a). And S10-1 utilized cypermethrin as its sole carbon and energy source

S10-1 is a gram-negative, anaerobic bacterium. The morphology of the S10-1 colonies, cul‐ tured for 10 days on MSM agar plate, were reddish-brown, smooth, circular, wet, nontrans‐ parent, glistening, and with entire margins (Figure 1a). The physiological and biochemical characteristics of S10-1 are shown in Table 1. SEM observations showed that the cells are ovoid to rod shaped (Figure 1b), sometimes even longer, measuring about 0.5–0.9 μm in width and 1.2–2.0 μm in length, and are motile by means of polar flagella (Figure 1c). Internal photosynthetic membranes appear as lamellae underlying and parallel to the cytoplasmic membrane (Figure 1d). The culture suspension was reddish-brown in color. In vivo absorption maxima of intact cells (Figure 1e) were recorded at 378, 455, 480, 510, 592, 806, and 865 nm, indicating the presence of bacteriochlorophyll a and carotenoids of the spheroidene series [31]. These morphological and biochemical properties are identical to

**3.1. Isolation and characterization of cypermethrin degrading bacterium**

in MSM. Thus strain S10-1 was selected for further detail investigation.

an authentic cypermethrin standard.

**3. Results and discussion**

the genus *Rhodobacter* [31].

**2.7. Detection of cypermethrin metabolites**

#### **2.4. Inoculum preparation**

Unless otherwise stated, the inoculants for this experiment were bacteria cultured in a 130 mL serum bottle containing 120 mL of PSB medium in a light incubator at 35℃ and 7500 lux. At the exponential phase (about 2–3 days), the cell pellets were harvested via centrifugation (5000×g, 10 min), washed 3 times with 50 mL of KH2PO4-K2HPO4 (0.15 mol/L, pH 7.0), and then suspended in the same phosphate buffer as the inoculants. In order to avoid the effects of hydrolysis and photolysis, each treatment was set in triplicate with non-inoculated samples as control under the same conditions and analyzed in the same manner. Samples for residual pesticide concentration analysis were collected from the cultures at regular intervals.

#### **2.5. Optimal conditions for degrading cypermethrin by S10-1**

To determine the optimal conditions for degrading cypermethrin by S10-1, single-factor test was designed in this study under different conditions. To confirm the effects of temperature on degradation, the media were placed in illuminating incubators at 10, 20, 25, 30, 35, and 40℃, respectively. To determine the effect of cypermethrin concentration on degradation, MSM media were added with cypermethrin ranging in concentration from 100 mg/L to 800 mg/L. The media were prepared at pH values from 4.0 to 11.0 buffers for the measurement of the effects of pH on degradation. All experiments were conducted in triplicate. The non-inoculated controls throughout the studied were implemented at the same condition in order to exclude the abiotic degradation affection.

#### **2.6. Extraction of cypermethrin for residue analysis**

The extraction and quantification of cypermethrin residue in the media was modified slightly from method described in Yin et al [27] and Liu et al [30]. At different time intervals, triplicate populations were sampled for cypermethrin concentration analysis. Cypermethrin was extracted three times from the media with 100 mL of hexane. The hexane extracts from the same samples were combined, dried with anhydrous sodium sulfate, and concentrated by exposure to nitrogen gas to near dryness on a rotary evaporator at room temperature, and then dissolved in 5 mL of hexane for GC detection. Before detection the residues were purified using hexane pre-poured Florisil® columns (Agilent SAMPLIQ Florisil®, USA) and 0.22 μm mem‐ branes (Millipore, USA), and were then recovered in 5 mL of hexane; finally, the residues were analyzed by performing GC. Preliminary experiments showed that the recovery of cyper‐ methrin in the above extraction and analysis procedures was >90%.

Residue analyses of cypermethrin degradation were performed using an Agilent 6890N GC system (Agilent Technologies, USA) equipped with an electron capture detector (μ-ECD); an HP-5 5% phenyl methyl siloxane capillary column (30 m × 320 μm × 0.25 μm; Agilent Tech‐ nologies, USA) was used for separation, with helium as the carrier gas (flow rate, 1 mL/min). Other GC parameters included an inlet temperature of 250℃ and a detector temperature of 300℃; initially, the oven temperature was 150℃ for 2.0 min, was ramped to 280℃ at 15℃/min, and then maintained at 280℃ for 5.0 min. The injection volume was 1.0 μL. Samples were introduced in split-less mode. Concentrations were determined by analyzing peak area with an authentic cypermethrin standard.

#### **2.7. Detection of cypermethrin metabolites**

Huada Genomic Company (Beijing, China). The sequences with the highest 16S rDNA partial sequence similarity were selected and compared by CLUSTAL W. Phylogenetic and molecular evolutionary analyses were conducted by MEGA 4.0 software with the Kimura 2-paremeter model and the neighbor joining algorithm [29]. Confidence estimates of branching order were

Unless otherwise stated, the inoculants for this experiment were bacteria cultured in a 130 mL serum bottle containing 120 mL of PSB medium in a light incubator at 35℃ and 7500 lux. At the exponential phase (about 2–3 days), the cell pellets were harvested via centrifugation (5000×g, 10 min), washed 3 times with 50 mL of KH2PO4-K2HPO4 (0.15 mol/L, pH 7.0), and then suspended in the same phosphate buffer as the inoculants. In order to avoid the effects of hydrolysis and photolysis, each treatment was set in triplicate with non-inoculated samples as control under the same conditions and analyzed in the same manner. Samples for residual

pesticide concentration analysis were collected from the cultures at regular intervals.

To determine the optimal conditions for degrading cypermethrin by S10-1, single-factor test was designed in this study under different conditions. To confirm the effects of temperature on degradation, the media were placed in illuminating incubators at 10, 20, 25, 30, 35, and 40℃, respectively. To determine the effect of cypermethrin concentration on degradation, MSM media were added with cypermethrin ranging in concentration from 100 mg/L to 800 mg/L. The media were prepared at pH values from 4.0 to 11.0 buffers for the measurement of the effects of pH on degradation. All experiments were conducted in triplicate. The non-inoculated controls throughout the studied were implemented at the same condition in order to exclude

The extraction and quantification of cypermethrin residue in the media was modified slightly from method described in Yin et al [27] and Liu et al [30]. At different time intervals, triplicate populations were sampled for cypermethrin concentration analysis. Cypermethrin was extracted three times from the media with 100 mL of hexane. The hexane extracts from the same samples were combined, dried with anhydrous sodium sulfate, and concentrated by exposure to nitrogen gas to near dryness on a rotary evaporator at room temperature, and then dissolved in 5 mL of hexane for GC detection. Before detection the residues were purified using hexane pre-poured Florisil® columns (Agilent SAMPLIQ Florisil®, USA) and 0.22 μm mem‐ branes (Millipore, USA), and were then recovered in 5 mL of hexane; finally, the residues were analyzed by performing GC. Preliminary experiments showed that the recovery of cyper‐

Residue analyses of cypermethrin degradation were performed using an Agilent 6890N GC system (Agilent Technologies, USA) equipped with an electron capture detector (μ-ECD); an

determined by bootstrap resampling analysis with 1000 replicates.

**2.5. Optimal conditions for degrading cypermethrin by S10-1**

**2.4. Inoculum preparation**

4 Biodegradation of Hazardous and Special Products

the abiotic degradation affection.

**2.6. Extraction of cypermethrin for residue analysis**

methrin in the above extraction and analysis procedures was >90%.

Metabolites were isolated from the culture filtrates of the organism grown in cypermethrin (100 mg/L) by extraction with acetonitrile, before and after acidification to pH 2 with 2 M HCl, and the residue obtained was dissolved in hexane [22]. The metabolites were identified and analyzed using the GC/MS system (Agilent 7890A/5975, Agilent Technologies, USA) equipped with electron ionization (EI). EI (70 eV) was performed with a trap current of 100 mA and a source temperature of 200℃. Full scan spectra were acquired at m/z 45–500 at 2 sec per scan. The metabolites were confirmed by standard MS, data collection and processing were performed using Agilent MSD ChemStation software containing the Agilent chemical library.

#### **3. Results and discussion**

#### **3.1. Isolation and characterization of cypermethrin degrading bacterium**

After repeated enrichment and purification processes, we obtained approximately 20 strains of organisms with different colony morphologies from the activated sludge samples. But the degradation experiments showed the isolate S10-1 possessed the relatively higher degradation, capacity of degrading cypermethrin (100 mg/L) by 90.4% after incubating 7 days at pH 7.0 and temperature 35℃ (Fig. 3a). And S10-1 utilized cypermethrin as its sole carbon and energy source in MSM. Thus strain S10-1 was selected for further detail investigation.

S10-1 is a gram-negative, anaerobic bacterium. The morphology of the S10-1 colonies, cul‐ tured for 10 days on MSM agar plate, were reddish-brown, smooth, circular, wet, nontrans‐ parent, glistening, and with entire margins (Figure 1a). The physiological and biochemical characteristics of S10-1 are shown in Table 1. SEM observations showed that the cells are ovoid to rod shaped (Figure 1b), sometimes even longer, measuring about 0.5–0.9 μm in width and 1.2–2.0 μm in length, and are motile by means of polar flagella (Figure 1c). Internal photosynthetic membranes appear as lamellae underlying and parallel to the cytoplasmic membrane (Figure 1d). The culture suspension was reddish-brown in color. In vivo absorption maxima of intact cells (Figure 1e) were recorded at 378, 455, 480, 510, 592, 806, and 865 nm, indicating the presence of bacteriochlorophyll a and carotenoids of the spheroidene series [31]. These morphological and biochemical properties are identical to the genus *Rhodobacter* [31].

**Items Results Items Results Items Results**

Motility +b M. R reaction - Succinate utilization +

Hydrogen sulfide + Citrate utilization + Mannitol utilization - V−P reaction - Acid from carbohydrates - Glycerol utilization + Gelatin liquefaction + Indole production - Pyruvate utilization + Catalase + Urease - Benzoate utilization - Oxidase + Pigment production + Ammonia utilization + Strach hydrolysis + Nitrate reduction + Tartrate -

A 1380-bp 16S rRNA fragment was amplified from the genomic DNA of S10-1 and sequenced (Genebank Accession NO. HM193898). Phylogenetic analysis of 16S rRNA revealed that S10-1 belonged to the genus *Rhodobacter sphaeroides* (Figure 2). S10-1 was temporarily identified as *R. sphaeroides* according to its morphology, colony and cultural properties, physiological and biochemical characteristics, absorption spectra (living cells), internal photosynthetic mem‐

Microbial belong to the genus *Rhodobacter*, which are known to play a major role in the treatment of organic wastewater, since they can utilize a broad range of organic compounds as carbon and energy sources; moreover, they are ubiquitous in fresh water, soil, wastewater, and activated sludge. Thus they have been selected for the treatment of many types of wastes [32-34], while *R. sphaeroides* appears to be a new bacterium that may participate in efficient degradation of cypermethrin. To our knowledge, there is not any information concerning the ability of *R. sphaeroides* to degrade cypermethrin and other SPs. However, reports showed that *R. sphaeroides* could effectively degrade pesticides including 2,4-d, quinalphos, monocroto‐

Cypermethrin was degraded by S10-1 during incubation temperatures ranging from 10℃ to 40℃. The cypermethrin residues were detected after 7 days' treatment. In cultures incubated at 10℃ and 20℃, the results show that the degradation rate were 15.3% and 23.8%. However, in cultures incubated at higher temperature, i.e. 25℃, 30℃ and 35℃, the degradation rate reached 70.4%, 87.4% and 90.4% within 7 days, respectively, but the degradation rate was only

Negative/Substrate not utilized;<sup>b</sup> Positive/Substrate utilized.

*Abbreviation*: VP-Vogues Proskauer; MR-Methyl Red.

brane, and phylogenetic analysis.

phos, captan and carbendazim [35].

**3.3. Effect of temperature on cypermethrin degradation in MSM**

**Table 1.** Physiological and biochemical characteristics of S10-1

**3.2. Phylogenetic analysis and identification of S10-1**

<sup>a</sup> 3% NaCl - Aerobic dark growth +

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

7

Gram stain -

Note: a

**Figure 1.** The characterization of strain S10-1. (a) The morphology of the S10-1 colonies, cultured for 10 days on MSM agar plate; (b) Scanning electron micrograph of strain S10-1 (10,000×); (c) Electron micrograph of negatively stained S10-1 cells showing polar flagella (40,000×); (d) Transmission electron micrograph of S10-1: a cross-section showing the photosynthetic membrane (*PM*) lying parallel to the cytoplasmic membrane (200,000×); (e) Absorption spectra of liv‐ ing S10-1 cells.

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil http://dx.doi.org/10.5772/56339 7


Note: a Negative/Substrate not utilized;<sup>b</sup> Positive/Substrate utilized.

*Abbreviation*: VP-Vogues Proskauer; MR-Methyl Red.

(a) (b)

6 Biodegradation of Hazardous and Special Products

(c) (d)

(e)

ing S10-1 cells.

)

)

**Figure 1.** The characterization of strain S10-1. (a) The morphology of the S10-1 colonies, cultured for 10 days on MSM agar plate; (b) Scanning electron micrograph of strain S10-1 (10,000×); (c) Electron micrograph of negatively stained S10-1 cells showing polar flagella (40,000×); (d) Transmission electron micrograph of S10-1: a cross-section showing the photosynthetic membrane (*PM*) lying parallel to the cytoplasmic membrane (200,000×); (e) Absorption spectra of liv‐ **Table 1.** Physiological and biochemical characteristics of S10-1

#### **3.2. Phylogenetic analysis and identification of S10-1**

A 1380-bp 16S rRNA fragment was amplified from the genomic DNA of S10-1 and sequenced (Genebank Accession NO. HM193898). Phylogenetic analysis of 16S rRNA revealed that S10-1 belonged to the genus *Rhodobacter sphaeroides* (Figure 2). S10-1 was temporarily identified as *R. sphaeroides* according to its morphology, colony and cultural properties, physiological and biochemical characteristics, absorption spectra (living cells), internal photosynthetic mem‐ brane, and phylogenetic analysis.

Microbial belong to the genus *Rhodobacter*, which are known to play a major role in the treatment of organic wastewater, since they can utilize a broad range of organic compounds as carbon and energy sources; moreover, they are ubiquitous in fresh water, soil, wastewater, and activated sludge. Thus they have been selected for the treatment of many types of wastes [32-34], while *R. sphaeroides* appears to be a new bacterium that may participate in efficient degradation of cypermethrin. To our knowledge, there is not any information concerning the ability of *R. sphaeroides* to degrade cypermethrin and other SPs. However, reports showed that *R. sphaeroides* could effectively degrade pesticides including 2,4-d, quinalphos, monocroto‐ phos, captan and carbendazim [35].

#### **3.3. Effect of temperature on cypermethrin degradation in MSM**

Cypermethrin was degraded by S10-1 during incubation temperatures ranging from 10℃ to 40℃. The cypermethrin residues were detected after 7 days' treatment. In cultures incubated at 10℃ and 20℃, the results show that the degradation rate were 15.3% and 23.8%. However, in cultures incubated at higher temperature, i.e. 25℃, 30℃ and 35℃, the degradation rate reached 70.4%, 87.4% and 90.4% within 7 days, respectively, but the degradation rate was only

within 7 days. However at high concentration (400 to 800 mg/L), only about 30% was degraded within 7 days. It might be because of the fact that microbial degradation starts slowly and requires an acclimation period before rapid degradation occurs at high concentration. Similar results were reported by Lin et al [36] who reported that initial concentration of carfofuran was significantly efficiently degraded by *Pichia anomala* strain HQ-C-01 in contaminated soils. In non-inoculated controls at different initial concentrations, abiotic degradation was negligi‐

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

9

0 10 20 30 40 50 Temperature (℃ )

3 4 5 6 7 8 9 10 11 12 pH

7

14.874, 16.980, and 17.323 min, respectively.

Figure 3. Optimal conditions for degrading cypermethrin by S10-1. (a) Effect of temperature on the degradation of cypermethrin by S10-1; (b) Effect of the initial cypermethrin concentration on the degradation by S10-1; (c) (d) Effect of pH on the degradation of cypermethrin by S10-1. Error bars

The degradation metabolites of cypermethrin by strain S10-1 were extracted and identified by GC/MS using Agilent MSD ChemStation software containing the Agilent chemical library. GC/MS analysis of the metabolites showed the presence of 4 products. These compounds corresponded with cyclopropanemethanol (Figure 4a), 5-methoxy-2-nitrobenzoic acid (Figure 4b), 3,5 dimethoxybenzamide (Figure 4c), and 5-aminoisophthalic acid (Figure 4d). The retention times of these compounds were 13.609,

Previous studies had reported about the biodegradation pathway of SPs [21, 39, 40]. In the molecular structure of SPs there is an ester bond which is not as firm as other chemical bonds. Literature indicated that the first step in the microbial degradation and detoxification of SPs is the hydrolysis of its carboxyl ester linkage [23, 36, 41]. However, the chemical bond broken of cypermethrin metabolites are not detected as that described in a previous study. It is evident from our GC/MS results that S10-1 degraded cypermethrin by reductive dechlorination, oxidation or/and hydrolysis to transform to other 4 metabolites. The cypermethrin degradation pathway appeared to be different to the initial steps of SPs degradation by *Ochrobactrum lupine*, *Pseudomonas aeruginosa*, *Pseudomonas aeruginosa* and *Achromobacter* sp. [21, 22, 42, 43]. Moreover, no 3-phenoxybenzoic acid (3-PBA) was detected in the metabolites by GC-MS after 7 days of treatment, while 3-PBA was generally regarded as the major metabolite after hydrolysis of SPs in soil and water [21, 36, 42-44]. Owing to its antimicrobial activities [23, 45] and transient GC/MS detectable peak

represent standard deviation (SD) from the mean. Error bars smaller than symbols are not depicted.

0 200 400 600 800 1000 Cypermethrin concentration (mg/L)

**3.6. Identification of cypermethrin degradation metabolites** 

**Figure 3.** Optimal conditions for degrading cypermethrin by S10-1. (a) Effect of temperature on the degradation of cy‐ permethrin by S10-1; (b) Effect of the initial cypermethrin concentration on the degradation by S10-1; (c) (d) Effect of pH on the degradation of cypermethrin by S10-1. Error bars represent standard deviation (SD) from the mean. Error bars

ble throughout the studies.

(a)

Degradation rate (%)

Degradation rate (%)

(b)

Degradation rate (%)

(c)

smaller than symbols are not depicted.

**Figure 2.** Phylogenetic tree constructed by the neighbor-joining method based on 16S rDNA sequences of S10-1 and related strains. Bootstrap values are given at branching points. The sequence of *Arthrobacter* spp. (AY628689) was se‐ lected as an out group. The tree was constructed using the neighbor-joining method. Bootstrap values at nodes were calculated using 1,000 replicates (only values >70% are indicated). The GeneBank accession numbers for 16S rRNA gene sequences are shown in parentheses.

61% when incubated at temperature 40℃ for 7 days. The best temperature for degradation was 35℃ (Figure 3a). Similar results were reported by Lin et al [36] who reported temperature significantly influenced cypermethrin degradation by *Streptomyces* sp. strain HU-S-01. Our results also reveal that cypermethrin degradation occurred at 30–35℃ indicating strain S10-1 preferred relatively high temperature condition. These results were consistent with previous findings of Chen et al [21]. It is possible that some key enzyme(s) responsible for cypermethrin degradation have their optimum enzymatic activity over such range of temperature. In noninoculated controls at different temperatures, abiotic degradation was negligible throughout the studies.

#### **3.4. Effect of initial concentration on cypermethrin degradation in MSM**

Cypermethrin degradation at different initial concentrations by strain S10-1 was investigated. The cypermethrin degradation rates were found to be 90.4%, 60.3%, 38.4%, 32.3%, and 28.7% at concentrations of 100, 200, 400, 600, and 800 mg/L, respectively (Figure 3b). At low cyper‐ methrin concentration ranging from 100 to 200 mg/L, the degradation rate reached above 60% within 7 days. However at high concentration (400 to 800 mg/L), only about 30% was degraded within 7 days. It might be because of the fact that microbial degradation starts slowly and requires an acclimation period before rapid degradation occurs at high concentration. Similar results were reported by Lin et al [36] who reported that initial concentration of carfofuran was significantly efficiently degraded by *Pichia anomala* strain HQ-C-01 in contaminated soils. In non-inoculated controls at different initial concentrations, abiotic degradation was negligi‐ ble throughout the studies.

61% when incubated at temperature 40℃ for 7 days. The best temperature for degradation was 35℃ (Figure 3a). Similar results were reported by Lin et al [36] who reported temperature significantly influenced cypermethrin degradation by *Streptomyces* sp. strain HU-S-01. Our results also reveal that cypermethrin degradation occurred at 30–35℃ indicating strain S10-1 preferred relatively high temperature condition. These results were consistent with previous findings of Chen et al [21]. It is possible that some key enzyme(s) responsible for cypermethrin degradation have their optimum enzymatic activity over such range of temperature. In noninoculated controls at different temperatures, abiotic degradation was negligible throughout

**Figure 2.** Phylogenetic tree constructed by the neighbor-joining method based on 16S rDNA sequences of S10-1 and related strains. Bootstrap values are given at branching points. The sequence of *Arthrobacter* spp. (AY628689) was se‐ lected as an out group. The tree was constructed using the neighbor-joining method. Bootstrap values at nodes were calculated using 1,000 replicates (only values >70% are indicated). The GeneBank accession numbers for 16S rRNA

 *Rhodopseudomonas palustris(AB498819) Rhodopseudomonas palustris(AB498821) Rhodopseudomonas palustris(D84187)*

 *Rhodobacter sphaeroides (GU573899) Rhodopseudomonas sp. (AB251404) Rhodopseudomonas sp. (AF095928)*

 *Bradyrhizobium japonicum(AB513462) Bradyrhizobium japonicum(AB513466) Rhodobacter azotoformans(D70847) Rhodobacter sp.(AF515782) Ectothiorhodospira marina(X93476) Ectothiorhodospira variabilis (AM943121)*

 *Afipia genosp. (U87766) Afipia genosp.(U87768)*

Arthrobacter sp.(AY628689)

 *S10-1*

100

54

49

100

54 98

100

43

100

8 Biodegradation of Hazardous and Special Products

0.12 0.10 0.08 0.06 0.04 0.02 0.00

100

100

100 100

Cypermethrin degradation at different initial concentrations by strain S10-1 was investigated. The cypermethrin degradation rates were found to be 90.4%, 60.3%, 38.4%, 32.3%, and 28.7% at concentrations of 100, 200, 400, 600, and 800 mg/L, respectively (Figure 3b). At low cyper‐ methrin concentration ranging from 100 to 200 mg/L, the degradation rate reached above 60%

**3.4. Effect of initial concentration on cypermethrin degradation in MSM**

the studies.

gene sequences are shown in parentheses.

Figure 3. Optimal conditions for degrading cypermethrin by S10-1. (a) Effect of temperature on the degradation of cypermethrin by S10-1; (b) Effect of the initial cypermethrin concentration on the degradation by S10-1; (c) (d) Effect of pH on the degradation of cypermethrin by S10-1. Error bars represent standard deviation (SD) from the mean. Error bars smaller than symbols are not depicted. **Figure 3.** Optimal conditions for degrading cypermethrin by S10-1. (a) Effect of temperature on the degradation of cy‐ permethrin by S10-1; (b) Effect of the initial cypermethrin concentration on the degradation by S10-1; (c) (d) Effect of pH on the degradation of cypermethrin by S10-1. Error bars represent standard deviation (SD) from the mean. Error bars smaller than symbols are not depicted.

**3.6. Identification of cypermethrin degradation metabolites** 

The degradation metabolites of cypermethrin by strain S10-1 were extracted and identified by GC/MS using Agilent MSD ChemStation software containing the Agilent chemical library. GC/MS analysis of the metabolites showed the presence of 4 products. These compounds corresponded with cyclopropanemethanol (Figure 4a), 5-methoxy-2-nitrobenzoic acid (Figure 4b), 3,5 dimethoxybenzamide (Figure 4c), and 5-aminoisophthalic acid (Figure 4d). The retention times of these compounds were 13.609,

Previous studies had reported about the biodegradation pathway of SPs [21, 39, 40]. In the molecular structure of SPs there is an ester bond which is not as firm as other chemical bonds. Literature indicated that the first step in the microbial degradation and detoxification of SPs is the hydrolysis of its carboxyl ester linkage [23, 36, 41]. However, the chemical bond broken of cypermethrin metabolites are not detected as that described in a previous study. It is evident from our GC/MS results that S10-1 degraded cypermethrin by reductive dechlorination, oxidation or/and hydrolysis to transform to other 4 metabolites. The cypermethrin degradation pathway appeared to be different to the initial steps of SPs degradation by *Ochrobactrum lupine*, *Pseudomonas aeruginosa*, *Pseudomonas aeruginosa* and *Achromobacter* sp. [21, 22, 42, 43]. Moreover, no 3-phenoxybenzoic acid (3-PBA) was detected in the metabolites by GC-MS after 7 days of treatment, while 3-PBA was generally regarded as the major metabolite after hydrolysis of SPs in soil and water [21, 36, 42-44]. Owing to its antimicrobial activities [23, 45] and transient GC/MS detectable peak

7

14.874, 16.980, and 17.323 min, respectively.

#### **3.5. Effect of pH on cypermethrin degradation in MSM**

The pH is also an important factor, which significantly effects the degrading ability of bacteria capable of degrading toxicities [37, 38]. To determine the effect of pH on degradation, MSM medium prepared with different pH buffers, fortified with 100 mg/L cypermethrin, and incubated at 35℃ and 7500 lux. Eight different pH (4.0, 5.0, 6.0, 7.0, 8.0, 9.0, 10.0, 11.0) were tested in the optimization experiment. The result showed that the degradation rate were 18.5%, 26.7%, 57.5%, 90.4%, 60.3%, 38.4%, 32.3%, 28.7%, respectively (Figure 3c). The optimal initial pH value for degradation was between 6.0 and 8.0. Results revealed that S10-1 was able to degrade cypermethrin over a wide range of pH. Similar results were reported by Zhang et al [36] who reported that initial pHs were significantly efficiently degraded by two *Serratia* spp., and rapid degradation of cypermethrin at high pH while it was relatively low at acidic pH. In non-inoculated controls at different pH conditions, abiotic degradation was negligible throughout the studies.

considerable interest in studying of degrading toxic compounds [35, 47]. The structural components of this metabolically diverse organism and their modes of integrated regulation are encoded by a genome of ∼4.5 Mb in size [46]. Moreover, its large inventory of transport and chemotaxis genes also implies that *Rhodobacter* is adept at sensing and acquiring diverse

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

11

)

**Figure 4.** GC/MS spectra of four main metabolites produced during cypermethrin degradation by strain S10-1. (a) cyclo‐ propanemethanol; (b) 5-methoxy-2-nitrobenzoic acid; (c) 3,5-dimethoxybenzamide; (d) 5-aminoisophthalic acid.

*R. sphaeroides* strain S10-1 was isolated from an activated sludge sample collected from the wastewater treatment pool of a pesticide plant. It can utilize cypermethrin as sole source of carbon, nitrogen and energy. The optimal temperature and pH for biodegradation of cyper‐ methrin by strain S10-1 were 35℃and pH 7.0, and the degradation rate reached 90.4% within 7 days under the optimal conditions. Four metabolic compounds were detected, hinting that there are complex redox reactions are involved in the cypermethrin degradation process.

In conclusion, our results indicated that strain S10-1 could be a good choice for the bioremedia‐ tion of cypermethrin contaminated water and soil. However, further studies such as its

compounds from its environment [48-50].

(a) (b)

(c) (d)

**4. Conclusion**

#### **3.6. Identification of cypermethrin degradation metabolites**

The degradation metabolites of cypermethrin by strain S10-1 were extracted and identified by GC/MS using Agilent MSD ChemStation software containing the Agilent chemical library. GC/ MS analysis of the metabolites showed the presence of 4 products. These compounds corre‐ sponded with cyclopropanemethanol (Figure 4a), 5-methoxy-2-nitrobenzoic acid (Figure 4b), 3,5-dimethoxybenzamide (Figure 4c), and 5-aminoisophthalic acid (Figure 4d). The retention times of these compounds were 13.609, 14.874, 16.980, and 17.323 min, respectively.

Previous studies had reported about the biodegradation pathway of SPs [21, 39, 40]. In the molecular structure of SPs there is an ester bond which is not as firm as other chemical bonds. Literature indicated that the first step in the microbial degradation and detoxification of SPs is the hydrolysis of its carboxyl ester linkage [23, 36, 41]. However, the chemical bond broken of cypermethrin metabolites are not detected as that described in a previous study. It is evident from our GC/MS results that S10-1 degraded cypermethrin by reductive dechlorination, oxidation or/and hydrolysis to transform to other 4 metabolites. The cypermethrin degradation pathway appeared to be different to the initial steps of SPs degradation by *Ochrobactrum lupine*, *Pseudomonas aeruginosa*, *Pseudomonas aeruginosa* and *Achromobacter* sp. [21, 22, 42, 43]. Moreover, no 3-phenoxybenzoic acid (3-PBA) was detected in the metabolites by GC-MS after 7 days of treatment, while 3-PBA was generally regarded as the major metabolite after hydrolysis of SPs in soil and water [21, 36, 42-44]. Owing to its antimicrobial activities [23, 45] and transient GC/MS detectable peak [21, 45], biodegradation of 3-PBA was rarely reported. Chen et al reported that fenvalerate was degraded by hydrolysis of the carboxylester linkage to yield 3-PBA, and then the intermediate was further utilized for bacterial growth by strain ZS-S-01, finally resulted in complete mineralization [42]. So, we speculated that carboxyles‐ terases and oxidoreductases involved in degradation of cypermethrin by strain S10-1, that needed to be testified by further experiments.

On the other hand, *R. sphaeroides* are metabolically flexible and under different situations they can grow chemoheterotrophically, chemoautotrophically, photoheterotrophically, and photoautotrophically [46]. Because of this multiplicity of growth modes there has been considerable interest in studying of degrading toxic compounds [35, 47]. The structural components of this metabolically diverse organism and their modes of integrated regulation are encoded by a genome of ∼4.5 Mb in size [46]. Moreover, its large inventory of transport and chemotaxis genes also implies that *Rhodobacter* is adept at sensing and acquiring diverse compounds from its environment [48-50].

**Figure 4.** GC/MS spectra of four main metabolites produced during cypermethrin degradation by strain S10-1. (a) cyclo‐ propanemethanol; (b) 5-methoxy-2-nitrobenzoic acid; (c) 3,5-dimethoxybenzamide; (d) 5-aminoisophthalic acid.

#### **4. Conclusion**

**3.5. Effect of pH on cypermethrin degradation in MSM**

10 Biodegradation of Hazardous and Special Products

**3.6. Identification of cypermethrin degradation metabolites**

needed to be testified by further experiments.

throughout the studies.

The pH is also an important factor, which significantly effects the degrading ability of bacteria capable of degrading toxicities [37, 38]. To determine the effect of pH on degradation, MSM medium prepared with different pH buffers, fortified with 100 mg/L cypermethrin, and incubated at 35℃ and 7500 lux. Eight different pH (4.0, 5.0, 6.0, 7.0, 8.0, 9.0, 10.0, 11.0) were tested in the optimization experiment. The result showed that the degradation rate were 18.5%, 26.7%, 57.5%, 90.4%, 60.3%, 38.4%, 32.3%, 28.7%, respectively (Figure 3c). The optimal initial pH value for degradation was between 6.0 and 8.0. Results revealed that S10-1 was able to degrade cypermethrin over a wide range of pH. Similar results were reported by Zhang et al [36] who reported that initial pHs were significantly efficiently degraded by two *Serratia* spp., and rapid degradation of cypermethrin at high pH while it was relatively low at acidic pH. In non-inoculated controls at different pH conditions, abiotic degradation was negligible

The degradation metabolites of cypermethrin by strain S10-1 were extracted and identified by GC/MS using Agilent MSD ChemStation software containing the Agilent chemical library. GC/ MS analysis of the metabolites showed the presence of 4 products. These compounds corre‐ sponded with cyclopropanemethanol (Figure 4a), 5-methoxy-2-nitrobenzoic acid (Figure 4b), 3,5-dimethoxybenzamide (Figure 4c), and 5-aminoisophthalic acid (Figure 4d). The retention times of these compounds were 13.609, 14.874, 16.980, and 17.323 min, respectively. Previous studies had reported about the biodegradation pathway of SPs [21, 39, 40]. In the molecular structure of SPs there is an ester bond which is not as firm as other chemical bonds. Literature indicated that the first step in the microbial degradation and detoxification of SPs is the hydrolysis of its carboxyl ester linkage [23, 36, 41]. However, the chemical bond broken of cypermethrin metabolites are not detected as that described in a previous study. It is evident from our GC/MS results that S10-1 degraded cypermethrin by reductive dechlorination, oxidation or/and hydrolysis to transform to other 4 metabolites. The cypermethrin degradation pathway appeared to be different to the initial steps of SPs degradation by *Ochrobactrum lupine*, *Pseudomonas aeruginosa*, *Pseudomonas aeruginosa* and *Achromobacter* sp. [21, 22, 42, 43]. Moreover, no 3-phenoxybenzoic acid (3-PBA) was detected in the metabolites by GC-MS after 7 days of treatment, while 3-PBA was generally regarded as the major metabolite after hydrolysis of SPs in soil and water [21, 36, 42-44]. Owing to its antimicrobial activities [23, 45] and transient GC/MS detectable peak [21, 45], biodegradation of 3-PBA was rarely reported. Chen et al reported that fenvalerate was degraded by hydrolysis of the carboxylester linkage to yield 3-PBA, and then the intermediate was further utilized for bacterial growth by strain ZS-S-01, finally resulted in complete mineralization [42]. So, we speculated that carboxyles‐ terases and oxidoreductases involved in degradation of cypermethrin by strain S10-1, that

On the other hand, *R. sphaeroides* are metabolically flexible and under different situations they can grow chemoheterotrophically, chemoautotrophically, photoheterotrophically, and photoautotrophically [46]. Because of this multiplicity of growth modes there has been *R. sphaeroides* strain S10-1 was isolated from an activated sludge sample collected from the wastewater treatment pool of a pesticide plant. It can utilize cypermethrin as sole source of carbon, nitrogen and energy. The optimal temperature and pH for biodegradation of cyper‐ methrin by strain S10-1 were 35℃and pH 7.0, and the degradation rate reached 90.4% within 7 days under the optimal conditions. Four metabolic compounds were detected, hinting that there are complex redox reactions are involved in the cypermethrin degradation process.

In conclusion, our results indicated that strain S10-1 could be a good choice for the bioremedia‐ tion of cypermethrin contaminated water and soil. However, further studies such as its interactions with environment, toxicological aspects, degradation enzymes, biochemical and genetic aspects are still needed before the application in actual field-scale bioremediation.

worker exposure to pyrethroid insecticides. Analytical and Bioanalytical Chemistry.

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

13

[6] Nauen, R, Zimmer, C. T, Andrews, M, Slater, R, Bass, C, Ekbom, B, Gustafsson, G, Hansen, L. M, Kristensen, M, & Zebitz, C. P. W. Target-site resistance to pyrethroids in European populations of pollen beetle, Meligethes aeneus F.. Pesticide Biochemis‐

[7] Umina, P. A, Weeks, A. R, Roberts, J, & Jenkins, S. Peter Mangano, G., Lord, A., and Micic, S. The current status of pesticide resistance in Australian populations of the redlegged earth mite (Halotydeus destructor). Pest Management Science. (2012). ,

[8] Ding, G, Shi, R, Gao, Y, Zhang, Y, Kamijima, M, Sakai, K, Wang, G, Feng, C, & Tian, Y. Pyrethroid pesticide exposure and risk of childhood acute lymphocytic leukemia

[9] Soderlund, D. M. Molecular mechanisms of pyrethroid insecticide neurotoxicity: re‐

[10] Alonso, M. B, Feo, M. L, Corcellas, C, Vidal, L. G, Bertozzi, C. P, Marigo, J, Secchi, E. R, Bassoi, M, Azevedo, A. F, & Dorneles, P. R. Pyrethroids: A new threat to marine

[11] Tsuji, R, Yamada, T, & Kawamura, S. Mammal toxicology of synthetic pyrethroids.

[12] Kulkarni, G, & Joshi, P. Cypermethrin and fenvalerate induced protein alterations in freshwater crab Barytelphusa cunicularis (westwood). Recent Research in Science

[13] Saha, S, & Kaviraj, A. Acute toxicity of synthetic pyrethroid cypermethrin to some freshwater organisms. Bulletin of Environmental Contamination and Toxicology.

[14] Anderson, R. L. Toxicity of synthetic pyrethroids to freshwater invertebrates. Envi‐

[15] Datta, M, & Kaviraj, A. Acute toxicity of the synthetic pyrethroid pesticide fenvaler‐ ate to some air breathing fishes. Toxicological and Environmental Chemistry.(2011). ,

[16] Anjum, R, Rahman, M, Masood, F, & Malik, A. Bioremediation of pesticides from soil and wastewater. Environmental Protection Strategies for Sustainable Development.

[17] Mugni, H, Demetrio, P, Bulus, G, Ronco, A, & Bonetto, C. Effect of aquatic vegetation on the persistence of cypermethrin toxicity in water. Bulletin of Environmental Con‐

in Shanghai. Environmental Science and Technology. (2012). DOI:es303362a.

cent advances. Archives of Toxicology. (2012). , 86(2), 165-181.

mammals?. Environment International. (2012). , 47, 99-106.

ronmental Toxicology and Chemistry.(2009). , 8(5), 403-410.

tamination and Toxicology.(2011). , 86(1), 23-27.

Topics in Current Chemistry. (2011). , 314, 83-111.

and Technology. (2011). , 3(12), 7-10.

(2008). , 80(1), 49-52.

93(10), 2034-2039.

(2012).

(2011). , 401(4), 1285-1293.

68(6), 889-896.

try and Physiology. (2012). , 103(3), 173-180.

#### **Acknowledgements**

This work was funded by the earmarked fund for Modern Agro-industry Technology Research System; the National Natural Science Foundation of China (No. 31071753), the key project of Hunan Provincial Education Department (11CY016), the key project of Shaoyang Municipal Science and Technology (J1107), and the 12th Five-Year key discipline of of Shaoyang Univer‐ sity.

#### **Author details**

L. B. Yin1 , L. Z. Zhao1 , Y. Liu2 , D. Y. Zhang2 , S. B. Zhang2 and K. Xiao1

1 Department of Biological and Chemical Engineering, Shaoyang University, Shaoyang, Hunan, People's Republic of China

2 Hunan Plant Protection Institute, Changsha, Hunan, People's Republic of China

#### **References**


worker exposure to pyrethroid insecticides. Analytical and Bioanalytical Chemistry. (2011). , 401(4), 1285-1293.

interactions with environment, toxicological aspects, degradation enzymes, biochemical and genetic aspects are still needed before the application in actual field-scale bioremediation.

This work was funded by the earmarked fund for Modern Agro-industry Technology Research System; the National Natural Science Foundation of China (No. 31071753), the key project of Hunan Provincial Education Department (11CY016), the key project of Shaoyang Municipal Science and Technology (J1107), and the 12th Five-Year key discipline of of Shaoyang Univer‐

, S. B. Zhang2

1 Department of Biological and Chemical Engineering, Shaoyang University, Shaoyang,

[1] Vischetti, C, Monaci, E, Coppola, L, Marinozzi, M, & Casucci, C. Evaluation of bio‐ massbed system in bio-cleaning water contaminated by fungicides applied in vine‐ yard. International Journal of Environmental Analytical Chemistry. (2012). , 92(8),

[2] Jaiswal, M, Chauhan, D, & Sankararamakrishnan, N. Copper chitosan nanocompo‐ site: synthesis, characterization, and application in removal of organophosphorous pesticide from agricultural runoff. Environmental Science and Pollution Research.

[3] Nkya, T. E, Akhouayri, I, Kisinza, W, & David, J. P. Impact of environment on mos‐ quito response to pyrethroid insecticides: facts, evidences and prospects. Insect Bio‐

[4] Elsheikha, H. M, Mcorist, S, & Geary, T. G. Antiparasitic drugs: mechanisms of action

[5] Ahn, K. C, Gee, S. J, Kim, H. J, Aronov, P. A, Vega, H, Krieger, R. I, & Hammock, B. D. Immunochemical analysis of 3-phenoxybenzoic acid, a biomarker of forestry

chemistry and Molecular Biology. http://dx.doi.org/10.1016/j.ibmb.(2012).

and resistance. Essentials of Veterinary Parasitology; (2011).

2 Hunan Plant Protection Institute, Changsha, Hunan, People's Republic of China

and K. Xiao1

**Acknowledgements**

12 Biodegradation of Hazardous and Special Products

sity.

**Author details**

, L. Z. Zhao1

Hunan, People's Republic of China

, Y. Liu2

, D. Y. Zhang2

L. B. Yin1

**References**

949-962.

(2012). , 19(6), 2055-2062.


[18] Riser-roberts, E. Remediation of petroleum contaminated soils: biological, physical, and chemical processes: (CRC); (1998).

[30] Liu, S, Yao, K, Jia, D, Zhao, N, Lai, W, & Yuan, H. A pretreatment method for HPLC analysis of cypermethrin in microbial degradation systems. Journal of Chromato‐

Isolation and Characterization of Cypermethrin Degrading Bacteria Screened from Contaminated Soil

http://dx.doi.org/10.5772/56339

15

[31] Vos, P, Garrity, G, Jones, D, Krieg, N. R, Ludwig, W, Rainey, F. A, Schleifer, K. H, & Whitman, W. B. Bergey's Manual of Systematic Bacteriology: The Firmicutes, Vol‐

[32] Ramana, C. V. Photoassimilation of aromatic compounds by Rhodobacter sphaer‐

[33] Yetis, M, Gündüz, U, Eroglu, I, Yücel, M, & Türker, L. Photoproduction of hydrogen from sugar refinery wastewater by Rhodobacter sphaeroides OU001. International

[34] Tao, Y, He, Y, Wu, Y, Liu, F, Li, X, Zong, W, & Zhou, Z. (2008). Characteristics of a new photosynthetic bacterial strain for hydrogen production and its application in wastewater treatment. International Journal of Hydrogen Energy.2008;, 33(3),

[35] Chalam, A, Sasikala, C, & Ramana, C. V. and Raghuveer Rao, P. Effect of pesticides on hydrogen metabolism of Rhodobacter sphaeroides and Rhodopseudomonas pal‐

[36] Lin, Q, Chen, S, Hu, M, Rizwan-ul-haq, M, Yang, L, & Li, H. (2011). Biodegradation of cypermethrin by a newly isolated actinomycetes HU-S-01 from wastewater sludge. International Journal of Environment and Science and Technology. 2011;,

[37] Howe, G. E, Marking, L. L, Bills, T. D, Rach, J. J, & Mayer, F. L. Effects of water tem‐ perature and pH on toxicity of terbufos, trichlorfon, 4-nitrophenol and 2,4-dinitro‐ phenol to the amphipod Gammarus pseudolimnaeus and rainbow trout (Oncorhynchus mykiss). Environmental Toxicology and Chemistry.(2009). , 13(1),

[38] Rowland, S, Jones, D, Scarlett, A, West, C, Hin, L. P, Boberek, M, Tonkin, A, Smith, B, & Whitby, C. Synthesis and toxicity of some metabolites of the microbial degradation of synthetic naphthenic acids. Science of the Total Environment. (2011). , 409(15),

[39] Wang, B, Ma, Y, Zhou, W, Zheng, J, Zhu, J, He, J, & Li, S. Biodegradation of synthetic pyrethroids by Ochrobactrum tritici strain pyd-1. World Journal of Microbiology and

[40] Zhang, S, Yin, L, Liu, Y, Zhang, D, Luo, X, Cheng, J, Cheng, F, & Dai, J. Cometabolic biotransformation of fenpropathrin by Clostridium species strain ZP3. Biodegrada‐

oides OU5. Ph.D Thesis. University of Hyderabad Idia; (2012).

Journal of Hydrogen Energy.(2000). , 25(11), 1035-1041.

ustris. FEMS Microbiology Ecology.(1996). , 19(1), 1-4.

graphic Science. (2012). , 50(6), 469-476.

ume 3, (Springer); (2009). , 3

963-973.

8(1), 45-56.

51-66.

2936-2941.

Biotechnology.(2011). , 27(10), 2315-2324.

tion.(2011). , 22(5), 869-875.


[30] Liu, S, Yao, K, Jia, D, Zhao, N, Lai, W, & Yuan, H. A pretreatment method for HPLC analysis of cypermethrin in microbial degradation systems. Journal of Chromato‐ graphic Science. (2012). , 50(6), 469-476.

[18] Riser-roberts, E. Remediation of petroleum contaminated soils: biological, physical,

[19] Marttinen, S, Kettunen, R, Sormunen, K, Soimasuo, R, & Rintala, J. Screening of physical-chemical methods for removal of organic material, nitrogen and toxicity

[20] Naveen, D, Majumder, C, Mondal, P, & Shubha, D. Biological treatment of cyanide containing wastewater. Research Journal of Chemical Sciences. (2011). , 1(7), 15-21.

[21] Chen, S, Hu, M, Liu, J, Zhong, G, Yang, L, Rizwan-ul-haq, M, & Han, H. Biodegrada‐ tion of beta-cypermethrin and 3-phenoxybenzoic acid by a novel Ochrobactrum lupi‐

[22] Zhang, C, Wang, S, & Yan, Y. Isomerization and biodegradation of beta-cypermeth‐ rin by Pseudomonas aeruginosa CH7 with biosurfactant production. Bioresource

[23] Chen, S, Geng, P, Xiao, Y, & Hu, M. Bioremediation of β-cypermethrin and 3-phenox‐ ybenzaldehyde contaminated soils using Streptomyces aureus HP-S-01. Applied Mi‐

[24] Kontur, W. S, Ziegelhoffer, E. C, Spero, M. A, Imam, S, Noguera, D. R, & Donohue, T. J. Pathways involved in reductant distribution during photobiological H2 production by Rhodobacter sphaeroides. Applied and Environmental Microbiology.(2011). ,

[25] Sonhom, R, Thepsithar, C, & Jongsareejit, B. High level production of 5-aminolevu‐ linic acid by Propionibacterium acidipropionici grown in a low-cost medium. Bio‐

[26] Kang, Z, Wang, Y, Wang, Q, & Qi, Q. Metabolic engineering to improve 5-aminole‐

[27] Yin, L, Li, X, Liu, Y, Zhang, D, Zhang, S, & Luo, X. Biodegradation of cypermethrin by Rhodopseudomonas palustris GJ-22 isolated from activated sludge. Fresenius En‐

[28] Mirnejad, R, Babavalian, H, Moghaddam, M. M, Khodi, S, & Shakeri, F. Rapid DNA extraction of bacterial genome using laundry detergents and assessment of the effi‐ ciency of DNA in downstream process using polymerase chain reaction. African

[29] Haws, D. C, Hodge, T. L, & Yoshida, R. Optimality of the neighbor joining algorithm and faces of the balanced minimum evolution polytope. Bulletin of Mathematical Bi‐

vulinic acid production. Bioengineered. (2011). , 2(6), 342-345.

from low strength landfill leachates. Chemosphere. (2002). , 46(6), 851-858.

ni DG-S-01. Journal of Hazardous Materials. (2011). , 187(1), 433-440.

and chemical processes: (CRC); (1998).

14 Biodegradation of Hazardous and Special Products

technology. (2011). , 102(14), 7139-7146.

technology letters. (2012). , 34(9), 1667-1672.

vironmental Bulletin. (2012). , 21(2), 397-405.

Journal of Biotechnology. (2012). , 11(1), 173-178.

ology. (2011). , 73(11), 2627-2648.

77(20), 7425-7429.

crobiology and Biotechnology. (2012). , 94(2), 505-515.


[41] Tallur, P. N, Megadi, V. B, & Ninnekar, H. Z. Biodegradation of cypermethrin by Mi‐ crococcus sp. strain CPN 1. Biodegradation.(2008). , 19(1), 77-82.

**Chapter 2**

**Chlorophyll Biodegradation**

Additional information is available at the end of the chapter

One of the most important biochemical processes is photosynthesis and life depends on the conversion of the solar energy into a chemical one. It occurs in photoautotrophic or‐ ganisms. The chlorophyll *a* is supported by chlorophyll *b*, carotenes and xanthophylls in the light capture. The chlorophyll *a* and accessory pigments are found in cell organelles called chloroplasts. The conversion of light energy into the chemical one can be described thermodynamically. The entropy of photosynthesis yields the energy of 27.32 kJ mol-1 and is a source of energy for the plants. The atmospheric oxygen is a product of the oxygenic photosynthesis, the product of the first phase process known as light – dependant reac‐ tions. In those reactions, the chlorophylls, in the so called reaction centre, function as an absorbent of a photon initiating the electron transport chain, with the loss of an electron. During the electron flow the energy is produced and the reduced form of the nicotina‐ mide adenine dinucleotide phosphate (NADPH) and the adenosine triphosphate (ATP) are formed. The chlorophyll molecule regains the electron from water. The other phase of photosynthesis is called the light – independent reactions or the Calvin cycle. In those re‐ actions the enzyme ribulose-1, 5-bisphosphate carboxylase/oxygenase (RuBisCO) captures the CO2 from the atmosphere and during the Calvin cycle glucose is formed. The light-ab‐ sorbing green pigments, the chlorophylls, are biodegraded under various conditions: with the leaf senescence in autumn, under stress conditions, such as, drought, flooding, chill‐ ing, mechanical wounding, etc., under the exposure to the ethylene and after the treat‐ ment of plants with herbicides. In autumn deciduous trees shed tones of leaves. The green leaves change the colour due to the chlorophyll biodegradation. The peculiarity of

the chlorophyll biodegradation in autumnal leaves will be described.

© 2013 Djapic; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

Nina Djapic

**1. Introduction**

http://dx.doi.org/10.5772/56155


#### **Chapter 2**

### **Chlorophyll Biodegradation**

#### Nina Djapic

[41] Tallur, P. N, Megadi, V. B, & Ninnekar, H. Z. Biodegradation of cypermethrin by Mi‐

[42] Chen, S, Yang, L, Hu, M, & Liu, J. Biodegradation of fenvalerate and 3-phenoxyben‐ zoic acid by a novel Stenotrophomonas sp. strain ZS-S-01 and its use in bioremedia‐ tion of contaminated soils. Applied Microbiology and Biotechnology.(2011). , 90(2),

[43] Chen, S, Zhang, Y, Hu, M, Geng, P, Li, Y, & An, G. Bioremediation of β-cypermethrin and 3-phenoxybenzoic acid in soils. In Water Resource and Environmental Protec‐

[44] Mccoy, M. R, Yang, Z, Fu, X, Ahn, K. C, Gee, S. J, Bom, D. C, Zhong, P, Chang, D, & Hammock, B. D. Monitoring of total type II pyrethroid pesticides in Citrus oils and water by converting to a common product phenoxybenzoic acid. Journal of Agricul‐

[45] Chen, S, Hu, Q, Hu, M, Luo, J, Weng, Q, & Lai, K. Isolation and characterization of a fungus able to degrade pyrethroids and phenoxybenzaldehyde. Bioresource Technol‐

[46] Mackenzie, C, Choudhary, M, Larimer, F. W, Predki, P. F, Stilwagen, S, Armitage, J. P, Barber, R. D, Donohue, T. J, Hosler, J. P, & Newman, J. E. The home stretch, a first analysis of the nearly completed genome of Rhodobacter sphaeroides 2.4. 1. Photo‐

[47] Barber, R. D, & Donohue, T. J. Function of a glutathione-dependent formaldehyde dehydrogenase in Rhodobacter sphaeroides formaldehyde oxidation and assimila‐

[48] Armitage, J. P, & Schmitt, R. Bacterial chemotaxis: Rhodobacter sphaeroides and Si‐ norhizobium meliloti variations on a theme? Microbiology Reading.(1997). , 143,

[49] Porter, S. L, Wadhams, G. H, & Armitage, J. P. Signal processing in complex chemo‐

[50] Kojadinovic, M, Sirinelli, A, Wadhams, G. H, & Armitage, J. P. New motion analysis system for characterization of the chemosensory response kinetics of Rhodobacter sphaeroides under different growth conditions. Applied and Environmental Microbi‐

taxis pathways. Nature Reviews Microbiology.(2011). , 9(3), 153-165.

tion (ISWREP), International Symposium on, IEEE), (2011). , 3, 1717-1721.

crococcus sp. strain CPN 1. Biodegradation.(2008). , 19(1), 77-82.

755-767.

16 Biodegradation of Hazardous and Special Products

ogy.(2011). , 3.

3671-3682.

tural and Food Chemistry.(2012). , 3.

synthesis Research.(2001). , 70(1), 19-41.

tion. Biochemistry.(1998). , 37(2), 530-537.

ology.(2011). , 77(12), 4082-4088.

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56155

#### **1. Introduction**

One of the most important biochemical processes is photosynthesis and life depends on the conversion of the solar energy into a chemical one. It occurs in photoautotrophic or‐ ganisms. The chlorophyll *a* is supported by chlorophyll *b*, carotenes and xanthophylls in the light capture. The chlorophyll *a* and accessory pigments are found in cell organelles called chloroplasts. The conversion of light energy into the chemical one can be described thermodynamically. The entropy of photosynthesis yields the energy of 27.32 kJ mol-1 and is a source of energy for the plants. The atmospheric oxygen is a product of the oxygenic photosynthesis, the product of the first phase process known as light – dependant reac‐ tions. In those reactions, the chlorophylls, in the so called reaction centre, function as an absorbent of a photon initiating the electron transport chain, with the loss of an electron. During the electron flow the energy is produced and the reduced form of the nicotina‐ mide adenine dinucleotide phosphate (NADPH) and the adenosine triphosphate (ATP) are formed. The chlorophyll molecule regains the electron from water. The other phase of photosynthesis is called the light – independent reactions or the Calvin cycle. In those re‐ actions the enzyme ribulose-1, 5-bisphosphate carboxylase/oxygenase (RuBisCO) captures the CO2 from the atmosphere and during the Calvin cycle glucose is formed. The light-ab‐ sorbing green pigments, the chlorophylls, are biodegraded under various conditions: with the leaf senescence in autumn, under stress conditions, such as, drought, flooding, chill‐ ing, mechanical wounding, etc., under the exposure to the ethylene and after the treat‐ ment of plants with herbicides. In autumn deciduous trees shed tones of leaves. The green leaves change the colour due to the chlorophyll biodegradation. The peculiarity of the chlorophyll biodegradation in autumnal leaves will be described.

© 2013 Djapic; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

#### **2. Chlorophyll biodegradation**

The chlorophyll *a* and chlorophyll *b* biodegradation pathways consist of a great number of steps and up to now only the steps where the chlorophylls' biodegradation products have a chromophore that can absorb the ultra-violet (UV)/visible light are known. Each step can be considered separately. In the cell, every step is coordinated, highly regulated and most steps are enzymatically catalyzed. The first group of reactions includes the loss of magnesium, the loss of phytol and the modifications on the periphery of the chlorophyll nucleus in which the aromatic tetrapyrrole macrocycle remains intact [1]. The starting investigations on chloro‐ phylls and chlorophylls' biodegradation products were done by Richard Willstaetter and Arthur Stoll who have discovered the nature of the chlorophylls [2]. Ivan Parfen'ievitch Borodin was the first who obtained crystalline chlorophyll [2]. Finally, Mikhail Semenovitch Tswett, after the formulation of the chromatography method, proved that the chlorophyll was a mixture of two green pigments "chlorophyllins alfa and beta" and that Ivan Parfen'ievitch Borodin's pigment is different from natural chlorophylls [3, 4, 5, 6]. Richard Willstaetter and Arthur Stoll showed that green leaves contain an enzyme chlorophyllase which hydrolyses the phytyl ester of the chlorophylls' propionate group to yield chlorophyllide *a* [2, 7, 8]. In the presence of alcohol, the crystalline substance formed by transesterification, is methyl chloro‐ phyllide [9]. Richard Willstaetter and Arthur Stoll determined the basic structure of chloro‐ phyll *a* and *b* and showed that Ivan Parfen'ievitch Borodin's crystalline pigments were chlorophyllide *a* and *b* formed from natural chlorophylls by the hydrolysis by the enzyme chlorophyllase [10].

The interconversion of chlorophyll *b* to chlorophyll *a* and vice versa occurs in oxygenic photosynthetic organisms [11]. The suggested chlorophyll cycle is depicted in Figure 1. The cycle starts with the reduction of chlorophyll *b* by chlorophyll *b* reductase, an NADPH dependant enzyme, to form 71 –hydroxyl chlorophyll *a*. The 71 –hydroxyl chlorophyll *a* is a stable intermediate and it was isolated from higher plants. The next reduction step is catalyzed by ferredoxin dependant 71 –hydroxyl chlorophyll *a* reductase to form chlorophyll *a*. It is also suggested that chlorophyll *a* interconverts to chlorophyll *b* by an oxygen dependant enzyme chlorophyll *a* oxygenase. The enzyme oxidizes chlorophyll *a* to 71 –hydroxyl chlorophyll *a* which is further oxidized to chlorophyll *b* by 71 –hydroxyl chlorophyll *a* dehydrogenase.

The chlorophyll biodegradation continues with dechelating the chlorophyllide *a* molecule. The enzyme involved in demetallation is Mg–dechelatase. A catalytic cofactor called Mg–deche‐ lating substance (MDS) is associated with the activity of Mg–dechelatase [13]. When the magnesium is expelled from the core of chlorophyllide *a*, the remaining structure is called

Fdred

Chl *a*

H2O

71

Fdred

O2

Fdox

Fdox H2O

71 - hydroxy Chl *a* reductase

hlide *a* Chlide *b*


71

Chl *a* oxygenase

 - hydroxyl Chl *a* reductase

71


71

 - hydroxyl Chl *a*  dehydrogenase


H2O

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 19

NADPH

NADP+

Chl *b* reductase 71

NADP+

Chl *b*

Chl *b* reductase NADPH

chlorophyllase

Phytol

The next group of reactions involves the cleavage of the macrocyclic aromatic ring system yielding open chain tetrapyrrolic compound, the *seco*–phytoporhyrins, in which side chains can be further modified. In further chlorophyll biodegradation, pheophorbide *a* (1) methene bridge at the C4–C5 position is oxidized by the pheophorbide *a* oxygenase (P*a*O) to yield the 4,5-dioxo-*seco*-pheophorbide *a* chlorophyll biodegradation product (2). The P*a*O is an iron– dependant monooxygenase. The electrons required for the redox cycle are supplied from the

The proposed mechanism is based on the single oxygen attachment to the double bond at the C4=C5 position forming regioselective intermediate by the cleavage of the oxirane ring.

pheophorbide *a* (1) (Figure 1).

H2O

Phytol

chlorophyllase

C

2H+

Mg2+

Mg dechelatase (MDS)

Pheide a **1**

**Figure 1.** The chlorophyll cycle

reduced ferredoxin (Figure 2).

The bioconversion of chlorophyll *b* to chlorophyll *a* undergoes through a 71 –hydroxyl inter‐ mediate. The chlorophyll interconversion cycle can be expanded to the following catabolic steps toward the chlorophyllide *a* (Figure 1). The chlorophyll *a* is enzymatically hydrolysed to chlorophyllide *a*. The chlorophyll *b* is enzymatically hydrolysed to chlorophyllide *b* and undergoes through the formation of 71 –hydroxyl chlorophyllide *a* which is enzymatically reduced to the chlorophyllide *a* [12].

Richard Willstaetter observed that the ash from the crude chlorophylls is rich in magnesia [2]. The magnesium is cleaved from the chlorophylls by a treatment with mild acids. The magne‐ sium coordination bond is stable against alkali [9]. On the other hand, treatment with hydrox‐ ide saponifies the chlorophylls' phytol side chain yielding chlorophyllides.

**Figure 1.** The chlorophyll cycle

**2. Chlorophyll biodegradation**

18 Biodegradation of Hazardous and Special Products

chlorophyllase [10].

dependant enzyme, to form 71

which is further oxidized to chlorophyll *b* by 71

undergoes through the formation of 71

reduced to the chlorophyllide *a* [12].

by ferredoxin dependant 71

The chlorophyll *a* and chlorophyll *b* biodegradation pathways consist of a great number of steps and up to now only the steps where the chlorophylls' biodegradation products have a chromophore that can absorb the ultra-violet (UV)/visible light are known. Each step can be considered separately. In the cell, every step is coordinated, highly regulated and most steps are enzymatically catalyzed. The first group of reactions includes the loss of magnesium, the loss of phytol and the modifications on the periphery of the chlorophyll nucleus in which the aromatic tetrapyrrole macrocycle remains intact [1]. The starting investigations on chloro‐ phylls and chlorophylls' biodegradation products were done by Richard Willstaetter and Arthur Stoll who have discovered the nature of the chlorophylls [2]. Ivan Parfen'ievitch Borodin was the first who obtained crystalline chlorophyll [2]. Finally, Mikhail Semenovitch Tswett, after the formulation of the chromatography method, proved that the chlorophyll was a mixture of two green pigments "chlorophyllins alfa and beta" and that Ivan Parfen'ievitch Borodin's pigment is different from natural chlorophylls [3, 4, 5, 6]. Richard Willstaetter and Arthur Stoll showed that green leaves contain an enzyme chlorophyllase which hydrolyses the phytyl ester of the chlorophylls' propionate group to yield chlorophyllide *a* [2, 7, 8]. In the presence of alcohol, the crystalline substance formed by transesterification, is methyl chloro‐ phyllide [9]. Richard Willstaetter and Arthur Stoll determined the basic structure of chloro‐ phyll *a* and *b* and showed that Ivan Parfen'ievitch Borodin's crystalline pigments were chlorophyllide *a* and *b* formed from natural chlorophylls by the hydrolysis by the enzyme

The interconversion of chlorophyll *b* to chlorophyll *a* and vice versa occurs in oxygenic photosynthetic organisms [11]. The suggested chlorophyll cycle is depicted in Figure 1. The cycle starts with the reduction of chlorophyll *b* by chlorophyll *b* reductase, an NADPH

–hydroxyl chlorophyll *a*. The 71

–hydroxyl chlorophyll *a* reductase to form chlorophyll *a*. It is also

–hydroxyl chlorophyll *a* dehydrogenase.

–hydroxyl chlorophyllide *a* which is enzymatically

stable intermediate and it was isolated from higher plants. The next reduction step is catalyzed

suggested that chlorophyll *a* interconverts to chlorophyll *b* by an oxygen dependant enzyme

mediate. The chlorophyll interconversion cycle can be expanded to the following catabolic steps toward the chlorophyllide *a* (Figure 1). The chlorophyll *a* is enzymatically hydrolysed to chlorophyllide *a*. The chlorophyll *b* is enzymatically hydrolysed to chlorophyllide *b* and

Richard Willstaetter observed that the ash from the crude chlorophylls is rich in magnesia [2]. The magnesium is cleaved from the chlorophylls by a treatment with mild acids. The magne‐ sium coordination bond is stable against alkali [9]. On the other hand, treatment with hydrox‐

chlorophyll *a* oxygenase. The enzyme oxidizes chlorophyll *a* to 71

The bioconversion of chlorophyll *b* to chlorophyll *a* undergoes through a 71

ide saponifies the chlorophylls' phytol side chain yielding chlorophyllides.

–hydroxyl chlorophyll *a* is a

–hydroxyl chlorophyll *a*

–hydroxyl inter‐

The chlorophyll biodegradation continues with dechelating the chlorophyllide *a* molecule. The enzyme involved in demetallation is Mg–dechelatase. A catalytic cofactor called Mg–deche‐ lating substance (MDS) is associated with the activity of Mg–dechelatase [13]. When the magnesium is expelled from the core of chlorophyllide *a*, the remaining structure is called pheophorbide *a* (1) (Figure 1).

The next group of reactions involves the cleavage of the macrocyclic aromatic ring system yielding open chain tetrapyrrolic compound, the *seco*–phytoporhyrins, in which side chains can be further modified. In further chlorophyll biodegradation, pheophorbide *a* (1) methene bridge at the C4–C5 position is oxidized by the pheophorbide *a* oxygenase (P*a*O) to yield the 4,5-dioxo-*seco*-pheophorbide *a* chlorophyll biodegradation product (2). The P*a*O is an iron– dependant monooxygenase. The electrons required for the redox cycle are supplied from the reduced ferredoxin (Figure 2).

The proposed mechanism is based on the single oxygen attachment to the double bond at the C4=C5 position forming regioselective intermediate by the cleavage of the oxirane ring.

1

**<sup>2</sup>** C82

Non - enzymatic

NH HN

<sup>O</sup> MeO2C

NH HN

<sup>O</sup> MeO2C

COOH

NH HN

OHC O

Fdox

Fdred

COOH

NH HN

O OHC

H

hydroxylation

N HN

COOH

tautomerization Non - enzymatic

COOH

COOH

ene bridge forming the chlorophyll biodegradation products 8, 9 and 10.

**8 9**

MeO2C

**5 6**

NADPH

NADP+

MeO2C

NH HN

O

**Figure 3.** The chlorophyll biodegradation pathway in Hamamelidaceae and Vitaceae autumnal leaves. The ethyl side chain of the 4, 5-dioxo-*seco*-pheophorbide *a* (2) is enzymatically hydroxylated to the compound 3 and after the pro‐ posed Baeyer-Villiger oxidation the urobilinogenic chlorophyll biodegradation product (4) is formed. All chlorophyll biodegradaton products under acid conditions form the thermodynamically more stable compounds (5, 6, 7). The re‐ duction of the chlorophyll biodegradation compounds 5, 6 and 7 proceeds via the reduction of the "western" meth‐

NH HN

OHC O

Fdox

Fdred

MeO2C

NH HN

O

NADPH

NADP+

NH HN

OHC O

O

NH HN

OHC O

OH

proposed Baeyer-Villiger oxidation

**3 4**

OH

OH

H H

5 15

NH HN

O O

OH

21

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155

N HN

COOH

tautomerization Non - enzymatic

COOH

COOH

MeO2C

MeO2C

**7**

NH HN

O

**10**

NH HN

O O

Fdox

Fdred

MeO2C

O

NADPH

NADP+

NH HN

NH HN

O O

O

tautomerization

OH

OH

**Figure 2.** The proposed mechanism of the pheophorbide *a* (1) oxidation, by the P*a*O to the 4,5-dioxo- *seco*-pheophor‐ bide *a* (2).

Subsequently, the oxiran ring is opened hydrolytically. The retro–aldol reaction proceeds and the rearrangement of protons form the 4,5-dioxo-*seco*-pheophorbide *a* (2) (Figure 2).

The amount of accumulated chlorophyll biodegradation products within the same plant species varies. The quantity of the chlorophyll biodegradation products in senescent leaves most probably depends on the time of the collection, seasonal climate, developmental stage of the plant, enzymes, etc. One of the reasons for the chlorophyll biodegradation in biological systems is due to the tendency of the organism to decrease the level of photodynamically active chlorophyll before the programmed cell death [14].

#### **3. Chlorophyll biodegradation in Hamamelidaceae and Vitaceae autumnal leaves**

The autonomous induction of leaf senescence occurs in autumn. The major consequence of leaf senescence is chlorophyll biodegradation. The crude methanol extracts of Hamamelidaceae and Vitaceae autumnal leaves' plant species were analyzed on the reversed phase (RP): RP– C8 and RP–C4 analytical columns by liquid chromatography–mass spectrometry (LC–MS) and their structure was determined by mass and nuclear magnetic resonance (NMR) spectra [15, 16, 17]. The results obtained gave insight in the chlorophyll biodegradation in plant species of Hamamelidaceae and Vitaceae autumnal leaves (Figure 3). The chlorophyll biodegradation pathway continues from the 4, 5-dioxo-*seco*-pheophorbide *a* (2) to the formation of the hydroxylated ethyl side chain chlorophyll biodegradation product (3). The enzyme catalyzing the hydroxylation of the ethyl side chain is still unknown.

Subsequently, the oxiran ring is opened hydrolytically. The retro–aldol reaction proceeds and

**Figure 2.** The proposed mechanism of the pheophorbide *a* (1) oxidation, by the P*a*O to the 4,5-dioxo- *seco*-pheophor‐

The amount of accumulated chlorophyll biodegradation products within the same plant species varies. The quantity of the chlorophyll biodegradation products in senescent leaves most probably depends on the time of the collection, seasonal climate, developmental stage of the plant, enzymes, etc. One of the reasons for the chlorophyll biodegradation in biological systems is due to the tendency of the organism to decrease the level of photodynamically active

**3. Chlorophyll biodegradation in Hamamelidaceae and Vitaceae autumnal**

The autonomous induction of leaf senescence occurs in autumn. The major consequence of leaf senescence is chlorophyll biodegradation. The crude methanol extracts of Hamamelidaceae and Vitaceae autumnal leaves' plant species were analyzed on the reversed phase (RP): RP– C8 and RP–C4 analytical columns by liquid chromatography–mass spectrometry (LC–MS) and their structure was determined by mass and nuclear magnetic resonance (NMR) spectra [15, 16, 17]. The results obtained gave insight in the chlorophyll biodegradation in plant species of Hamamelidaceae and Vitaceae autumnal leaves (Figure 3). The chlorophyll biodegradation pathway continues from the 4, 5-dioxo-*seco*-pheophorbide *a* (2) to the formation of the hydroxylated ethyl side chain chlorophyll biodegradation product (3). The enzyme catalyzing

the rearrangement of protons form the 4,5-dioxo-*seco*-pheophorbide *a* (2) (Figure 2).

chlorophyll before the programmed cell death [14].

the hydroxylation of the ethyl side chain is still unknown.

**leaves**

bide *a* (2).

H

N N

D C

**1**

E

MeO2C

COOH

1

O

O2

2e-

H2O <sup>P</sup>*a*<sup>O</sup>

Fe2+ Fe3+

Fdox Fdred

NADPH

NADP+

N HN

D C

E

MeO2C

**2** COOH

O

NH HN

OHC <sup>O</sup>

<sup>A</sup> <sup>B</sup>

NH N

<sup>10</sup> <sup>20</sup>

A B

5

20 Biodegradation of Hazardous and Special Products

**Figure 3.** The chlorophyll biodegradation pathway in Hamamelidaceae and Vitaceae autumnal leaves. The ethyl side chain of the 4, 5-dioxo-*seco*-pheophorbide *a* (2) is enzymatically hydroxylated to the compound 3 and after the pro‐ posed Baeyer-Villiger oxidation the urobilinogenic chlorophyll biodegradation product (4) is formed. All chlorophyll biodegradaton products under acid conditions form the thermodynamically more stable compounds (5, 6, 7). The re‐ duction of the chlorophyll biodegradation compounds 5, 6 and 7 proceeds via the reduction of the "western" meth‐ ene bridge forming the chlorophyll biodegradation products 8, 9 and 10.

The urobilinogenic chlorophyll biodegradaton products isolated from autumnal leaves of *Parrotia persica* and *Hamamelis virginiana*, Hamamelidaceae significantly differ from other isolated chlorophyll biodegradation products in one functional group [15, 16]. The aldehyde group at C5 position is absent and chlorophyll biodegradation product's structure refers to the urobilinogen. The aldehyde group at the C5 (3) position might be oxidized by Baeyer– Villiger monooxygenase. The carbon numeration differs from all previously isolated chloro‐ phyll biodegradation products and is as in urobilinogen (Figure 3) [15, 16].

The proposed oxidation step is by the Baeyer–Villiger monooxygenase, from the C82 -hydroxy‐ lated chlorophyll biodegradation product (3) to the urobilinogenic chlorophyll biodegradation product (4). The oxidation results in the formation of an additional chiral centre at the position C4. The absolute configuration of all chiral centres is unknown. It is proposed that this enzyme can convert the chlorophyll biodegradation product's aldehyde group into the corresponding dehydro pyrrolidin–2–one of the urobilinogenic chlorophyll biodegradation product mediat‐ ed by the hydroperoxide intermediate. The nucleophilic addition of the hydroperoxide to the C5 oxo group affords peroxide intermediate. After the rearrangement, the insertion of the oxygen is made. After the ester hydrolysis, the alcohol is formed [17, 18, 19]. After the acidic catalyzed tautomerization, the urobilinogenic chlorophyll biodegradation product is formed. Another conceivable mechanism that can explain the formation of the urobilinogenic chloro‐ phyll biodegradation product is by the oxidative decarboxylation of an acid intermediate generated by the oxidation of the aldehyde group.

The chlorophyll biodegradation products (2, 3, 4) form the thermodynamically more stable compounds (5, 6, 7) after the acidic catalyzed tautomerization, [20]. The reduction of the compounds 5, 6 and 7 proceed via the reduction of the "western" methene bridge. It is catalyzed by a reductase. The reductase is a ferredoxin dependant enzyme with no require‐ ments of cofactors such as flavin or metals. After the reduction of the chlorophyll biodegra‐ dation products 5, 6, 7 the chlorophyll biodegradation products 8, 9 and 10 are formed.

Further chlorophyll biodegradation products were, up to now, not detected. The further chlorophyll biodegradation products lose the chromophore that can absorb the UV light, they become colourless, and were not identified up to now.

#### **4. Chlorophyll biodegradation in Solanaceae autumnal leaves**

In autumnal leaveas of the Solanaceae family: tobacco (*Nicotiana tabacum*), potatoe (*Solanum tuberosum*), *Atropa belladonna*, etc. chlorophyll biodegradation products are present. In Sola‐ naceae plant species the 4, 5-dioxo-*seco*-pheophorbide *a* (2) precedes, in further chlorophyll biodegradation, the hydroxylation at the C82 position and then the glycosylation at the C82 position (11) [21]. The glycosyl bond is most probably introduced by glycosyltransferase. The Solanaceae chlorophyll biodegradation product (11) form a thermodynamically stable aromatic ring D under the acidic conditions (12) (Figure 4). The reduction of the thermody‐ namically more stable compound (12) proceeds via the reduction of the "western" methene

bridge. The reductase catalyzed reaction proceeds and the glycosylated chlorophyll biode‐

**Figure 4.** The chlorophyll biodegradation pathway in Solanaceae autumnal leaves, known up to now.

The detection of other chlorophyll biodegradation products present in Solanaceae autumnal leaves will extend the chlorophyll biodegradation pathway, known up to now. The isolation of the sufficient quantity of the Solanaceae glycosylated chlorophyll biodegradation products

gradation product 13 is formed.

C

**2**

82

<sup>H</sup> <sup>H</sup>

**5 6**

**8 9**

Non - enzymatic

F

dox

F

dred NAD

NAD

P+

P

H

hydroxylation

tautomerization Non - enzymatic

tautomerization

F

dox

F

dred

NADP

NADP+

H

**3**

Glycosylation

N HN

MeO2C O

**11**

H

NH HN

MeO2C O

NH HN

MeO2C O

**13**

COOH

NH HN

OHC

O

**12**

Fdox

Fdred NADPH

NADP+

COOH

NH HN

OHC

O

Non - enzymatic tautomerization

COOH

NH HN

OHC

O

OGlc

23

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155

OGlc

OGlc

The urobilinogenic chlorophyll biodegradaton products isolated from autumnal leaves of *Parrotia persica* and *Hamamelis virginiana*, Hamamelidaceae significantly differ from other isolated chlorophyll biodegradation products in one functional group [15, 16]. The aldehyde group at C5 position is absent and chlorophyll biodegradation product's structure refers to the urobilinogen. The aldehyde group at the C5 (3) position might be oxidized by Baeyer– Villiger monooxygenase. The carbon numeration differs from all previously isolated chloro‐

lated chlorophyll biodegradation product (3) to the urobilinogenic chlorophyll biodegradation product (4). The oxidation results in the formation of an additional chiral centre at the position C4. The absolute configuration of all chiral centres is unknown. It is proposed that this enzyme can convert the chlorophyll biodegradation product's aldehyde group into the corresponding dehydro pyrrolidin–2–one of the urobilinogenic chlorophyll biodegradation product mediat‐ ed by the hydroperoxide intermediate. The nucleophilic addition of the hydroperoxide to the C5 oxo group affords peroxide intermediate. After the rearrangement, the insertion of the oxygen is made. After the ester hydrolysis, the alcohol is formed [17, 18, 19]. After the acidic catalyzed tautomerization, the urobilinogenic chlorophyll biodegradation product is formed. Another conceivable mechanism that can explain the formation of the urobilinogenic chloro‐ phyll biodegradation product is by the oxidative decarboxylation of an acid intermediate

The chlorophyll biodegradation products (2, 3, 4) form the thermodynamically more stable compounds (5, 6, 7) after the acidic catalyzed tautomerization, [20]. The reduction of the compounds 5, 6 and 7 proceed via the reduction of the "western" methene bridge. It is catalyzed by a reductase. The reductase is a ferredoxin dependant enzyme with no require‐ ments of cofactors such as flavin or metals. After the reduction of the chlorophyll biodegra‐ dation products 5, 6, 7 the chlorophyll biodegradation products 8, 9 and 10 are formed.

Further chlorophyll biodegradation products were, up to now, not detected. The further chlorophyll biodegradation products lose the chromophore that can absorb the UV light, they

In autumnal leaveas of the Solanaceae family: tobacco (*Nicotiana tabacum*), potatoe (*Solanum tuberosum*), *Atropa belladonna*, etc. chlorophyll biodegradation products are present. In Sola‐ naceae plant species the 4, 5-dioxo-*seco*-pheophorbide *a* (2) precedes, in further chlorophyll

position (11) [21]. The glycosyl bond is most probably introduced by glycosyltransferase. The Solanaceae chlorophyll biodegradation product (11) form a thermodynamically stable aromatic ring D under the acidic conditions (12) (Figure 4). The reduction of the thermody‐ namically more stable compound (12) proceeds via the reduction of the "western" methene

position and then the glycosylation at the C82

**4. Chlorophyll biodegradation in Solanaceae autumnal leaves**


phyll biodegradation products and is as in urobilinogen (Figure 3) [15, 16].

generated by the oxidation of the aldehyde group.

22 Biodegradation of Hazardous and Special Products

become colourless, and were not identified up to now.

biodegradation, the hydroxylation at the C82

The proposed oxidation step is by the Baeyer–Villiger monooxygenase, from the C82

**Figure 4.** The chlorophyll biodegradation pathway in Solanaceae autumnal leaves, known up to now.

bridge. The reductase catalyzed reaction proceeds and the glycosylated chlorophyll biode‐ gradation product 13 is formed.

The detection of other chlorophyll biodegradation products present in Solanaceae autumnal leaves will extend the chlorophyll biodegradation pathway, known up to now. The isolation of the sufficient quantity of the Solanaceae glycosylated chlorophyll biodegradation products

All chromatograms obtained revealed the presence of three peaks, the UV detection at λ=312 nm. One peak had no Gaussian peak resolution (*Gb*-1), other two peaks coeluted with no

methanol on RP C4 – analytical column (Figure 6). All peaks had the *m/z* 635. The ESIMS of the

**Figure 6.** The chromatogram of *Gingko biloba* male methanol autumnal leaves' crude extract. The LC conditions: Col‐ umn: Nucleosil 100-5 C4 4x250 mm. The mobile phase: 90% v/v water (0.1% TFA):methanol to 0% v/v water

The semi-preparative separation on one HPLC instrument on the semi-preparative RP-C<sup>4</sup> column with water (0.1% TFA): methanol solvent mixture gradient elution revealed the chromatogram (extracted at λ=312 nm) with complete peak distortion of *Ginkgo biloba* chlorophyll biodegradation products (Figure 8). The peak distortion was probably due to compounds' secondary equilibrium. It appears that the conversion rate of *Ginkgo biloba* compounds increased during the separation under the applied separation conditions. The

C with gradient elution water (0.1% TFA):

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 25

separation to the baseline (*Gb*-2 and *Gb*-3) at the 25<sup>o</sup>

*Gb*-3 eluting at the 37.8 min. is depicted in Figure 7.

(0.1%TFA):methanol in 80 minutes. Flow rate: 0.2 ml/min. UV detection at λ=312.

**Figure 7.** The ESIMS of the *Gb*-3 eluting at the 37.8 min.

**Figure 5.** The proton spectrum of the compound 9.

will provide the information about the conformation of the sugar part and the information regarding the linkage between the aglycone and the sugar, such as, in the case of the antho‐ cynins [22, 23, 24, 25].

#### **5. Chlorophyll biodegradation in** *Ginkgo biloba* **autumnal leaves**

Crude methanol extract of autumnal leaves' of *Ginkgo biloba*, Ginkgoaceae male plant were analyzed by LC-ESI-MS and LC/ESI-MS chromatograms and spectra obtained revealed the presence of the chlorophyll biodegradation products. Of hundreds of samples analyzed, only in one chromatogram obtained the traces the chlorophyll biodegradation product 9 with the molecular ion at *m/z* 645 for the molecular formula C35H40N4O8 was observed. Its proton NMR spectrum is depicted in Figure 5. Integration of the proton NMR spectrum revealed the presence of 33 protons. The signal of one proton on the heteroatom was still not exchanged with deuterium and was present in the low field region. The aldehyde H-5 proton signal was present in the low field region. Three low field *doublet of doublets* sets in the spectrum indicated the presence of an AMX spin-splitting pattern. The proton signal H-15 was overlapped with the HDO signal. Propionyl side chain H-171 protons came in a narrow δ range along with the methylene bridge H-20 protons. The H-172 protons, next to the electron withdrawing carboxyl group, had chemical shifts in the upfield region compared to H-171 protons. The most prominent peak in the proton spectrum was the signal assigned to the methoxycarbonyl group at the position 82 . The methyl groups and other protons were assigned from the "1D difference" NOE spectra. In the COSY spectrum, the well resolved downfield 32 HM vinyl proton was assigned to the geminal 32 HA proton tracing further to the vicinal 31 HX proton. This vinyl isolated spin system terminated at the 31 HX proton.

All chromatograms obtained revealed the presence of three peaks, the UV detection at λ=312 nm. One peak had no Gaussian peak resolution (*Gb*-1), other two peaks coeluted with no separation to the baseline (*Gb*-2 and *Gb*-3) at the 25<sup>o</sup> C with gradient elution water (0.1% TFA): methanol on RP C4 – analytical column (Figure 6). All peaks had the *m/z* 635. The ESIMS of the *Gb*-3 eluting at the 37.8 min. is depicted in Figure 7.

**Figure 6.** The chromatogram of *Gingko biloba* male methanol autumnal leaves' crude extract. The LC conditions: Col‐ umn: Nucleosil 100-5 C4 4x250 mm. The mobile phase: 90% v/v water (0.1% TFA):methanol to 0% v/v water (0.1%TFA):methanol in 80 minutes. Flow rate: 0.2 ml/min. UV detection at λ=312.

**Figure 7.** The ESIMS of the *Gb*-3 eluting at the 37.8 min.

will provide the information about the conformation of the sugar part and the information regarding the linkage between the aglycone and the sugar, such as, in the case of the antho‐

9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 2.0 Chemical Shift (ppm)

3 1 H A

3 1 H X

OH

NH HN

MeO2C O

**9**

**Figure 5.** The proton spectrum of the compound 9.

COOH

17

24 Biodegradation of Hazardous and Special Products

1

NH HN

O OHC

5

10

3 2 H M

Crude methanol extract of autumnal leaves' of *Ginkgo biloba*, Ginkgoaceae male plant were analyzed by LC-ESI-MS and LC/ESI-MS chromatograms and spectra obtained revealed the presence of the chlorophyll biodegradation products. Of hundreds of samples analyzed, only in one chromatogram obtained the traces the chlorophyll biodegradation product 9 with the molecular ion at *m/z* 645 for the molecular formula C35H40N4O8 was observed. Its proton NMR spectrum is depicted in Figure 5. Integration of the proton NMR spectrum revealed the presence of 33 protons. The signal of one proton on the heteroatom was still not exchanged with deuterium and was present in the low field region. The aldehyde H-5 proton signal was present in the low field region. Three low field *doublet of doublets* sets in the spectrum indicated the presence of an AMX spin-splitting pattern. The proton signal H-15 was overlapped with

group, had chemical shifts in the upfield region compared to H-171 protons. The most prominent peak in the proton spectrum was the signal assigned to the methoxycarbonyl group

NOE spectra. In the COSY spectrum, the well resolved downfield 32 HM vinyl proton was assigned to the geminal 32 HA proton tracing further to the vicinal 31 HX proton. This vinyl

HX proton.

. The methyl groups and other protons were assigned from the "1D difference"

protons came in a narrow δ range along with the

H-17 2 (2H)

> H A-20

H-8 1 (2H)H -17

H


B

H -17

1

B

1

A

H-10 (2H) H-13 2-OCH 3

> H-13 2

H-8

2(2H)

H-1

protons, next to the electron withdrawing carboxyl

**5. Chlorophyll biodegradation in** *Ginkgo biloba* **autumnal leaves**

cynins [22, 23, 24, 25].

H-5


the HDO signal. Propionyl side chain H-171

methylene bridge H-20 protons. The H-172

isolated spin system terminated at the 31

at the position 82

The semi-preparative separation on one HPLC instrument on the semi-preparative RP-C<sup>4</sup> column with water (0.1% TFA): methanol solvent mixture gradient elution revealed the chromatogram (extracted at λ=312 nm) with complete peak distortion of *Ginkgo biloba* chlorophyll biodegradation products (Figure 8). The peak distortion was probably due to compounds' secondary equilibrium. It appears that the conversion rate of *Ginkgo biloba* compounds increased during the separation under the applied separation conditions. The

**Figure 8.** The chromatogram of *Ginkgo biloba* autumnal leaves, UV detection at λ=312 nm when the separation was done on one older model of the HPLC instrument

The compound eluting at 42 min. is assigned as the *Gb*-1, at the 44 min. as the *Gb*-2 and at the 45 min. as the *Gb*-3. The determination of *Ginkgo biloba* chlorophyll biodegradation products was done by mass spectrometry and NMR spectroscopy. The High Resolution Electron Spray Ionisation Mass Spectrometry (HRESIMS) of the *Gb*-1 showed a molecular ion at *m/z* 679.2350 for the molecular formula C33H37N4O9Na2 [M+2Na]2+, calculated *m /z* 679.2350, Δ 0.00 ppm. The

**Figure 10.** The chromatogram of *Ginkgo biloba* leaves' methanol crude extract. The LC conditions: Column: Nucleosil 120-7 C4 10x250 mm. The mobile phase: 90% v/v water (acetate buffer): 10% v/v methanol to 0% v/v water (acetate

0 10 20 30 40 50 Time [min]

657.2531, Δ + 0.15 ppm. The formic acid was added to the sample and the HRESIMS was recorded. The mass spectrum obtained revealed the presence of *m/z* 679.2351 and *m/z* 657.2530. The *Gb*-2 had a molecular ion at *m/z* 679.2350 for the molecular formula C33H37N4O9Na2 [M +2Na]2+, calculated *m /z* 679.2750, Δ 0.00 ppm. The other molecular ion had *m/z* 701.2172 for the molecular formula C33H36N4O9Na3 [M+3Na]3+, calculated *m/z* 701.2170, Δ– 0.15 ppm. The addition of formic acid to the sample revealed the presence of the peaks *m/z* 657.2531, *m/z* 679.2347 and *m/z* 701.2166. For the *Gb*-3 compound, the Electron Spray Ionisation Mass Spectrometry (ESIMS) revealed the presence of three peaks, *m /z* 679.24, 701.22 and 723.20 when the sample was dissolved in methanol. The sample dissolved in methanol with formic acid

The proton NMR spectra were recorded. In the *Gb*-1 proton spectrum signals for the aldehyde proton were present, two well resolved *singlets* and one *pseudo triplet* (Figure 11). The *Gb*-2 and

The formyl group can have two conformations NO – *Z* (14) and NO – *E* (15) (Figure 14) [26, 27]. In all previously isolated *seco* – phytoporphyrins such conformational change has not been

The vinyl group proton signals were of unresolved line shape (Figure 11, 12 and 13). The *doublet of doublets* were fairly broad at the room temperature. The vicinal proton of the vinyl group showed two superpositioned *doublets of doublets*. Similar to the proton spectrum of the chlorophyll biodegradation product 9, in the low field, three *doublet of doublets* sets in the proton spectrum indicated the presence of an AMX spin-splitting pattern (Figure 5, 11, 12 and 13). In the low field two sharp *singlets* were recorded (Figure 11). They were or impurities or the

, calculated *m /z*

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 27

other ion had *m/z* 657.2530 for the molecular formula C33H37N4O9Na [M+Na]+

buffer): 100% v/v methanol at 300C in 90 minutes. Flow rate: 1.2 ml/min. UV detection at 312 nm.

revealed the presence of three peaks *m /z* 657.25, 679.24 and 701.22.

*Gb*-3 had the same signals in the low field (Figure 12 and 13).

observed.

Intens. mAU

**Figure 9.** The chromatogram of *Ginkgo biloba* leaves' methanol crude extract. The LC conditions: Column: Nucleosil 120-7 C4 10x250 mm. The mobile phase: 90% v/v water (acetate buffer): 10% v/v methanol to 0% v/v water (acetate buffer): 100% v/v methanol at 220C in 90 minutes. Flow rate: 1.2 ml/min. UV detection at 312 nm.

peaks became smaller spreading over a long elution time (Figure 8). The UV spectrum of the broad peak from 40 to 42 min. revealed the presence of a chromophore with the absorption band of 317 nm. The *Ginkgo biloba* chlorophyll biodegradation products' peaks were broad and highly distorted.

The separation investigations continued on semi-preparative RP C4 using the latest model of one LC instrument. The solvent mixture used was water (acetate buffer (pH 3.75)): methanol. The temperature was changed (Figure 9 and 10). At 30o C the separation between the peaks was observed (Figure 10). The isolation of *Ginkgo biloba* chlorophyll biodegradation products was done as depicted in Figure 10. Three collected fractions were evaporated under reduced pressure (t<40o C) and MS and NMR spectra were recorded.

**Figure 10.** The chromatogram of *Ginkgo biloba* leaves' methanol crude extract. The LC conditions: Column: Nucleosil 120-7 C4 10x250 mm. The mobile phase: 90% v/v water (acetate buffer): 10% v/v methanol to 0% v/v water (acetate buffer): 100% v/v methanol at 300C in 90 minutes. Flow rate: 1.2 ml/min. UV detection at 312 nm.

The compound eluting at 42 min. is assigned as the *Gb*-1, at the 44 min. as the *Gb*-2 and at the 45 min. as the *Gb*-3. The determination of *Ginkgo biloba* chlorophyll biodegradation products was done by mass spectrometry and NMR spectroscopy. The High Resolution Electron Spray Ionisation Mass Spectrometry (HRESIMS) of the *Gb*-1 showed a molecular ion at *m/z* 679.2350 for the molecular formula C33H37N4O9Na2 [M+2Na]2+, calculated *m /z* 679.2350, Δ 0.00 ppm. The other ion had *m/z* 657.2530 for the molecular formula C33H37N4O9Na [M+Na]+ , calculated *m /z* 657.2531, Δ + 0.15 ppm. The formic acid was added to the sample and the HRESIMS was recorded. The mass spectrum obtained revealed the presence of *m/z* 679.2351 and *m/z* 657.2530. The *Gb*-2 had a molecular ion at *m/z* 679.2350 for the molecular formula C33H37N4O9Na2 [M +2Na]2+, calculated *m /z* 679.2750, Δ 0.00 ppm. The other molecular ion had *m/z* 701.2172 for the molecular formula C33H36N4O9Na3 [M+3Na]3+, calculated *m/z* 701.2170, Δ– 0.15 ppm. The addition of formic acid to the sample revealed the presence of the peaks *m/z* 657.2531, *m/z* 679.2347 and *m/z* 701.2166. For the *Gb*-3 compound, the Electron Spray Ionisation Mass Spectrometry (ESIMS) revealed the presence of three peaks, *m /z* 679.24, 701.22 and 723.20 when the sample was dissolved in methanol. The sample dissolved in methanol with formic acid revealed the presence of three peaks *m /z* 657.25, 679.24 and 701.22.

The proton NMR spectra were recorded. In the *Gb*-1 proton spectrum signals for the aldehyde proton were present, two well resolved *singlets* and one *pseudo triplet* (Figure 11). The *Gb*-2 and *Gb*-3 had the same signals in the low field (Figure 12 and 13).

peaks became smaller spreading over a long elution time (Figure 8). The UV spectrum of the broad peak from 40 to 42 min. revealed the presence of a chromophore with the absorption band of 317 nm. The *Ginkgo biloba* chlorophyll biodegradation products' peaks were broad and

**Figure 9.** The chromatogram of *Ginkgo biloba* leaves' methanol crude extract. The LC conditions: Column: Nucleosil 120-7 C4 10x250 mm. The mobile phase: 90% v/v water (acetate buffer): 10% v/v methanol to 0% v/v water (acetate

0 10 20 30 40 50 Time [min]

Minutes

**Figure 8.** The chromatogram of *Ginkgo biloba* autumnal leaves, UV detection at λ=312 nm when the separation was

10.00 20.00 30.00 40.00 50.00 60.00 70.00 80.00 90.00

The separation investigations continued on semi-preparative RP C4 using the latest model of one LC instrument. The solvent mixture used was water (acetate buffer (pH 3.75)): methanol.

was observed (Figure 10). The isolation of *Ginkgo biloba* chlorophyll biodegradation products was done as depicted in Figure 10. Three collected fractions were evaporated under reduced

C) and MS and NMR spectra were recorded.

buffer): 100% v/v methanol at 220C in 90 minutes. Flow rate: 1.2 ml/min. UV detection at 312 nm.

C the separation between the peaks

The temperature was changed (Figure 9 and 10). At 30o

highly distorted.

UV Chromatogram, 312 nm

0

500

1000

1500

2000 Intens. mAU AU

0.00 0.20 0.40 0.60 0.80 1.00 1.20 1.40

done on one older model of the HPLC instrument

26 Biodegradation of Hazardous and Special Products

pressure (t<40o

The formyl group can have two conformations NO – *Z* (14) and NO – *E* (15) (Figure 14) [26, 27]. In all previously isolated *seco* – phytoporphyrins such conformational change has not been observed.

The vinyl group proton signals were of unresolved line shape (Figure 11, 12 and 13). The *doublet of doublets* were fairly broad at the room temperature. The vicinal proton of the vinyl group showed two superpositioned *doublets of doublets*. Similar to the proton spectrum of the chlorophyll biodegradation product 9, in the low field, three *doublet of doublets* sets in the proton spectrum indicated the presence of an AMX spin-splitting pattern (Figure 5, 11, 12 and 13). In the low field two sharp *singlets* were recorded (Figure 11). They were or impurities or the

**Figure 11.** The enlarged proton spectrum of *Gb*-1 in the low field region

**Figure 12.** The enlarged proton spectrum of *Gb*-2 in the low field region

proton at the "western" methylene bridge. In the *Gb*-2 chlorophyll biodegradation product's proton spectrum in the vinyl region the proposed "western" methylene bridge proton gave one *singlet* (Figure 12). In the *Gb*-3 compound's low field proton spectrum only vinyl signals were recorded (Figure 13).

In all three *Gb* chlorophyll biodegradation products methoxy group of the five membered ring E was not present (Figure 15, 16 and 17).

5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 Chemical Shift (ppm)

6.55 6.50 6.45 6.40 6.35 6.30 6.25 6.20 6.15 6.10 6.05 6.00 5.95 5.90 5.85 5.80 5.75 5.70 5.65 5.60 5.55 5.50 5.45 5.40 5.35 5.30 5.25 5.20 Chemical Shift (ppm)

Vinyl group Vinyl group Vinyl group

**Figure 15.** The proton spectrum of the *Gb*-1 in the high field region

9.45 9.40 9.35 9.30 9.25 9.20 Chemical Shift (ppm)

**Figure 13.** The enlarged proton spectrum of *Gb*-3 in the low field region

H

NO - *Z* **14**

N

<sup>C</sup> <sup>H</sup>

O

**Figure 14.** The conformational change of the pyrrole's aldehyde group

Aldehyde group

*Gb* - 1

N H

NO - *E* **15**

<sup>C</sup> <sup>O</sup>

H

*Gb* - 3

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 29

**Figure 13.** The enlarged proton spectrum of *Gb*-3 in the low field region

**Figure 14.** The conformational change of the pyrrole's aldehyde group

**Figure 15.** The proton spectrum of the *Gb*-1 in the high field region

proton at the "western" methylene bridge. In the *Gb*-2 chlorophyll biodegradation product's proton spectrum in the vinyl region the proposed "western" methylene bridge proton gave one *singlet* (Figure 12). In the *Gb*-3 compound's low field proton spectrum only vinyl signals

6.55 6.50 6.45 6.40 6.35 6.30 6.25 6.20 6.15 6.10 6.05 6.00 5.95 5.90 5.85 5.80 5.75 5.70 5.65 5.60 5.55 5.50 5.45 5.40 5.35 5.30 5.25 5.20 Chemical Shift (ppm)

Vinyl group

*Gb* - 1

In all three *Gb* chlorophyll biodegradation products methoxy group of the five membered ring

were recorded (Figure 13).

E was not present (Figure 15, 16 and 17).

9.45 9.40 9.35 9.30 9.25 9.20 Chemical Shift (ppm)

**Figure 11.** The enlarged proton spectrum of *Gb*-1 in the low field region

**Figure 12.** The enlarged proton spectrum of *Gb*-2 in the low field region

Vinyl group Vinyl group

28 Biodegradation of Hazardous and Special Products

Aldehyde group

**Figure 16.** The proton spectrum of the *Gb*-2 in the high field region

**Figure 17.** The proton spectrum of the *Gb*-3 in the high field region

The signals of the ethylene protons were found in all spectra recorded by the residual water peak. From the residual water peak to the high field region the presence of one *doublet* and one *doublet of doublets* was present. Those signals are, yet, not assigned. The methyl proton region is not similar to any previous chlorophyll biodegration products' high field region (Figure 15, 16 and 17). The chlorophyll biodegradation products found in *Ginkgo biloba* autumnal leaves are different from all previously isolated ones. The vinyl proton-proton connectivity in the COSY spectrum was easily observed (Figure 18). The *Gb*-3 COSY spectrum recorded was the same as the COSY spectrum of the chlorophyll biodegradation product 9 in the low field region. The establishment of other scalar coupled protons was not determined (Figure 19).

The dynamic NMR spectra were not recorded, yet, such as, they were done in case of the

9.5 9.0 8.5 8.0 7.5 7.0 6.5 6.0 5.5 5.0 F2 Chemical Shift (ppm)

4.0

31

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155

4.5

5.0

5.5

6.0

6.5

7.0

F1 Chemical Shift (ppm)

7.5

8.0

8.5

9.0

9.5

The *Ginkgo biloba* chlorophyll biodegradation products were, also, detected in the *Ginkgo*

The screening for the presence of the chlorophyll biodegradation products was done in ferns. Of dozens of ferns' autumnal leaves' methanol extracts analyzed only in *Athyrium filix femina*, Dryopteridaceae the *Ginkgo biloba* chlorophyll biodegradation products were detected (Figure 21). The cornifer plant – *Larix kaempferi*, Pinaceae autumnal leaves' methanol crude

polycyclic quinones, substituted tetrahydropyrimidines, etc. [28, 29].

**Figure 18.** The COSY spectrum of the *Gb*-3 with the spectral expansion in the vinyl region

*biloba* female autumnal leaves' methanol crude extract (Figure 20).

**Figure 18.** The COSY spectrum of the *Gb*-3 with the spectral expansion in the vinyl region

5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 Chemical Shift (ppm)

5.0 4.5 4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 Chemical Shift (ppm)

The signals of the ethylene protons were found in all spectra recorded by the residual water peak. From the residual water peak to the high field region the presence of one *doublet* and one *doublet of doublets* was present. Those signals are, yet, not assigned. The methyl proton region is not similar to any previous chlorophyll biodegration products' high field region (Figure 15, 16 and 17). The chlorophyll biodegradation products found in *Ginkgo biloba* autumnal leaves are different from all previously isolated ones. The vinyl proton-proton connectivity in the COSY spectrum was easily observed (Figure 18). The *Gb*-3 COSY spectrum recorded was the same as the COSY spectrum of the chlorophyll biodegradation product 9 in the low field region.

The establishment of other scalar coupled protons was not determined (Figure 19).

**Figure 16.** The proton spectrum of the *Gb*-2 in the high field region

30 Biodegradation of Hazardous and Special Products

**Figure 17.** The proton spectrum of the *Gb*-3 in the high field region

*Gb* - 2

*Gb* - 3

The dynamic NMR spectra were not recorded, yet, such as, they were done in case of the polycyclic quinones, substituted tetrahydropyrimidines, etc. [28, 29].

The *Ginkgo biloba* chlorophyll biodegradation products were, also, detected in the *Ginkgo biloba* female autumnal leaves' methanol crude extract (Figure 20).

The screening for the presence of the chlorophyll biodegradation products was done in ferns. Of dozens of ferns' autumnal leaves' methanol extracts analyzed only in *Athyrium filix femina*, Dryopteridaceae the *Ginkgo biloba* chlorophyll biodegradation products were detected (Figure 21). The cornifer plant – *Larix kaempferi*, Pinaceae autumnal leaves' methanol crude

**Figure 19.** The spectral magnification of the *Gb*-3 COSY spectrum in the high field region

extract was analyzed and the *Ginkgo biloba* chlorophyll biodegradation products' were detected (Figure 21).

In the *Gingko biloba*, *Larix kaempferi* and *Athyrium filix femina* autumnal leaves the chlorophyll

**Figure 21.** The chromatogram of *Larix kaempferi* and *Athyrium filix femina* autumnal leaves' methanol crude extract. The LC conditions: Column: Nucleosil 100-5 C4 4x250 mm. The mobile phase: 90% v/v water (TFA): methanol to 0%

**Figure 20.** The chromatogram of the *Gingko biloba* female autumnal leaves' methanol crude extract. The LC condi‐ tions: Column: Nucleosil 100-5 C4 4x250 mm. The mobile phase: 90% v/v water (0.1%TFA): methanol to 0% v/v water

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 33

(0.1%TFA):methanol in 80 minutes. Flow rate: 0.2 ml/min. UV detection at λ=312.

Further chlorophyll biodegradation products are colourless and were not identified, up to

The ancient Egyptians used the ethylene to gas figs in order to stimulate ripening. The ancient Chinese used incense combustion to enhance the ripening of pears. In the 19th century, when the coal gas was used for street illumination, the trees in the vicinity of the street lamps showed extensive defoliation [30]. In 1901, Dimitry Nikolayevich Neljubov, observed that dark-grown pea seedlings growing in the laboratory exhibited symptoms that were later termed the triple response [31]. Dimitry Nikolayevich Neljubov identified ethylene, which was present in the

biodegradation product 3 undergoes the process that still has not been determined.

**6. The influence of ethylene on chlorophyll degradation**

v/v water (TFA):methanol in 70 minutes. Flow rate: 0.2 ml/min. UV detection at λ=312.

now.

The chlorophyll biodegradation product 3 in Hamamelidaceae and Vitaceae autumnal leaves undergoes the oxidation of the aldehyde group at the C5 position forming the urobilinogenic chlorophyll biodegradation product 4.

In the Solanaceae autumnal leaves the chlorophyll biodegradation product 3 undergoes the glycosylation at the C82 position forming the glycosylated chlorophyll biodegradation product 11.

**Figure 20.** The chromatogram of the *Gingko biloba* female autumnal leaves' methanol crude extract. The LC condi‐ tions: Column: Nucleosil 100-5 C4 4x250 mm. The mobile phase: 90% v/v water (0.1%TFA): methanol to 0% v/v water (0.1%TFA):methanol in 80 minutes. Flow rate: 0.2 ml/min. UV detection at λ=312.

**Figure 21.** The chromatogram of *Larix kaempferi* and *Athyrium filix femina* autumnal leaves' methanol crude extract. The LC conditions: Column: Nucleosil 100-5 C4 4x250 mm. The mobile phase: 90% v/v water (TFA): methanol to 0% v/v water (TFA):methanol in 70 minutes. Flow rate: 0.2 ml/min. UV detection at λ=312.

In the *Gingko biloba*, *Larix kaempferi* and *Athyrium filix femina* autumnal leaves the chlorophyll biodegradation product 3 undergoes the process that still has not been determined.

Further chlorophyll biodegradation products are colourless and were not identified, up to now.

#### **6. The influence of ethylene on chlorophyll degradation**

extract was analyzed and the *Ginkgo biloba* chlorophyll biodegradation products' were detected

**Figure 19.** The spectral magnification of the *Gb*-3 COSY spectrum in the high field region

4.0 3.5 3.0 2.5 2.0 1.5 1.0 F2 Chemical Shift (ppm)

1.0

1.5

2.0

2.5

F1 Chemical Shift (ppm)

3.0

3.5

4.0

4.5

The chlorophyll biodegradation product 3 in Hamamelidaceae and Vitaceae autumnal leaves undergoes the oxidation of the aldehyde group at the C5 position forming the urobilinogenic

In the Solanaceae autumnal leaves the chlorophyll biodegradation product 3 undergoes the glycosylation at the C82 position forming the glycosylated chlorophyll biodegradation

(Figure 21).

product 11.

chlorophyll biodegradation product 4.

32 Biodegradation of Hazardous and Special Products

The ancient Egyptians used the ethylene to gas figs in order to stimulate ripening. The ancient Chinese used incense combustion to enhance the ripening of pears. In the 19th century, when the coal gas was used for street illumination, the trees in the vicinity of the street lamps showed extensive defoliation [30]. In 1901, Dimitry Nikolayevich Neljubov, observed that dark-grown pea seedlings growing in the laboratory exhibited symptoms that were later termed the triple response [31]. Dimitry Nikolayevich Neljubov identified ethylene, which was present in the laboratory air from the coal gas, as the molecule causing the response [30]. Dimitry Nikolaye‐ vich Neljubov observation can be compared to Leonhard Euler's observation on seven bridges of Koenigsberg, which led to the formation of the Theory of Graphs [32]. Dimitry Nikolayevich Neljubov's observation led to the recognition of ethylene as a plant hormone [30]. In 1910, Herbert Henry Cousins reported that gas evaporation from stored oranges caused the premature ripening of bananas [33]. It was likely that the oranges used were infected with the fungus *Penicillium* which produces large amounts of ethylene. Herbert Henry Cousins showed that plants can produce their own ethylene [33]. In 1917, Doubt concluded that ethylene stimulates abscission [34]. In 1934, Gane reported that plants synthesize ethylene [35]. In 1935, Crocker proposed that ethylene is the plant hormone responsible for fruit ripening [36]. Ethylene, as well, has numerous effects on plant species and organs [30]. The peel of the citrus fruits requires ethylene to induce chlorophyll degradation [37]. The exposure of green leaves to ethylene can induce senescence only in leaves that have reached a defined age [38]. The biochemistry, genetics and physiology of ripening has been extensively studied in economi‐ cally important fruit crops and the results obtained permitted the formation of the ethylene biosynthesis pathway, signaling and control of gene expression [39]. There is still much to be discovered about these processes at the biochemical and genetic levels.

[6] Tswett, M. S. in Chromophylls in Plant and Animal Worlds (1910). Warsaw Universi‐

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 35

[7] Stoll, A. in Ueber Chlorophyllase und die Chlorophyllide, Dissertation, Eidg. Techni‐

[8] Willstaetter, R, & Stoll, A. Untersuchungen uber Chlorophyll. XIX. Ueber die Chloro‐

[9] Ružicka, L. Biographical Memoirs of Fellows of the Royal Society (1972). , 18-566.

[10] KrasnovskyJr., AA. Chlorophyll isolation, structure and function: major landmarks of the early history of research in the Russion Empire and the Soviet Union. Photo‐

[11] Willows, R. D. Biosynthesis of chlorophylls from protoporphyrin IX, Nat. Prod. Rep.

[12] Ruediger, W. Biosynthesis of chlorophyll b and the chlorophyll cycle. Photosynthesis

[13] Shioi, Y, Watanabe, K, & Takamiya, K. Enzymatic conversion of Pheophorbide a to the precursor of Pyropheophorbide a in leaves of Chenopodium album, Plant and

[14] Peoples, M. B, & Dalling, M. J. in Senescence and Aging in Plants (Eds.: L. D. Noo‐

[15] Djapic, N, & Pavlovic, M. Chlorophyll Catabolite from Parrotia Persica Autumnal

[16] Djapic, N, Pavlovic, M, Arsovski, S, & Vujic, G. Chlorophyl Biodegradation Product from Hamamelis virginiana Autumnal Leaves. Rev. Chim. (Bucuresti) (2009). , 60(4),

[17] Djapic, N, Djuric, A, & Pavlovic, A. Chlorophyll biodegradation in Vitis vinifera var. Pinot noir autumnal leaves. Research Journal of Agricultural Sciences (2009). , 41(2),

[18] Donoghue, N. A, Norris, D. B, & Trudgill, P. W. The purification and properties of cyclohexanone oxygenase from Nocardia globerula CL1 and Acinetobacter NCIB

[19] Kelly, D. R. A proposal for the origin of stereoselectivity in enzime catalysed Baeyer-

[20] Ðapic, N. Behaviour of Fothergilla gardenii chlorophyll catabolite under acidic con‐

Villiger reactions, Tetrahedron: Asymmetry (1996). , 7-1149.

dén, A. C. Leopold), Academic press, New York, San Diego, (1988). , 181.

Leaves. Rev. Chim. (Bucuresti) (2008). , 59(8), 878-882.

ty, Warsaw (in Russian).

sche Hochschule, Zuerich, (1912).

synthesis Research(2003). , 76-389.

(2003). , 20-327.

398-402.

256-260.

Research (2002). , 74-187.

Cell Physiol. (1996). , 37-1143.

9871, Eur. J. Biochem. (1976). , 63-175.

ditions. Kragujevac J. Sci. (2012). , 34-79.

[21] Djapic, N. private communication. (2012).

phyllide, Liebigs Ann. Chem. (1912). , 387-317.

#### **Author details**

Nina Djapic\*

Address all correspondence to: djapic@tfzr.uns.ac.rs

Technical faculty "Mihajlo Pupin", University of Novi Sad, Zrenjanin, Serbia

#### **References**


[6] Tswett, M. S. in Chromophylls in Plant and Animal Worlds (1910). Warsaw Universi‐ ty, Warsaw (in Russian).

laboratory air from the coal gas, as the molecule causing the response [30]. Dimitry Nikolaye‐ vich Neljubov observation can be compared to Leonhard Euler's observation on seven bridges of Koenigsberg, which led to the formation of the Theory of Graphs [32]. Dimitry Nikolayevich Neljubov's observation led to the recognition of ethylene as a plant hormone [30]. In 1910, Herbert Henry Cousins reported that gas evaporation from stored oranges caused the premature ripening of bananas [33]. It was likely that the oranges used were infected with the fungus *Penicillium* which produces large amounts of ethylene. Herbert Henry Cousins showed that plants can produce their own ethylene [33]. In 1917, Doubt concluded that ethylene stimulates abscission [34]. In 1934, Gane reported that plants synthesize ethylene [35]. In 1935, Crocker proposed that ethylene is the plant hormone responsible for fruit ripening [36]. Ethylene, as well, has numerous effects on plant species and organs [30]. The peel of the citrus fruits requires ethylene to induce chlorophyll degradation [37]. The exposure of green leaves to ethylene can induce senescence only in leaves that have reached a defined age [38]. The biochemistry, genetics and physiology of ripening has been extensively studied in economi‐ cally important fruit crops and the results obtained permitted the formation of the ethylene biosynthesis pathway, signaling and control of gene expression [39]. There is still much to be

discovered about these processes at the biochemical and genetic levels.

Technical faculty "Mihajlo Pupin", University of Novi Sad, Zrenjanin, Serbia

[1] Brown, S. B, & Houghton, J. D. Hendry GAF. Chlorophylls. In: Scheer H. (ed.) CRC

[2] Robinson, R. Richard Willstätter. (1872). Obituary Notices of Fellows of the Royal So‐

[3] Tswett, M. S. O novoy kategorii adsorbtsionnykh yavleny i o primenenii ikh k bio‐ kkhimicheskomu analizu. Trudy Varhavskago Obshchestva estevoispytatelei Otd Bi‐

[4] Tswett, M. S. Physikalisch- chemische Studien ueber das Chlorophyll. Die Adsorptio‐

[5] Tswett, M. S. Adsorptionsanalyse und chromatographische Methode. Anwendung

auf die Chemie des Chlorophylls, Ber. Deutsch. Bot. Ges. (1906). , 24-384.

Address all correspondence to: djapic@tfzr.uns.ac.rs

Press, Boca Raton, FL; (1991).

ciety 1953; , 8(22), 609-634.

nen, Ber. Deutsch. Bot. Ges. (1906). , 24-316.

ol. (1903). , 14-20.

**Author details**

34 Biodegradation of Hazardous and Special Products

Nina Djapic\*

**References**


[22] Andersen, Ø. M, Aksnes, D. W, Nerdal, W, & Johansen, O-P. Structure elucidation of cyanidin-3-sambubioside and assignments of the 1 H and 13C NMR resonances through two-dimensional shift-correlated NMR technique, Phytochemical Analysis, (1991). , 2(4), 175-183.

[35] Gane, R. Production of ethylene by some fruits. Nature (1934). , 134-1008.

peel of citrus fuits, Plant Physiol. (1981). , 68-854.

and Molecular Biology (2006). , 29(3), 508-515.

tal Botany (2005). , 56(421), 2915-2923.

7-231.

[36] Crocker, W, Hitchcock, A. E, & Zimmerman, P. W. Similarities in the effects of ethyl‐ ene and the plant auxins. Contributions of the Boyce Thompson Institute (1935). ,

Chlorophyll Biodegradation http://dx.doi.org/10.5772/56155 37

[37] Purvis, A. C, & Barmore, C. R. Involvement of ethylene in chlorophyll degradation in

[38] Jing, H-C. Schippers JHM., Hille J., Dijkwel PP. Ethylene-induced leaf senescence de‐ pends on age-related changes and OLD genes in Arabidopsis. Journal of Experimen‐

[39] Chaves ALSCelso de Mello-Farias P. Ethylene and fruit ripening: From illumination gas to the control of gene expression, more than a century of discoveries. Genetics


[35] Gane, R. Production of ethylene by some fruits. Nature (1934). , 134-1008.

[22] Andersen, Ø. M, Aksnes, D. W, Nerdal, W, & Johansen, O-P. Structure elucidation of

[23] Pedersen, A. T, Andersen, Ø. M, Aksnes, D. W, & Nerdal, W. Anomeric sugar config‐ uration of anthocyanin O-pyranoside determined from heteronuclear one-bond cou‐

[24] Andersen, Ø. M, Opheim, S, Aksnes, D. W, & Frøystein, N. Å. Structure of petanin, an acylated anthocyanin isolated from Solanum tuberosum using homo- and heteronuclear two-dimensional nuclear magnetic resonance techniques, Phytochemical

[25] Nerdal, W, Pedersen, A. T, & Andersen, Ø. M. Two-dimensional nuclear Overhauser Enhancement NMR Experiments on Pelargonidin-glucopyranoside, an Anthocyanin

[26] Farnier, M, Drakenberg, T, & Berger, S. Etude, par RMN, des conformations d'alde‐ hydes α et β pyrroliques C-substitues. Stereospecificite des couplages lointains. Tet‐

[27] Farnier, M, & Drakenberg, T. Nuclear Magnetic Resonance Conformational Studies of C-Substituted Pyrrolecarbaldehydes. Part 1. Substituent Effects on Aldehyde Con‐ formations as shown by Long Range Coupling Constants. J. C. S. Perkin II (1975). ,

[28] Smirnov, A, Fulton, D. B, Andreotti, A, & Petrich, J. W. Exploring ground-state heter‐ ogeneity of hypericin and hypocrellin A and B: Dynamic and 2D ROESY NMR study.

[29] Garcia, M. B, Grilli, S, Lunazzi, L, Mazzanti, A, & Orelli, L. R. Conformational Stud‐ ies by Dynamic NMR. 84.1 Structure, Conformation, and Stereodynamics of the Atro‐

[30] Taiz, L, & Zeiger, E. Plant physiology. 3rd edition. Sunderland: Sinauer Associates,

[31] Neljubow, D. Ueber die horizentale Nutation der Stengel von Pisum sativum und ei‐

[32] Alexanderson, G. L. About the cover: Euler and Koenigsberg's bridges: A historical view. Bulletin (New Series) of the American Mathematical Society (2006). , 43(4),

[33] Cousins, H. H. Agricultural Experiments: Citrus. Annual Report of the Jamaican De‐

[34] Doubt, S. L. The response of plants to illuminating gas. Botan. Gaz. (1917). , 63-209.

pisomers of N-Aryl tetrahydropyrimidines. J. Org. Chem. (2001). , 66-6679.

Journal of the American Chemical Society (1999). , 35(121), 7979-7988.

niger anderen Pflanzen. Beih Bot Zentralbl (1901). , 10-128.

partment of Agriculture (1910). , 7-15.

through two-dimensional shift-correlated NMR technique, Phytochemical Analysis,

H and 13C NMR resonances

cyanidin-3-sambubioside and assignments of the 1

pling constants, Phytochemical Analysis (1995). , 6(6), 313-316.

of Low Molecular Mass. Acta Chemica Scandinavica, (1992). , 3.

(1991). , 2(4), 175-183.

36 Biodegradation of Hazardous and Special Products

Analysis (1991). , 2(5), 230-236.

4-333.

Inc., MA. (2002).

567-573.

rahedron Letters, (1973). , 14(6), 429-432.


**Chapter 3**

**Bioremediation of Agricultural Land Damaged by**

Bioremediation is the process of using naturally occurring microbes to digest and convert unwanted material into harmless substances. In this chapter, an innovative study has been carried out by using compost and specific bacteria (Halo Bacteria) to restore the high saline

A disaster is the tragedy of a natural or human-made hazard (a hazard is a situation which poses a level of threat to life, health, property, or environment) that negatively affects society or environment. A natural disaster is a consequence when a natural hazard (e.g., volcanic eruption or earthquake) affects humans. Tsunamis and earthquakes are two of the most dangerous and yet most common hazards to affect population centers and economic infra‐ structures worldwide. Generally, tsunami flooding results from a train of long-period waves that can rapidly travel long distances from where they were generated by deep-ocean earth‐ quakes, submarine landslides, volcanic eruptions, or asteroid impacts [1, 2, 3]. Due to tsunami the sea water carry sediments along with salt itself. There have been many studies on recent and ancient tsunami deposits. These include descriptions of tsunami deposits in coastal lake, estuary, lagoon, bay floor and shelf environments and even the farmland [4,5]. The mega earthquake and consequent tsunami had caused a great damage to not only human life and infrastructure but also the agricultural land and the crops in Tohoku region, Japan. The after math of the tsunami has created many problems to environment and geo-environment of these affected areas. Soil pollution and high salinity which caused the farmland unusable for cultivation is one of the major geo-environmental problems. The objective of this study is to get an idea about the extent of soil chemical properties change due to tsunami and to apply

> © 2013 Moqsud and Omine; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

> © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

and reproduction in any medium, provided the original work is properly cited.

soil damaged by Tsunami occurred on 11th March, 2011 at the Tohoku Area in Japan.

bioremediation approach to salinity control of the agricultural land.

**Tsunami**

M. Azizul Moqsud and K. Omine

http://dx.doi.org/10.5772/56595

**1. Introduction**

Additional information is available at the end of the chapter

## **Bioremediation of Agricultural Land Damaged by Tsunami**

M. Azizul Moqsud and K. Omine

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56595

#### **1. Introduction**

Bioremediation is the process of using naturally occurring microbes to digest and convert unwanted material into harmless substances. In this chapter, an innovative study has been carried out by using compost and specific bacteria (Halo Bacteria) to restore the high saline soil damaged by Tsunami occurred on 11th March, 2011 at the Tohoku Area in Japan.

A disaster is the tragedy of a natural or human-made hazard (a hazard is a situation which poses a level of threat to life, health, property, or environment) that negatively affects society or environment. A natural disaster is a consequence when a natural hazard (e.g., volcanic eruption or earthquake) affects humans. Tsunamis and earthquakes are two of the most dangerous and yet most common hazards to affect population centers and economic infra‐ structures worldwide. Generally, tsunami flooding results from a train of long-period waves that can rapidly travel long distances from where they were generated by deep-ocean earth‐ quakes, submarine landslides, volcanic eruptions, or asteroid impacts [1, 2, 3]. Due to tsunami the sea water carry sediments along with salt itself. There have been many studies on recent and ancient tsunami deposits. These include descriptions of tsunami deposits in coastal lake, estuary, lagoon, bay floor and shelf environments and even the farmland [4,5]. The mega earthquake and consequent tsunami had caused a great damage to not only human life and infrastructure but also the agricultural land and the crops in Tohoku region, Japan. The after math of the tsunami has created many problems to environment and geo-environment of these affected areas. Soil pollution and high salinity which caused the farmland unusable for cultivation is one of the major geo-environmental problems. The objective of this study is to get an idea about the extent of soil chemical properties change due to tsunami and to apply bioremediation approach to salinity control of the agricultural land.

© 2013 Moqsud and Omine; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

The sea water inundated the large areas of agricultural land causing the excessive saline in the soil.

### **2. Salinity of soil**

Salinity of soils is the condition of soils that have a high salt content. The predominant salt is normally sodium chloride (NaCl). As a result, saline soils are therefore also *sodic soils* but there may be sodic soils those are not saline, but alkaline. Salty soils are a common feature and an environmental problem in irrigated lands in arid and semi-arid regions all over the world. They have poor or little crop production. The problems are often associated with high water tables, caused by a lack of natural subsurface drainage to the underground. Poor subsurface drainage may be caused by insufficient transport capacity of the aquifer or because water cannot exit the aquifer. Worldwide, the major factor in the development of saline soils is a lack of precipitation. Most naturally saline soils are found in (semi)arid regions and climates of the globe.

**Figure 2.** Salt accumulated on the surface of the tsunami sediment in Tohoku area, Japan

soil structure without considering potential impacts on plant health.

**3. Tohoku region Pacific Coast earthquake in 2011**

11 March 2011.The location of the epicenter (38.3220

land causing the excessive saline in the soil.

Figure 2 shows the salt accumulation on the surface of the soil after the tsunami in Tohuku area in Japan. For the case of tsunami, a vast area of the land area goes under sea water. And the accumulation of salt after the tsunami water caused a serious damage to the geo-environ‐ ment. Soil water salinity can affect soil physical properties by causing fine particles to bind together into aggregates. This process is known as flocculation and is beneficial in terms of soil aeration, root penetration, and root growth. Although increasing soil solution salinity has a positive effect on soil aggregation and stabilization, at high levels salinity can have negative and potentially lethal effects on plants. As a result, salinity cannot be increased to maintain

The great east Japan Earthquake (Higashi Nihon Daishinsai in Japanese) was a magnitude 9.0 undersea mega thrust earthquake off the coast of Japan that occurred at 14:46:23 JST on Friday,

70 kilometers east of the Oshika Peninsula of Tohoku and the hypocenter at an underwater depth of approximately 32 km. It was the most powerful known earthquake to have hit Japan, and one of the five most powerful earthquakes in the world overall since modern recordkeeping began in 1900. The earthquake triggered extremely destructive tsunami waves of around 40 m in Miyako, Iwate prefecture and in some cases travelling up to 10 km inland. In addition to loss of life and destruction of infrastructure, the tsunami caused a number of nuclear accidents in the power plant in Fukushima which caused evacuation zones affecting hundreds of thousands of residents. The sea water inundated the large areas of agricultural

N, 142.3690 E) of this earthquake is about

Bioremediation of Agricultural Land Damaged by Tsunami

http://dx.doi.org/10.5772/56595

41

**Figure 1.** Mechanism of salinity affected soil and plant interaction

Figure 1 shows the mechanisms of salinity affected soil and plant interaction. Salinity becomes a problem when enough salts accumulate in the root zone to negatively affect plant growth. Excess salts in the root zone hinder plant roots from withdrawing water from surrounding soil. This lowers the amount of water available to the plant, regardless of the amount of water actually in the root zone. For example, when plant growth is compared in two identical soils with the same moisture levels, one soil receiving salty water and the other receiving salt-free water, plants are able to use more water from the soil receiving salt-free water. Although the water is not held tighter to the soil in saline environments, the presence of salt in the water causes plants to exert more energy extracting water from the soil. The main point is that excess salinity in soil water can decrease plant available water and cause plant stress. So, high salinity of soil is very dangerous for the plant as most of the plants can not survive in that soil condition.

**Figure 2.** Salt accumulated on the surface of the tsunami sediment in Tohoku area, Japan

The sea water inundated the large areas of agricultural land causing the excessive saline in the

Salinity of soils is the condition of soils that have a high salt content. The predominant salt is normally sodium chloride (NaCl). As a result, saline soils are therefore also *sodic soils* but there may be sodic soils those are not saline, but alkaline. Salty soils are a common feature and an environmental problem in irrigated lands in arid and semi-arid regions all over the world. They have poor or little crop production. The problems are often associated with high water tables, caused by a lack of natural subsurface drainage to the underground. Poor subsurface drainage may be caused by insufficient transport capacity of the aquifer or because water cannot exit the aquifer. Worldwide, the major factor in the development of saline soils is a lack of precipitation. Most naturally saline soils are found in (semi)arid regions and climates of the

Figure 1 shows the mechanisms of salinity affected soil and plant interaction. Salinity becomes a problem when enough salts accumulate in the root zone to negatively affect plant growth. Excess salts in the root zone hinder plant roots from withdrawing water from surrounding soil. This lowers the amount of water available to the plant, regardless of the amount of water actually in the root zone. For example, when plant growth is compared in two identical soils with the same moisture levels, one soil receiving salty water and the other receiving salt-free water, plants are able to use more water from the soil receiving salt-free water. Although the water is not held tighter to the soil in saline environments, the presence of salt in the water causes plants to exert more energy extracting water from the soil. The main point is that excess salinity in soil water can decrease plant available water and cause plant stress. So, high salinity of soil is very dangerous for the plant as most of the plants can not survive in that soil condition.

soil.

globe.

**2. Salinity of soil**

40 Biodegradation of Hazardous and Special Products

**Figure 1.** Mechanism of salinity affected soil and plant interaction

Figure 2 shows the salt accumulation on the surface of the soil after the tsunami in Tohuku area in Japan. For the case of tsunami, a vast area of the land area goes under sea water. And the accumulation of salt after the tsunami water caused a serious damage to the geo-environ‐ ment. Soil water salinity can affect soil physical properties by causing fine particles to bind together into aggregates. This process is known as flocculation and is beneficial in terms of soil aeration, root penetration, and root growth. Although increasing soil solution salinity has a positive effect on soil aggregation and stabilization, at high levels salinity can have negative and potentially lethal effects on plants. As a result, salinity cannot be increased to maintain soil structure without considering potential impacts on plant health.

#### **3. Tohoku region Pacific Coast earthquake in 2011**

The great east Japan Earthquake (Higashi Nihon Daishinsai in Japanese) was a magnitude 9.0 undersea mega thrust earthquake off the coast of Japan that occurred at 14:46:23 JST on Friday, 11 March 2011.The location of the epicenter (38.3220 N, 142.3690 E) of this earthquake is about 70 kilometers east of the Oshika Peninsula of Tohoku and the hypocenter at an underwater depth of approximately 32 km. It was the most powerful known earthquake to have hit Japan, and one of the five most powerful earthquakes in the world overall since modern recordkeeping began in 1900. The earthquake triggered extremely destructive tsunami waves of around 40 m in Miyako, Iwate prefecture and in some cases travelling up to 10 km inland. In addition to loss of life and destruction of infrastructure, the tsunami caused a number of nuclear accidents in the power plant in Fukushima which caused evacuation zones affecting hundreds of thousands of residents. The sea water inundated the large areas of agricultural land causing the excessive saline in the soil.

#### **4. Soil investigation**

The field test of soil for its chemical analysis was conducted in Rikuzentakata city of Iwate prefecture, Japan. Fig. 3 shows the damaged area in Rikuzentakata city [9]. This city was one the major affected areas by the tsunami on 11th March, 2011. Fig. 4 shows the place of investigation on 5th May, 2011. Fig. 5 shows the place of soil investigation on 30th June, 2011.

The pH and EC (Electric Conductivity) of the damaged agricultural land were measured by the digital pH meter (Horiba, D-54SE). The electrical conductivity of the soil was also measured by using digital EC meter (Oakton, PCSTEST35). The salinity of the soil can be calculated by the value of electrical conductivity.

**Figure 3.** Map of Japan indicating the study area and soil investigation area

#### **5. Soil properties of Rikuzentaka area after tsunami**

The soil of the Rikuzentakata was very fertile. The farmers used to cultivate different types of crops including the corn and vegetables. The average moisture content of the soil was around 25 %. However, due to the tsunami water inundated the large areas of the Rikuzentakata, a huge amount of submarine sediments come along with the sea water and settled on the agricultural land along with different types of tsunami debris. The sediment of the tsunami also brought some kinds of toxic materials which were settled under the sea over a long period of time. The sediment is mainly a clay but some sandy particles were also found in different parts of the area.

**6. Bioremediation for restoration of saline soil**

**Figure 5.** The place of soil investigation on 30th June 2011 in Rikuzentakata area

H

Out of the damaged area

**Figure 4.** The place of soil investigation on 5 May 2011 in Rikuzentakata area

salty (saline) soils.

The aim of soil salinity control is to prevent soil degradation by salinization and reclaim already

A B C

Corn area

D

E

Sunflower area

F

Bioremediation of Agricultural Land Damaged by Tsunami

http://dx.doi.org/10.5772/56595

43

G

Various attempts are now carrying out to control the salinity of the agricultural land. The primary method of controlling soil salinity is to permit 10-20% of the irrigation water to leach

The salt concentration of the drainage water is normally 5 to 10 times higher than that of the irrigation water, thus salt export matches salt import and it will not accumulate. However, it

the soil, be drained and discharged through an appropriate drainage system.

**Figure 4.** The place of soil investigation on 5 May 2011 in Rikuzentakata area

**4. Soil investigation**

42 Biodegradation of Hazardous and Special Products

the value of electrical conductivity.

**Figure 3.** Map of Japan indicating the study area and soil investigation area

parts of the area.

**5. Soil properties of Rikuzentaka area after tsunami**

The soil of the Rikuzentakata was very fertile. The farmers used to cultivate different types of crops including the corn and vegetables. The average moisture content of the soil was around 25 %. However, due to the tsunami water inundated the large areas of the Rikuzentakata, a huge amount of submarine sediments come along with the sea water and settled on the agricultural land along with different types of tsunami debris. The sediment of the tsunami also brought some kinds of toxic materials which were settled under the sea over a long period of time. The sediment is mainly a clay but some sandy particles were also found in different

The field test of soil for its chemical analysis was conducted in Rikuzentakata city of Iwate prefecture, Japan. Fig. 3 shows the damaged area in Rikuzentakata city [9]. This city was one the major affected areas by the tsunami on 11th March, 2011. Fig. 4 shows the place of investigation on 5th May, 2011. Fig. 5 shows the place of soil investigation on 30th June, 2011.

The pH and EC (Electric Conductivity) of the damaged agricultural land were measured by the digital pH meter (Horiba, D-54SE). The electrical conductivity of the soil was also measured by using digital EC meter (Oakton, PCSTEST35). The salinity of the soil can be calculated by

**Figure 5.** The place of soil investigation on 30th June 2011 in Rikuzentakata area

#### **6. Bioremediation for restoration of saline soil**

The aim of soil salinity control is to prevent soil degradation by salinization and reclaim already salty (saline) soils.

Various attempts are now carrying out to control the salinity of the agricultural land. The primary method of controlling soil salinity is to permit 10-20% of the irrigation water to leach the soil, be drained and discharged through an appropriate drainage system.

The salt concentration of the drainage water is normally 5 to 10 times higher than that of the irrigation water, thus salt export matches salt import and it will not accumulate. However, it

**Figure 6.** Tsunami Sediment deposited on the vast area in tohoku, Japan

will take a long time and efforts for such kind of design of the salinity removal from the saline soil [7,8].

**Figure 8.** Compost by mixing rice bran, oil cakes, grinds of fish bones and water after 3 days

compost would supply some nutrients and organics to the soil.

**Table 1.** EC and pH of soil in different depths on 5th May 2011

Then, the compost containing the halo bacteria had been applied in the large areas of Riku‐ zentakata of Iwate Prefecture which was affected by sea water inundation due to tsunami. Due to bacterial activities, the salinity of the agricultural areas would have been reduced as well as

Bioremediation of Agricultural Land Damaged by Tsunami

http://dx.doi.org/10.5772/56595

45

**(a) EC (mS/cm)**

Depth 1 2 3 4 5 6 7 8 9 10

5cm 1.51 0.36 0.39 1.49 1.26 0.47 1.20 1.77 1.01 0.25

10cm 3.04 2.00 1.94 2.93 3.16 1.32 2.77 3.43 1.83 1.15

15cm 2.23 2.43 2.81 4.03 2.21 1.84 0.97 3.46 2.4 2.25

**(b) pH**

Depth 1 2 3 4 5 6 7 8 9 10

5cm 7.21 8.47 8.06 7.55 7.22 8.39 7.71 7.62 7.19 8.27

10cm 5.75 5.68 6.5 5.78 5.71 6.57 5.98 5.6 6.74 6.52

15cm 6.22 5.48 5.69 5.2 6.13 5.84 7.12 5.64 5.77 5.82

In this study, an innovative idea has been taken for reducing the salt concentration from the soil of the agricultural areas by bioremediation. By using the halo bacteria with the compost the bioremediation was carried out.

It was possible to increase a volume of the compost by mixing rice bran, oil cakes, grinds of fish bones and water in a specific ratio. Total 30 kg compost was made. Further, it had increased up to 300 kg. After mixing each material, temperature of the compost increased at 48 ˚C for 2 days and turned over for aeration (Figs. 7 and 8).

**Figure 7.** Preparing compost by mixing rice bran, oil cakes, grinds of fish bones and water

**Figure 8.** Compost by mixing rice bran, oil cakes, grinds of fish bones and water after 3 days

will take a long time and efforts for such kind of design of the salinity removal from the saline

In this study, an innovative idea has been taken for reducing the salt concentration from the soil of the agricultural areas by bioremediation. By using the halo bacteria with the compost

It was possible to increase a volume of the compost by mixing rice bran, oil cakes, grinds of fish bones and water in a specific ratio. Total 30 kg compost was made. Further, it had increased up to 300 kg. After mixing each material, temperature of the compost increased at 48 ˚C for 2

soil [7,8].

the bioremediation was carried out.

44 Biodegradation of Hazardous and Special Products

days and turned over for aeration (Figs. 7 and 8).

**Figure 6.** Tsunami Sediment deposited on the vast area in tohoku, Japan

**Figure 7.** Preparing compost by mixing rice bran, oil cakes, grinds of fish bones and water

Then, the compost containing the halo bacteria had been applied in the large areas of Riku‐ zentakata of Iwate Prefecture which was affected by sea water inundation due to tsunami. Due to bacterial activities, the salinity of the agricultural areas would have been reduced as well as compost would supply some nutrients and organics to the soil.


**Table 1.** EC and pH of soil in different depths on 5th May 2011


March 2012.The effect of bioremediation is understood in the field, however, the exact amount of the bioremediation is difficult to measure in the field as the natural environmental effects

0 10 20 30 40 50 60 70

Curing time (day)

 

Bioremediation of Agricultural Land Damaged by Tsunami

http://dx.doi.org/10.5772/56595

47

 Rikuzentakata soil Souma soil

**EC (mS/cm)** Date A B C D E F G 30/9/2011 0.174 0.213 0.211 0.249 0.186 0.203 0.226 12/3/2012 0.016 0.017 0.018 0.143 0.017 0.016 0.017

Some substances known to have toxic properties have been introduced into the environment through man-made activities. These substances range in degree of toxicity and danger to human health. Many of these substances either immediately or ultimately come in contact with or are sequestered by soil. Conventional methods to remove, reduce, or mitigate toxic sub‐ stances introduced into soil or ground water via anthropogenic activities and processes include pump and treat systems, soil vapor extraction, incineration, and containment. Utility of each of these conventional methods of treatment of contaminated soil and/or water suffers from

The mechanism of bioremediation of the salt affected agricultural land is illustrated in the Figure 10. It is seen that the bacteria mixing with compost help to release the salt element from the soil surface due to the microbial activities. When the effects of bioremediation were taken place then the salt removed easily from the surface with the help of rain water or snow melting.

have the influence on the soil properties in the wide areas of the Rikuzentakata.

0 0.5 1 1.5 2 2.5 3

**Figure 9.** Relationship between electric conductivity and curing period on soil with salt

EC (mS/cm)

EC (mS/cm)

**Table 3.** Comparison of electric conductivity of the soils before and after bioremediation

recognizable drawbacks and may involve some level of risk.

**8. Mechanisms of bioremediation of salinity affected soil**

**Table 2.** EC and pH of soil in different depths on 30th June, 2011

#### **7. Laboratory test of Bioremediation with the compost**

In order to confirm the effect of the compost, an artificial saline soil was made by mixing natural salt. The original EC of the soil is 2.85 mS/cm. The compost with Halo bacteria of 1g and rice bran of 15 g were mixed into the soil of 500 cm3 . The rice bran is a material for accelerate growth of the bacteria. After the incubation, a value of EC is measured. Fig. 9 shows the result of EC for the soil mixed with the compost. It is clear that EC of the soil with salt is decreased by mixing the compost with Halo bacteria. It is estimated that 25% of salt can be reduced by mixing the compost for 7 days in this bioremediation.

The bioremediation of the saline soil is possible by using the home made compost. The way of making the compost is easy and less costly. So, this method of remediation can be used in many developing countries in the world. The benefit of this bioremediation is that the cost during the process of bioremediation is very low compare to other method of salinity man‐ agement of agricultural soil currently used.

Soil investigation at the site was performed on March 2012. Table 3 shows comparison of electric conductivity of the soils before and after the restoration by the compost. Due to the rail fall and vegetation of sunflower, the salt concentration decreased gradually and the highest EC at the site was 0.25 mS/cm on September 2011. The value of EC decreased furthermore on

**Figure 9.** Relationship between electric conductivity and curing period on soil with salt

March 2012.The effect of bioremediation is understood in the field, however, the exact amount of the bioremediation is difficult to measure in the field as the natural environmental effects have the influence on the soil properties in the wide areas of the Rikuzentakata.


**Table 3.** Comparison of electric conductivity of the soils before and after bioremediation

**Table 2.** EC and pH of soil in different depths on 30th June, 2011

46 Biodegradation of Hazardous and Special Products

bran of 15 g were mixed into the soil of 500 cm3

the compost for 7 days in this bioremediation.

agement of agricultural soil currently used.

**7. Laboratory test of Bioremediation with the compost**

In order to confirm the effect of the compost, an artificial saline soil was made by mixing natural salt. The original EC of the soil is 2.85 mS/cm. The compost with Halo bacteria of 1g and rice

of the bacteria. After the incubation, a value of EC is measured. Fig. 9 shows the result of EC for the soil mixed with the compost. It is clear that EC of the soil with salt is decreased by mixing the compost with Halo bacteria. It is estimated that 25% of salt can be reduced by mixing

The bioremediation of the saline soil is possible by using the home made compost. The way of making the compost is easy and less costly. So, this method of remediation can be used in many developing countries in the world. The benefit of this bioremediation is that the cost during the process of bioremediation is very low compare to other method of salinity man‐

Soil investigation at the site was performed on March 2012. Table 3 shows comparison of electric conductivity of the soils before and after the restoration by the compost. Due to the rail fall and vegetation of sunflower, the salt concentration decreased gradually and the highest EC at the site was 0.25 mS/cm on September 2011. The value of EC decreased furthermore on

. The rice bran is a material for accelerate growth

Some substances known to have toxic properties have been introduced into the environment through man-made activities. These substances range in degree of toxicity and danger to human health. Many of these substances either immediately or ultimately come in contact with or are sequestered by soil. Conventional methods to remove, reduce, or mitigate toxic sub‐ stances introduced into soil or ground water via anthropogenic activities and processes include pump and treat systems, soil vapor extraction, incineration, and containment. Utility of each of these conventional methods of treatment of contaminated soil and/or water suffers from recognizable drawbacks and may involve some level of risk.

#### **8. Mechanisms of bioremediation of salinity affected soil**

The mechanism of bioremediation of the salt affected agricultural land is illustrated in the Figure 10. It is seen that the bacteria mixing with compost help to release the salt element from the soil surface due to the microbial activities. When the effects of bioremediation were taken place then the salt removed easily from the surface with the help of rain water or snow melting.

in our bioremediation of saline soil, we can easily provide the sufficient nutrients as well as to restore the salinity affected soil for a vast area generally affected by tsunami in a reasonable

Bioremediation of Agricultural Land Damaged by Tsunami

http://dx.doi.org/10.5772/56595

49

The mega earthquake and consequent huge tsunami has done a great damage to the entire areas of the pacific regions in Tohoku, Japan. The sea water which covered the agricultural lands in these areas has created a critical situation for the farmers. The farmers have lost not only the crops they were cultivating but also the soil of the agricultural field had been seriously damaged by the sea water, salinity and other pollutants. The pH value and EC value of the soil in these areas are considered as the higher value in terms of safer limit for the regular crops. To reclaim this saline soil, compost containing the Halo bacteria had been applied as an approach of bioremediation. The Halo bacteria used the excessive salts from the soil and consequently can reduce the salinity problem which was proved in the laboratory test. This compost can also provide necessary nutrients to the soil and plant. So, bioremediation by compost to restore the tsunami damaged saline soil proved to be an efficient and can be applicable in other parts of the world especially developing countries which are suffering by

[1] Morton, R.A., G. Gelfenbaum and B.E. Jaffe (2007) Physical criteria for distinguishing sandy tsunami and storm deposits using modern examples. *Sedimentary Geology*, vol.

[2] Katsudanso Kenkyu (Active Fault Res. Jpn.)12, pp 1-23 (in Japanese with English ab‐

[3] Minoura K.,Gusiakov, V.G.,Kurbatov,A., Takeuti,S.,Svendsen, J.I., Bondevik, S.,Od, T.(1996). Tsunami sedimentation associated with the 1923 kamchatka earthquake.

cost.

**10. Conclusions**

the sea level rise problems.

and K. Omine2

*Sedimentary Geology,* Vol.106, pp.145-154.

1 Department of Civil Engineering, Yamaguchi University, Japan

2 Department of Civil Engineering, Nagasaki University, Japan

**Author details**

M. Azizul Moqsud1

**References**

200,,pp.184-207.

stract).

**Figure 10.** Mechanism of salt removal from tsunami affected soil by bioremediation

The salt in the soil particle can be easily removed by this bioremediation. So this mechanism to remove the salt from the saline soil can be used not only for the tsunami affected areas but also in the coastal areas in different countries in where the sea level rise caused by global warming is a real threatening matter for the local population.

#### **9. Salinity, plant growth and bioremediation in tsunami affected area**

Salinity is an important land degradation problem. Soil salinity can be reduced by leaching soluble salts out of soil with excess irrigation water. Soil salinity control involves watertable control and flushing in combination with tile drainage or another form of subsurface drainage however, if the vast area is affected by salinity then it is really difficult to treat that soil as the fresh waster needed to wash the soil will need a huge amount of money. So bioremediation of saline soil is a better option as the compost can supply some nutrients as well to the soil. High levels of soil salinity can be tolerated if salt-tolerant plants are grown. Sensitive crops lose their vigor already in slightly saline soils, most crops are negatively affected by (moderately) saline soils, and only salinity resistant crops can stay alive in severely saline soils.

A high salt level interferes with the germination of new seeds. Salinity acts like drought on plants, preventing roots from performing their osmotic activity where water and nutrients move from an area of low concentration into an area of high concentration. Therefore, because of the salt levels in the soil, water and nutrients cannot move into the plant roots.

As soil salinity levels increase, the stress on germinating seedlings also increases. Perennial plants seem to handle salinity better than annual plants. In some cases, salinity also has a toxic effect on plants because of the high concentration of certain salts in the soil. Salinity prevents the plants from uptaking the proper balance of nutrients they require for healthy growth. So, in our bioremediation of saline soil, we can easily provide the sufficient nutrients as well as to restore the salinity affected soil for a vast area generally affected by tsunami in a reasonable cost.

#### **10. Conclusions**

Compost with salt tolerance bacteria

48 Biodegradation of Hazardous and Special Products

Other useful microorganisms

Salt tolerance bacteria

**Figure 10.** Mechanism of salt removal from tsunami affected soil by bioremediation

warming is a real threatening matter for the local population.

Ca


**N a N a** K **N a**


**N a**

Na

The salt in the soil particle can be easily removed by this bioremediation. So this mechanism to remove the salt from the saline soil can be used not only for the tsunami affected areas but also in the coastal areas in different countries in where the sea level rise caused by global

**9. Salinity, plant growth and bioremediation in tsunami affected area**

soils, and only salinity resistant crops can stay alive in severely saline soils.

of the salt levels in the soil, water and nutrients cannot move into the plant roots.

Salinity is an important land degradation problem. Soil salinity can be reduced by leaching soluble salts out of soil with excess irrigation water. Soil salinity control involves watertable control and flushing in combination with tile drainage or another form of subsurface drainage however, if the vast area is affected by salinity then it is really difficult to treat that soil as the fresh waster needed to wash the soil will need a huge amount of money. So bioremediation of saline soil is a better option as the compost can supply some nutrients as well to the soil. High levels of soil salinity can be tolerated if salt-tolerant plants are grown. Sensitive crops lose their vigor already in slightly saline soils, most crops are negatively affected by (moderately) saline

A high salt level interferes with the germination of new seeds. Salinity acts like drought on plants, preventing roots from performing their osmotic activity where water and nutrients move from an area of low concentration into an area of high concentration. Therefore, because

As soil salinity levels increase, the stress on germinating seedlings also increases. Perennial plants seem to handle salinity better than annual plants. In some cases, salinity also has a toxic effect on plants because of the high concentration of certain salts in the soil. Salinity prevents the plants from uptaking the proper balance of nutrients they require for healthy growth. So,

Soil particle

> Bacteria absorbing salt Organic acid excreted from bacteria chelating salt

NO3 -

Na

SO4 2-

Mg

K **N a**

Soil after adding salt tolerance bacteria

Soil solution Air in a soil

Na

Ca+


Soil particle

K

**N a**

**N a**

**N a+**

Mg2+

**N a**

Mg

Na

CO2 O2

The mega earthquake and consequent huge tsunami has done a great damage to the entire areas of the pacific regions in Tohoku, Japan. The sea water which covered the agricultural lands in these areas has created a critical situation for the farmers. The farmers have lost not only the crops they were cultivating but also the soil of the agricultural field had been seriously damaged by the sea water, salinity and other pollutants. The pH value and EC value of the soil in these areas are considered as the higher value in terms of safer limit for the regular crops. To reclaim this saline soil, compost containing the Halo bacteria had been applied as an approach of bioremediation. The Halo bacteria used the excessive salts from the soil and consequently can reduce the salinity problem which was proved in the laboratory test. This compost can also provide necessary nutrients to the soil and plant. So, bioremediation by compost to restore the tsunami damaged saline soil proved to be an efficient and can be applicable in other parts of the world especially developing countries which are suffering by the sea level rise problems.

#### **Author details**


#### **References**


[4] Bourgeois J., T.A. Hansen, P.L. Kauffman,E.G. (1988). A tsunami deposit at the Creta‐ ceous-Tertiary boundary in Texas. Science, Vol. 241, pp. 567-570.

**Chapter 4**

**Microbial Degradation of Some Halogenated**

Yu-Huei Peng and Yang-hsin Shih

position methods are now in great request.

nervous system, liver and blood cells [5].

http://dx.doi.org/10.5772/56306

**1. Introduction**

Additional information is available at the end of the chapter

**Compounds: Biochemical and Molecular Features**

Due to the advance of organic and synthetic chemistry and many applications of man-made organic compounds, lots of xenobiotic chemicals are produced and benefit our life. However, some of them are persistent in the environment and their toxicities are accumulated through the food webs. Due to the potential hazard for human and the ecosystem, the regulations on the usage of these persistent organic pollutants (POPs) and the development of safe decom‐

Tetrachloroethene (PCE) and trichloroethene (TCE) (Table 1) have been widely used as dry cleaning solvents and degreasing agents. Due to poor disposal practices and accidental release, they are within the most-abundant groundwater contaminants. Exposure to PCE is injurious to epidermis, kidney and nervous system [1]. It has been classified as a probably carcinogen [2]. Exposure to TCE leads to acute effects on liver, kidney, central nervous, and endocrine systems. It is also associated with several types of cancers based upon epidemiological research [3]. PCE and TCE are regulated in U.S.A. to a maximum contaminant level of 5 ppb. The use of PCE and TCE in the food and pharmaceutical industries has been banned across much of the world since the 1970s. However, these chemicals are still used as a degreasing agent for other demand. Besides, the mono-chlorinated ethene, vinyl chloride (VC), is known as carcinogen that causes liver cancer [4] and the *cis*-1,2-dichloroethene (cDCE) is harmful to

Polybrominated diphenyl ethers (PBDEs), composed by two phenol rings and linked by one oxygen atom (Table 1), allow maximum ten bromide atom incorporated on the phenol rings to form 209 possible congeners. They have been widely used as flame retardants in many products over more than three decades. Their usage has protected both human lives and their properties from fire damage. PBDEs disrupt the balance of thyroid hormone, lead to repro‐

and reproduction in any medium, provided the original work is properly cited.

© 2013 Peng and Shih; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,


### **Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features**

Yu-Huei Peng and Yang-hsin Shih

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56306

#### **1. Introduction**

[4] Bourgeois J., T.A. Hansen, P.L. Kauffman,E.G. (1988). A tsunami deposit at the Creta‐

[5] Dawson A.G., Long, D., Smith, D.E., (1988). The steregga slides; evidence from east‐

[6] Yamazaki T., Yamaoka,M.Shiki,T.(1989). Miocene offshore tractive current-worked conlomerates-Tsubutegaura, Chiba Peninsula, Central Japan. In: Taira,A.,Masu‐ da,F(Eds). Sedimentary Facies in the Active Plate Margin, TERRAPUB, Tokyo, pp.

[7] Shiki T and Yamazaki,T.,(1996). Tsunami-induced conglomerates in Miocene upper bathyal deposits, Chiba Peninsula, Central Japan. *Sedimentary Geology*. Vol 104, pp.

[8] Wright C.,Mella, A. (1963). Modifications to the soil pattern of south-central Chile re‐ sulting from seismic and associated phenomena during the period May to August

ceous-Tertiary boundary in Texas. Science, Vol. 241, pp. 567-570.

1960. Bull.Seismol.Soc.Am.Vol 53,pp. 1367-1402.

[9] The Geospatial Information Authority of Japan.

483-494.

50 Biodegradation of Hazardous and Special Products

175-188.

ern Scotland for possible tsunamis. Marine Geology. Vol.99,pp.265-287.

Due to the advance of organic and synthetic chemistry and many applications of man-made organic compounds, lots of xenobiotic chemicals are produced and benefit our life. However, some of them are persistent in the environment and their toxicities are accumulated through the food webs. Due to the potential hazard for human and the ecosystem, the regulations on the usage of these persistent organic pollutants (POPs) and the development of safe decom‐ position methods are now in great request.

Tetrachloroethene (PCE) and trichloroethene (TCE) (Table 1) have been widely used as dry cleaning solvents and degreasing agents. Due to poor disposal practices and accidental release, they are within the most-abundant groundwater contaminants. Exposure to PCE is injurious to epidermis, kidney and nervous system [1]. It has been classified as a probably carcinogen [2]. Exposure to TCE leads to acute effects on liver, kidney, central nervous, and endocrine systems. It is also associated with several types of cancers based upon epidemiological research [3]. PCE and TCE are regulated in U.S.A. to a maximum contaminant level of 5 ppb. The use of PCE and TCE in the food and pharmaceutical industries has been banned across much of the world since the 1970s. However, these chemicals are still used as a degreasing agent for other demand. Besides, the mono-chlorinated ethene, vinyl chloride (VC), is known as carcinogen that causes liver cancer [4] and the *cis*-1,2-dichloroethene (cDCE) is harmful to nervous system, liver and blood cells [5].

Polybrominated diphenyl ethers (PBDEs), composed by two phenol rings and linked by one oxygen atom (Table 1), allow maximum ten bromide atom incorporated on the phenol rings to form 209 possible congeners. They have been widely used as flame retardants in many products over more than three decades. Their usage has protected both human lives and their properties from fire damage. PBDEs disrupt the balance of thyroid hormone, lead to repro‐

© 2013 Peng and Shih; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

ductive toxicity, hepatic toxicity, immunotoxicity and developmental neurotoxicity in mammals [6, 7]. The toxicity of PBDEs and their metabolites are due to elevated free radicals, DNA damages, cell cycle blockage and apoptosis rate [8, 9]. Among the congeners of PBDEs, the usage of penta-BDEs and octa-BDEs has been banned in the European Union and several states of the USA; tetra-BDE to hepta-BDE have also been classified as POPs and the production of decabromodiphenyl ether (DBDE) will cease in 2013 in USA. However, the concentration of PBDEs in environment remains exponentially increasing because of the consequence of long-term usage [10]. Due to their ubiquitous distribution in the environment, potential toxicity, tendency for bioaccumulation, and the increased accumulation amount in the environment, the fates of PBDEs in the nature is serious concern for public health.

and molecular point of view. We also attempt to compare the advantages and drawbacks of

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

53

Biodegradation of chloroethenes, PBDEs and HBCD occurs in various environmental or living samples [31, 32]. In the environment, microorganisms play major roles in the degradation reactions; while intrinsic detoxicification systems in plants and animals bodies metabolize these compounds [31, 33]. In this article, the diverse and complex microbial degradation

Biotransformation of chloroethenes, PBDEs and HBCD in aerobic or anaerobic environments has been demonstrated (Fig. 1). In aerobic environment, chloroethenes and PBDEs are metabolized with the generation of energy or degraded cometabolically without energy-yield. In anaerobic condition, they are reduced through the energy yielded from the oxidation of electron donors, i.e. reductive dehalogenation or dehalorespiration. Biotransformation of HBCD might mediate through hydrolytic dehalogenation, which may occur either in aerobic or anaerobic conditions. Detail for the each type of reaction and the degraders will be described

Under aerobic conditions, chloroethenes and PBDEs can be oxidized both cometabolically and metabolically (Fig. 1, left part). Metabolic degradation indicates the use of the above com‐ pounds as growth substrate. Chloroethenes and PBDEs more easily undergo aerobic trans‐

Metabolic degradation of cDCE and VC as sole carbon and energy source has been reported by many bacteria, such as the *Pseudomonas* sp. and *Bacillus* sp. [34, 35]. Using cDCE as auxiliary substrate for growth is much less [35] and not shown in TCE and PCE. After oxidative transformation, the auxiliary substrate may be mineralized or the carbon atoms may be incorporated into biomass. Microbial growth can be confirmed by monitoring the stable

On the contrary, cometabolic degradation of chloroethenes occurs fortuitously during the degradation of growth dependent substrates (auxiliary substrates), such as methane, ammo‐ nia, or aromatic hydrocarbons. Even cDCE can be cometabolized when VC is metabolic degraded [34]. Cometabolic degradation of TCE, DCE and VC is common [37, 38]. So far *Pseudomonas stutzeri* OX1 is the only one that could aerobically cometabolize PCE [39]. Therefore, without primary substrate supplement, intrinsic bioremediation with air or nutrients injection alone could not enhance the aerobic cometabolic mechanism and would not

PBDEs could be degraded into phenol or catechols by aerobic microbial through hydroxylation or bond cleavage [33] (Table 2). *Sphingomonas* sp. SS3 and SS33 can transform mono- or di-

the combined approaches which may apply to field remediation.

machineries are presented and compared.

in the following sections.

**2.1. Aerobic oxidative degradation**

formation with less numbers of halogen substituent.

isotope fractionation and is suitable for field assessment [36].

cause the microbial degradation of PCE and TCE contaminated sites.

**2. Biodegradation of chloroethenes, PBDEs and HBCD**

Hexabromocyclododecane (HBCD), a brominated aliphatic cyclic hydrocarbon (Table 1), is another widely used brominated flame retardant (BFR). It has animal thyroidal and develop‐ mental toxicity. The toxicity is due to altering the expression and function of metabolic enzymes, increasing hormone turn over and apoptosis [11, 12]. HBCD has been detected widely in biota and abiotic samples [11]. Due to its persistent, bioaccumulative, and toxic properties, HBCD has been proposed as a Substance of Very High Concern under the REACH regulations [13] and included on the USEPA's lists of Chemicals of Concern [14]; It is also under screening-level risk assessments to determine if it meets criteria of compounds in the Stock‐ holm Convention and in the UN-ECE Protocol on POPs [15].

Most traditional remediation methods are not suitable for degrading chloroethenes, PBDEs and HBCD. For example, dehalogenation processes under oxidative, alkaline, or irradiation conditions are high cost of energy and treatment reagents [16, 17]. Pyrolysis is only limited for specific contaminated media with high heat conductance [18]. The generation of hazardous by-products is also a problem [19] It has been reported that PBDEs and HBCD could be photodegraded [20]; however, pollutants accumulated in the soils, sediments, water bodies are not easy approach to light. Recently, the permeable reactive barrier made by zerovalent iron offers a new direction for halogenated compounds remediation [21]. Electrons offered from iron can reduce the halogenated compounds through reductive dehalogenation.

Zerovalent iron is cheaper than above processing methods, and the shortage of low efficiency is compensated by newly developed nanotechnology. Nanoscale zerovalent metals can degrade chloroethenes, PBDEs, and other contaminants with a fast kinetics and high efficiency [22-26]. There are still some limitations by using nano-metals, such as: toxic by-products generation due to incomplete dehalogenation [23], potential hazardous effect from nanopar‐ ticles [27], and large requirement for metals. Therefore, it comes to be one of the recent trends in developing nanomaterials with high efficiency and low environmental impact, and combining with other treatment technologies.

Microbiological approaches produce less intervention to the environment and are less expensive than physical or chemical methods. Biodegradation of chloroethenes has been extensively investigated and reviewed [28-30]. Bioremediation for PBDEs and HBCD are just at the beginning. The main objective of this review is to summarize current knowledge of microbial degradation of chloroethene, PBDEs and HBCD, especially from the biochemical and molecular point of view. We also attempt to compare the advantages and drawbacks of the combined approaches which may apply to field remediation.

#### **2. Biodegradation of chloroethenes, PBDEs and HBCD**

Biodegradation of chloroethenes, PBDEs and HBCD occurs in various environmental or living samples [31, 32]. In the environment, microorganisms play major roles in the degradation reactions; while intrinsic detoxicification systems in plants and animals bodies metabolize these compounds [31, 33]. In this article, the diverse and complex microbial degradation machineries are presented and compared.

Biotransformation of chloroethenes, PBDEs and HBCD in aerobic or anaerobic environments has been demonstrated (Fig. 1). In aerobic environment, chloroethenes and PBDEs are metabolized with the generation of energy or degraded cometabolically without energy-yield. In anaerobic condition, they are reduced through the energy yielded from the oxidation of electron donors, i.e. reductive dehalogenation or dehalorespiration. Biotransformation of HBCD might mediate through hydrolytic dehalogenation, which may occur either in aerobic or anaerobic conditions. Detail for the each type of reaction and the degraders will be described in the following sections.

#### **2.1. Aerobic oxidative degradation**

ductive toxicity, hepatic toxicity, immunotoxicity and developmental neurotoxicity in mammals [6, 7]. The toxicity of PBDEs and their metabolites are due to elevated free radicals, DNA damages, cell cycle blockage and apoptosis rate [8, 9]. Among the congeners of PBDEs, the usage of penta-BDEs and octa-BDEs has been banned in the European Union and several states of the USA; tetra-BDE to hepta-BDE have also been classified as POPs and the production of decabromodiphenyl ether (DBDE) will cease in 2013 in USA. However, the concentration of PBDEs in environment remains exponentially increasing because of the consequence of long-term usage [10]. Due to their ubiquitous distribution in the environment, potential toxicity, tendency for bioaccumulation, and the increased accumulation amount in the

environment, the fates of PBDEs in the nature is serious concern for public health.

holm Convention and in the UN-ECE Protocol on POPs [15].

52 Biodegradation of Hazardous and Special Products

reduce the halogenated compounds through reductive dehalogenation.

combining with other treatment technologies.

Hexabromocyclododecane (HBCD), a brominated aliphatic cyclic hydrocarbon (Table 1), is another widely used brominated flame retardant (BFR). It has animal thyroidal and develop‐ mental toxicity. The toxicity is due to altering the expression and function of metabolic enzymes, increasing hormone turn over and apoptosis [11, 12]. HBCD has been detected widely in biota and abiotic samples [11]. Due to its persistent, bioaccumulative, and toxic properties, HBCD has been proposed as a Substance of Very High Concern under the REACH regulations [13] and included on the USEPA's lists of Chemicals of Concern [14]; It is also under screening-level risk assessments to determine if it meets criteria of compounds in the Stock‐

Most traditional remediation methods are not suitable for degrading chloroethenes, PBDEs and HBCD. For example, dehalogenation processes under oxidative, alkaline, or irradiation conditions are high cost of energy and treatment reagents [16, 17]. Pyrolysis is only limited for specific contaminated media with high heat conductance [18]. The generation of hazardous by-products is also a problem [19] It has been reported that PBDEs and HBCD could be photodegraded [20]; however, pollutants accumulated in the soils, sediments, water bodies are not easy approach to light. Recently, the permeable reactive barrier made by zerovalent iron offers a new direction for halogenated compounds remediation [21]. Electrons offered from iron can

Zerovalent iron is cheaper than above processing methods, and the shortage of low efficiency is compensated by newly developed nanotechnology. Nanoscale zerovalent metals can degrade chloroethenes, PBDEs, and other contaminants with a fast kinetics and high efficiency [22-26]. There are still some limitations by using nano-metals, such as: toxic by-products generation due to incomplete dehalogenation [23], potential hazardous effect from nanopar‐ ticles [27], and large requirement for metals. Therefore, it comes to be one of the recent trends in developing nanomaterials with high efficiency and low environmental impact, and

Microbiological approaches produce less intervention to the environment and are less expensive than physical or chemical methods. Biodegradation of chloroethenes has been extensively investigated and reviewed [28-30]. Bioremediation for PBDEs and HBCD are just at the beginning. The main objective of this review is to summarize current knowledge of microbial degradation of chloroethene, PBDEs and HBCD, especially from the biochemical Under aerobic conditions, chloroethenes and PBDEs can be oxidized both cometabolically and metabolically (Fig. 1, left part). Metabolic degradation indicates the use of the above com‐ pounds as growth substrate. Chloroethenes and PBDEs more easily undergo aerobic trans‐ formation with less numbers of halogen substituent.

Metabolic degradation of cDCE and VC as sole carbon and energy source has been reported by many bacteria, such as the *Pseudomonas* sp. and *Bacillus* sp. [34, 35]. Using cDCE as auxiliary substrate for growth is much less [35] and not shown in TCE and PCE. After oxidative transformation, the auxiliary substrate may be mineralized or the carbon atoms may be incorporated into biomass. Microbial growth can be confirmed by monitoring the stable isotope fractionation and is suitable for field assessment [36].

On the contrary, cometabolic degradation of chloroethenes occurs fortuitously during the degradation of growth dependent substrates (auxiliary substrates), such as methane, ammo‐ nia, or aromatic hydrocarbons. Even cDCE can be cometabolized when VC is metabolic degraded [34]. Cometabolic degradation of TCE, DCE and VC is common [37, 38]. So far *Pseudomonas stutzeri* OX1 is the only one that could aerobically cometabolize PCE [39]. Therefore, without primary substrate supplement, intrinsic bioremediation with air or nutrients injection alone could not enhance the aerobic cometabolic mechanism and would not cause the microbial degradation of PCE and TCE contaminated sites.

PBDEs could be degraded into phenol or catechols by aerobic microbial through hydroxylation or bond cleavage [33] (Table 2). *Sphingomonas* sp. SS3 and SS33 can transform mono- or dihalogenated DEs for growth [40, 41]. In addition, *Sphingomonas* sp. PH-07 could break down several lower-bronimated BDE congeners (up to tri-BDEs) [42]. Other PBDEs degradation bacteria are reported [43]: *Rhodococcus jostii* RHA1 and *Burkholderia xenovorans* LB400 transform several lower-brominated BDE congeners (up to penta- and hexa-BDEs); *Rhodococcus* sp. RR1 transforms di- and mono-BDEs and the *Pseudonocardia dioxanivorans* CB1190 only degrades mono-BDEs. The transformation by-products include phenol, catechol, halophenol and halocatehol, indicating nonspecific attractions. These degraders might transform PBDEs through cometabolic reactions because auxiliary substrates such as diphenyl ether are supplemented. The *Lysinibacillus fusiformis* strain DB-1, cometabolically debrominate DBDE with the metabolism of lactate, pyruvate and acetate, is isolated [44].

Reductive debromination of PBDEs has been reported through pure strains (Table 2) or mixed cultures. Most of the debromination processes require TCE to be co-substrate. 20 mixed microcosms can degrade octa-BDE mixture to hexa- to mono-BDEs within 2 months [50]. *Sulfurospirillum multivorans* could debrominate DBDE into hepta- and octa-BDEs after 2 months of incubation. *D. ethenogenes* strain 195 could debrominate the octa-BDE mixtures into hepta- to di-BDEs after 6 months of incubation [51]. *Dehalococcoides* sp. Strain DG could degrade octa-BDE mixture into terta- and penta-BDEs or transform penta-BDE mixture into terta-BDE [52]. Several dechlorinating bacteria, *Desulfitobacterium hafniense* PCP-1, *Dehalobacter restric‐ tus* PER-K23, *Desulfitobacterium chlororespirans* Co23 and *Desulfitobacterium dehalogenans*JW/IU-DCI debrominate the octa-BDE mixture and the most frequently detected congeners, penta 99 and tetra 47 when PCP, PCE, 3-chloro-4-hydroxybenzoate, or 3-chloro-4-hydroxyphenylace‐ tate are applied as co-substrates [53]. Some mixture cultures do not need halogenated com‐ pounds to stimulate PBDEs transformation [50]. Recently, a lactate-dependent bacterium, *Acetobacterium* sp. strain AG, was isolated and can transform penta-BDE mixtures without other halogenated electron acceptors [52]. We also found that the cometabolism with glucose facilitated the biodegradation of mono-BDE, in terms of kinetics and efficiency in one anaerobic

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

55

In a mix microcosm, anaerobic environment necessary for dehalorespiration could be estab‐ lished by other symbiotic microorganisms. In our previous study, the mono-BDE is trans‐ formed to diphenyl ether in an aerobic culture from sewage sediment, indicating an anaerobic debromination reaction occurred. The enriched *Clostridiales* specie shown in the denatured

The metabolic pathway of VC is much clearer than that of cDCE. Alkene monooxygenase (AkMO) involves in the initial epoxidation step. The encoded genes (*etnABCD*) and the structures have been identified. Downstream events of the transformation are mediated through coenzyme M transferase (encoded by *etnE* gene), alcohol/aldehyde dehydrogenase, CoA transferase and CoM reductase/carboxylase. The final product, acyl-CoA, is then metabolized through TCA cycle [56]. Proteomic and transcriptomic analyses have confirmed

Aerobic cometabolic degradation of chloroethenes is supposed through several kinds of oxygenases: toluene monooxygenase, toluene dioxygenase, phenol monooxygenase and methane monooxygenase [57]. *P. stutzeri* OX1 depletes PCE and releases chloride irons when toluene is applied as an auxiliary substrate [39]. PCE, DCE, and VC could be transformed by the purified toluene-*o*-xylene monooxygenase (ToMO). ToMO is a four-component enzyme which consists a catalytic oxygen-bridged dinuclear center encoded by *touABE*, a NADH ferredoxin oxidoreductase (encoded by *touF*), a mediating protein (encoded by *touD*), and a Rieske-type ferredoxin (encoded by *touC*). The *touA~F* genes cloned into *E. Coli* could make it

Different dioxygenases are supposed to involve in aerobic degradation of lower numbered PBDEs. 1,2-dioxygenase is involving in the initial dihydroxylation step when mono-halogen‐

gradient gel electrophoresis (DGGE) may responsible for such reaction [55].

the roles of above enzymes in aerobic VC transformation process.

sludge in Taiwan [54].

**2.3. Degradation enzymes**

to be PCE-degradable.

So far, there is only one degrader been reported can transform HBCD: *Pseudomonas* sp. HB01 [45]. Since bromide atom was not detected after degradation reaction, such transformation might not through haloelimination. In general, cometabolism requires supplement of auxiliary substrates and there is no energy yielded. Therefore, microorganisms do not favor proceed this kind of reaction. Besides, the dehalogenation reaction is usually incomplete, resulting accumulation of toxic intermediates. The contribution on the bioremediation from cometabo‐ lism is limited [30].

#### **2.2. Anaerobic reductive dehalogenation**

Reductive dehalogenation is an anaerobic respiration process. Electron donors are oxi‐ dized and the halogenated compounds are reduced through accepting the electrons. The free energy generated from this reaction supports the growth of microbial degraders. Hy‐ drogen atom replaces the halogen atoms one after another resulting in the dehalogenation sequence from higher-numbered compounds to lower-numbered ones. Contrary to aero‐ bic degradation, the potential for reductive dehalogenation increases with the number of halogenated substituent [29].

Hydrogen gas is generated primary by fermentative and acetogenic bacteria (Fig.1, right part). The dehalorespiration bacteria compete with hydrogenotrophs, such as sulphate-reducers, nitrate-reducers, methanogens, acetogens, and other reducers [46, 47]. Except hydrogen, other electron donors also can be used for reductive dehalogenation, ex. *Sulfurospirillum multivor‐ ans* can also use pyruvate and formate [48].

Several mixed cultures and pure strains are known to reductively transform chloroethenes. Mixed culture could cooperate and transform PCE to ethene. The pure strains belong to different genus, such as Bacillus, Dehalobacter, Dehalococcoides, Desulfitobacterium, Geo‐ bacter, and Sulfurospirillum (Table 2). Most of them only dechlorinate PCE and TCE to cDCE. Only *Dehalococcoides ethenogenes* strain 195 can reductively dechlorinate PCE to ethene. The accumulated hazardous DCE and VC is a major obstacle in bioremediation of chloroethene contaminated sites. *Dehalococcoides* sp. strain BAV1, dechlorinates DCE and VC and cometa‐ bolizes PCE and TCE; the accumulated toxic compounds can be transformed into benign ethene [49].

Reductive debromination of PBDEs has been reported through pure strains (Table 2) or mixed cultures. Most of the debromination processes require TCE to be co-substrate. 20 mixed microcosms can degrade octa-BDE mixture to hexa- to mono-BDEs within 2 months [50]. *Sulfurospirillum multivorans* could debrominate DBDE into hepta- and octa-BDEs after 2 months of incubation. *D. ethenogenes* strain 195 could debrominate the octa-BDE mixtures into hepta- to di-BDEs after 6 months of incubation [51]. *Dehalococcoides* sp. Strain DG could degrade octa-BDE mixture into terta- and penta-BDEs or transform penta-BDE mixture into terta-BDE [52]. Several dechlorinating bacteria, *Desulfitobacterium hafniense* PCP-1, *Dehalobacter restric‐ tus* PER-K23, *Desulfitobacterium chlororespirans* Co23 and *Desulfitobacterium dehalogenans*JW/IU-DCI debrominate the octa-BDE mixture and the most frequently detected congeners, penta 99 and tetra 47 when PCP, PCE, 3-chloro-4-hydroxybenzoate, or 3-chloro-4-hydroxyphenylace‐ tate are applied as co-substrates [53]. Some mixture cultures do not need halogenated com‐ pounds to stimulate PBDEs transformation [50]. Recently, a lactate-dependent bacterium, *Acetobacterium* sp. strain AG, was isolated and can transform penta-BDE mixtures without other halogenated electron acceptors [52]. We also found that the cometabolism with glucose facilitated the biodegradation of mono-BDE, in terms of kinetics and efficiency in one anaerobic sludge in Taiwan [54].

In a mix microcosm, anaerobic environment necessary for dehalorespiration could be estab‐ lished by other symbiotic microorganisms. In our previous study, the mono-BDE is trans‐ formed to diphenyl ether in an aerobic culture from sewage sediment, indicating an anaerobic debromination reaction occurred. The enriched *Clostridiales* specie shown in the denatured gradient gel electrophoresis (DGGE) may responsible for such reaction [55].

#### **2.3. Degradation enzymes**

halogenated DEs for growth [40, 41]. In addition, *Sphingomonas* sp. PH-07 could break down several lower-bronimated BDE congeners (up to tri-BDEs) [42]. Other PBDEs degradation bacteria are reported [43]: *Rhodococcus jostii* RHA1 and *Burkholderia xenovorans* LB400 transform several lower-brominated BDE congeners (up to penta- and hexa-BDEs); *Rhodococcus* sp. RR1 transforms di- and mono-BDEs and the *Pseudonocardia dioxanivorans* CB1190 only degrades mono-BDEs. The transformation by-products include phenol, catechol, halophenol and halocatehol, indicating nonspecific attractions. These degraders might transform PBDEs through cometabolic reactions because auxiliary substrates such as diphenyl ether are supplemented. The *Lysinibacillus fusiformis* strain DB-1, cometabolically debrominate DBDE

So far, there is only one degrader been reported can transform HBCD: *Pseudomonas* sp. HB01 [45]. Since bromide atom was not detected after degradation reaction, such transformation might not through haloelimination. In general, cometabolism requires supplement of auxiliary substrates and there is no energy yielded. Therefore, microorganisms do not favor proceed this kind of reaction. Besides, the dehalogenation reaction is usually incomplete, resulting accumulation of toxic intermediates. The contribution on the bioremediation from cometabo‐

Reductive dehalogenation is an anaerobic respiration process. Electron donors are oxi‐ dized and the halogenated compounds are reduced through accepting the electrons. The free energy generated from this reaction supports the growth of microbial degraders. Hy‐ drogen atom replaces the halogen atoms one after another resulting in the dehalogenation sequence from higher-numbered compounds to lower-numbered ones. Contrary to aero‐ bic degradation, the potential for reductive dehalogenation increases with the number of

Hydrogen gas is generated primary by fermentative and acetogenic bacteria (Fig.1, right part). The dehalorespiration bacteria compete with hydrogenotrophs, such as sulphate-reducers, nitrate-reducers, methanogens, acetogens, and other reducers [46, 47]. Except hydrogen, other electron donors also can be used for reductive dehalogenation, ex. *Sulfurospirillum multivor‐*

Several mixed cultures and pure strains are known to reductively transform chloroethenes. Mixed culture could cooperate and transform PCE to ethene. The pure strains belong to different genus, such as Bacillus, Dehalobacter, Dehalococcoides, Desulfitobacterium, Geo‐ bacter, and Sulfurospirillum (Table 2). Most of them only dechlorinate PCE and TCE to cDCE. Only *Dehalococcoides ethenogenes* strain 195 can reductively dechlorinate PCE to ethene. The accumulated hazardous DCE and VC is a major obstacle in bioremediation of chloroethene contaminated sites. *Dehalococcoides* sp. strain BAV1, dechlorinates DCE and VC and cometa‐ bolizes PCE and TCE; the accumulated toxic compounds can be transformed into benign

with the metabolism of lactate, pyruvate and acetate, is isolated [44].

lism is limited [30].

**2.2. Anaerobic reductive dehalogenation**

54 Biodegradation of Hazardous and Special Products

*ans* can also use pyruvate and formate [48].

halogenated substituent [29].

ethene [49].

The metabolic pathway of VC is much clearer than that of cDCE. Alkene monooxygenase (AkMO) involves in the initial epoxidation step. The encoded genes (*etnABCD*) and the structures have been identified. Downstream events of the transformation are mediated through coenzyme M transferase (encoded by *etnE* gene), alcohol/aldehyde dehydrogenase, CoA transferase and CoM reductase/carboxylase. The final product, acyl-CoA, is then metabolized through TCA cycle [56]. Proteomic and transcriptomic analyses have confirmed the roles of above enzymes in aerobic VC transformation process.

Aerobic cometabolic degradation of chloroethenes is supposed through several kinds of oxygenases: toluene monooxygenase, toluene dioxygenase, phenol monooxygenase and methane monooxygenase [57]. *P. stutzeri* OX1 depletes PCE and releases chloride irons when toluene is applied as an auxiliary substrate [39]. PCE, DCE, and VC could be transformed by the purified toluene-*o*-xylene monooxygenase (ToMO). ToMO is a four-component enzyme which consists a catalytic oxygen-bridged dinuclear center encoded by *touABE*, a NADH ferredoxin oxidoreductase (encoded by *touF*), a mediating protein (encoded by *touD*), and a Rieske-type ferredoxin (encoded by *touC*). The *touA~F* genes cloned into *E. Coli* could make it to be PCE-degradable.

Different dioxygenases are supposed to involve in aerobic degradation of lower numbered PBDEs. 1,2-dioxygenase is involving in the initial dihydroxylation step when mono-halogen‐ ated DEs to be degraded [40]. Downstream degradation processes are supposed through phenol hydroxylases and catechol 1,2-dioxygenase. The transformation by-products range from phenol, catechol, halophenol and halocatechol, indicating nonspecific attack reactions [40, 41]. 2,3-dioxygenase is responsible to dihydroxylate lower numbered PBDEs and their similar chemicals such as DE in the close species *Sphingomonas sp.* PH-07 [42]. The range of PBDEs transformed by *R. jostii* RHA1 depends on the types of growth substrate. The enzymes responsible for degradation are inducible [43]. The expression of biphenyl dioxygenase (BPDO) and ethylbenzene dioxygenase (EBDO) are upregulated during PBDEs degradation. Ectopically expression of these enzymes in closed bacteria that bears no PBDEs degradation activity could transform PBDEs. EBDO depleted mono- through penta-BDEs and BPDO only depleted mono-, di- and one tetra-BDEs. [58].

The localization of chloroethene RDases is supposed in the membrane, where they could accept electrons from proton producing hydrogenase via menaquinone. The membranebound characteristics of RDases has been proved, such as PceA of *D. ethenogenes*. The lo‐ calization of constitutively expressed PceA in *S. multivorans* was initially found in the cytoplasmic fraction [65]. John et al. used freeze-fracture replica immunogold labeling technique and found it would be at the cytoplasmic membrane when cells grown on pyr‐

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

57

The major catalysis reaction of RDases is directed by subunit A, encoded by reductive dehalogenase homologous A (*rdhA*) genes. Over 650 *rdhA* genes have been identified based upon genomic sequence annotation or homologous cloning [67]. However, most of them are not yet been functional characterized. It is common for one dehalorespiration bacterium baring multicopy of *rdhA* genes in the genome. Besides *tceA* and *pceA* genes, there is still 17 RDase genes with unknown function in the genome of *D. ethenogenes* strain 195 [68]. Whether the roles

Most *rdhA* genes are organized with genes encoding for accessory proteins. The *pce-*gene cluster from *D. hafniense* strain Y51 constitutes *pceA* followed by *pceB*, *pceC* and *pceT* [69]. PceT is the trigger factor involving in folding newly synthesized polypeptides. It interacts with the Tat motif of PceA, thus solubilizing and stabilizing PceA polypeptide proceeding downstream maturation and transportation processes [62]. PceB protein contains three transmembrane domains and is assumed as a membrane anchor protein of PceA. PceC contains six-trans‐ membrane domains, an FMN binding domain, and a C-terminal polyferredoxin-like domain. It is similar to the membrane-binding transcription regulators [28, 60]. More examples for the organization of different RDase gene clusters are presented in [28]. *RdhA* and *rdhB* genes usually locate adjacent and are the basic components of the *rdh* gene cluster. They would co-

The expression and silence of RDases during dechlorination reaction is dynamic and regulated. It could be monitored through the amount of RNA or protein. *Dehalococcoides* sp. strain MB bares 7 RDase genes. Only *dceA6* is highly expressed when PCE and TCE are transformed into tDCE. Transcription regulation protein binding site related to gene expression is detected in the upstream of *dceA6* gene [70]. A shotgun metagenome microarray is created to investigate gene transcription in a mixed culture. *rdhA14* and *rdhB14*, are the only two with higher transcript levels during VC degradation, while another 4 *rdhA* genes has higher transcript levels in the absence of VC [71]. The absolute quantification of RDase proteins during the dechlorination process is performed by using nano-liquid chromatography-tandem mass spectrometry in two PCE/TCE degradation consortia. Within 5 selected RDases, only the quantities of PceA and TceA are detectable [72]. The regulation on the expression of *rdh* genes during dechlorination reaction or steady state is not clear. How the physiological environments affect the gene expression is also unclear. Uncover these questions would be helpful for

**2.5. Genomic structure and transcription regulation of reductive dehalogenases**

of these genes are relevant to dehalogenation remains unclear.

express in order to perform dehalorespiration together.

environmental monitoring and remediation.

uvate or formate as electron donors [66].

The structures of HBCD and hexachlorocyclohexanes (HCHs) are quiet similar. Heeb et al. purified the HCH-converting haloalkane dehalogenase LinB, from *Sphigobium indicum* B90A and applied the enzyme for HBCD degradation. LinB transforms HBCD into pentabromocy‐ clododecanols (PBCDOHs) and further tetrabromocyclododecadiols (TBCDDOHs) [59]. Whether LinB or other haloalkane dehalogenase are the de novo HBCD degradation enzyme is unknown. What enzyme responsible for HBCD degradation in *Pseudomonas* sp. HB01 is also waited to be uncovered.

Reductive dechlorination reactions are catalyzed by reductive dehalogenases (RDases) The purified PCE RDase, PceA, has proved to transform PCE and TCE to cDCE [60]. The function of TCE RDase, TceA, in transforming TCE to ethene has also been identified [61]. VcrA and BvcA catalyze the transformation of DCE to ethene [62, 63]. In addition to chloroethenes, RDases also could reduce other chlorinated compounds.

RDases which could debrominate PBDEs have not yet been identified. However, some PBDEs degradation bacteria also could transform chloroethene (Table 2), such as *Dehalococcoides* sp., *Desulfitobacterium* sp., and *Sulfurospirillum* sp. Whether these microorganisms use chloroethene RDases to transform PBDEs is unknown. It is also possible that enzymes with different degradation activities or substrate specificity within single degrader may cooperatively transform different PBDEs congeners.

#### **2.4. The structure and function of reductive dehalogenase**

Most RDases presented similar features and conserved motifs [28, 29]. In the N-terminus, RDases possess a putative signal sequence containing the twin-arginine translocation (Tat) motif. Such motif is presented in secretary proteins to be transported across the cytoplasmic membrane through the Tat export system. It is proposed that newly synthesized RDase proteins is folded with cofactors (corrinoid and iron-sulfur clusters) in the cytoplasm with the aids of chaperone proteins. The Tat sequence is then proteolytically cleaved during the maturation process. In the C-terminus, two iron-sulfur cluster binding motifs are presented. The Fe-S clusters cooperate with corrinoid, transfering electrons from upstream donors to chloroethenes and thus catalyze the dehalogenation reaction [28].

The localization of chloroethene RDases is supposed in the membrane, where they could accept electrons from proton producing hydrogenase via menaquinone. The membranebound characteristics of RDases has been proved, such as PceA of *D. ethenogenes*. The lo‐ calization of constitutively expressed PceA in *S. multivorans* was initially found in the cytoplasmic fraction [65]. John et al. used freeze-fracture replica immunogold labeling technique and found it would be at the cytoplasmic membrane when cells grown on pyr‐ uvate or formate as electron donors [66].

#### **2.5. Genomic structure and transcription regulation of reductive dehalogenases**

ated DEs to be degraded [40]. Downstream degradation processes are supposed through phenol hydroxylases and catechol 1,2-dioxygenase. The transformation by-products range from phenol, catechol, halophenol and halocatechol, indicating nonspecific attack reactions [40, 41]. 2,3-dioxygenase is responsible to dihydroxylate lower numbered PBDEs and their similar chemicals such as DE in the close species *Sphingomonas sp.* PH-07 [42]. The range of PBDEs transformed by *R. jostii* RHA1 depends on the types of growth substrate. The enzymes responsible for degradation are inducible [43]. The expression of biphenyl dioxygenase (BPDO) and ethylbenzene dioxygenase (EBDO) are upregulated during PBDEs degradation. Ectopically expression of these enzymes in closed bacteria that bears no PBDEs degradation activity could transform PBDEs. EBDO depleted mono- through penta-BDEs and BPDO only

The structures of HBCD and hexachlorocyclohexanes (HCHs) are quiet similar. Heeb et al. purified the HCH-converting haloalkane dehalogenase LinB, from *Sphigobium indicum* B90A and applied the enzyme for HBCD degradation. LinB transforms HBCD into pentabromocy‐ clododecanols (PBCDOHs) and further tetrabromocyclododecadiols (TBCDDOHs) [59]. Whether LinB or other haloalkane dehalogenase are the de novo HBCD degradation enzyme is unknown. What enzyme responsible for HBCD degradation in *Pseudomonas* sp. HB01 is also

Reductive dechlorination reactions are catalyzed by reductive dehalogenases (RDases) The purified PCE RDase, PceA, has proved to transform PCE and TCE to cDCE [60]. The function of TCE RDase, TceA, in transforming TCE to ethene has also been identified [61]. VcrA and BvcA catalyze the transformation of DCE to ethene [62, 63]. In addition to chloroethenes,

RDases which could debrominate PBDEs have not yet been identified. However, some PBDEs degradation bacteria also could transform chloroethene (Table 2), such as *Dehalococcoides* sp., *Desulfitobacterium* sp., and *Sulfurospirillum* sp. Whether these microorganisms use chloroethene RDases to transform PBDEs is unknown. It is also possible that enzymes with different degradation activities or substrate specificity within single degrader may cooperatively

Most RDases presented similar features and conserved motifs [28, 29]. In the N-terminus, RDases possess a putative signal sequence containing the twin-arginine translocation (Tat) motif. Such motif is presented in secretary proteins to be transported across the cytoplasmic membrane through the Tat export system. It is proposed that newly synthesized RDase proteins is folded with cofactors (corrinoid and iron-sulfur clusters) in the cytoplasm with the aids of chaperone proteins. The Tat sequence is then proteolytically cleaved during the maturation process. In the C-terminus, two iron-sulfur cluster binding motifs are presented. The Fe-S clusters cooperate with corrinoid, transfering electrons from upstream donors to

depleted mono-, di- and one tetra-BDEs. [58].

56 Biodegradation of Hazardous and Special Products

RDases also could reduce other chlorinated compounds.

**2.4. The structure and function of reductive dehalogenase**

chloroethenes and thus catalyze the dehalogenation reaction [28].

transform different PBDEs congeners.

waited to be uncovered.

The major catalysis reaction of RDases is directed by subunit A, encoded by reductive dehalogenase homologous A (*rdhA*) genes. Over 650 *rdhA* genes have been identified based upon genomic sequence annotation or homologous cloning [67]. However, most of them are not yet been functional characterized. It is common for one dehalorespiration bacterium baring multicopy of *rdhA* genes in the genome. Besides *tceA* and *pceA* genes, there is still 17 RDase genes with unknown function in the genome of *D. ethenogenes* strain 195 [68]. Whether the roles of these genes are relevant to dehalogenation remains unclear.

Most *rdhA* genes are organized with genes encoding for accessory proteins. The *pce-*gene cluster from *D. hafniense* strain Y51 constitutes *pceA* followed by *pceB*, *pceC* and *pceT* [69]. PceT is the trigger factor involving in folding newly synthesized polypeptides. It interacts with the Tat motif of PceA, thus solubilizing and stabilizing PceA polypeptide proceeding downstream maturation and transportation processes [62]. PceB protein contains three transmembrane domains and is assumed as a membrane anchor protein of PceA. PceC contains six-trans‐ membrane domains, an FMN binding domain, and a C-terminal polyferredoxin-like domain. It is similar to the membrane-binding transcription regulators [28, 60]. More examples for the organization of different RDase gene clusters are presented in [28]. *RdhA* and *rdhB* genes usually locate adjacent and are the basic components of the *rdh* gene cluster. They would coexpress in order to perform dehalorespiration together.

The expression and silence of RDases during dechlorination reaction is dynamic and regulated. It could be monitored through the amount of RNA or protein. *Dehalococcoides* sp. strain MB bares 7 RDase genes. Only *dceA6* is highly expressed when PCE and TCE are transformed into tDCE. Transcription regulation protein binding site related to gene expression is detected in the upstream of *dceA6* gene [70]. A shotgun metagenome microarray is created to investigate gene transcription in a mixed culture. *rdhA14* and *rdhB14*, are the only two with higher transcript levels during VC degradation, while another 4 *rdhA* genes has higher transcript levels in the absence of VC [71]. The absolute quantification of RDase proteins during the dechlorination process is performed by using nano-liquid chromatography-tandem mass spectrometry in two PCE/TCE degradation consortia. Within 5 selected RDases, only the quantities of PceA and TceA are detectable [72]. The regulation on the expression of *rdh* genes during dechlorination reaction or steady state is not clear. How the physiological environments affect the gene expression is also unclear. Uncover these questions would be helpful for environmental monitoring and remediation.

#### **2.6. The dynamic of degrading population and the evolution of degradation ability**

collages set biosparging wells for injection substrate and air into TCE contaminated area. Above 95% TCE was removed through cometabolic reactions because the elevated chemical oxygen demand (COD), microbial population, oxidation-reduction potential (ORP) and specific degrading genes after the supplement of substrate [78]. Shortage of auxiliary substrates or accumulation of toxic intermediates also decrease the dehalogenation effect, combined

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

59

Sequential anaerobic/aerobic biodegradation is one of the approaches to accelerate the degradation of recalcitrant halogenated compounds. Anaerobic degraders could target the higher-numbered halogenated compounds. Aerobic degraders only process lower-numbered halogenated compounds. They could transform the by-products produced from anaerobic degradation to antoxic compounds through metabolic or cometabolic reactions. Integration of

Chloroethene and PBDEs can be depleted by microscale or nanoscale zerovalent metals [79, 80]. Preliminary dehalogenation of highly halogenated compounds by the reduced metal to generate less halogenated byproducts those are susceptible for microbial degradation. Therefore, the integration of zerovalent metals with biodegradation promotes the dehaloge‐ nation efficiency of each type of remediation methods. Reductive debromination of DBDE with nanoscale ZVI (nZVI) results various intermediates ranging from nona-BDEs to tri-BDEs. The known aerobic PBDEs degrader, *Sphingomonas* sp. strain PH-07, which is able to grow in the presence of nZVI, aerobically mineralizes the low brominated-DEs (tri-BDEs – mono-BDEs) from nZVI treatment [81]. The interactions between metals and microbes are complicated and delicate. H2 generated from the oxidation of metals promotes the growth of some dehalores‐ piration microbes. Some microorganisms could reduce oxidized metals for multiple runs of reductive reactions or degrade target compounds through cometabolism. Co-incubation with ZVI, microbes in the DBDE-degrading anaerobic sludges hinders the accessibility of MZVI to DBDE and reduced the removal ability in initial stage. However, the synergistic effect in DBDE degradation appears later on. According to the analysis of the microbial community change, co-incubation with MZVI leads to the enrichment of heterotrophic microbial populations bearing nitrate- or iron-reducing activities. The interaction between MZVI and microbes contributed to the synergistic effect [54]. Not only is the growth of microbes affected by metals, but also the expression of functional RDases. Bare nZVI down-regulate the expression of *tceA* and *vcrA* genes while coated particles up-regulate their expression [82]. In addition to the reduced metals, combining the electro-fenton process in aerobic degradation is also a newly evolved and potential way in bioremediation. Application of electrolysis also stimulate the

There are advantages by using each type of remediation approach, while there are also limitations and drawbacks. Combining biodegradation with other abiotic/biotic degradation approaches could overcome their weakness and accelerate the degradation efficiency. The recalcitrant halogenated compounds could be completely mineralized. The impact on envi‐ ronment might also be minimized. Integration of different approaches is a new direction for

remediation methods may recover the above drawbacks.

these two systems makes it possible for complete mineralization.

microbial reductive dechlorination and oxidative activities [83].

future investigation.

The complete dehalogenation requires different microorganisms which bear various functions in degradation or growth support. Besides, it competes with methanogens and other reducers for H2. The snapshot of the microbial composition stands for specific ecological condition. The dynamic of composition reveals the effect of various remediation treatments and the interac‐ tion between microorganisms. The microbial compositions when co-incubated with zerovalent iron (ZVI) are analyzed by DGGE. The enrichment of iron-reducing bacteria would support the reduction activity of iron for multiple rounds of reactions; the enrichment of nitratereducing bacteria also facilitates the cometabolic dehalogenation. These may due to the synergistic effect [54]. Terminal restriction fragment length polymorphism (TRFLP) analysis is also used to analyze the microbial compositions [73]. The resolution limitation of these techniques makes underestimating the complexity of a community. Therefore, new technique is needed to detect specific microbes that are responsible for a key biodegradation process while present in the communities in low numbers. The 16S rRNA genes within a community could be analyzed by recently evolved pyrosequencing or phylogenetic microarray (Phylo‐ Chip). PhyloChip composes ten thousands unique 16S rRNA genes. The microbial composi‐ tions in TCE contaminated groundwater that is biostimulated or bioaugmented are analyzed. The increase of methanogens at late treatment stage coincident with the increase in methane concentration [74].

There is no close phylogenic relation among diverse dehalorespration degraders. Horizontal gene transfer (HGT) though transposable elements, transmissible plasmids or phage infections is assumed for such convergent evolution. Phylogenetic analyses of the sequences of *rdh* operon and the adjacent genomic structures support HGT. The *pceABC* operon in *D. haf‐ niense* strain TCE1 has been shown to be presented in a circularized transposable element, Tn-Dha1 [75].The single-copy transfer messenger RNA gene (*ssrA*) essential in bacteria is a common target for mobile element. Integration of mobile element results in the duplication of *ssrA* gene around transported gene cluster. Many strain-specific *rdhA* genes collocates within such structures and in a region of high genomic variability between *Dehalococcoides* strains [76]. According to the metagenomic sequence analysis, one prophage element is located adjacent to *tceAB* genes in the *Dehalococcoides*-containing consortium, KB-1. The failure in detecting *tceA* gene expression in virus and the more closed transposase genes indicating higher possibility for HGT through transposable elements [71]. It seems that dehalorespiration degraders do no acquire RDase genes through single way. This would increase the diversity of degraders and function of RDase, which is advantageous for remediation.

#### **3. Integration of biodegradation with other remediation methods**

The degradation rate of natural attenuation is slower than chemical or physical treatments. Biostimulation or bioaugement are common strategies for bioremediation of chloroethene [30] Chemical supplements such as potassium permanganate or oxygen injection which can increase oxygen concentration are benefit for microbial dechlorination [77]. Kuo and his collages set biosparging wells for injection substrate and air into TCE contaminated area. Above 95% TCE was removed through cometabolic reactions because the elevated chemical oxygen demand (COD), microbial population, oxidation-reduction potential (ORP) and specific degrading genes after the supplement of substrate [78]. Shortage of auxiliary substrates or accumulation of toxic intermediates also decrease the dehalogenation effect, combined remediation methods may recover the above drawbacks.

**2.6. The dynamic of degrading population and the evolution of degradation ability**

concentration [74].

58 Biodegradation of Hazardous and Special Products

The complete dehalogenation requires different microorganisms which bear various functions in degradation or growth support. Besides, it competes with methanogens and other reducers for H2. The snapshot of the microbial composition stands for specific ecological condition. The dynamic of composition reveals the effect of various remediation treatments and the interac‐ tion between microorganisms. The microbial compositions when co-incubated with zerovalent iron (ZVI) are analyzed by DGGE. The enrichment of iron-reducing bacteria would support the reduction activity of iron for multiple rounds of reactions; the enrichment of nitratereducing bacteria also facilitates the cometabolic dehalogenation. These may due to the synergistic effect [54]. Terminal restriction fragment length polymorphism (TRFLP) analysis is also used to analyze the microbial compositions [73]. The resolution limitation of these techniques makes underestimating the complexity of a community. Therefore, new technique is needed to detect specific microbes that are responsible for a key biodegradation process while present in the communities in low numbers. The 16S rRNA genes within a community could be analyzed by recently evolved pyrosequencing or phylogenetic microarray (Phylo‐ Chip). PhyloChip composes ten thousands unique 16S rRNA genes. The microbial composi‐ tions in TCE contaminated groundwater that is biostimulated or bioaugmented are analyzed. The increase of methanogens at late treatment stage coincident with the increase in methane

There is no close phylogenic relation among diverse dehalorespration degraders. Horizontal gene transfer (HGT) though transposable elements, transmissible plasmids or phage infections is assumed for such convergent evolution. Phylogenetic analyses of the sequences of *rdh* operon and the adjacent genomic structures support HGT. The *pceABC* operon in *D. haf‐ niense* strain TCE1 has been shown to be presented in a circularized transposable element, Tn-Dha1 [75].The single-copy transfer messenger RNA gene (*ssrA*) essential in bacteria is a common target for mobile element. Integration of mobile element results in the duplication of *ssrA* gene around transported gene cluster. Many strain-specific *rdhA* genes collocates within such structures and in a region of high genomic variability between *Dehalococcoides* strains [76]. According to the metagenomic sequence analysis, one prophage element is located adjacent to *tceAB* genes in the *Dehalococcoides*-containing consortium, KB-1. The failure in detecting *tceA* gene expression in virus and the more closed transposase genes indicating higher possibility for HGT through transposable elements [71]. It seems that dehalorespiration degraders do no acquire RDase genes through single way. This would increase the diversity of degraders and

function of RDase, which is advantageous for remediation.

**3. Integration of biodegradation with other remediation methods**

The degradation rate of natural attenuation is slower than chemical or physical treatments. Biostimulation or bioaugement are common strategies for bioremediation of chloroethene [30] Chemical supplements such as potassium permanganate or oxygen injection which can increase oxygen concentration are benefit for microbial dechlorination [77]. Kuo and his

Sequential anaerobic/aerobic biodegradation is one of the approaches to accelerate the degradation of recalcitrant halogenated compounds. Anaerobic degraders could target the higher-numbered halogenated compounds. Aerobic degraders only process lower-numbered halogenated compounds. They could transform the by-products produced from anaerobic degradation to antoxic compounds through metabolic or cometabolic reactions. Integration of these two systems makes it possible for complete mineralization.

Chloroethene and PBDEs can be depleted by microscale or nanoscale zerovalent metals [79, 80]. Preliminary dehalogenation of highly halogenated compounds by the reduced metal to generate less halogenated byproducts those are susceptible for microbial degradation. Therefore, the integration of zerovalent metals with biodegradation promotes the dehaloge‐ nation efficiency of each type of remediation methods. Reductive debromination of DBDE with nanoscale ZVI (nZVI) results various intermediates ranging from nona-BDEs to tri-BDEs. The known aerobic PBDEs degrader, *Sphingomonas* sp. strain PH-07, which is able to grow in the presence of nZVI, aerobically mineralizes the low brominated-DEs (tri-BDEs – mono-BDEs) from nZVI treatment [81]. The interactions between metals and microbes are complicated and delicate. H2 generated from the oxidation of metals promotes the growth of some dehalores‐ piration microbes. Some microorganisms could reduce oxidized metals for multiple runs of reductive reactions or degrade target compounds through cometabolism. Co-incubation with ZVI, microbes in the DBDE-degrading anaerobic sludges hinders the accessibility of MZVI to DBDE and reduced the removal ability in initial stage. However, the synergistic effect in DBDE degradation appears later on. According to the analysis of the microbial community change, co-incubation with MZVI leads to the enrichment of heterotrophic microbial populations bearing nitrate- or iron-reducing activities. The interaction between MZVI and microbes contributed to the synergistic effect [54]. Not only is the growth of microbes affected by metals, but also the expression of functional RDases. Bare nZVI down-regulate the expression of *tceA* and *vcrA* genes while coated particles up-regulate their expression [82]. In addition to the reduced metals, combining the electro-fenton process in aerobic degradation is also a newly evolved and potential way in bioremediation. Application of electrolysis also stimulate the microbial reductive dechlorination and oxidative activities [83].

There are advantages by using each type of remediation approach, while there are also limitations and drawbacks. Combining biodegradation with other abiotic/biotic degradation approaches could overcome their weakness and accelerate the degradation efficiency. The recalcitrant halogenated compounds could be completely mineralized. The impact on envi‐ ronment might also be minimized. Integration of different approaches is a new direction for future investigation.

#### **4. Conclusion**

The current knowledge of microbial degradation of chloroethenes, PBDEs and HBCD, has been summarized and reviewed. The biodegradation of these halogenated compounds through aerobic oxidation, aerobic cometabolization, or reductive dehalogenation are introduced. The correspondent enzymes are discussed from the biochemical and molecular point of view. The structure and function of RDases, as well as gene expression regulation and genomic evolution are the major focus. Integration and sequential anaerobic/aerobic biodegradation or (elec‐ tro)chemical/microbial degradation are suggested for overcoming the disadvantages of single type of treatment. It is possible to completely mineralize these halogenated pollutants by the combination of bio- and abiotic processes and shows promise for site remediation in natural settings and in engineered systems.

Molecular weight 165/131 249.1 ~ 959.2 641.7

Epidermis, liver and kidney damage, immune- and neuro-toxicity, reproductive and endocrine effects, probably cancer

Chemical oxidation ,

a: the toxicity of HBCD is based upon investigation in animal model.

Hexa-BDE to mono-

PCE Cl-

Tri-, di-, and mono-

BDE,

*Pseudomonas sp. HB01* HBCD PBCDOHs,

*Rhodococcus jostii RHA1*Penta-BDE to mono-BDE

BDE

bromo-DE

*Sphingomonas sp. SS3* Fluoro-, chloro-, and

reduced metals (Fe, Fe/Pd)

irradiation,

150/1280 4.8 ~0.02 0.003

neurotoxicity

Pyrolysis, photolysis,

Biological degradationAerobic/anaerobic Aerobic/anaerobic Aerobic/anaerobic(?)

**Table 1.** Physicochemical properties, biological impacts and degradation routes of chloroethenes, PBDEs and HBCD.

**Substrate End-products Genes c references**

DBDE ND [44]

Mono-BDE ND [43]

Hydroxylated-BDE BPDO- *bphAa*,

TBCDDOHs

*Rhodococcus sp. RR1* Di-, and mono-BDE ND [43]

Catechol, dibromophenol, dihydroxy mono- and dibromo-BDE

Phenol, catechol, Halophenol and Halocatehol

Hydroxylated-BDE [43]

, ToMO-*touABCDEF* [39]

EBDO- *etbAa1*, *etbAc*

Disrupt the balance of thyroid hormone, reproductive, hepatic, and immunotoxicity, developmental

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

reduced metals (Fe, Fe/Pd),

Thyroidal and developmental toxicity

http://dx.doi.org/10.5772/56306

61

Pyrolysis, photolysis

[45]

[43, 84]

[42]

[40]

Water solubility (mg

Abiotic degradation

l -1)

Toxicitya

*Aerobic Burkholderia xenovorans LB400*

*strain DB-1*

*OX1*

*PH-07*

*Pseudonocardia dioxanivorans CB1190*

*Lysinibacillus fusiformis*

*Pseudomonas stutzeri*

*Sphingomonas sp.*

**Figure 1.** Aerobic degradation and anaerobic reductive dehalogenation reactions of chloeoethenes, PBDEs and HBCD. Circle dot indicates above compounds; star indicates reductive dehalogenation driven by the oxidation of electron do‐ nors or occurring cometabolically with other dehalorespiration process; gray arrow indicates the hydrolytic dehaloge‐ nation of HBCD.

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features http://dx.doi.org/10.5772/56306 61


**4. Conclusion**

nation of HBCD.

settings and in engineered systems.

60 Biodegradation of Hazardous and Special Products

The current knowledge of microbial degradation of chloroethenes, PBDEs and HBCD, has been summarized and reviewed. The biodegradation of these halogenated compounds through aerobic oxidation, aerobic cometabolization, or reductive dehalogenation are introduced. The correspondent enzymes are discussed from the biochemical and molecular point of view. The structure and function of RDases, as well as gene expression regulation and genomic evolution are the major focus. Integration and sequential anaerobic/aerobic biodegradation or (elec‐ tro)chemical/microbial degradation are suggested for overcoming the disadvantages of single type of treatment. It is possible to completely mineralize these halogenated pollutants by the combination of bio- and abiotic processes and shows promise for site remediation in natural

**Figure 1.** Aerobic degradation and anaerobic reductive dehalogenation reactions of chloeoethenes, PBDEs and HBCD. Circle dot indicates above compounds; star indicates reductive dehalogenation driven by the oxidation of electron do‐ nors or occurring cometabolically with other dehalorespiration process; gray arrow indicates the hydrolytic dehaloge‐

**PCE/TCE PBDEs HBCD**

O

Br Br

Br Br

Br

Br

Br

Br

Cl Cl

Cl Cl

or H

**Table 1.** Physicochemical properties, biological impacts and degradation routes of chloroethenes, PBDEs and HBCD.



**Acknowledgements**

**Author details**

**References**

Yu-Huei Peng and Yang-hsin Shih\*

\*Address all correspondence to: yhs@ntu.edu.tw

[2] Tetrachloroethylene. IARC monograph 2007.

[5] 1,2-dichloroethene. ATSDR 1997.

Chemosphere 2008; 70(4) 640-647.

591-599.

gy 2011; 63(5) 413-417.

[4] Toxicologica Review of Vinyl Chloride. USEPA 2000.

The authors thank National Science Council (NSC), Taiwan, ROC for financial support.

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

63

Department of Agricultural Chemistry, National Taiwan University, Taipei, Taiwan

[1] Toxicological Profile for Tetrachloroethylene (Update). ATSDR 1997.

[3] Toxicological Profile for Trichloroethylene (Update). ATSDR 1997.

[6] Costa L. G., Giordano G. Developmental neurotoxicity of polybrominated diphenyl

[7] Tseng L. H. et al. Developmental exposure to decabromodiphenyl ether (PBDE 209): effects on thyroid hormone and hepatic enzyme activity in male mouse offspring.

[8] An J. et al. The cytotoxic effects of synthetic 6-hydroxylated and 6-methoxylated pol‐ ybrominated diphenyl ether 47 (BDE47). Environmental Toxicology 2011; 26(6)

[9] Yan C. et al. The involvement of ROS overproduction and mitochondrial dysfunction in PBDE-47-induced apoptosis on Jurkat cells. Experimental and Toxicologic Patholo‐

[10] Gauthier L. T. et al. Temporal trends and spatial distribution of non-polybrominated diphenyl ether flame retardants in the eggs of colonial populations of Great Lakes

herring gulls. Environmental Science & Technology 2009; 43(2) 312-317.

ether (PBDE) flame retardants. Neurotoxicology 2007; 28(6) 1047-1067.

a penta- BDE mixture: hexa-, penta-, and tera-BDEs.

b Octa-BDE mixture: nona-, octa-, hepta-, and hexa-BDEs.

c Genes that only relevant to degradation of chloroethens, PBDEs and HBCD are listed.

ND: data not shown. BPDO: biphenyl dioxygenase. EBDO: ethylbenzene dioxygenase. PBCDOHs: pentabromocyclodo‐ decanols. TBCDDOHs: tetrabromocyclododecadiols.

**Table 2.** Selected bacteria which degrade chloroethens, PBDEs and HBCD.

#### **Acknowledgements**

*Sphingomonas sp. SS33* Di-, and mono-;

62 Biodegradation of Hazardous and Special Products

*Anaerobic*

*strain AG*

*strain DG*

*strain MB*

*Co23*

*SZ*

*Acetobacterium sp.*

*Dehalococcoides sp. strain BAV1*

*Dehalococcoides sp.*

*Dehalococcoides ethenogenes strain 195*

*Dehalococcoides sp.*

*Desulfitobacterium chlororespirans strain*

*Desulfitobacterium dehalogenans strain JW/IU-DC1*

*Desulfitobacterium hafniense PCP-1*

*Sulfurospirillum multivorans*

*Geobacter lovleyi strain*

fluoro-, chloro-, and

Phenol, catechol, di-,

[41]

[52]

[52]

*pceA, tceA* [51, 67, 89]

*pceA* [51, 67]

and monohalophenol, di-, and mono-halocatehol

Penta-BDE mixturea Tetra-, tri-, and di-

BDEs

*Bacillus sp JSK1* PCE *Cis-*DCE [85] *Dehalobacter restrictus* PCE, TCE *Cis-*DCE *pceA,* [67, 86]

Ethene

BDEs Tetra- BDE

Ethene

DCE

BDEs

c Genes that only relevant to degradation of chloroethens, PBDEs and HBCD are listed.

**Table 2.** Selected bacteria which degrade chloroethens, PBDEs and HBCD.

Octa- and hepta-

ND: data not shown. BPDO: biphenyl dioxygenase. EBDO: ethylbenzene dioxygenase. PBCDOHs: pentabromocyclodo‐

Tetra- and penta-

Tetra-, penta-, hexaand Penta-BDEs

DCE, VC Ethene *bvcA* [49, 87]

PCE, TCE *Trans*-DCE [90]

Octa-BDE ND [53]

Octa-BDE ND [53]

PCE, TCE cDCE [92]

ND [53, 67, 91]

bromo-DE

TCE

VC

PCE, TCE, Octa-BDE,

PCE, TCE DBDE

a penta- BDE mixture: hexa-, penta-, and tera-BDEs. b Octa-BDE mixture: nona-, octa-, hepta-, and hexa-BDEs.

decanols. TBCDDOHs: tetrabromocyclododecadiols.

Octa-BDE mixtureb Penta- BDE mixturea

PCE, TCE, cis -DCE, and

Octa-BDE mixtureb

The authors thank National Science Council (NSC), Taiwan, ROC for financial support.

#### **Author details**

Yu-Huei Peng and Yang-hsin Shih\*

\*Address all correspondence to: yhs@ntu.edu.tw

Department of Agricultural Chemistry, National Taiwan University, Taipei, Taiwan

#### **References**


[11] Marvin C. H. et al. Hexabromocyclododecane: Current Understanding of Chemistry, Environmental Fate and Toxicology and Implications for Global Management. Envi‐ ronmental Science & Technology 2011; 45(20) 8613-8623.

[26] Shih Y. H. et al. Reduction of hexachlorobenzene by nanoscale zero-valent iron: ki‐ netics, pH effect, and degradation mechanism. Separation and purification technolo‐

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

65

[27] Lee C. et al. Bactericidal effect of zero-valent iron nanoparticles on Escherichia coli.

[28] Futagami T. et al. Biochemical and genetic bases of dehalorespiration. Chemical re‐

[29] Smidt H., de Vos W. M. Anaerobic microbial dehalogenation. Annual review of mi‐

[30] Tiehm A., Schmidt K. R. Sequential anaerobic/aerobic biodegradation of chloroe‐ thenes - aspects of field application. Current Opinion in Biotechnology 2011; 22(3)

[31] Huang H. et al. In vitro biotransformation of PBDEs by root crude enzyme extracts: Potential role of nitrate reductase (NaR) and glutathione S-transferase (GST) in their

[32] Davis J. W. et al. The transformation of hexabromocyclododecane in aerobic and anaerobic soils and aquatic sediments. Water Research 2005; 39(6) 1075-1084.

[33] Hakk H., Letcher R. J. Metabolism in the toxicokinetics and fate of brominated flame

[34] Tiehm A. et al. Growth kinetics and stable carbon isotope fractionation during aero‐ bic degradation of cis-1,2-dichloroethene and vinyl chloride. Water Research 2008;

[35] Olaniran A. O. et al. Aerobic biodegradation of dichloroethenes by indigenous bacte‐ ria isolated from contaminated sites in Africa. Chemosphere 2008; 73(1) 24-29.

[36] Schmidt K. R. et al. Aerobic biodegradation of cis-1,2-dichloroethene as sole carbon source: Stable carbon isotope fractionation and growth characteristics. Chemosphere

[37] Zhang Y., Tay J. H. Co-metabolic degradation activities of trichloroethylene by phe‐ nol-grown aerobic granules. Journal of Biotechnology 2012; 162(2-3) 274-282.

[38] Frascari D. et al. A kinetic study of chlorinated solvent cometabolic biodegradation by propane-grown Rhodococcus sp. PB1. Biochemical Engineering Journal 2008;

[39] Ryoo D. et al. Aerobic degradation of tetrachloroethylene by toluene-o-xylene mono‐ oxygenase of Pseudomonas stutzeri OX1. Nat Biotech 2000; 18(7) 775-778.

retardants--a review. Environmental international 2003; 29(6) 801-828.

Environmental Science & Technology 2008; 42(13) 4927-4933.

debromination. Chemosphere (2012)., 90(6), 1885-1892.

gy (2011)., 76(3), 268-274.

cord 2008; 8(1) 1-12.

42(10–11) 2431-2438.

2010; 78(5) 527-532.

42(2) 139-147.

415-421.

crobiology (2004)., 58, 43-73.


[26] Shih Y. H. et al. Reduction of hexachlorobenzene by nanoscale zero-valent iron: ki‐ netics, pH effect, and degradation mechanism. Separation and purification technolo‐ gy (2011)., 76(3), 268-274.

[11] Marvin C. H. et al. Hexabromocyclododecane: Current Understanding of Chemistry, Environmental Fate and Toxicology and Implications for Global Management. Envi‐

[12] Deng J. et al. Hexabromocyclododecane-induced developmental toxicity and apopto‐

[15] Summary of a Proposal to List Hexabromocyclododecane in Annex A to the Conven‐

[16] Honning J. et al. Role of diffusion in chemical oxidation of PCE in a dual permeabili‐

[17] Weber R. et al. PCB destruction in subcritical and supercritical water--evaluation of PCDF formation and initial steps of degradation mechanisms. Environmental Science

[18] Lin Y. M. et al. Emissions of Polybrominated Diphenyl Ethers during the Thermal Treatment for Electric Arc Furnace Fly Ash. Aerosol and Air Quality Research 2012;

[19] Barontini F. et al. Thermal stability and decomposition products of hexabromocyclo‐ dodecane. Industrial & Engineering Chemistry Research 2001; 40(15) 3270-3280. [20] Shih Y.-h., Wang C.-K. Photolytic degradation of polybromodiphenyl ethers under UV-lamp and solar irradiations. Journal of Hazardous Materials 2009; 165(1-3) 34-38.

[21] Beitinger E. Permeable treatment walls - Design, construction, and cost. in: NATO/ CCMS Pilot Study: Evaluation of Demonstrated and Emerging Technologies for the

[22] He F., Zhao D. Y. Hydrodechlorination of trichloroethene using stabilized Fe-Pd nanoparticles: Reaction mechanism and effects of stabilizers, catalysts and reaction

[23] Shih Y. H., Tai Y. T. Reaction of decabrominated diphenyl ether by zerovalent iron

[24] Su Y.-f. et al. Effects of various ions on the dechlorination kinetics of hexachloroben‐

[25] Shih Y.-h. et al. Pentachlorophenol reduction by Pd/Fe bimetallic nanoparticles: Ef‐ fects of copper, nickel, and ferric cations. Applied Catalysis B: Environmental 2011;

zene by nanoscale zero-valent iron. Chemosphere 2012; 88(11) 1346-1352.

Treatment of Contaminated land and Grooundwater (Phase III). 1998.

conditions. Applied Catalysis B-Environmental 2008; 84(3-4) 533-540.

nanoparticles. Chemosphere 2010; 78(10) 1200-1206.

ty system. Environmental Science & Technology 2007; 41(24) 8426-8432.

ronmental Science & Technology 2011; 45(20) 8613-8623.

sis in zebrafish embryos. Aquat Toxicol 2009; 93(1) 29-36.

[14] Hexabromocyclododecane (HBCD) Action Plan. USEPA (2010).

[13] HBCD Factsheet. BSEF 2009.

64 Biodegradation of Hazardous and Special Products

tion. POPRC 2009.

12(2) 237-250.

105(1-2) 24-29.

& Technology 2002; 36(8) 1839-1844.


[40] Schmidt S. et al. Biodegradation of diphenyl ether and its monohalogenated deriva‐ tives by Sphingomonas sp. strain SS3. Applied and Environmental Microbiology 1992; 58(9) 2744-50.

[53] Robrock K. R. et al. Pathways for the anaerobic microbial debromination of polybro‐ minated diphenyl ethers. Environmental Science & Technology 2008; 42(8) 2845-52.

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

67

[54] Shih Y. H. et al. Synergistic effect of microscale zerovalent iron particles combined with anaerobic sludges on the degradation of decabromodiphenyl ether. Bioresource

[55] Chen C. Y. et al. Microbial degradation of 4-monobrominated diphenyl ether in an aerobic sludge and the DGGE analysis of diversity. Journal of Environmental Science and Health Part B-Pesticides Food Contaminants and Agricultural Wastes 2010; 45(5)

[56] Mattes T. E. et al. Aerobic biodegradation of the chloroethenes: pathways, enzymes,

[57] Furukawa K. Oxygenases and dehalogenases: molecular approaches to efficient deg‐ radation of Chlorinated environmental pollutants. Bioscience, Biotechnology, and Bi‐

[58] Robrock K. R. et al. Biphenyl and Ethylbenzene Dioxygenases of Rhodococcus jostii RHA1 Transform PBDEs. biotechnology and bioengineering 2011; 108(2) 313-321. [59] Heeb N. V. et al. Biotransformation of Hexabromocyclododecanes (HBCDs) with LinB—An HCH-Converting Bacterial Enzyme. Environmental Science & Technology

[60] Suyama A. et al. Molecular Characterization of the PceA Reductive Dehalogenase of Desulfitobacterium sp. Strain Y51. Journal of Bacteriology 2002; 184(13) 3419-3425. [61] Magnuson J. K. et al. Trichloroethene reductive dehalogenase from Dehalococcoides ethenogenes: Sequence of tceA and substrate range characterization. Applied and

[62] Maillard J. et al. Redundancy and specificity of multiple trigger factor chaperones in

[63] Tang S. et al. Functional Characterization of Reductive Dehalogenases Using Blue Native Polyacrylamide Gel Electrophoresis. Applied and Environmental Microbiolo‐

[64] Maillard J. et al. Characterization of the corrinoid iron-sulfur protein tetrachloroe‐ thene reductive dehalogenase of Dehalobacter restrictus. Applied and Environmen‐

[65] John M. et al. Growth substrate dependent localization of tetrachloroethene reduc‐ tive dehalogenase in Sulfurospirillum multivorans. Archives of Microbiology 2006;

ecology, and evolution. FEMS Microbiology Reviews 2010; 34(4) 445-475.

Technology (2012)., 108, 14-20.

ochemistry 2006; 70(10) 2335-2348.

Environmental Microbiology 2000; 66(12) 5141-5147.

Desulfitobacteria. Microbiology-Sgm (2011)., 157, 2410-2421.

2012; 46(12) 6566-6574.

gy (2012)., 79(3), 974-981.

186(2) 99-106.

tal Microbiology 2003; 69(8) 4628-4638.

379-385.


[53] Robrock K. R. et al. Pathways for the anaerobic microbial debromination of polybro‐ minated diphenyl ethers. Environmental Science & Technology 2008; 42(8) 2845-52.

[40] Schmidt S. et al. Biodegradation of diphenyl ether and its monohalogenated deriva‐ tives by Sphingomonas sp. strain SS3. Applied and Environmental Microbiology

[41] Schmidt S. et al. Biodegradation and Transformation of 4,4'-Dihalodiphenyl and 2,4- Dihalodiphenyl Ethers by Sphingomonas Sp Strain Ss33. Applied and Environmental

[42] Kim Y. M. et al. Biodegradation of diphenyl ether and transformation of selected bro‐ minated congeners by Sphingomonas sp. PH-07. Applied microbiology and biotech‐

[43] Robrock K. R. et al. Aerobic biotransformation of polybrominated diphenyl ethers (PBDEs) by bacterial isolates. Environmental Science & Technology 2009; 43(15)

[44] Deng D. et al. Aerobic debromination of deca-BDE: Isolation and characterization of an indigenous isolate from a PBDE contaminated sediment. International Biodeterio‐

[45] Yamada T. et al. Isolation of Pseudomonas sp. strain HB01 which degrades the per‐ sistent brominated flame retardant gamma-hexabromocyclododecane. Bioscience, Bi‐

[46] Aulenta F. et al. Competition for H2 between sulfate reduction and dechlorination in butyrate-fed anaerobic cultures. Process Biochemistry (2008)., 43(2), 161-168.

[47] Conrad M. E. et al. Field evidence for co-metabolism of trichloroethene stimulated by addition of electron donor to groundwater. Environmental Science & Technology

[48] John M. et al. Retentive Memory of Bacteria: Long-Term Regulation of Dehalorespi‐ ration in Sulfurospirillum multivorans. Journal of Bacteriology 2009; 191(5)

[49] He J. Z. et al. Detoxification of vinyl chloride to ethene coupled to growth of an anae‐

[50] Lee L. K., He J. Reductive Debromination of Polybrominated Diphenyl Ethers by Anaerobic Bacteria from Soils and Sediments. Applied and environmental microbiol‐

[51] He J. Z. et al. Microbial reductive debromination of polybrominated diphenyl ethers

[52] Ding C. et al. Isolation of Acetobacterium sp. strain AG that reductively debromi‐ nates octa- and penta-brominated diphenyl ether technical mixtures. Applied and

(PBDEs). Environmental Science & Technology 2006; 40(14) 4429-4434.

Environmental Microbiology (2012)., doi:10.1128/AEM.02919-12

1992; 58(9) 2744-50.

66 Biodegradation of Hazardous and Special Products

Microbiology 1993; 59(11) 3931-3933.

ration & Biodegradation 2011; 65(3) 465-469.

robic bacterium. Nature 2003; 424(6944) 62-65.

otechnology, and Biochemistry 2009; 73(7) 1674-1678.

nology 2007; 77(1) 187-194.

(2010)., 44(12), 4697-4704.

ogy 2010; 76(3) 794-802.

1650-1655.

5705-5711.


[66] Regeard C. et al. Development of degenerate and specific PCR primers for the detec‐ tion and isolation of known and putative chloroethene reductive dehalogenase genes. J Microbiol Methods 2004; 56(1) 107-18.

[79] Kim Y.-M. et al. Degradation of polybrominated diphenyl ethers by a sequential treatment with nanoscale zero valent iron and aerobic biodegradation. Journal of

Microbial Degradation of Some Halogenated Compounds: Biochemical and Molecular Features

http://dx.doi.org/10.5772/56306

69

[80] Xiu Z. M. et al. Effect of Bare and Coated Nanoscale Zerovalent Iron on tceA and vcrA Gene Expression in Dehalococcoides spp. Environmental Science & Technology

[81] Lohner S. T., Tiehm A. Application of Electrolysis to Stimulate Microbial Reductive PCE Dechlorination and Oxidative VC Biodegradation. Environmental Science &

[82] Robrock K. R., Mohn, W.W., Eltis, L.D., Alvarez-Cohen, L. Biphenyl and ethylben‐ zene dioxygenases of Rhodococcus jostii RHA1 transform PBDEs. biotechnology and

[83] Kalimuthu K. et al. Reductive dechlorination of perchloroethylene by bacillus sp JSK1 isolated from dry cleaning industrial sludge. Carpathian Journal of Earth and

[84] Holliger C. et al. A highly purified enrichment culture couples the reductive dech‐ lorination of tetrachloroethene to growth. Applied and Environmental Microbiology

[85] Krajmalnik-Brown R. et al. Genetic identification of a putative vinyl chloride reduc‐ tase in Dehalococcoides sp. strain BAV1. Applied and Environmental Microbiology

[86] MaymoGatell X. et al. Isolation of a bacterium that reductively dechlorinates tetra‐

[87] Cheng D., He J. Z. Isolation and Characterization of "Dehalococcoides" sp Strain MB, Which Dechlorinates Tetrachloroethene to trans-1,2-Dichloroethene. Applied and En‐

[88] Villemur R. et al. Occurrence of several genes encoding putative reductive dehaloge‐ nases in Desulfitobacterium hafniense/frappieri and Dehalococcoides ethenogenes.

[89] Sung Y. et al. Geobacter lovleyi sp. nov. Strain SZ, a Novel Metal-Reducing and Tet‐ rachloroethene-Dechlorinating Bacterium. Applied and Environmental Microbiology

chloroethene to ethene. Science 1997; 276(5318) 1568-1571.

vironmental Microbiology 2009; 75(18) 5910-5918.

Canadian journal of microbiology 2002; 48(8) 697-706.

Chemical Technology & Biotechnology 2012; 87(2) 216-224.

2010; 44(19) 7647-7651.

1993; 59(9) 2991-2997.

2004; 70(10) 6347-51.

2006; 72(4) 2775-2782.

Technology 2009; 43(18) 7098-7104.

bioengineering 2010; 108(2) 313-321.

Environmental Sciences 2011; 6(1) 165-170.


[79] Kim Y.-M. et al. Degradation of polybrominated diphenyl ethers by a sequential treatment with nanoscale zero valent iron and aerobic biodegradation. Journal of Chemical Technology & Biotechnology 2012; 87(2) 216-224.

[66] Regeard C. et al. Development of degenerate and specific PCR primers for the detec‐ tion and isolation of known and putative chloroethene reductive dehalogenase

[67] West K. A. et al. Comparative genomics of "Dehalococcoides ethenogenes" 195 and an enrichment culture containing unsequenced "Dehalococcoides" strains. Applied

[68] Furukawa K. et al. Biochemical and molecular characterization of a tetrachloroethene dechlorinating Desulfitobacterium sp strain Y51: a review. Journal of Industrial Mi‐

[69] Chow W. L. et al. Identification and transcriptional analysis of trans-DCE-producing reductive dehalogenases in Dehalococcoides species. ISME journal 2010; 4(8)

[70] Waller A. S. et al. Transcriptional Analysis of a Dehalococcoides-Containing Microbi‐ al Consortium Reveals Prophage Activation. Applied and Environmental Microbiol‐

[71] Werner J. J. et al. Absolute quantification of Dehalococcoides proteins: enzyme bioin‐ dicators of chlorinated ethene dehalorespiration. Environmental microbiology 2009;

[72] Révész S. et al. Bacterial community changes in TCE biodegradation detected in mi‐ crocosm experiments. International Biodeterioration & Biodegradation 2006; 58(3–4)

[73] Lee P. K. H. et al. Phylogenetic Microarray Analysis of a Microbial Community Per‐ forming Reductive Dechlorination at a TCE-Contaminated Site. Environmental Sci‐

[74] Maillard J. et al. Isolation and characterization of Tn-Dha1, a transposon containing the tetrachloroethene reductive dehalogenase of Desulfitobacterium hafniense strain

[75] Sahl J. W. et al. Coupling permanganate oxidation with microbial dechlorination of

[76] Kuo Y. C. et al. Remediation of TCE-contaminated groundwater using integrated bi‐ osparging and enhanced bioremediation system. Research Journal of Chemistry and

[77] Liu Y. Q. et al. TCE dechlorination rates, pathways, and efficiency of nanoscale iron particles with different properties. Environmental Science & Technology 2005; 39(5)

[78] Keum Y. S., Li Q. X. Reductive debromination of polybrominated diphenyl ethers by zerovalent iron. Environmental Science & Technology 2005; 39(7) 2280-2286.

genes. J Microbiol Methods 2004; 56(1) 107-18.

and Environmental Microbiology 2008; 74(11) 3533-3540.

crobiology & Biotechnology 2005; 32(11-12) 534-541.

1020-1030.

ogy 2012; 78(4) 1178-1186.

68 Biodegradation of Hazardous and Special Products

ence & Technology 2011; 46(2) 1044-1054.

Environment 2012; 16(2) 37-47.

TCE1. Environmental microbiology 2005; 7(1) 107-117.

tetrachloroethene. Water Environ Res. 2007; 79(1) 5-12.

11(10) 2687-2697.

239-247.

1338-1345.


**Chapter 5**

**Biodegradation of Melanoidin from Distillery Effluent:**

Bioremediation is the process that deals with the microbial degradation of hazardous com‐ pounds from environment. The process of bioremediation is the natural process of biodegra‐ dation, which can degrade the pollutants and sometimes can completely oxidize the compound. Microorganisms play the vital role in the process of bioremediation and biode‐ gradation because of their great metabolic diversity, which includes the ability to metabolise these pollutants [1]. The degradation of toxic compounds to less harmful forms with the use of biological systems is called as bioremediation [2]. Bioremediation is limited in the number of toxic material, it can handle but where applicable it is cost effective and ecofriendly [3]. Today, water resources have been the most exploited of the natural systems, most of our water bodies are seriously polluted due to rapid population growth, industrial proliferation, urbanizations, increasing living standards and wide spheres of human activities. Many rivers of the world receive heavy flux due to industrial effluents [4]. The wastewater consisting of substances varying from simple nutrients to highly toxic hazardous chemicals, which when used for irrigation caused both beneficial and damaging effects to various crops including

The awful consequence of increasing population along with an rapidly increase in the industrial field is a major concern of pollution in the environment. Water is necessary for irrigation to increase the productivity in agriculture. Industrial effluents already pollute most of the Indian rivers. Many industries are playing the crucial role in water pollution such as textile industries, dairy industries and distillery *etc*. Distilleries are the major agro-based industries, which utilize molasses as raw material for the production of rectified spirit. In addition to rectified spirit, distilleries also produce ethanol, which can be mixed with diesel and used as biofuel and helps in reducing import of crude oil thereby saving foreign exchange

and reproduction in any medium, provided the original work is properly cited.

© 2013 Santal and Singh; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

**Role of Microbes and Their Potential Enzymes**

Anita Rani Santal and Nater Pal Singh

http://dx.doi.org/10.5772/56252

**1. Introduction**

vegetables [5].

Additional information is available at the end of the chapter

### **Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes**

Anita Rani Santal and Nater Pal Singh

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56252

#### **1. Introduction**

Bioremediation is the process that deals with the microbial degradation of hazardous com‐ pounds from environment. The process of bioremediation is the natural process of biodegra‐ dation, which can degrade the pollutants and sometimes can completely oxidize the compound. Microorganisms play the vital role in the process of bioremediation and biode‐ gradation because of their great metabolic diversity, which includes the ability to metabolise these pollutants [1]. The degradation of toxic compounds to less harmful forms with the use of biological systems is called as bioremediation [2]. Bioremediation is limited in the number of toxic material, it can handle but where applicable it is cost effective and ecofriendly [3]. Today, water resources have been the most exploited of the natural systems, most of our water bodies are seriously polluted due to rapid population growth, industrial proliferation, urbanizations, increasing living standards and wide spheres of human activities. Many rivers of the world receive heavy flux due to industrial effluents [4]. The wastewater consisting of substances varying from simple nutrients to highly toxic hazardous chemicals, which when used for irrigation caused both beneficial and damaging effects to various crops including vegetables [5].

The awful consequence of increasing population along with an rapidly increase in the industrial field is a major concern of pollution in the environment. Water is necessary for irrigation to increase the productivity in agriculture. Industrial effluents already pollute most of the Indian rivers. Many industries are playing the crucial role in water pollution such as textile industries, dairy industries and distillery *etc*. Distilleries are the major agro-based industries, which utilize molasses as raw material for the production of rectified spirit. In addition to rectified spirit, distilleries also produce ethanol, which can be mixed with diesel and used as biofuel and helps in reducing import of crude oil thereby saving foreign exchange

© 2013 Santal and Singh; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

[6]. In the year 1999, there were 285 distilleries in India producing 2.70 × 109 L of wastewater each year [7]. This number has gone up to 319 producing 3.25 × 109 L of alcohol and generating 4.04 × 1011 L of wastewater annually [8] and this effluent or spent wash is a major source of aquatic and soil pollution [9]. The spentwash is highly acidic in nature and has a variety of recalcitrant coloring compounds as melanoidins, phenolics and metal sulfides that are mainly responsible for the dark color of distillery effluent. The pH of spentwash increases from 4.5 to 8.5 during the anaerobic treatment process and finally it is called as post methanated distillery effluent (PMDE). The spentwash is a waste having very high biochemical oxygen demand (BOD; 35,000-40,000 mgL-1), chemical oxygen demand (COD; 90,000-1,10,000 mgL-1), total solids (TS; 82,480 mgL-1), nitrogen (2,200 mgL-1), phenolics (4.20 mgL-1) and sulphate (3,410 mgL-1). In addition to these contaminants, several heavy metals (Cd, Mn, Fe, Zn, Ni and Pb) are also present. The spentwash primarily undergoes anaerobic treatment process which converts a significant proportion (>50%) of the BOD and COD. However, different biochemical changes in the spentwash occur during anaerobic digestion. There is formation of significant amount of hydrogen sulfide (H2S) as a result of the reduction of oxidized sulfur compound. Sulfide binds with the heavy metals present in the effluent and forms a colloidal solution of metal sulfide colorant [10]. This massive quantity, approx. 40 billion litres of effluent, if disposed untreated can cause considerable stress on the watercourses leading to widespread damage to aquatic life. Moreover, the oxygen is also exhausted giving death of fishes and other aquatic life. This wastewater is extremely harmful to the plants. Cane molasses contained around 2% of dark brown pigment melanoidins [11]. Dark brown color hinders photosynthesis by blocking sunlight and is therefore deleterious to aquatic life [12-14]. It reduces soil alkalinity and inhibits seed germination. Further due to the possibility of complexation reaction of introduced melanoidins with metal ions, they could influence the biogeochemical cycle of many constituents in natural water [14], which are highly resistant to microbial attack. Hence, the wastewater requires pretreatment before its safe disposal into the environment [15, 16].

is eutrophication in natural water bodies, reduction of sunlight penetration leading to decreased photosynthetic activity and dissolved oxygen concentration in lakes, rivers or lagoons [19, 20]. Therefore, the degradation and decolorization of molasses melanoidin by chemical [21] and biological means [20, 22] has been attempted in order to characterize the

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

73

Melanoidins are generated through the maillard reaction. Maillard reaction is known to proceed non-enzymatically between amino and carbonyl groups of organic matter [13, 14, 17, 18, 23] and are closely related to humic substances in the natural environment [24]. There are two important pathways in the non-enzymatic browning of food i.e. caramelization and maillard reaction. That leads to the formation of volatile aromatic compounds, intermediate nonvolatile compounds, and brown pigments called melanoidins. Since they are highly resistant to microbial attack, the conventional biological processes are inapplicable to color

Melanoidins have physiologically positive effects such as anti-oxidative activity including strong scavenging activity against reactive oxygen species [13, 25]. Miyata *et al*. [26] in their studies attempt to utilize the strong oxidizing activity of a white rot fungus *Coriolus hirsutus* to decolorise melanoidins, which is a main color component of heat treatment liquor of sewage sludge. This reaction occurs in many situations, in foods [27], *in vivo*, and in soil. Melanoidins, which are the final by-products of the reaction, are brown nitrogen containing polymers that are difficult to decompose. The formation of melanoidins is affected by the reactants and their concentrations, types of catalysts and buffers, reaction temperature, time, pH value, water activity, presence of oxygen and metal ions. During heat treatment, the maillard reaction accompanied by formation of a class of compounds known as maillard products. The reaction proceeds effectively >50 ºC and is favoured at pH 4 to 7 [28]. Recently, the empirical formula of melanoidins has been suggested as C17-18H26-27O10N [13]. The basic structure of melanoidin is given in Figure 1 [29]. The molecular weight distribution is between 5-40 kDa. It consists of acidic, polymeric and highly dispersed colloids, which are negatively charged due to the dissociation of carboxylic acids and phenoilc groups [30]. Due to their antioxidant properties, melanoidins are toxic to many microorganisms involved in wastewater treatment [31]. Melanoidins and their degradation are generally recalcitrant to biodegradation due to their complex structure and xenobiotic nature and in some cases are both mutagenic and carcino‐ genic [14]. Melanoidins behave as anionic hydrophilic polymers, which can form stable complexes with metal cations and reported that ketone or hydroxyl groups of pyranone or pyridone residues act as donor groups in melanoidins and participate in the chelation with metals as melanoidins have net negative charge and therefore, different heavy metals (Cu2+, Cr3+, Fe3+, Zn2+, Pb2+, *etc*.) form large complex molecules with melanoidins, amino acids, proteins

Miura *et al*. [33] have reported the formation of colored compounds formed in the maillard reaction between D-xylose and glycine. These pigments turned brown with decomposition, so they had been postulated to be important intermediates in the generation of melanoidins. Hayase *et al.* [34] have reported the identification of the novel blue pigment designated as Blue-M1. These consist of four molecules of D-xylose and glycine, and also have methine proton

chemical structure and bioremediation aspects.

removal from melanoidin containing wastewater from distillery [13].

and sugars in acidic medium and get precipitated [14, 32].

This book chapter contributes and summarizes the literature available on the structure, nature and properties of melanoidins. Besides these, the important part of this chapter involves role of bacteria, fungi, yeast and algae helps in biodegradation of melanoidins and also the enzymes involved in the degradation of melanoidins. Therefore, this chapter has a major significance in the prevention of environmental pollution.

#### **2. Nature of melanoidins**

Melanoidins are natural, dark brown, complex biopolymers produced by non-enzymatic browning reactions called as Maillard amino-carbonyl reaction taking place between the amino and carbonyl groups in organic substances [17, 18]. It is considered that melanoidins extensively exist in food, drinks and wastewater released from distillery and fermentation industries. Melanoidins exist not only in various foods but also in various industrial wastes e.g. distillery and sugar mill wastes [16]. Melanoidins owing to their structural complexity, dark color and offensive odour pose threat to aquatic and terrestrial ecosystem. Currently, the most visible environmental problem caused by contamination with melanoidin compounds is eutrophication in natural water bodies, reduction of sunlight penetration leading to decreased photosynthetic activity and dissolved oxygen concentration in lakes, rivers or lagoons [19, 20]. Therefore, the degradation and decolorization of molasses melanoidin by chemical [21] and biological means [20, 22] has been attempted in order to characterize the chemical structure and bioremediation aspects.

[6]. In the year 1999, there were 285 distilleries in India producing 2.70 × 109 L of wastewater

4.04 × 1011 L of wastewater annually [8] and this effluent or spent wash is a major source of aquatic and soil pollution [9]. The spentwash is highly acidic in nature and has a variety of recalcitrant coloring compounds as melanoidins, phenolics and metal sulfides that are mainly responsible for the dark color of distillery effluent. The pH of spentwash increases from 4.5 to 8.5 during the anaerobic treatment process and finally it is called as post methanated distillery effluent (PMDE). The spentwash is a waste having very high biochemical oxygen demand (BOD; 35,000-40,000 mgL-1), chemical oxygen demand (COD; 90,000-1,10,000 mgL-1), total solids (TS; 82,480 mgL-1), nitrogen (2,200 mgL-1), phenolics (4.20 mgL-1) and sulphate (3,410 mgL-1). In addition to these contaminants, several heavy metals (Cd, Mn, Fe, Zn, Ni and Pb) are also present. The spentwash primarily undergoes anaerobic treatment process which converts a significant proportion (>50%) of the BOD and COD. However, different biochemical changes in the spentwash occur during anaerobic digestion. There is formation of significant amount of hydrogen sulfide (H2S) as a result of the reduction of oxidized sulfur compound. Sulfide binds with the heavy metals present in the effluent and forms a colloidal solution of metal sulfide colorant [10]. This massive quantity, approx. 40 billion litres of effluent, if disposed untreated can cause considerable stress on the watercourses leading to widespread damage to aquatic life. Moreover, the oxygen is also exhausted giving death of fishes and other aquatic life. This wastewater is extremely harmful to the plants. Cane molasses contained around 2% of dark brown pigment melanoidins [11]. Dark brown color hinders photosynthesis by blocking sunlight and is therefore deleterious to aquatic life [12-14]. It reduces soil alkalinity and inhibits seed germination. Further due to the possibility of complexation reaction of introduced melanoidins with metal ions, they could influence the biogeochemical cycle of many constituents in natural water [14], which are highly resistant to microbial attack. Hence, the wastewater requires pretreatment before its safe disposal into the environment [15, 16]. This book chapter contributes and summarizes the literature available on the structure, nature and properties of melanoidins. Besides these, the important part of this chapter involves role of bacteria, fungi, yeast and algae helps in biodegradation of melanoidins and also the enzymes involved in the degradation of melanoidins. Therefore, this chapter has a major significance

Melanoidins are natural, dark brown, complex biopolymers produced by non-enzymatic browning reactions called as Maillard amino-carbonyl reaction taking place between the amino and carbonyl groups in organic substances [17, 18]. It is considered that melanoidins extensively exist in food, drinks and wastewater released from distillery and fermentation industries. Melanoidins exist not only in various foods but also in various industrial wastes e.g. distillery and sugar mill wastes [16]. Melanoidins owing to their structural complexity, dark color and offensive odour pose threat to aquatic and terrestrial ecosystem. Currently, the most visible environmental problem caused by contamination with melanoidin compounds

L of alcohol and generating

each year [7]. This number has gone up to 319 producing 3.25 × 109

72 Biodegradation of Hazardous and Special Products

in the prevention of environmental pollution.

**2. Nature of melanoidins**

Melanoidins are generated through the maillard reaction. Maillard reaction is known to proceed non-enzymatically between amino and carbonyl groups of organic matter [13, 14, 17, 18, 23] and are closely related to humic substances in the natural environment [24]. There are two important pathways in the non-enzymatic browning of food i.e. caramelization and maillard reaction. That leads to the formation of volatile aromatic compounds, intermediate nonvolatile compounds, and brown pigments called melanoidins. Since they are highly resistant to microbial attack, the conventional biological processes are inapplicable to color removal from melanoidin containing wastewater from distillery [13].

Melanoidins have physiologically positive effects such as anti-oxidative activity including strong scavenging activity against reactive oxygen species [13, 25]. Miyata *et al*. [26] in their studies attempt to utilize the strong oxidizing activity of a white rot fungus *Coriolus hirsutus* to decolorise melanoidins, which is a main color component of heat treatment liquor of sewage sludge. This reaction occurs in many situations, in foods [27], *in vivo*, and in soil. Melanoidins, which are the final by-products of the reaction, are brown nitrogen containing polymers that are difficult to decompose. The formation of melanoidins is affected by the reactants and their concentrations, types of catalysts and buffers, reaction temperature, time, pH value, water activity, presence of oxygen and metal ions. During heat treatment, the maillard reaction accompanied by formation of a class of compounds known as maillard products. The reaction proceeds effectively >50 ºC and is favoured at pH 4 to 7 [28]. Recently, the empirical formula of melanoidins has been suggested as C17-18H26-27O10N [13]. The basic structure of melanoidin is given in Figure 1 [29]. The molecular weight distribution is between 5-40 kDa. It consists of acidic, polymeric and highly dispersed colloids, which are negatively charged due to the dissociation of carboxylic acids and phenoilc groups [30]. Due to their antioxidant properties, melanoidins are toxic to many microorganisms involved in wastewater treatment [31]. Melanoidins and their degradation are generally recalcitrant to biodegradation due to their complex structure and xenobiotic nature and in some cases are both mutagenic and carcino‐ genic [14]. Melanoidins behave as anionic hydrophilic polymers, which can form stable complexes with metal cations and reported that ketone or hydroxyl groups of pyranone or pyridone residues act as donor groups in melanoidins and participate in the chelation with metals as melanoidins have net negative charge and therefore, different heavy metals (Cu2+, Cr3+, Fe3+, Zn2+, Pb2+, *etc*.) form large complex molecules with melanoidins, amino acids, proteins and sugars in acidic medium and get precipitated [14, 32].

Miura *et al*. [33] have reported the formation of colored compounds formed in the maillard reaction between D-xylose and glycine. These pigments turned brown with decomposition, so they had been postulated to be important intermediates in the generation of melanoidins. Hayase *et al.* [34] have reported the identification of the novel blue pigment designated as Blue-M1. These consist of four molecules of D-xylose and glycine, and also have methine proton

**Figure 1.** Basic structure of melanoidins [29].

between two pyrrolpyrrole rings. In the initial stage of maillard reaction, disaccharides form a Schiff base with amino groups of amino acids and are subsequently transformed *via* the Amadori rearrangement product [35, 36].

mineral colloids, microbial cells, virus particles) and sometimes by coagulant aids (e.g. activated silica, bentonite, polyelectrolytes, starch) [41]. Coagulation of the wastewater containing recalcitrant pollutants is practiced through use of conventional salts of aluminium and iron as coagulants [42]. Migo *et al.* [43] used a commercial inorganic flocculent, a polymer of ferric hydroxysulfate with a chemical formula [Fe2 (OH)n (SO4)3-n/2]m for the treatment of

Biological treatment

Anaerobic treatment/ Aerobic treatment

Bacterial treatment [69-88] Fungal treatment [89-114]

Treatmentby other microorganisms

http://dx.doi.org/10.5772/56252

75

Micro alga [115] Cyanobacteria [116] Yeast [31]

 Reactors [53-67]

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

Melanoidin degradation

Peroxidases Oxidoreductases Cellulolytic enzymes cyanidase Proteases Amylases etc. [122-127]

Physico-chemical treatment Enzymatic treatment

**Figure 2.** Various methods of melanoidin degradation.

Adsorption [38] Coagulation [42] Flocculation [43] Oxidation process [44] Membrane treatment [46] Evaporation[47-49] Combustion [50]

A study comparing the efficiency in decolorising biologically pretreated molasses wastewater of different oxidation processes using ozone, single hydrogen peroxide, Fenton's reagent and ozone combined with hydrogen peroxide has been performed. Ozone treatment was able to reduce about 76% of color. A combination of ozone with a low concentration of hydrogen peroxide was able to increase the color removal efficiency up to 89%. Gel permeation chro‐ matography corroborated the reduction in the concentration of chromophore groups respon‐ sible for wastewater color. Single hydrogen peroxide and Fenton's reagent were not able to reduce the color. Bicarbonate ions were found to be strong inhibitors of decolorising reactions [44]. Kumaresan *et al*. [45] employed emulsion liquid membrane (ELM) technique in a batch process for spentwash treatment. Water-oil-water type of emulsion was used to separate and concentrate the solutes resulting in 86% and 97% decrease in COD and BOD, respectively. Electrodialysis has been explored for desalting spentwash using cation and anion exchange membranes resulting in 50-60% reduction in potassium content [46]. During treatment of winery and distillery wastewater by natural evaporation in ponds, formation of malodorous compounds induces harmful olfactory effects. The organic compounds are oxidised to CO2 and the nitrate is reduced to N2 (odourless compounds), without VFA formation. The preven‐

molasses wastewater.

#### **3. Degradation of melanoidin polymer**

Waste treatment methods aim the removal of unwanted compounds in wastewater for safe discharge into environment. Various technologies have been used for reducing the pollution load from distillery wastewater, which included physico-chemical treatment (Adsorption, Coagulation and flocculation, Oxidation process, Membrane treatment, Evaporation/combus‐ tion) and biological treatment (Figure 2) [37].

Adsorption techniques for wastewater treatment have become more popular in recent years. Among the physico-chemical processes, adsorption technology is considered to be one the most effective and proven technology having potential application in both water and waste‐ water treatment. Adsorption is a rapid phenomenon of passive sequestration and separation of adsorbate from aqueous/gaseous phase on to solid phase [38]. Decolorization of synthetic melanoidin using commercially available activated carbon as well as activated carbon produced from sugarcane bagasse was investigated by Bernardo *et al.* [39]. Activated carbon is a well known adsorbent due to its extended surface area, microporous structure, high adsorption capacity and high degree of surface reactivity. Previous studies on decolorization of molasses wastewater include adsorption on commercial as well as indigenously prepared activated carbons [40]. Coagulation involves the destabilization of colloidal particles (e.g. Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes http://dx.doi.org/10.5772/56252 75

**Figure 2.** Various methods of melanoidin degradation.

between two pyrrolpyrrole rings. In the initial stage of maillard reaction, disaccharides form a Schiff base with amino groups of amino acids and are subsequently transformed *via* the

R

R: H; glc: (glc)n

glc

R

R glc

Waste treatment methods aim the removal of unwanted compounds in wastewater for safe discharge into environment. Various technologies have been used for reducing the pollution load from distillery wastewater, which included physico-chemical treatment (Adsorption, Coagulation and flocculation, Oxidation process, Membrane treatment, Evaporation/combus‐

Adsorption techniques for wastewater treatment have become more popular in recent years. Among the physico-chemical processes, adsorption technology is considered to be one the most effective and proven technology having potential application in both water and waste‐ water treatment. Adsorption is a rapid phenomenon of passive sequestration and separation of adsorbate from aqueous/gaseous phase on to solid phase [38]. Decolorization of synthetic melanoidin using commercially available activated carbon as well as activated carbon produced from sugarcane bagasse was investigated by Bernardo *et al.* [39]. Activated carbon is a well known adsorbent due to its extended surface area, microporous structure, high adsorption capacity and high degree of surface reactivity. Previous studies on decolorization of molasses wastewater include adsorption on commercial as well as indigenously prepared activated carbons [40]. Coagulation involves the destabilization of colloidal particles (e.g.

Amadori rearrangement product [35, 36].

**Figure 1.** Basic structure of melanoidins [29].

glc

74 Biodegradation of Hazardous and Special Products

R

**3. Degradation of melanoidin polymer**

tion) and biological treatment (Figure 2) [37].

mineral colloids, microbial cells, virus particles) and sometimes by coagulant aids (e.g. activated silica, bentonite, polyelectrolytes, starch) [41]. Coagulation of the wastewater containing recalcitrant pollutants is practiced through use of conventional salts of aluminium and iron as coagulants [42]. Migo *et al.* [43] used a commercial inorganic flocculent, a polymer of ferric hydroxysulfate with a chemical formula [Fe2 (OH)n (SO4)3-n/2]m for the treatment of molasses wastewater.

A study comparing the efficiency in decolorising biologically pretreated molasses wastewater of different oxidation processes using ozone, single hydrogen peroxide, Fenton's reagent and ozone combined with hydrogen peroxide has been performed. Ozone treatment was able to reduce about 76% of color. A combination of ozone with a low concentration of hydrogen peroxide was able to increase the color removal efficiency up to 89%. Gel permeation chro‐ matography corroborated the reduction in the concentration of chromophore groups respon‐ sible for wastewater color. Single hydrogen peroxide and Fenton's reagent were not able to reduce the color. Bicarbonate ions were found to be strong inhibitors of decolorising reactions [44]. Kumaresan *et al*. [45] employed emulsion liquid membrane (ELM) technique in a batch process for spentwash treatment. Water-oil-water type of emulsion was used to separate and concentrate the solutes resulting in 86% and 97% decrease in COD and BOD, respectively. Electrodialysis has been explored for desalting spentwash using cation and anion exchange membranes resulting in 50-60% reduction in potassium content [46]. During treatment of winery and distillery wastewater by natural evaporation in ponds, formation of malodorous compounds induces harmful olfactory effects. The organic compounds are oxidised to CO2 and the nitrate is reduced to N2 (odourless compounds), without VFA formation. The preven‐ tive treatment of odours by nitrate addition was tested on an industrial scale in winery and distillery ponds [47]. Molasses spentwash containing 4% solids can be concentrated to a maximum of 40% solids in a quintuple-effect evaporation system with thermal vapor recom‐ pression [48, 49]. Combustion is also an effective method of on-site vinasse disposal as it is accompanied by production of potassium-rich ash [50] that can be used for land application.

widely applied for the treatment of industrial effluents. Effluents from alcohol producing industries are mostly highly polluted and therefore in principle very suitable for anaerobic treatment. Distilleries use different kinds of raw materials such as sugar cane juice, sugar cane molasses, sugar beet molasses, wine or corn for the production of alcohol [40]. The use of different materials and the different processes applied, resulted the production of wide variety of effluent. The process conditions under which good results of the anaerobic process are obtained depend heavily on the type of distillery effluent being treated. The choice of the right set of process parameters for every type of distillery effluent has shown to be of crucial importance for the anaerobic process. Bhoi *et al.* [53] worked on treatability studies of black liquor by upflow anaerobic sludge blanket reactor. This was deals with the feasibility studies of anaerobically treated black liquor along with the readily degradable synthetic wastewater (SWW) in a bench-scale upflow anaerobic sludge blanket reactor (UASBR). Treatment has been carried out in four phases of operation. In the first phase, 100% of SWW prepared from molasses was used. In the other phases of operation, 10%, 20%, and 30% of the SWW chemical oxygen demand (COD) was replaced by Black liquor COD successively. In each of the operation phases, the composition of the feed was not changed until COD reduction and gas production were stabilized. The entire operation took 509 days. The results indicate that the COD removal efficiency exhibited a decreasing trend with increase in the percentage of black liquor and apparently, UASB cannot be used as a treatment option for SWW containing more than 20% black liquor. Farhadian *et al.* [54] studied the treatment of strong beet sugar waste‐ water by an upflow anaerobic fixed bed (UAFB) at pilot plant scale. Reactors filled with standard industrial packing and inoculated with anaerobic culture (chicken manure, cow manure, anaerobic sludge digested from domestic wastewater) at 32-34 °C with 20 hrs hydraulic retention time (HRT) and influent COD ranging between 2000-8000 mgL-1 showed the efficiency of organic content reduction in the reactor ranged from 75% to 93%. The reactor filled with standard pall rings made of polypropylene with an effective surface area of 206

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

77

m-3 performed best in comparison to the reactor filled with cut polyethylene pipe 134

with PE while it was 10-16% in case of PVC when compared to standard pall rings. Distillery waste tertiary treatment in a laboratory stabilization pond was evaluated. Effluents from a combined laboratory-scale anaerobic filter-aerobic trickling filter process were used as the influent. The effect of the hydraulic retention time (HRT) in the range of 5 - 30 days was evaluated at influent total chemical oxygen demand (TCOD) in the range of 271 - 5286 mgL-1 (BOD5 in the range of 66-1212 mgL-1). TCOD and BOD5 removals of up to 54% and 74%, respectively, were obtained for the most concentrated influent used at 30 days of HRT. The effect of the HRT on TCOD and BOD5 removal efficiencies followed an exponential relation‐ ship. An empirical equation adequately described the effect of the organic surface-loading rate

The pond acted as an intermediate flow pattern between completely mixed flow and plugflow. It was found that a second-order model fit in well with the experimental results of organic matter removal rate and effluent substrate concentration. The second-order kinetic constant was found to be KS = 0.23 mg BOD5/mg TSS day. The proposed model accurately predicted the behaviour of the pond showing deviations between the experimental and theoretical values

m-3). There was 2-7% decrease in efficiency

m-3 and reactor filled with PVC packing (50 m2

m2

m2

on the process efficiency.

Although many techniques have been explored for the treatment of melanoidins but they all are cost effective and produce large amount of sludge. So, to overcome these problem biological methods plays a crucial role in treatment of the melanoidins.

#### **4. Biological methods for the treatment of distillery effluent**

Microorganisms (bacteria/fungi/actinomycetes) due to their inherent capacity to metabolize a variety of substrate have been utilized since long back for biodegradation of complex, toxic and recalcitrant compounds, which cause severe damage to environment. Thus, these organ‐ isms have been exploited for biodegradation and decolorization of melanoidin pigment present in industrial wastes especially from distillery and fermentation industry. Biological treatment of molasses wastewater undergoes two subdivisions which include anaerobic treatment and aerobic treatment.

Anaerobic treatment is an accepted practice, and various high rate reactor designs have been tried at pilot and full scale operation. Aerobic treatment of anaerobically treated effluent using different microbial populations has also been explored. Majority of biological treatment technologies remove color by either concentrating the color into sludge or by partial or complete breakdown of the color molecules. These methods are discussed in detail in the following section.

#### **4.1. Anaerobic treatment /reactors**

In the anaerobic treatment of wastewater, the energy is extracted from the waste components without the introduction of air or oxygen. The anaerobic treatment of wastewater has now emerged as an energy saving wastewater treatment technology. It involves biological process‐ esthattakeplaceintheabsenceofoxygenandinwhichorganicmaterialsaredegradedtoproduce biogas [51]. Molasses wastewater treatment using anaerobic process is a very promising reemerging technology, which presents interesting advantages as compared to classical aerobic treatment. It produces very little sludge, requires less energy and can be successfully operated at high organic loading rates; also, the biogas thus generated can be utilized for steam genera‐ tion in the boilers thereby meeting the energy demands of the unit [52].

In India, the traditional method for this type of wastewater treatment involved sequestration in anaerobic lagoons during the dry season, followed by release in to the river. Anaerobic lagoons afford the simplest form of treatment to distillery effluent. Drawbacks of such form of treatment are the requirement of large land area, odour nuisance, and chances of ground water pollution. Since the development of the UASB process in the 1970s, this process has been tive treatment of odours by nitrate addition was tested on an industrial scale in winery and distillery ponds [47]. Molasses spentwash containing 4% solids can be concentrated to a maximum of 40% solids in a quintuple-effect evaporation system with thermal vapor recom‐ pression [48, 49]. Combustion is also an effective method of on-site vinasse disposal as it is accompanied by production of potassium-rich ash [50] that can be used for land application.

Although many techniques have been explored for the treatment of melanoidins but they all are cost effective and produce large amount of sludge. So, to overcome these problem biological

Microorganisms (bacteria/fungi/actinomycetes) due to their inherent capacity to metabolize a variety of substrate have been utilized since long back for biodegradation of complex, toxic and recalcitrant compounds, which cause severe damage to environment. Thus, these organ‐ isms have been exploited for biodegradation and decolorization of melanoidin pigment present in industrial wastes especially from distillery and fermentation industry. Biological treatment of molasses wastewater undergoes two subdivisions which include anaerobic

Anaerobic treatment is an accepted practice, and various high rate reactor designs have been tried at pilot and full scale operation. Aerobic treatment of anaerobically treated effluent using different microbial populations has also been explored. Majority of biological treatment technologies remove color by either concentrating the color into sludge or by partial or complete breakdown of the color molecules. These methods are discussed in detail in the

In the anaerobic treatment of wastewater, the energy is extracted from the waste components without the introduction of air or oxygen. The anaerobic treatment of wastewater has now emerged as an energy saving wastewater treatment technology. It involves biological process‐ esthattakeplaceintheabsenceofoxygenandinwhichorganicmaterialsaredegradedtoproduce biogas [51]. Molasses wastewater treatment using anaerobic process is a very promising reemerging technology, which presents interesting advantages as compared to classical aerobic treatment. It produces very little sludge, requires less energy and can be successfully operated at high organic loading rates; also, the biogas thus generated can be utilized for steam genera‐

In India, the traditional method for this type of wastewater treatment involved sequestration in anaerobic lagoons during the dry season, followed by release in to the river. Anaerobic lagoons afford the simplest form of treatment to distillery effluent. Drawbacks of such form of treatment are the requirement of large land area, odour nuisance, and chances of ground water pollution. Since the development of the UASB process in the 1970s, this process has been

tion in the boilers thereby meeting the energy demands of the unit [52].

methods plays a crucial role in treatment of the melanoidins.

treatment and aerobic treatment.

76 Biodegradation of Hazardous and Special Products

**4.1. Anaerobic treatment /reactors**

following section.

**4. Biological methods for the treatment of distillery effluent**

widely applied for the treatment of industrial effluents. Effluents from alcohol producing industries are mostly highly polluted and therefore in principle very suitable for anaerobic treatment. Distilleries use different kinds of raw materials such as sugar cane juice, sugar cane molasses, sugar beet molasses, wine or corn for the production of alcohol [40]. The use of different materials and the different processes applied, resulted the production of wide variety of effluent. The process conditions under which good results of the anaerobic process are obtained depend heavily on the type of distillery effluent being treated. The choice of the right set of process parameters for every type of distillery effluent has shown to be of crucial importance for the anaerobic process. Bhoi *et al.* [53] worked on treatability studies of black liquor by upflow anaerobic sludge blanket reactor. This was deals with the feasibility studies of anaerobically treated black liquor along with the readily degradable synthetic wastewater (SWW) in a bench-scale upflow anaerobic sludge blanket reactor (UASBR). Treatment has been carried out in four phases of operation. In the first phase, 100% of SWW prepared from molasses was used. In the other phases of operation, 10%, 20%, and 30% of the SWW chemical oxygen demand (COD) was replaced by Black liquor COD successively. In each of the operation phases, the composition of the feed was not changed until COD reduction and gas production were stabilized. The entire operation took 509 days. The results indicate that the COD removal efficiency exhibited a decreasing trend with increase in the percentage of black liquor and apparently, UASB cannot be used as a treatment option for SWW containing more than 20% black liquor. Farhadian *et al.* [54] studied the treatment of strong beet sugar waste‐ water by an upflow anaerobic fixed bed (UAFB) at pilot plant scale. Reactors filled with standard industrial packing and inoculated with anaerobic culture (chicken manure, cow manure, anaerobic sludge digested from domestic wastewater) at 32-34 °C with 20 hrs hydraulic retention time (HRT) and influent COD ranging between 2000-8000 mgL-1 showed the efficiency of organic content reduction in the reactor ranged from 75% to 93%. The reactor filled with standard pall rings made of polypropylene with an effective surface area of 206 m2 m-3 performed best in comparison to the reactor filled with cut polyethylene pipe 134 m2 m-3 and reactor filled with PVC packing (50 m2 m-3). There was 2-7% decrease in efficiency with PE while it was 10-16% in case of PVC when compared to standard pall rings. Distillery waste tertiary treatment in a laboratory stabilization pond was evaluated. Effluents from a combined laboratory-scale anaerobic filter-aerobic trickling filter process were used as the influent. The effect of the hydraulic retention time (HRT) in the range of 5 - 30 days was evaluated at influent total chemical oxygen demand (TCOD) in the range of 271 - 5286 mgL-1 (BOD5 in the range of 66-1212 mgL-1). TCOD and BOD5 removals of up to 54% and 74%, respectively, were obtained for the most concentrated influent used at 30 days of HRT. The effect of the HRT on TCOD and BOD5 removal efficiencies followed an exponential relation‐ ship. An empirical equation adequately described the effect of the organic surface-loading rate on the process efficiency.

The pond acted as an intermediate flow pattern between completely mixed flow and plugflow. It was found that a second-order model fit in well with the experimental results of organic matter removal rate and effluent substrate concentration. The second-order kinetic constant was found to be KS = 0.23 mg BOD5/mg TSS day. The proposed model accurately predicted the behaviour of the pond showing deviations between the experimental and theoretical values of effluent substrate concentrations equal to or lower than 11% [55]. Sharma *et al.* [56] worked on the effect of nutrients supplementation on anaerobic sludge development and activity for treating distillery effluent. Startup of laboratory anaerobic reactors and treatment efficiency were investigated by supplementing the distillery effluent feed with macronutrients (Ca, P) and micronutrients (Ni, Fe and Co) under mesophilic conditions. Calcium and Phosphate were detrimental to the treatment efficiency and sludge granulation. Traces of salt of Iron, nickel and cobalt individually and in combinations improved the COD removal efficiency and sludge granulation process.

*brevibacter, Methanothrix* and *Methanosarcina* [63]; however, the predominance of *Methano‐ thrix* in granular sludge is most essential for the establishment of a high performance UASB process. In this study, abundance of *Methanosarcina* sp. was observed whereas *Methanothrix* sp. was present to a lesser extent thereby indicating that the latter was more sensitive to refractile compounds. Influence of reactor configuration on fermentative hydrogen (H2) production and substrate degradation was evaluated employing anaerobic mixed consortia. Reactors were operated at acidophilic (pH 6.0) condition employing designed synthetic

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

of 24 hrs at 28 ± 2 °C. Experimental data enumerated the influence of reactor configuration on both H2 production and wastewater treatment. Biofilm reactor (28.98 mmol H2/day; 1.25 Kg

mmol H2/g-CODL-day (19.32 mmol H2/g-CODR/day) and 5.03 mmol H2/g-CODL/day (16.10 mmol H2/g-CODR/day) were observed during stabilized phase of operation of biofilm and suspended growth reactors respectively. Higher concentration of VFA generation was observed in the biofilm reactor. Both the configurations recorded higher acetate concentration over other soluble metabolites indicating the dominance of acid-forming metabolic pathway during the H2 production process [64]. A lab-scale anaerobic hybrid (combining sludge blanket and filter) reactor was operated in a Continuous mode to study anaerobic biodegradation of distillery effluent. The study demonstrated that at optimum hydraulic retention time (HRT),

reactor was 79%. The anaerobic reduction of sulfate increases sulfide concentration, which inhibited the metabolism of methanogens and reduced the performance of the reactors. The kinetics of biomass growth *i.e.* yield coefficient (Y, 0.0532) and decay coefficient (Kd, 0.0041 d- 1) was obtained using Lawrence and McCarty model. However, this model failed in determining the kinetics of substrate utilization. Bhatia *et al*. [65] model having inbuilt provision of process inhibition described the kinetics of substrate utilization, *i.e.* maximum rate of substrate utilization and inhibition coefficient values. Modeling of the reactor demon‐ strated that Parkin *et al.* [66], and Bhatia *et al.* [65] models, both, could be used to predict the effluent substrate concentration. However, Parkin *et al.* [66] model predicts effluent COD more precisely (within ± 2 %) than Bhatia *et al.* [65] model (within ± 5 - 20 %) of the experimental value. Karhadkar *et al*. [67] model predicted biogas yield within ± 5 % of the experimental value. Besides these anaerobic treatments another treatment i.e. aerobic treatment is required

Anaerobically treated distillery wastewater still contains high concentrations of organic pollutants and then cannot be discharged directly. The partially treated effluent has high BOD, COD and suspended solids. It can reduce the availability of essential mineral nutrients by trapping them into immobilized organic forms, and may produce phytotoxic substances during decomposition. Stringent regulations on discharge of colored effluent impede direct discharge of anaerobically treated effluent [52]. Therefore, aerobic treatment of sugarcane molasses wastewater has been mainly attempted for the decolorization of the major colorant,

/day) showed relatively efficient performance over the corresponding suspended

/day with a retention time

http://dx.doi.org/10.5772/56252

79

/day). Specific H2 yields of 6.96

d, the COD removal efficiency of the

wastewater as substrate at an organic loading rate of 3.4 Kg COD/m3

growth configuration (20.93 mmol H2/day; 1.08 Kg COD/m3

5 days and organic loading rate (OLR), 8.7 kg COD / m3

for the safe disposal of effluent in the environment.

**4.2. Aerobic treatment**

COD/m3

Driessen *et al.* [57] studied on the anaerobic treatment of distillery effluent with the UASB process Special attention was given to the treatment of effluents from sugar cane based distilleries with very high COD concentration of 60000 to 160000 mg COD. Despite expected toxicity problems. On the other hand, two identical UASB reactors operated in parallel as duplicates for 327 days for the treatment of malt whisky pot ale and achieved COD reductions of up to 90% for influent concentrations of 3526-52126 mgL-1. When the OLRs of 15 kg m-3 day and above were used, the COD removal efficiency dropped to less than 20%. A mesophilic two stage system consisting of an anaerobic filter (AF) and an UASB reactor was found suitable for anaerobic digestion of distillery waste, enabling better conditions for the methanogenic phase [58]. Immobilization of bacteria in biofilm and on bioflocs is a crucial step in anaerobic degradation because of advantages such as higher activities, higher COD removal percent at short hydraulic retention times and better tolerance to disturbances such as toxic and organic shock loadings. At the same time there were certain disadvantages as well because in addition to some readily biodegradable matter, vinasses contain compounds like phenols, which were toxic to bacteria and inhibit the digestion. Also, due to seasonal nature of many of these industries and the absence of microorganisms in vinasses capable of carrying out anaerobic digestion, long incubation periods are required for the start-up stage. Besides, other opera‐ tional problems in anaerobic digestion such as low growth rate of anaerobic bacteria and the loss of biomass in systems with high hydraulic rates frequently does not achieve a satisfactory purification of vinasses [59]. The formation of H2S in anaerobic reactors is the result of the reduction of oxidized sulphur compounds. Methanogenic bacteria can tolerate sulphide concentration up to 1000 mgL-1 total sulphide. A complete loss of methane production occurred at 200 mgL-1 of un-ionized H2S during digestion of flocculent sludge. Anaerobic contact process incorporating an ultra filtration (UF) unit was used to treat distillery wastewater characterized by high and low carbon to nitrogen concentrations. This treatment system showed methane yield of up to 0.6 m3 kg-1 VS and removed up to 80% of the volatile acids [60]. Two-phase anaerobic digestion of alcohol stillage proved to be superior to the single-phase process in terms of substrate loading rate and methane yield, without affecting the treatment perform‐ ance [61]. While maintaining BOD and COD reduction of 85% and 65% respectively, the twophase achieved methane yield three times that of single-phase system.

Harada *et al*. [62] reported 39-67% COD removal, with a corresponding BOD removal of over 80%. The results suggested that the wastewater contained high concentration of refractile compounds; this, in turn, affected the microbial population in the sludge granules. Generally, the predominant genera of methanogens in granular sludge are *Methanobacterium, Methano‐* *brevibacter, Methanothrix* and *Methanosarcina* [63]; however, the predominance of *Methano‐ thrix* in granular sludge is most essential for the establishment of a high performance UASB process. In this study, abundance of *Methanosarcina* sp. was observed whereas *Methanothrix* sp. was present to a lesser extent thereby indicating that the latter was more sensitive to refractile compounds. Influence of reactor configuration on fermentative hydrogen (H2) production and substrate degradation was evaluated employing anaerobic mixed consortia. Reactors were operated at acidophilic (pH 6.0) condition employing designed synthetic wastewater as substrate at an organic loading rate of 3.4 Kg COD/m3 /day with a retention time of 24 hrs at 28 ± 2 °C. Experimental data enumerated the influence of reactor configuration on both H2 production and wastewater treatment. Biofilm reactor (28.98 mmol H2/day; 1.25 Kg COD/m3 /day) showed relatively efficient performance over the corresponding suspended growth configuration (20.93 mmol H2/day; 1.08 Kg COD/m3 /day). Specific H2 yields of 6.96 mmol H2/g-CODL-day (19.32 mmol H2/g-CODR/day) and 5.03 mmol H2/g-CODL/day (16.10 mmol H2/g-CODR/day) were observed during stabilized phase of operation of biofilm and suspended growth reactors respectively. Higher concentration of VFA generation was observed in the biofilm reactor. Both the configurations recorded higher acetate concentration over other soluble metabolites indicating the dominance of acid-forming metabolic pathway during the H2 production process [64]. A lab-scale anaerobic hybrid (combining sludge blanket and filter) reactor was operated in a Continuous mode to study anaerobic biodegradation of distillery effluent. The study demonstrated that at optimum hydraulic retention time (HRT), 5 days and organic loading rate (OLR), 8.7 kg COD / m3 d, the COD removal efficiency of the reactor was 79%. The anaerobic reduction of sulfate increases sulfide concentration, which inhibited the metabolism of methanogens and reduced the performance of the reactors. The kinetics of biomass growth *i.e.* yield coefficient (Y, 0.0532) and decay coefficient (Kd, 0.0041 d- 1) was obtained using Lawrence and McCarty model. However, this model failed in determining the kinetics of substrate utilization. Bhatia *et al*. [65] model having inbuilt provision of process inhibition described the kinetics of substrate utilization, *i.e.* maximum rate of substrate utilization and inhibition coefficient values. Modeling of the reactor demon‐ strated that Parkin *et al.* [66], and Bhatia *et al.* [65] models, both, could be used to predict the effluent substrate concentration. However, Parkin *et al.* [66] model predicts effluent COD more precisely (within ± 2 %) than Bhatia *et al.* [65] model (within ± 5 - 20 %) of the experimental value. Karhadkar *et al*. [67] model predicted biogas yield within ± 5 % of the experimental value. Besides these anaerobic treatments another treatment i.e. aerobic treatment is required for the safe disposal of effluent in the environment.

#### **4.2. Aerobic treatment**

of effluent substrate concentrations equal to or lower than 11% [55]. Sharma *et al.* [56] worked on the effect of nutrients supplementation on anaerobic sludge development and activity for treating distillery effluent. Startup of laboratory anaerobic reactors and treatment efficiency were investigated by supplementing the distillery effluent feed with macronutrients (Ca, P) and micronutrients (Ni, Fe and Co) under mesophilic conditions. Calcium and Phosphate were detrimental to the treatment efficiency and sludge granulation. Traces of salt of Iron, nickel and cobalt individually and in combinations improved the COD removal efficiency and sludge

Driessen *et al.* [57] studied on the anaerobic treatment of distillery effluent with the UASB process Special attention was given to the treatment of effluents from sugar cane based distilleries with very high COD concentration of 60000 to 160000 mg COD. Despite expected toxicity problems. On the other hand, two identical UASB reactors operated in parallel as duplicates for 327 days for the treatment of malt whisky pot ale and achieved COD reductions of up to 90% for influent concentrations of 3526-52126 mgL-1. When the OLRs of 15 kg m-3 day and above were used, the COD removal efficiency dropped to less than 20%. A mesophilic two stage system consisting of an anaerobic filter (AF) and an UASB reactor was found suitable for anaerobic digestion of distillery waste, enabling better conditions for the methanogenic phase [58]. Immobilization of bacteria in biofilm and on bioflocs is a crucial step in anaerobic degradation because of advantages such as higher activities, higher COD removal percent at short hydraulic retention times and better tolerance to disturbances such as toxic and organic shock loadings. At the same time there were certain disadvantages as well because in addition to some readily biodegradable matter, vinasses contain compounds like phenols, which were toxic to bacteria and inhibit the digestion. Also, due to seasonal nature of many of these industries and the absence of microorganisms in vinasses capable of carrying out anaerobic digestion, long incubation periods are required for the start-up stage. Besides, other opera‐ tional problems in anaerobic digestion such as low growth rate of anaerobic bacteria and the loss of biomass in systems with high hydraulic rates frequently does not achieve a satisfactory purification of vinasses [59]. The formation of H2S in anaerobic reactors is the result of the reduction of oxidized sulphur compounds. Methanogenic bacteria can tolerate sulphide concentration up to 1000 mgL-1 total sulphide. A complete loss of methane production occurred at 200 mgL-1 of un-ionized H2S during digestion of flocculent sludge. Anaerobic contact process incorporating an ultra filtration (UF) unit was used to treat distillery wastewater characterized by high and low carbon to nitrogen concentrations. This treatment system showed methane yield of up to 0.6 m3 kg-1 VS and removed up to 80% of the volatile acids [60]. Two-phase anaerobic digestion of alcohol stillage proved to be superior to the single-phase process in terms of substrate loading rate and methane yield, without affecting the treatment perform‐ ance [61]. While maintaining BOD and COD reduction of 85% and 65% respectively, the two-

phase achieved methane yield three times that of single-phase system.

Harada *et al*. [62] reported 39-67% COD removal, with a corresponding BOD removal of over 80%. The results suggested that the wastewater contained high concentration of refractile compounds; this, in turn, affected the microbial population in the sludge granules. Generally, the predominant genera of methanogens in granular sludge are *Methanobacterium, Methano‐*

granulation process.

78 Biodegradation of Hazardous and Special Products

Anaerobically treated distillery wastewater still contains high concentrations of organic pollutants and then cannot be discharged directly. The partially treated effluent has high BOD, COD and suspended solids. It can reduce the availability of essential mineral nutrients by trapping them into immobilized organic forms, and may produce phytotoxic substances during decomposition. Stringent regulations on discharge of colored effluent impede direct discharge of anaerobically treated effluent [52]. Therefore, aerobic treatment of sugarcane molasses wastewater has been mainly attempted for the decolorization of the major colorant, melanoidins, and for reduction of the COD and BOD. A large number of microorganisms such as bacteria (pure and mixed culture), cyanobacteria, yeast and fungi have been isolated in recent years and are capable of degrading melanoidins and thus decolorising the molasses wastewater. The aerobic methods have been described below.

#### *4.2.1. Bacterial treatment*

Numerous bacteria capable of melanoidins decolorization have been reported. Spent wash is pollution intensive wastewater generated by distilleries. Its dark brown color is due to recalcitrant melanoidin pigments. Bacterial cultures were screened for their ability to degrade these pigments.

Recently Lactic acid bacteria (Lactobacillus coryniformis, Lactobacillus sakei, Lactobacillus plantarum, Weisella soli, Pediococcus parvulus, Pediococcus pentosaceus) were used to decolorise the melanoidins. The isolate Lactobacillus plantarum exhibit 44% decourisation of melanoidins [68]. An effort has also been made by the Kryzwonos [69] using the consortia of genus Bacilli to decolorize the melanoidins. Two mixed bacterial cultures of the genus Bacillus (C1 and C2) were tested for color removal ability. Similarly Yadav and Chandra [70] developed the consortia Proteus mirabilis (IITRM5; FJ581028), Bacillus sp. (IITRM7; FJ581030), Raoultella planticola (IITRM15; GU329705) and Enterobacter sakazakii (IITRM16, FJ581031) in the ratio of 4:3:2:1. This consortia was responsible for the 75% decolorization of melanoidins within 10 days (Figure 3).

The isolate *Alcaligenes faecalis* strain SAG5 showed 72.6 % decolorization of melanoidins at optimum pH (7.5) and temperature (37 °C) on 5th day of cultivation. The toxicity evaluation with mung bean (*Vigna radiata*) revealed that the raw distillery effluent was environmentally highly toxic as compared to biologically treated distillery effluent, which indicated that the effluent after bacterial treatment is environmentally safe [71]. The degradation of synthetic and natural melanoidins was studied by using the axenic and mixed bacterial consortium (*Bacillus licheniformis* (RNBS1), *Bacillus* sp. (RNBS3) and *Alcaligenes* sp. (RNBS4). Results have revealed that the mixed consortium was more effective compared to axenic culture decolorising 73.7% and 69.8%% synthetic and natural melanoidins whereas axenic cultures RNBS1, RNBS3 and RNBS4 decolorized 65.88%, 62.5% and 66.1% synthetic and 52.6%, 48.9% and 59.6% natural melanoidins, respectively. The HPLC analysis of degraded samples has shown reduction in peak areas compared to controls, suggesting that decrease in color intensity might be largely attributed to the degradation of melanoidins by isolated bacteria [72]. Tondee and Sirianun‐ tapiboon [73] isolated *Lactobacillus plantarum* No. PV71-1861 from pickle samples in Thailand. The strain showed the highest melanoidin pigment (MP) decolorization yield of 68.12% with MP solution containing glucose 2%, yeast extract 0.4%, KH2PO4 0.1%, MgSO47H2O 0.05% and initial pH 6 under static condition at 30 °C within 7 days. But, it showed low growth and MP decolorization yields under aerobic conditions. Gel filtration chromatograms of the MP solutions showed that the small molecular weight fraction of MP solution was decolorized by the strain when the large molecular weight fraction still remained in the effluent. For appli‐ cation, the strain could apply to treat anaerobic treated-molasses wastewater (T-MWW) with high removal efficiency. The highest MP removal efficiencies and growth yield of 76.6% and

2.6 mg/ml, respectively, were observed with the T-MWW within 7 days of culture, and the

**Figure 3.** Melanoidin decolorization activity by different bacterial cultures. Bacterial consortium I: *Proteus mirabilis* (IITRM5; FJ581028), *Bacillus* sp. (IITRM7; FJ581030), *Raoultella planticola* (IITRM15; GU329705) and Enterobacter saka‐ *zakii* (IITRM16, FJ581031) in the ratio of 4:3:2:1[70]; Bacterial consortium II: *Bacillus licheniformis* (RNBS1), *Bacillus* sp. (RNBS3) and *Alcaligenes* sp. (RNBS4) [72]; Bacterial consortium III: *Pseudomonas aeruginosa* PAO1, *Stenotrophomonas maltophila* and *Proteus mirabilis* [15]; Bacterial consortium IV: *Bacillus thuringiensis* (MTCC 4714), *Bacillus brevis* (MTCC

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

81

The identification of culturable bacteria by 16S rDNA based approach showed that the consor‐ tium composed of *Klebsiella oxytoca*, *Serratia mercescens*, *Citrobacter* sp. and unknown bacteri‐ um decolorise the synthetic melanoidins. In the context of academic study, prevention on the difficultiesofprovidingeffluentaswellasitsvariationsincompositions,severalsyntheticmedia prepared with respectto color and CODcontents basedon analysis ofmolasses wastewater, *i.e*., Viandox sauce (13.5%, v/v), caramel (30%, w/v), beet molasses wastewater (41.5%, v/v) and sugarcane molasses wastewater (20%, v/v) were used for decolorization using consortium with color removal 9.5%, 1.13%, 8.02% and 17.5%, respectively, within 2 days. However, Viandox sauce was retained for further study. The effect of initial pH and Viandox concentration on decolorization and growth of bacterial consortium were further determined. The highest decolorization of 18.3% was achieved at pH 4 after 2 day of incubation [74]. Mohana *et al.* [15] isolated the microorganisms that were capable of decolorizing anaerobically treated distillery

effluent pH of the system was decreased to lower than 4.0 after 2-3 days operation.

4716) and *Bacillus* sp. (MTCC 6506) [16].

melanoidins, and for reduction of the COD and BOD. A large number of microorganisms such as bacteria (pure and mixed culture), cyanobacteria, yeast and fungi have been isolated in recent years and are capable of degrading melanoidins and thus decolorising the molasses

Numerous bacteria capable of melanoidins decolorization have been reported. Spent wash is pollution intensive wastewater generated by distilleries. Its dark brown color is due to recalcitrant melanoidin pigments. Bacterial cultures were screened for their ability to degrade

Recently Lactic acid bacteria (Lactobacillus coryniformis, Lactobacillus sakei, Lactobacillus plantarum, Weisella soli, Pediococcus parvulus, Pediococcus pentosaceus) were used to decolorise the melanoidins. The isolate Lactobacillus plantarum exhibit 44% decourisation of melanoidins [68]. An effort has also been made by the Kryzwonos [69] using the consortia of genus Bacilli to decolorize the melanoidins. Two mixed bacterial cultures of the genus Bacillus (C1 and C2) were tested for color removal ability. Similarly Yadav and Chandra [70] developed the consortia Proteus mirabilis (IITRM5; FJ581028), Bacillus sp. (IITRM7; FJ581030), Raoultella planticola (IITRM15; GU329705) and Enterobacter sakazakii (IITRM16, FJ581031) in the ratio of 4:3:2:1. This consortia was responsible for the 75% decolorization of melanoidins within 10

The isolate *Alcaligenes faecalis* strain SAG5 showed 72.6 % decolorization of melanoidins at optimum pH (7.5) and temperature (37 °C) on 5th day of cultivation. The toxicity evaluation with mung bean (*Vigna radiata*) revealed that the raw distillery effluent was environmentally highly toxic as compared to biologically treated distillery effluent, which indicated that the effluent after bacterial treatment is environmentally safe [71]. The degradation of synthetic and natural melanoidins was studied by using the axenic and mixed bacterial consortium (*Bacillus licheniformis* (RNBS1), *Bacillus* sp. (RNBS3) and *Alcaligenes* sp. (RNBS4). Results have revealed that the mixed consortium was more effective compared to axenic culture decolorising 73.7% and 69.8%% synthetic and natural melanoidins whereas axenic cultures RNBS1, RNBS3 and RNBS4 decolorized 65.88%, 62.5% and 66.1% synthetic and 52.6%, 48.9% and 59.6% natural melanoidins, respectively. The HPLC analysis of degraded samples has shown reduction in peak areas compared to controls, suggesting that decrease in color intensity might be largely attributed to the degradation of melanoidins by isolated bacteria [72]. Tondee and Sirianun‐ tapiboon [73] isolated *Lactobacillus plantarum* No. PV71-1861 from pickle samples in Thailand. The strain showed the highest melanoidin pigment (MP) decolorization yield of 68.12% with MP solution containing glucose 2%, yeast extract 0.4%, KH2PO4 0.1%, MgSO47H2O 0.05% and initial pH 6 under static condition at 30 °C within 7 days. But, it showed low growth and MP decolorization yields under aerobic conditions. Gel filtration chromatograms of the MP solutions showed that the small molecular weight fraction of MP solution was decolorized by the strain when the large molecular weight fraction still remained in the effluent. For appli‐ cation, the strain could apply to treat anaerobic treated-molasses wastewater (T-MWW) with high removal efficiency. The highest MP removal efficiencies and growth yield of 76.6% and

wastewater. The aerobic methods have been described below.

*4.2.1. Bacterial treatment*

80 Biodegradation of Hazardous and Special Products

these pigments.

days (Figure 3).

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes http://dx.doi.org/10.5772/56252 81

**Figure 3.** Melanoidin decolorization activity by different bacterial cultures. Bacterial consortium I: *Proteus mirabilis* (IITRM5; FJ581028), *Bacillus* sp. (IITRM7; FJ581030), *Raoultella planticola* (IITRM15; GU329705) and Enterobacter saka‐ *zakii* (IITRM16, FJ581031) in the ratio of 4:3:2:1[70]; Bacterial consortium II: *Bacillus licheniformis* (RNBS1), *Bacillus* sp. (RNBS3) and *Alcaligenes* sp. (RNBS4) [72]; Bacterial consortium III: *Pseudomonas aeruginosa* PAO1, *Stenotrophomonas maltophila* and *Proteus mirabilis* [15]; Bacterial consortium IV: *Bacillus thuringiensis* (MTCC 4714), *Bacillus brevis* (MTCC 4716) and *Bacillus* sp. (MTCC 6506) [16].

2.6 mg/ml, respectively, were observed with the T-MWW within 7 days of culture, and the effluent pH of the system was decreased to lower than 4.0 after 2-3 days operation.

The identification of culturable bacteria by 16S rDNA based approach showed that the consor‐ tium composed of *Klebsiella oxytoca*, *Serratia mercescens*, *Citrobacter* sp. and unknown bacteri‐ um decolorise the synthetic melanoidins. In the context of academic study, prevention on the difficultiesofprovidingeffluentaswellasitsvariationsincompositions,severalsyntheticmedia prepared with respectto color and CODcontents basedon analysis ofmolasses wastewater, *i.e*., Viandox sauce (13.5%, v/v), caramel (30%, w/v), beet molasses wastewater (41.5%, v/v) and sugarcane molasses wastewater (20%, v/v) were used for decolorization using consortium with color removal 9.5%, 1.13%, 8.02% and 17.5%, respectively, within 2 days. However, Viandox sauce was retained for further study. The effect of initial pH and Viandox concentration on decolorization and growth of bacterial consortium were further determined. The highest decolorization of 18.3% was achieved at pH 4 after 2 day of incubation [74]. Mohana *et al.* [15] isolated the microorganisms that were capable of decolorizing anaerobically treated distillery

spent wash. A bacterial consortium DMC comprising of three bacterial cultures was selected on the basis of rapid effluent decolorization and degradation, which exhibited 67 ± 2% decoloriza‐ tion within 24 hrs and 51 ± 2% chemical oxygen demand reduction within 72 hrs when incubat‐ ed at 37 o C under static condition in effluent supplemented with glucose 0.5%, KH2PO4 0.1%, KCl 0.05% and MgSO4.7H2O 0.05%. Addition of organic or inorganic nitrogen sources did not support decolorization. The cultures were identified as *Pseudomonas aeruginosa* PAO1, *Stenotro‐ phomonas maltophila* and *Proteus mirabilis* by the 16S rDNA analysis. Kumar and Chandra [16] studiedonthedecolorizationof syntheticmelanoidins (*i.e*.,GGA,GAA, SGAandSAA) by three *Bacillus* isolates *Bacillus thuringiensis* (MTCC 4714), *Bacillus brevis* (MTCC 4716) and *Bacillus sp.* (MTCC 6506). Significant reduction in the values of physicochemical parameters was noticed along with the decolorization of all four melanoidins (10%, v/v). *B. thuringiensis* (MTCC 4714) causedmaximumdecolorizationfollowedby*B.brevis*(MTCC4716)and*Bacillus*sp.(MTCC6506). A mixed culture comprised of these three strains was capable of decolorizing all four melanoi‐ dins. The medium that contained glucose as sole carbon source showed 15% more decoloriza‐ tion than that containing both carbon and nitrogen sources. Melanoidin SGA was maximally decolorized (50%) while melanoidins GAA was decolorized least in the presence of glucose as a sole energy source. Acetogenic bacteria are capable of oxidative decomposition of melanoi‐ dins. Adikane *et al.* [75] worked decolorization of molasses spent wash (MSW) in absence of any additional carbon or nitrogen source using soil as inoculum. A decolorization of 69% was obtained using 10% (w/v) soil and 12.5% (v/v) MSW after 7 days incubation. Optimized parameters including days 6 days, pH 6, MSW 12.5% and soil concentration 40%, were ob‐ tained for maximum decolorization. A decolorization of 81% was achieved using 10% soil and 12.5% MSW after 18 days incubation in absence of any media supplement. Nearly 12% reduc‐ tion in decolorization activity of the soil sample was observed over a period of 12 months when stored at 6 o C. It could be concluded that the decolorization of MSW might be achieved using soil as inoculum without addition of chemical amendments. On the other hand, *Bacillus megaterium* SW3 and *Bacillus subtilis* SW8 were used for decolorization and bioremediation of DMSW. Both bacterial isolates were grown well in 7.5% (v/v) diluted digested spent wash supplementedwithglucoseandureaasareadilyavailablecarbonandnitrogensource.Optimum condition for growth and DMSW decolorization were at pH 7.2, temperature 37°C, glucose 5% (w/v), and urea 2% (w/v) in a minimal salt medium. The maximum decolorization (51%, 52.5% and 57%) and COD reduction (53%, 54.5% and 59%) was found after 5 days of incubation under the optimized condition were achieved for cultures SW3, SW8 and consortia (SW3 + SW8) respectively [76].

Ghosh *et al.* [78] worked on enrichment and identification of bacteria capable of reducing COD of anaerobically treated molasses spent wash. The isolates were grouped into six haplotypes by amplified ribosomal DNA restriction analysis (ARDRA) and BOX-PCR. They showed maximum similarity to six genera *viz. Pseudomonas, Enterobacter, Stenotrophomona, Aeromonas, Acinetobacter* and *Klebsiella.* The extent of COD (44%) reduced collectively by the six strains was equal to that reduced individually by *Aeromonas, Acinetobacter, Pseudomonas* and *Enterobact‐ er.* With spent wash as sole carbon source, the COD reducing strains grew faster at 37 o

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

C. Sirianuntapiboon *et al.* [79] isolated 170 strains of acetogenic bacteria, a strain No.BP103

pigments medium containing glucose 3.0%, yeast extract 0.5%, KH2PO4 0.1% and MgSO4.7H2O 0.05% and the pH adjusted to 6.0. In addition this strain could decolorise 32.3% and 73.5% of molasses pigments in stillage and anaerobically treated molasses wastewater, both supple‐ menting glucose 3.0% yeast extract 0.5%, KH2PO4 0.1% and MgSO4.7H2O 0.05%. However without nutrients supplement, the decolorization yields were 9.75% and 44.3% respectively. In a replacement culture system involving six time replacement (30days), the strain No. BP103 Showed a constant decolorization yield of 72.0% to 84.0% and caused decreases of biological oxidation demand and chemical oxygen demand values of approximately 58.5% to 82.2% and 35.5% to 71.2%, respectively. Under a periodical feeding system, the decolorization yield was 30.0 to 45.0%. Cibis *et al*. [80] achieved biodegradation of potato slops (distillation residue) by

Savant *et al.* [81] isolated a novel acid-tolerant, hydrogenotrophic methanogenic isolate ATMT, using enrichment culture technique at pH 5 using slurry from an acidogenic digester running on alcohol distillery waste. The original pH of the slurry was 5.7 and the volatile fatty acid concentration was 9000 ppm. Cells of isolate ATMT were Gram-positive, non-motile and 0.3-0.5 μm in size. They did not form spores. The isolate could grow in the pH range 5.0-7.5, with

and biochemical characteristics of the isolate, together with the 16S rDNA sequence analysis, clearly revealed that the isolate could be accommodated within any of the existing species of the genus *Methanobrevibacter.* Therefore, it is proposed that a novel species of the genus *Methanobrevibacter* should be created for this isolate, *Methanobrevibacter acididurans* sp. nov., and the type strain is strain ATMT. Ghosh *et al.* [82] studied on the application of two bacterial strains, *Pseudomonas putida* U and *Aeromonas s*train Ema, in a two-stage column bioreactor to reduce COD and the color of anaerobic treated spent wash under aerobic conditions. *P.*

Dahiya *et al.* [83] worked on decolorization of molasses wastewater by cells of *Pseudomonas*

bilised cells could be reused for decolorization activity. However, in subsequent cycles, this was found to decrease from 76% to 50% and from 50% to 24%. Decolorization activity was regenerated from 30% to 45% by recultivating the immobilised cells in a fresh growth medium. Cellulose carrier coated with collagen was found to be most effective carrier, which produced the highest decolorization activity of 94% in a day process. This carrier could be reused with

*fluorescens*. Decolorization was 76% under nonsterile conditions in four days at 30 o

showed the highest decolorization yield when cultivated at 30 o

a mixed population of bacteria under thermophilic conditions up to 60 o

maximum growth at pH 6.0. The optimum temperature for growth was 35 o

77% was achieved under non-optimal conditions.

*putida* reduced the COD and color by 44.4% and 60%.

30 o

C than

83

C for 5 days in molasses

http://dx.doi.org/10.5772/56252

C. A COD removal of

C. Morphological

C. Immo‐

Chavan *et al.* [77] isolated a *Pseudomonas* sp. and studied for degradation of recalcitrant melanoidin pigment. The optimum conditions for decolorization were pH 6.8 -7.2, temperature 30-35 ºC and glucose 0.4% (w/v). The maximum decolorization up to 56% and 63% reduction in COD of the spent wash could be achieved after 72 hrs by the microbial treatment. Spectro‐ photometric and HPLC analysis of treated effluent confirmed biodegradation of melanoidin pigments by the isolate. This approach could be used to develop a cost-effective, eco-friendly biotechnology package for the bioremediation of spent wash before its disposal.

Ghosh *et al.* [78] worked on enrichment and identification of bacteria capable of reducing COD of anaerobically treated molasses spent wash. The isolates were grouped into six haplotypes by amplified ribosomal DNA restriction analysis (ARDRA) and BOX-PCR. They showed maximum similarity to six genera *viz. Pseudomonas, Enterobacter, Stenotrophomona, Aeromonas, Acinetobacter* and *Klebsiella.* The extent of COD (44%) reduced collectively by the six strains was equal to that reduced individually by *Aeromonas, Acinetobacter, Pseudomonas* and *Enterobact‐ er.* With spent wash as sole carbon source, the COD reducing strains grew faster at 37 o C than 30 o C. Sirianuntapiboon *et al.* [79] isolated 170 strains of acetogenic bacteria, a strain No.BP103 showed the highest decolorization yield when cultivated at 30 o C for 5 days in molasses pigments medium containing glucose 3.0%, yeast extract 0.5%, KH2PO4 0.1% and MgSO4.7H2O 0.05% and the pH adjusted to 6.0. In addition this strain could decolorise 32.3% and 73.5% of molasses pigments in stillage and anaerobically treated molasses wastewater, both supple‐ menting glucose 3.0% yeast extract 0.5%, KH2PO4 0.1% and MgSO4.7H2O 0.05%. However without nutrients supplement, the decolorization yields were 9.75% and 44.3% respectively. In a replacement culture system involving six time replacement (30days), the strain No. BP103 Showed a constant decolorization yield of 72.0% to 84.0% and caused decreases of biological oxidation demand and chemical oxygen demand values of approximately 58.5% to 82.2% and 35.5% to 71.2%, respectively. Under a periodical feeding system, the decolorization yield was 30.0 to 45.0%. Cibis *et al*. [80] achieved biodegradation of potato slops (distillation residue) by a mixed population of bacteria under thermophilic conditions up to 60 o C. A COD removal of 77% was achieved under non-optimal conditions.

spent wash. A bacterial consortium DMC comprising of three bacterial cultures was selected on the basis of rapid effluent decolorization and degradation, which exhibited 67 ± 2% decoloriza‐ tion within 24 hrs and 51 ± 2% chemical oxygen demand reduction within 72 hrs when incubat‐

KCl 0.05% and MgSO4.7H2O 0.05%. Addition of organic or inorganic nitrogen sources did not support decolorization. The cultures were identified as *Pseudomonas aeruginosa* PAO1, *Stenotro‐ phomonas maltophila* and *Proteus mirabilis* by the 16S rDNA analysis. Kumar and Chandra [16] studiedonthedecolorizationof syntheticmelanoidins (*i.e*.,GGA,GAA, SGAandSAA) by three *Bacillus* isolates *Bacillus thuringiensis* (MTCC 4714), *Bacillus brevis* (MTCC 4716) and *Bacillus sp.* (MTCC 6506). Significant reduction in the values of physicochemical parameters was noticed along with the decolorization of all four melanoidins (10%, v/v). *B. thuringiensis* (MTCC 4714) causedmaximumdecolorizationfollowedby*B.brevis*(MTCC4716)and*Bacillus*sp.(MTCC6506). A mixed culture comprised of these three strains was capable of decolorizing all four melanoi‐ dins. The medium that contained glucose as sole carbon source showed 15% more decoloriza‐ tion than that containing both carbon and nitrogen sources. Melanoidin SGA was maximally decolorized (50%) while melanoidins GAA was decolorized least in the presence of glucose as a sole energy source. Acetogenic bacteria are capable of oxidative decomposition of melanoi‐ dins. Adikane *et al.* [75] worked decolorization of molasses spent wash (MSW) in absence of any additional carbon or nitrogen source using soil as inoculum. A decolorization of 69% was obtained using 10% (w/v) soil and 12.5% (v/v) MSW after 7 days incubation. Optimized parameters including days 6 days, pH 6, MSW 12.5% and soil concentration 40%, were ob‐ tained for maximum decolorization. A decolorization of 81% was achieved using 10% soil and 12.5% MSW after 18 days incubation in absence of any media supplement. Nearly 12% reduc‐ tion in decolorization activity of the soil sample was observed over a period of 12 months when

C under static condition in effluent supplemented with glucose 0.5%, KH2PO4 0.1%,

C. It could be concluded that the decolorization of MSW might be achieved using

soil as inoculum without addition of chemical amendments. On the other hand, *Bacillus megaterium* SW3 and *Bacillus subtilis* SW8 were used for decolorization and bioremediation of DMSW. Both bacterial isolates were grown well in 7.5% (v/v) diluted digested spent wash supplementedwithglucoseandureaasareadilyavailablecarbonandnitrogensource.Optimum condition for growth and DMSW decolorization were at pH 7.2, temperature 37°C, glucose 5% (w/v), and urea 2% (w/v) in a minimal salt medium. The maximum decolorization (51%, 52.5% and 57%) and COD reduction (53%, 54.5% and 59%) was found after 5 days of incubation under the optimized condition were achieved for cultures SW3, SW8 and consortia (SW3 + SW8)

Chavan *et al.* [77] isolated a *Pseudomonas* sp. and studied for degradation of recalcitrant melanoidin pigment. The optimum conditions for decolorization were pH 6.8 -7.2, temperature 30-35 ºC and glucose 0.4% (w/v). The maximum decolorization up to 56% and 63% reduction in COD of the spent wash could be achieved after 72 hrs by the microbial treatment. Spectro‐ photometric and HPLC analysis of treated effluent confirmed biodegradation of melanoidin pigments by the isolate. This approach could be used to develop a cost-effective, eco-friendly

biotechnology package for the bioremediation of spent wash before its disposal.

ed at 37 o

82 Biodegradation of Hazardous and Special Products

stored at 6 o

respectively [76].

Savant *et al.* [81] isolated a novel acid-tolerant, hydrogenotrophic methanogenic isolate ATMT, using enrichment culture technique at pH 5 using slurry from an acidogenic digester running on alcohol distillery waste. The original pH of the slurry was 5.7 and the volatile fatty acid concentration was 9000 ppm. Cells of isolate ATMT were Gram-positive, non-motile and 0.3-0.5 μm in size. They did not form spores. The isolate could grow in the pH range 5.0-7.5, with maximum growth at pH 6.0. The optimum temperature for growth was 35 o C. Morphological and biochemical characteristics of the isolate, together with the 16S rDNA sequence analysis, clearly revealed that the isolate could be accommodated within any of the existing species of the genus *Methanobrevibacter.* Therefore, it is proposed that a novel species of the genus *Methanobrevibacter* should be created for this isolate, *Methanobrevibacter acididurans* sp. nov., and the type strain is strain ATMT. Ghosh *et al.* [82] studied on the application of two bacterial strains, *Pseudomonas putida* U and *Aeromonas s*train Ema, in a two-stage column bioreactor to reduce COD and the color of anaerobic treated spent wash under aerobic conditions. *P. putida* reduced the COD and color by 44.4% and 60%.

Dahiya *et al.* [83] worked on decolorization of molasses wastewater by cells of *Pseudomonas fluorescens*. Decolorization was 76% under nonsterile conditions in four days at 30 o C. Immo‐ bilised cells could be reused for decolorization activity. However, in subsequent cycles, this was found to decrease from 76% to 50% and from 50% to 24%. Decolorization activity was regenerated from 30% to 45% by recultivating the immobilised cells in a fresh growth medium. Cellulose carrier coated with collagen was found to be most effective carrier, which produced the highest decolorization activity of 94% in a day process. This carrier could be reused with 50% of the decolorization activity retained until the seventh day. Jain *et al.* [84] studied on degradation of anaerobically digested distillery wastewater by three bacterial strains, *viz. Xanthomonas fragariae, Bacillus megaterium* and *Bacillus cereus* in free and immobilized form, isolated from the activated sludge of a distillery wastewater treatment plant. The removal of COD and color with all the three strains increased with time up to 48 hrs and only marginal increase in COD and color removal efficiency was observed beyond this period up to 72 hrs. After this period, removal efficiency remained fairly constant up to 120 hrs. The maximum COD and color removal efficiency varied from 66 to 81 % and 65 to 75 % respectively for both free and immobilized cells of all the three strains. The strain *Bacillus cereus* showed the maximum efficiency of COD (81 %) and color (75 %) removal out of the three strains.

activity (MDA). Strains were isolated from stored wastewater of an alcohol fermentation involving molasses. One of them, strain W-NS, showed high and stable MDA, and was identical with *Lactobacillus hilgardii*. The decolorization yield of this strain under the optimal

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

85

Increasing attention has been directed towards utilizing microbial activity for decolorization of molasses wastewater. Several reports have indicated that some fungi in particular have such a potential [89]. One of the most studied fungus having ability to degrade and decolorise distillery effluent is *Aspergillus* such as *Aspergillus fumigatus* G-2-6, *Aspergillus niger*, *Aspergillus niveus*, *Aspergillus fumigatus* UB260 brought about an average of 69-75% decolorization along with 70-90% COD reduction [22, 90-92]. Previous studies showed that the white-rot fungus *Phanerochaete chrysosporium* could remove color and total phenols from the sugar refinery effluent [93]. Ohmomo *et al.* [94] reported the continuous decolorization of molasses waste‐ water in a bubbling column reactor with *Coriolus versicolor* immobilized within Ca-alginate gel. Later, Ohmomo *et al*. [95] used autoclaved mycelium of *Aspergilus oryzae* Y-2-32 that adsorbed lower weight fractions of melanoidins and degree of adsorption was influenced by the kind of sugars used for cultivation. The wine distilleries produce large volume of waste‐ water having phenolic compounds, which give a high inhibitory and antibacterial activity to this wastewater, thus slowing down the anaerobic digestion process. Partial elimination of these phenolics compounds was obtained by using *Geotrichum candidum* [96]. *Rhizoctonia* sp. D90 decolorized molasses melanoidins medium and a synthetic melanoidin medium by 87.5% and 84.5%, respectively, under experimental conditions [97] (Figure 4). Murata *et al.* [98] isolated a fungus *Streptomyces werraensis* TT 14 from soil that decolorized the model melanoi‐ din, the decolorization rate being 64% in the optimal medium of pH - 5.5 and 45% in a synthetic medium. There was virtually no difference in the UV-VIS absorption spectra of the microbially treated melanoidin and control. The peaks of the gel permeation chromatogram for the treated melanoidin and for the control showed the same retention times, but lower molecular weight compounds increased in the decolorized melanoidin. Kida *et al.* [99] *Aspergillus awamori* var. *kawachi* has been used for production of single cell protein from Japanese distillery wastewater

Fujita *et al.* [100] also considered the decolorization of melanoidin present in an effluent with *Coriolus hirsutus*. The aim of this study was to find a suitable industrial biological treatment processforefficientdecolorizationofsugarrefineryeffluent.White-rotfungihavebeenreported tobe capableofdecolorizingmelanoidincontainingwastewaters.Previous studies showedthat the white-rot fungus *Phanerochaete chrysosporium* can remove color and total phenols from the sugar refinery effluent [93]. Several studies have been reported for decolorizing the distillery effluent using fungi. Fumi *et al.* [101] worked on the optimisation of long term activated sludge treatment of winery wastewater. The results obtained in work carried out in order to verify the overallefficiencyoffull-scale,long-term,activated- sludgetreatmentofwinerywastewater.The analytical data showed the high removal of COD during the whole experimentation period and withvariousworkingparameters.Sludgeproductionwas lowerthanthatproducedbyordinary

conditions was improved to 40% by immobilization of cells within Ca- alginate gel.

*4.2.2. Fungal treatment*

after aerobic cultivation.

Masaharu *et al*. [85] studied on the screening of acid-forming facultative anaerobes for their decolorizing ability with the distilled wastewater (DWDA) produced during the alcoholic fermentation of cane molasses was carried out. On the primary screening, 113 strains, which were isolated from enrichment cultures of various plant and animal samples or soil samples showed decolorizing activity on modified GYP medium containing 10% (v/v) distilled wastewater. Strain SM-3 was selected as the best strain. This strain decolorized 31.4% of molasses pigment at a concentration of 10% (v/v) distilled wastewater supplemented with 1% glucose within 5 days at 30 o C under semi-anaerobic conditions. The main DWDA was found in the cell-free extract and cell membrane fractions. The strain was identified as *Lactobacillus casei*. Kambe *et al.* [86] screened out various microorganisms for their ability to decolorise molasses wastewater under thermophilic and anaerobic conditions. Strain MD-32, which was newly isolated from a soil sample, was selected as the best strain. From taxonomical studies, the strain was concluded to belong to the genus *Bacillus*, most closely resembling *B. smithii*. The strain decolorized 35.5% of molasses pigment within 20 days at 55 o C under anaerobic conditions but no decolorization activity was observed when it was cultivated aerobically.

At all the concentration tested molasses pigment was effectively decolorized by MD-32, with decolorization yields of around 15% within 2 days. The molecular weight distribution as determined by gel filtration chromatography revealed that the decolorization of molasses pigment by the isolated strain is accompanied by a decrease in not only small molecules but also large ones. Sirianuntapiboon [87] isolated acid forming bacteria from vegetables, fruits and fermented food samples. They were primary screened to select the strains which had decolorization activity. The results showed that 50 isolates had decolorization activity on solid medium (clear zone). Among these, the strains No. BP103 and No. 13A gave the highest decolorization activity in the liquid medium which contained molasses pigments. Fructose was the most suitable carbon source for both strains for decolorizing molasses pigments. The optimal concentration of fructose in the media for highest decolorizing activity was 2.0%. For the effects of nitrogen source on decolorization activity, the organic nitrogen compounds were the best nitrogen sources for both strains. The decolorization activity of the strain No. 13A and No. BP103 were 80.5% and 82.0%, respectively when the nitrogen source was yeast extract. At the optimal conditions, the decolorizing activity of the strains No. 13A and BP103 were 90.5% and 96.7%, respectively. Strain No. 13A and No. BP103 were identified as *Acetobacter acetii.* Ohmomo *et al.*[88] performed screening of facultative anaerobes with melanoidin decolorizing activity (MDA). Strains were isolated from stored wastewater of an alcohol fermentation involving molasses. One of them, strain W-NS, showed high and stable MDA, and was identical with *Lactobacillus hilgardii*. The decolorization yield of this strain under the optimal conditions was improved to 40% by immobilization of cells within Ca- alginate gel.

#### *4.2.2. Fungal treatment*

50% of the decolorization activity retained until the seventh day. Jain *et al.* [84] studied on degradation of anaerobically digested distillery wastewater by three bacterial strains, *viz. Xanthomonas fragariae, Bacillus megaterium* and *Bacillus cereus* in free and immobilized form, isolated from the activated sludge of a distillery wastewater treatment plant. The removal of COD and color with all the three strains increased with time up to 48 hrs and only marginal increase in COD and color removal efficiency was observed beyond this period up to 72 hrs. After this period, removal efficiency remained fairly constant up to 120 hrs. The maximum COD and color removal efficiency varied from 66 to 81 % and 65 to 75 % respectively for both free and immobilized cells of all the three strains. The strain *Bacillus cereus* showed the

maximum efficiency of COD (81 %) and color (75 %) removal out of the three strains.

glucose within 5 days at 30 o

84 Biodegradation of Hazardous and Special Products

Masaharu *et al*. [85] studied on the screening of acid-forming facultative anaerobes for their decolorizing ability with the distilled wastewater (DWDA) produced during the alcoholic fermentation of cane molasses was carried out. On the primary screening, 113 strains, which were isolated from enrichment cultures of various plant and animal samples or soil samples showed decolorizing activity on modified GYP medium containing 10% (v/v) distilled wastewater. Strain SM-3 was selected as the best strain. This strain decolorized 31.4% of molasses pigment at a concentration of 10% (v/v) distilled wastewater supplemented with 1%

in the cell-free extract and cell membrane fractions. The strain was identified as *Lactobacillus casei*. Kambe *et al.* [86] screened out various microorganisms for their ability to decolorise molasses wastewater under thermophilic and anaerobic conditions. Strain MD-32, which was newly isolated from a soil sample, was selected as the best strain. From taxonomical studies, the strain was concluded to belong to the genus *Bacillus*, most closely resembling *B. smithii*.

conditions but no decolorization activity was observed when it was cultivated aerobically.

At all the concentration tested molasses pigment was effectively decolorized by MD-32, with decolorization yields of around 15% within 2 days. The molecular weight distribution as determined by gel filtration chromatography revealed that the decolorization of molasses pigment by the isolated strain is accompanied by a decrease in not only small molecules but also large ones. Sirianuntapiboon [87] isolated acid forming bacteria from vegetables, fruits and fermented food samples. They were primary screened to select the strains which had decolorization activity. The results showed that 50 isolates had decolorization activity on solid medium (clear zone). Among these, the strains No. BP103 and No. 13A gave the highest decolorization activity in the liquid medium which contained molasses pigments. Fructose was the most suitable carbon source for both strains for decolorizing molasses pigments. The optimal concentration of fructose in the media for highest decolorizing activity was 2.0%. For the effects of nitrogen source on decolorization activity, the organic nitrogen compounds were the best nitrogen sources for both strains. The decolorization activity of the strain No. 13A and No. BP103 were 80.5% and 82.0%, respectively when the nitrogen source was yeast extract. At the optimal conditions, the decolorizing activity of the strains No. 13A and BP103 were 90.5% and 96.7%, respectively. Strain No. 13A and No. BP103 were identified as *Acetobacter acetii.* Ohmomo *et al.*[88] performed screening of facultative anaerobes with melanoidin decolorizing

The strain decolorized 35.5% of molasses pigment within 20 days at 55 o

C under semi-anaerobic conditions. The main DWDA was found

C under anaerobic

Increasing attention has been directed towards utilizing microbial activity for decolorization of molasses wastewater. Several reports have indicated that some fungi in particular have such a potential [89]. One of the most studied fungus having ability to degrade and decolorise distillery effluent is *Aspergillus* such as *Aspergillus fumigatus* G-2-6, *Aspergillus niger*, *Aspergillus niveus*, *Aspergillus fumigatus* UB260 brought about an average of 69-75% decolorization along with 70-90% COD reduction [22, 90-92]. Previous studies showed that the white-rot fungus *Phanerochaete chrysosporium* could remove color and total phenols from the sugar refinery effluent [93]. Ohmomo *et al.* [94] reported the continuous decolorization of molasses waste‐ water in a bubbling column reactor with *Coriolus versicolor* immobilized within Ca-alginate gel. Later, Ohmomo *et al*. [95] used autoclaved mycelium of *Aspergilus oryzae* Y-2-32 that adsorbed lower weight fractions of melanoidins and degree of adsorption was influenced by the kind of sugars used for cultivation. The wine distilleries produce large volume of waste‐ water having phenolic compounds, which give a high inhibitory and antibacterial activity to this wastewater, thus slowing down the anaerobic digestion process. Partial elimination of these phenolics compounds was obtained by using *Geotrichum candidum* [96]. *Rhizoctonia* sp. D90 decolorized molasses melanoidins medium and a synthetic melanoidin medium by 87.5% and 84.5%, respectively, under experimental conditions [97] (Figure 4). Murata *et al.* [98] isolated a fungus *Streptomyces werraensis* TT 14 from soil that decolorized the model melanoi‐ din, the decolorization rate being 64% in the optimal medium of pH - 5.5 and 45% in a synthetic medium. There was virtually no difference in the UV-VIS absorption spectra of the microbially treated melanoidin and control. The peaks of the gel permeation chromatogram for the treated melanoidin and for the control showed the same retention times, but lower molecular weight compounds increased in the decolorized melanoidin. Kida *et al.* [99] *Aspergillus awamori* var. *kawachi* has been used for production of single cell protein from Japanese distillery wastewater after aerobic cultivation.

Fujita *et al.* [100] also considered the decolorization of melanoidin present in an effluent with *Coriolus hirsutus*. The aim of this study was to find a suitable industrial biological treatment processforefficientdecolorizationofsugarrefineryeffluent.White-rotfungihavebeenreported tobe capableofdecolorizingmelanoidincontainingwastewaters.Previous studies showedthat the white-rot fungus *Phanerochaete chrysosporium* can remove color and total phenols from the sugar refinery effluent [93]. Several studies have been reported for decolorizing the distillery effluent using fungi. Fumi *et al.* [101] worked on the optimisation of long term activated sludge treatment of winery wastewater. The results obtained in work carried out in order to verify the overallefficiencyoffull-scale,long-term,activated- sludgetreatmentofwinerywastewater.The analytical data showed the high removal of COD during the whole experimentation period and withvariousworkingparameters.Sludgeproductionwas lowerthanthatproducedbyordinary

Another fungus, *Coriolus* No.20, belonging to the class basidiomycetes, can remove color substances from molasses wastewater (MWW). In the case of treatment of MWW solution, the decrease was 82.5% in darkness under optimal conditions. This decolorization was through the adsorption of melanoidin to the mycelium and the yield of adsorption was 9.67 and 12.2 mg g-1 of mycelium as dry basic in the case of autoclaved mycelium and resting mycelium, respectively. The resting mycelium showed the highest adsorption yield (16.32 mg g-1 of mycelium) when the melanoidins solution was supplemented with 20% glucose solution. In the fed-batch system, the strain showed a constant decolorization yield of 75% during four times replacements (32 days) in both untreated and treated MWW solutions. In the continuous feed system for decolorization of untreated MWW solution, l0% of fresh untreated MWW solution was added every day after seven days of cultivation. The decolorization rate was constant (37.5%) during 12 days of operation. In the continuous decolorization of concentrated untreated MWW solution, the strain showed a constant decolorization yield (about 70%) during three times feeding of fresh concentrated untreated MWW solution. But I0 ml of 20% glucose solution had to be added after 13 days cultivation to keep the reducing sugar concen‐

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

87

Miyata *et al.* [26] worked on Manganese-Independent and Dependent decolorization of melanoidinusingfungi.*Coriolushirsutus*pelletsweregrowninamelanoidin-containingmedium produced extracellular H2O2 up to 43 μm. However, nonenzymatic decolorization of melanoi‐ din by H2O2, which has been previously reported, seemed very little in the fungal culture. The culture fluid contained two extracellular peroxidases, a manganese-independent peroxidase (MIP) and manganese peroxidase (MnP). Since both MIP and MnP showed melanoidindecolorizing activities in the presence of H2O2, it was resulted that melanoidin decolorization in the*C.hirsutus* cultureinvolvedtheproductionofextracellularH2O2 andtheperoxidases.Miyata *et al.* [104] studied on the combined use of activated sludge and the fungus *Coriolus hirsutus*. A white rotfungus,*Coriolushirsutus*, exhibitedastrongabilitytodecolorisemelanoidinincultures not supplemented with nitrogenous nutrients. Addition of peptone to the cultures lowered the ability of the fungus to decolorise melanoidin, but that of inorganic nitrogens, ammonium and nitrate did not bring about any marked reduction in the ability. It suggested an inhibitory effect of organic nitrogen on melanoidin decolorization. Therefore, for enhancing the decolorization of melanoidin in wastewaters by the fungus, activated sludge pretreatment of the wastewa‐ ters was expected to be effective, *i.e.* activated sludge is capable of converting available organ‐ ic N into inorganic N. To confirm this, waste sludge heat treatment liquor (HTL), wastewater from a sewage treatment plant, was pretreated with activated sludge. In practice, pretreat‐ ment of HTL under appropriate conditions accelerated the fungal decolorization of HTL. In the pretreated HTL, the fungus was shown to produce a high level of manganese-independent peroxidase (MIP). Addition of Mn(I1) to the pretreated HTL caused a further increase in the decolorization efficiency of the fungus and a marked increase in the manganese peroxidase (MnP) activity. Consequently, the increases in MIP and MnP activities were considered to play

an important role in the enhanced ability of *C. hirsutus* to decolorise HTL.

Dahiya *et al.* [105] worked on decolorization of synthetic and spentwash melanoidins using the white-rot fungus *Phanerochaete chrysosporium* JAG-40. This fungus was isolated from the

tration in the culture broth at more than 1.0% [103].

**Figure 4.** Melanoidin decolorization activity by different fungi.

activated-sludgeplants.Miranda*etal.*[90]workedoncoloreliminationfrommolasseswastewa‐ ter by *Aspergillus niger*. Color elimination by *Aspergillus niger*fromwastewater was studied.The influences of the nutrient concentrations, initial pH and carbon source on this color elimina‐ tion were analyzed. During the batch process, through all experiments the maximal color eliminationwas attainedafter 3or 4days inthe culture.Batchprocesses showedamaximal color elimination of 69% when MgSO4, KH2PO4, NH4NO3 and a carbon source were added to the wastewater. The continuous process, with the same nutrient concentrations, showed less color removalandthedecolorizationactivitywasmaintainedforonly4days.Benito*etal*.[102] studied on decolorization of wastewater from an alcoholic fermentation process with *Trametes versicol‐ or*. Tests varying the concentrations of carbon source, nutrients, initial pH and mycelia were carried out in order to analyze their influence on the percentage decolorization, COD removal and decrease of ammonium content of the wastewater. In this way, 82% color elimination, 77% COD removal and 36% N-NH: decrease was attained.

Another fungus, *Coriolus* No.20, belonging to the class basidiomycetes, can remove color substances from molasses wastewater (MWW). In the case of treatment of MWW solution, the decrease was 82.5% in darkness under optimal conditions. This decolorization was through the adsorption of melanoidin to the mycelium and the yield of adsorption was 9.67 and 12.2 mg g-1 of mycelium as dry basic in the case of autoclaved mycelium and resting mycelium, respectively. The resting mycelium showed the highest adsorption yield (16.32 mg g-1 of mycelium) when the melanoidins solution was supplemented with 20% glucose solution. In the fed-batch system, the strain showed a constant decolorization yield of 75% during four times replacements (32 days) in both untreated and treated MWW solutions. In the continuous feed system for decolorization of untreated MWW solution, l0% of fresh untreated MWW solution was added every day after seven days of cultivation. The decolorization rate was constant (37.5%) during 12 days of operation. In the continuous decolorization of concentrated untreated MWW solution, the strain showed a constant decolorization yield (about 70%) during three times feeding of fresh concentrated untreated MWW solution. But I0 ml of 20% glucose solution had to be added after 13 days cultivation to keep the reducing sugar concen‐ tration in the culture broth at more than 1.0% [103].

Miyata *et al.* [26] worked on Manganese-Independent and Dependent decolorization of melanoidinusingfungi.*Coriolushirsutus*pelletsweregrowninamelanoidin-containingmedium produced extracellular H2O2 up to 43 μm. However, nonenzymatic decolorization of melanoi‐ din by H2O2, which has been previously reported, seemed very little in the fungal culture. The culture fluid contained two extracellular peroxidases, a manganese-independent peroxidase (MIP) and manganese peroxidase (MnP). Since both MIP and MnP showed melanoidindecolorizing activities in the presence of H2O2, it was resulted that melanoidin decolorization in the*C.hirsutus* cultureinvolvedtheproductionofextracellularH2O2 andtheperoxidases.Miyata *et al.* [104] studied on the combined use of activated sludge and the fungus *Coriolus hirsutus*. A white rotfungus,*Coriolushirsutus*, exhibitedastrongabilitytodecolorisemelanoidinincultures not supplemented with nitrogenous nutrients. Addition of peptone to the cultures lowered the ability of the fungus to decolorise melanoidin, but that of inorganic nitrogens, ammonium and nitrate did not bring about any marked reduction in the ability. It suggested an inhibitory effect of organic nitrogen on melanoidin decolorization. Therefore, for enhancing the decolorization of melanoidin in wastewaters by the fungus, activated sludge pretreatment of the wastewa‐ ters was expected to be effective, *i.e.* activated sludge is capable of converting available organ‐ ic N into inorganic N. To confirm this, waste sludge heat treatment liquor (HTL), wastewater from a sewage treatment plant, was pretreated with activated sludge. In practice, pretreat‐ ment of HTL under appropriate conditions accelerated the fungal decolorization of HTL. In the pretreated HTL, the fungus was shown to produce a high level of manganese-independent peroxidase (MIP). Addition of Mn(I1) to the pretreated HTL caused a further increase in the decolorization efficiency of the fungus and a marked increase in the manganese peroxidase (MnP) activity. Consequently, the increases in MIP and MnP activities were considered to play an important role in the enhanced ability of *C. hirsutus* to decolorise HTL.

activated-sludgeplants.Miranda*etal.*[90]workedoncoloreliminationfrommolasseswastewa‐ ter by *Aspergillus niger*. Color elimination by *Aspergillus niger*fromwastewater was studied.The influences of the nutrient concentrations, initial pH and carbon source on this color elimina‐ tion were analyzed. During the batch process, through all experiments the maximal color eliminationwas attainedafter 3or 4days inthe culture.Batchprocesses showedamaximal color elimination of 69% when MgSO4, KH2PO4, NH4NO3 and a carbon source were added to the wastewater. The continuous process, with the same nutrient concentrations, showed less color removalandthedecolorizationactivitywasmaintainedforonly4days.Benito*etal*.[102] studied on decolorization of wastewater from an alcoholic fermentation process with *Trametes versicol‐ or*. Tests varying the concentrations of carbon source, nutrients, initial pH and mycelia were carried out in order to analyze their influence on the percentage decolorization, COD removal and decrease of ammonium content of the wastewater. In this way, 82% color elimination, 77%

COD removal and 36% N-NH: decrease was attained.

**Figure 4.** Melanoidin decolorization activity by different fungi.

86 Biodegradation of Hazardous and Special Products

Dahiya *et al.* [105] worked on decolorization of synthetic and spentwash melanoidins using the white-rot fungus *Phanerochaete chrysosporium* JAG-40. This fungus was isolated from the soil samples saturated with spilled molasses collected from a sugar mill. This isolate decolor‐ ized synthetic and natural melanoidins present in spentwash in liquid fermentation; up to 80% in 6 days at 30 o C under aerobic conditions. A large inoculum size stimulated fungal biomass production, but this gave less decolorization of pigment; 5% w/v (dry weight) mycelial suspension was found optimum for maximum decolorization in melanoidin medium supple‐ mented with glucose and peptone. Gel-filtration chromatography showed that larger molec‐ ular weight fractions of melanoidins were decolorized more rapidly than small molecular weight fractions. Malandra *et al*. [106] studied the microorganisms associated with a rotating biological contactor (RBC) treating winery wastewater. One of the yeast isolates was able to reduce the COD of synthetic wastewater by 95% and 46% within 24 hrs under aerated and non aerated conditions, respectively. Kwak *et al*. [107] worked on effect of reaction pH on the photodegradation of model melanoidins. The wastewaters of molasses-based alcohol distill‐ eries contain brown colored melanoidin pigments that are one of the major pollutants.

about 90% of the molasses pigment in 10 days and at the same time caused an about 80% decrease in the biological oxygen demand value when glucose 2.5%, NaNO3 0.2%, KH2PO4 0.1% and MgSO4.7H2O 0.05% were added to the molasses wastewater from the stillage as nutrients. But without supplementation of the nutrients, the decolorization yield was only 17.5%. Furthermore, this strain showed a decolorization yield of about 70% in 11 days and caused a decrease in the biological oxygen demand value of about 90% in 15 days under nonsterile conditions. In the fed-batch system, this strain showed a constant decolorization yield of about 80% and caused a decrease in the biological oxygen demand value of about 70%.

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

89

Pant and Adholeya [8] isolated two fungal strains producing ligninolytic enzymes and having the potential to decolorise distillery effluent from the soil of a distillery effluent contaminated site. DNA was isolated from the pure cultures of these fungi and polymerase chain reaction (PCR) amplification of their internal transcribed spacer (ITS) region of nuclear ribosomal DNA was carried out. Further, the DNA was sequenced and the comparison of generated sequence with database led to their identification as *Penicillium pinophilum* TERI DB1 and *Alternaria gaisen* TERI DB6, respectively. These two isolates along with one isolate of *Pleurotus florida* EM 1303 were assessed for their ligninolytic enzyme activity in culture filtrate as well as after solid state fermentation on two substrates wheat straw and corncob powder. Ergosterol was measured to assess the growth of fungi on solid media. Both *P. pinophilum* TERI DB1 and *A. gaisen* TERI DB6 were found to produce laccase, manganese-dependent peroxidases (MnP) and lignin peroxidases (LiP). The immobilized fungal biomass was used for decolorization of the post biomethanated wastewater from the distillery. Reduction in color up to the magnitude of 86%, 50% and 47% was observed with *P. florida, P. pinophilum* and *A. gaisen* respectively. In another study, the fungi *Trichoderma viride* showed the highest decolorization yield (53.5%) when cultivated at 30 ºC for 7 days in medium contained the molasses which was diluted to 40 gL-1 in distilled water. The other *Trichoderma* species and *Penicillium* sp. also gave similar results of 40% - 45%. Decolorization yield was increased by adding peptone and yeast extract to the production medium except *Penicillium* sp. When the pH decreased below 5.0 after the incubation, the decolorization yield increased. Although reducing sugar in culture broth decreased with decreasing color intensity, there was no connection between protein utilization and decolorizing activity [113]. Two fungi (*Emericella nidulans* var. lata and *Neurospora intermedia)* have the capability to decolorise melanoidins. Maximum color was removed at pH

C, stirring 125 rpm, dextrose (0.05%) and sodium nitrate (0.025%) by both

fungi. After optimization, there was two-fold increase in color removal from 38% to 62% (DF3) and 31% to 64% (DF4) indicating significance of Taguchi approach in decolorization of

The treatment of anaerobically treated 10% distillery effluent using the microalga *Chlorella vulgaris* followed by *Lemna minuscula* resulted in 52% reduction in color [115]. In another study, Kalavathi *et al*. [116] examined the degradation of 5% melanoidin by the marine cyanobacte‐ rium *Oscillatoria boryana* BDU 92181. The organism was found to release hydrogen peroxide, hydroxyl ions and molecular oxygen during photosynthesis resulting in 60% decolorization

3, temperature 30 o

distillery mill effluent [114].

*4.2.3. Treatment by other microorganisms*

Ragukumar *et al.* [108] reported decolorization of such intensely brown colored molasses spent wash (MSW) by *Flavodon flavus,* a white-rot basidiomycete fungus isolated from a marine habitat. They attempted to improve the process of decolorization of MSW by this fungus by immobilization. Polyurethane foam-immobilized-fungus decolorized 10% diluted MSW by 60% and 73% by day 5 and 7 respectively. The immobilized fungus could be effectively used for a minimum of 3 cycles repeatedly to decolorise MSW. Besides decolorization, the fungus also removed the toxicity of MSW. Raghukumar and Rivonkar, [109] studied on the decolor‐ ization of molasses spent wash by the white rot fungus *Flavodon flavus* isolated from decom‐ posting leaves of a sea grass, decolorized pigment in molasses spentwash (MSW) by 80% after 8 days of incubation, when used at concentrations of 10% and 50%. Decolorizing activity was also present in the media prepared with half strength sea water. Decolorizing activity was also seen in low nitrogen medium, nutrient rich medium and in sugar cane bagasse medium. The percentage decolorization of MSW was highest when glucose or sucrose was used as a carbon source in the low nitrogen medium

On the other hand, *Phanerochaete chrysosporium* was used for the vinasse degradation under two different growth conditions. Vinasse was treated by *P. chrysosporium* in a liquid inoculum form, during 32 days at room temperature (approximately 25 °C) and at 39 °C. The chemical oxygen demand (COD), total phenol concentration and color removal were measured and there was a decrease in COD, phenolic concentration and color of 47.4%, 54.7% and 45.1% respectively, at roomtemperature andadecrease in54.2%, 59.4% and56.8% respectively at 39 °C [110].Thakkar *et al.* [111] studied the biocatalytic decolorization of molasses by *Phanerochaete chrysosporium*. Bioremediation potential of *Phanerochaete chrysosporium* strains NCIM 1073, NCIM 1106 and NCIM1197todecolorisemolassesinsolidandliquidmolassesmediawasstudied.Strainsvaried in the pattern of molasses decolorization on solid medium by giant colony method. Under submerged cultivation conditions, strain NCIM 1073 did not decolorise molasses while, strains NCIM 1106 and NCIM 1197 could decolorise molasses up to 82% and 76%, respectively. Under stationary cultivation conditions, none of the strains could decolorise molasses.

Srianuntapiboon *et al*. [112] worked on three kinds of molasses wastewater collected from the stillage of an alcohol factory, and their chemical properties. *Mycelia sterilia* D 90 decolorized about 90% of the molasses pigment in 10 days and at the same time caused an about 80% decrease in the biological oxygen demand value when glucose 2.5%, NaNO3 0.2%, KH2PO4 0.1% and MgSO4.7H2O 0.05% were added to the molasses wastewater from the stillage as nutrients. But without supplementation of the nutrients, the decolorization yield was only 17.5%. Furthermore, this strain showed a decolorization yield of about 70% in 11 days and caused a decrease in the biological oxygen demand value of about 90% in 15 days under nonsterile conditions. In the fed-batch system, this strain showed a constant decolorization yield of about 80% and caused a decrease in the biological oxygen demand value of about 70%.

Pant and Adholeya [8] isolated two fungal strains producing ligninolytic enzymes and having the potential to decolorise distillery effluent from the soil of a distillery effluent contaminated site. DNA was isolated from the pure cultures of these fungi and polymerase chain reaction (PCR) amplification of their internal transcribed spacer (ITS) region of nuclear ribosomal DNA was carried out. Further, the DNA was sequenced and the comparison of generated sequence with database led to their identification as *Penicillium pinophilum* TERI DB1 and *Alternaria gaisen* TERI DB6, respectively. These two isolates along with one isolate of *Pleurotus florida* EM 1303 were assessed for their ligninolytic enzyme activity in culture filtrate as well as after solid state fermentation on two substrates wheat straw and corncob powder. Ergosterol was measured to assess the growth of fungi on solid media. Both *P. pinophilum* TERI DB1 and *A. gaisen* TERI DB6 were found to produce laccase, manganese-dependent peroxidases (MnP) and lignin peroxidases (LiP). The immobilized fungal biomass was used for decolorization of the post biomethanated wastewater from the distillery. Reduction in color up to the magnitude of 86%, 50% and 47% was observed with *P. florida, P. pinophilum* and *A. gaisen* respectively. In another study, the fungi *Trichoderma viride* showed the highest decolorization yield (53.5%) when cultivated at 30 ºC for 7 days in medium contained the molasses which was diluted to 40 gL-1 in distilled water. The other *Trichoderma* species and *Penicillium* sp. also gave similar results of 40% - 45%. Decolorization yield was increased by adding peptone and yeast extract to the production medium except *Penicillium* sp. When the pH decreased below 5.0 after the incubation, the decolorization yield increased. Although reducing sugar in culture broth decreased with decreasing color intensity, there was no connection between protein utilization and decolorizing activity [113]. Two fungi (*Emericella nidulans* var. lata and *Neurospora intermedia)* have the capability to decolorise melanoidins. Maximum color was removed at pH 3, temperature 30 o C, stirring 125 rpm, dextrose (0.05%) and sodium nitrate (0.025%) by both fungi. After optimization, there was two-fold increase in color removal from 38% to 62% (DF3) and 31% to 64% (DF4) indicating significance of Taguchi approach in decolorization of distillery mill effluent [114].

#### *4.2.3. Treatment by other microorganisms*

soil samples saturated with spilled molasses collected from a sugar mill. This isolate decolor‐ ized synthetic and natural melanoidins present in spentwash in liquid fermentation; up to 80%

production, but this gave less decolorization of pigment; 5% w/v (dry weight) mycelial suspension was found optimum for maximum decolorization in melanoidin medium supple‐ mented with glucose and peptone. Gel-filtration chromatography showed that larger molec‐ ular weight fractions of melanoidins were decolorized more rapidly than small molecular weight fractions. Malandra *et al*. [106] studied the microorganisms associated with a rotating biological contactor (RBC) treating winery wastewater. One of the yeast isolates was able to reduce the COD of synthetic wastewater by 95% and 46% within 24 hrs under aerated and non aerated conditions, respectively. Kwak *et al*. [107] worked on effect of reaction pH on the photodegradation of model melanoidins. The wastewaters of molasses-based alcohol distill‐ eries contain brown colored melanoidin pigments that are one of the major pollutants.

Ragukumar *et al.* [108] reported decolorization of such intensely brown colored molasses spent wash (MSW) by *Flavodon flavus,* a white-rot basidiomycete fungus isolated from a marine habitat. They attempted to improve the process of decolorization of MSW by this fungus by immobilization. Polyurethane foam-immobilized-fungus decolorized 10% diluted MSW by 60% and 73% by day 5 and 7 respectively. The immobilized fungus could be effectively used for a minimum of 3 cycles repeatedly to decolorise MSW. Besides decolorization, the fungus also removed the toxicity of MSW. Raghukumar and Rivonkar, [109] studied on the decolor‐ ization of molasses spent wash by the white rot fungus *Flavodon flavus* isolated from decom‐ posting leaves of a sea grass, decolorized pigment in molasses spentwash (MSW) by 80% after 8 days of incubation, when used at concentrations of 10% and 50%. Decolorizing activity was also present in the media prepared with half strength sea water. Decolorizing activity was also seen in low nitrogen medium, nutrient rich medium and in sugar cane bagasse medium. The percentage decolorization of MSW was highest when glucose or sucrose was used as a carbon

On the other hand, *Phanerochaete chrysosporium* was used for the vinasse degradation under two different growth conditions. Vinasse was treated by *P. chrysosporium* in a liquid inoculum form, during 32 days at room temperature (approximately 25 °C) and at 39 °C. The chemical oxygen demand (COD), total phenol concentration and color removal were measured and there was a decrease in COD, phenolic concentration and color of 47.4%, 54.7% and 45.1% respectively, at roomtemperature andadecrease in54.2%, 59.4% and56.8% respectively at 39 °C [110].Thakkar *et al.* [111] studied the biocatalytic decolorization of molasses by *Phanerochaete chrysosporium*. Bioremediation potential of *Phanerochaete chrysosporium* strains NCIM 1073, NCIM 1106 and NCIM1197todecolorisemolassesinsolidandliquidmolassesmediawasstudied.Strainsvaried in the pattern of molasses decolorization on solid medium by giant colony method. Under submerged cultivation conditions, strain NCIM 1073 did not decolorise molasses while, strains NCIM 1106 and NCIM 1197 could decolorise molasses up to 82% and 76%, respectively. Under

stationary cultivation conditions, none of the strains could decolorise molasses.

Srianuntapiboon *et al*. [112] worked on three kinds of molasses wastewater collected from the stillage of an alcohol factory, and their chemical properties. *Mycelia sterilia* D 90 decolorized

C under aerobic conditions. A large inoculum size stimulated fungal biomass

in 6 days at 30 o

88 Biodegradation of Hazardous and Special Products

source in the low nitrogen medium

The treatment of anaerobically treated 10% distillery effluent using the microalga *Chlorella vulgaris* followed by *Lemna minuscula* resulted in 52% reduction in color [115]. In another study, Kalavathi *et al*. [116] examined the degradation of 5% melanoidin by the marine cyanobacte‐ rium *Oscillatoria boryana* BDU 92181. The organism was found to release hydrogen peroxide, hydroxyl ions and molecular oxygen during photosynthesis resulting in 60% decolorization of distillery effluent. In addition, this study suggested that cyanobacteria could use melanoidin as a better nitrogen source than carbon. Further, cyanobacteria also excrete colloidal substances like lipopolysaccharides, proteins, polyhydroxybutyrate (PHB), polyhydroxy-alkanoates (PHA), *etc*. These compounds possess COO- and ester sulphate (OSO3-) groups that can form complexes with cationic sites thereby resulting in flocculation of organic matter in the effluent. It was observed that the strain *Oscillatoria* resulted in almost complete color removal (96%) whereas *Lyngbya* and *Synechocystis* were less effective resulting in 81 and 26% color reduction, respectively [117]. The consortium of the three strains showed a maximum decolorization of 98%. This was attributed to adsorption in the initial stages followed by degradation of organic compounds which dominated in the subsequent stages. Sirianuntapiboon *et al.* [31] worked on decolorization of molasses wastewater by *Citeromyces* sp. WR-43-6. 205 yeast strains isolated from Thai-fruit samples were screened, and isolated strain No. WR-43-6 showed the highest decolorization yield (68.91%) when cultivated at 30o C for 8 days in a molasses pigments solution containing glucose 2.0%, sodium nitrate 0.1% and KH2PO4 0.1%, the pH being adjusted to 6.0. This potent strain was identified as *Citeromyces* sp. and showed highest removal efficiencies on stillage from an alcohol factory (U-MWW). The color intensity, chemical oxygen demand (COD) and biochemical oxygen demand (BOD) removal efficiencies were 75%, almost 100 and 76%, respectively. In a periodical feeding system, *Citeromyces* sp. WR-43-6 showed an almost constant decolorization yield of 60-70% over 8 days feeding of 10% fresh medium. In a replacement culture system, *Citeromyces* sp. WR-43-6 also gave a constant decolorization yield (about 75%) during four times replacement. Yeast *Citeromyces* was used for treating MWW and high and stable removal efficiencies in both color intensity and organic matter were obtained.

the sludge volume and the ease and simplicity of controlling the process [120]. Recent research has focused on the development of enzymatic processes for the treatment of wastewaters, solid wastes, hazardous wastes and soils in recognition of these potential advantages [121]. A large number of enzymes (e.g. peroxidases, oxidoreductases, cellulolytic enzymes cyanidase, proteases, amylases, etc.) from a variety of different sources have been reported to play an

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

91

Although the enzymatic system related with decolorization of melanoidins is yet to be completely understood, it seems greatly connected with fungal ligninolytic mechanisms. The white-rot fungi have a complex enzymatic system which is extracellular and non-specific, and under nutrient-limiting conditions is capable of degrading lignolytic compounds, melanoi‐ dins, and polyaromatic compounds that cannot be degraded by other microorganisms [102]. A large number of enzymes from a variety of different plants and microorganisms have been reported to play an important role in an array of waste treatment applications. Several studies regarding degradation of melanoidins, humic acids and related compounds using basidio‐ mycetes have also suggested a participation of at least one laccase enzyme in fungi belonging to *Trametes* genus. The role of enzymes other than laccase or peroxidases in the decolorization of melanoidins by *Trametes* strain was reported during the 1980s. Several reports claimed that intracellular sugar-oxidase- type enzymes (sorbose- oxidase or glucose-oxidase) had melanoi‐ din-decolorizing activities. It was suggested that melanoidins were decolorized by the active oxygen (O2; H2O2) produced by the reaction with sugar oxidases [128]. Decolorization by microbial methods includes the enzymatic breakdown of melanoidin and flocculation by microbially secreted substances. Ohmomo *et al*. [129] used *C. versicolor* Ps4a, which decolorized molasses wastewater 80% in darkness under optimum conditions. Decolorization activity involved two types of intracellular enzymes, sugar-dependent and sugar-independent. One of these enzymes required no sugar and oxygen for appearance of the activity and could decolorise MWW up to 20% in darkness and 11-17% of synthetic melanoidins. Thus, the participation of these H2O2 producing enzymes as a part of the complex enzymatic system for melanoidin degradation by fungi should be taken into account while designing any treatment strategy. One of the more complete enzymatic studies regarding melanoidin decolorization was reported by Miyata *et al.* [26]. Color removal of synthetic melanoidin by *C. hirsutus* involved the participation of peroxidases (MnP and MIP) and the extracellular H2O2 produced by glucose-oxidase, without disregard of a partial participation of fungal laccase. The manga‐ nese peroxidase (MnP) has also been reported in bacteria as extracellular enzyme for decol‐ orization of melanoidin [72]. The involvement of MnP and laccase in white rot fungus for degradation of various biopolymers (lignin and tannin) has also been reported [130, 131]. But, the detail role of MnP and laccase in bacteria for decolorization of melanoidin has not been

Mansur *et al.* [132] obtained a maximum decolorization of around 60% on day 8 after inocu‐ lating with fungus *Trametes* sp. Here effluent was added at a final concentration of 20% (v/v) after 5 days of fungal growth, the time at which high levels of laccase activity were detected in the extracellular mycelium. The white-rot basidiomycete *T. versicolor* is an active degrader of humic acids as well as of melanoidins. A melanoidin mineralizing 47 kDa extracellular

important role in an array of waste treatment applications [122-127].

fully investigated.

Moriya *et al*. [118] used two flocculant strains of yeast, *Hansenula fabianii* and *Hansenula anomala* for treatment of wastewater from beet molasses-spirits production and achieved 25.9% and 28.5% removal of TOC respectively from wastewater without dilution. Dilution of wastewater was not favourable for practical treatment of wastewater due to the longer treatment time and higher energy cost. Shojaosadati *et al.* [119] optimized the growth condi‐ tions for single cell protein (SCP) production and COD reduction by the use of *Hansenula* sp. in sugar beet stillage. They concluded that production of SCP from stillage is one of the most promising options. Besides, the yeast was also found to utilize lactate and acetate that are inhibitory to ethanol production. As a result, the treated effluent could be used as dilution water for fermentation thereby reducing the residual stillage volume by 70%.

#### **5. Role of enzymes in effluent decolorization**

The enzymatic treatment falls between the physicochemical and biological treatment process‐ es. It has technological advantages and requires economical considerations to apply it on a large scale. It has some potential advantages over the conventional treatment. These includes: applicability to biorefractory compounds; operation either at high or low contaminant concentrations; operation over a wide range of pH, temperature and salinity; absence of shock loading effects; absence of delays associated with the acclimatization of biomass; reduction in the sludge volume and the ease and simplicity of controlling the process [120]. Recent research has focused on the development of enzymatic processes for the treatment of wastewaters, solid wastes, hazardous wastes and soils in recognition of these potential advantages [121]. A large number of enzymes (e.g. peroxidases, oxidoreductases, cellulolytic enzymes cyanidase, proteases, amylases, etc.) from a variety of different sources have been reported to play an important role in an array of waste treatment applications [122-127].

of distillery effluent. In addition, this study suggested that cyanobacteria could use melanoidin as a better nitrogen source than carbon. Further, cyanobacteria also excrete colloidal substances like lipopolysaccharides, proteins, polyhydroxybutyrate (PHB), polyhydroxy-alkanoates (PHA), *etc*. These compounds possess COO- and ester sulphate (OSO3-) groups that can form complexes with cationic sites thereby resulting in flocculation of organic matter in the effluent. It was observed that the strain *Oscillatoria* resulted in almost complete color removal (96%) whereas *Lyngbya* and *Synechocystis* were less effective resulting in 81 and 26% color reduction, respectively [117]. The consortium of the three strains showed a maximum decolorization of 98%. This was attributed to adsorption in the initial stages followed by degradation of organic compounds which dominated in the subsequent stages. Sirianuntapiboon *et al.* [31] worked on decolorization of molasses wastewater by *Citeromyces* sp. WR-43-6. 205 yeast strains isolated from Thai-fruit samples were screened, and isolated strain No. WR-43-6 showed the highest

solution containing glucose 2.0%, sodium nitrate 0.1% and KH2PO4 0.1%, the pH being adjusted to 6.0. This potent strain was identified as *Citeromyces* sp. and showed highest removal efficiencies on stillage from an alcohol factory (U-MWW). The color intensity, chemical oxygen demand (COD) and biochemical oxygen demand (BOD) removal efficiencies were 75%, almost 100 and 76%, respectively. In a periodical feeding system, *Citeromyces* sp. WR-43-6 showed an almost constant decolorization yield of 60-70% over 8 days feeding of 10% fresh medium. In a replacement culture system, *Citeromyces* sp. WR-43-6 also gave a constant decolorization yield (about 75%) during four times replacement. Yeast *Citeromyces* was used for treating MWW and high and stable removal efficiencies in both color intensity and organic matter were

Moriya *et al*. [118] used two flocculant strains of yeast, *Hansenula fabianii* and *Hansenula anomala* for treatment of wastewater from beet molasses-spirits production and achieved 25.9% and 28.5% removal of TOC respectively from wastewater without dilution. Dilution of wastewater was not favourable for practical treatment of wastewater due to the longer treatment time and higher energy cost. Shojaosadati *et al.* [119] optimized the growth condi‐ tions for single cell protein (SCP) production and COD reduction by the use of *Hansenula* sp. in sugar beet stillage. They concluded that production of SCP from stillage is one of the most promising options. Besides, the yeast was also found to utilize lactate and acetate that are inhibitory to ethanol production. As a result, the treated effluent could be used as dilution

The enzymatic treatment falls between the physicochemical and biological treatment process‐ es. It has technological advantages and requires economical considerations to apply it on a large scale. It has some potential advantages over the conventional treatment. These includes: applicability to biorefractory compounds; operation either at high or low contaminant concentrations; operation over a wide range of pH, temperature and salinity; absence of shock loading effects; absence of delays associated with the acclimatization of biomass; reduction in

water for fermentation thereby reducing the residual stillage volume by 70%.

**5. Role of enzymes in effluent decolorization**

C for 8 days in a molasses pigments

decolorization yield (68.91%) when cultivated at 30o

90 Biodegradation of Hazardous and Special Products

obtained.

Although the enzymatic system related with decolorization of melanoidins is yet to be completely understood, it seems greatly connected with fungal ligninolytic mechanisms. The white-rot fungi have a complex enzymatic system which is extracellular and non-specific, and under nutrient-limiting conditions is capable of degrading lignolytic compounds, melanoi‐ dins, and polyaromatic compounds that cannot be degraded by other microorganisms [102]. A large number of enzymes from a variety of different plants and microorganisms have been reported to play an important role in an array of waste treatment applications. Several studies regarding degradation of melanoidins, humic acids and related compounds using basidio‐ mycetes have also suggested a participation of at least one laccase enzyme in fungi belonging to *Trametes* genus. The role of enzymes other than laccase or peroxidases in the decolorization of melanoidins by *Trametes* strain was reported during the 1980s. Several reports claimed that intracellular sugar-oxidase- type enzymes (sorbose- oxidase or glucose-oxidase) had melanoi‐ din-decolorizing activities. It was suggested that melanoidins were decolorized by the active oxygen (O2; H2O2) produced by the reaction with sugar oxidases [128]. Decolorization by microbial methods includes the enzymatic breakdown of melanoidin and flocculation by microbially secreted substances. Ohmomo *et al*. [129] used *C. versicolor* Ps4a, which decolorized molasses wastewater 80% in darkness under optimum conditions. Decolorization activity involved two types of intracellular enzymes, sugar-dependent and sugar-independent. One of these enzymes required no sugar and oxygen for appearance of the activity and could decolorise MWW up to 20% in darkness and 11-17% of synthetic melanoidins. Thus, the participation of these H2O2 producing enzymes as a part of the complex enzymatic system for melanoidin degradation by fungi should be taken into account while designing any treatment strategy. One of the more complete enzymatic studies regarding melanoidin decolorization was reported by Miyata *et al.* [26]. Color removal of synthetic melanoidin by *C. hirsutus* involved the participation of peroxidases (MnP and MIP) and the extracellular H2O2 produced by glucose-oxidase, without disregard of a partial participation of fungal laccase. The manga‐ nese peroxidase (MnP) has also been reported in bacteria as extracellular enzyme for decol‐ orization of melanoidin [72]. The involvement of MnP and laccase in white rot fungus for degradation of various biopolymers (lignin and tannin) has also been reported [130, 131]. But, the detail role of MnP and laccase in bacteria for decolorization of melanoidin has not been fully investigated.

Mansur *et al.* [132] obtained a maximum decolorization of around 60% on day 8 after inocu‐ lating with fungus *Trametes* sp. Here effluent was added at a final concentration of 20% (v/v) after 5 days of fungal growth, the time at which high levels of laccase activity were detected in the extracellular mycelium. The white-rot basidiomycete *T. versicolor* is an active degrader of humic acids as well as of melanoidins. A melanoidin mineralizing 47 kDa extracellular protein corresponding to the major mineralizing enzyme system from *T. versicolor* was isolated by Dehorter and Blondeau [133]. This Mn2+ dependent enzyme system required oxygen and was described to be as peroxidase. Uniform, small and spongy pellets of the fungus *T. versicolor* were used as inoculum for color removal using different nutrients such as ammonium nitrate, manganese phosphate, magnesium sulphate and potassium phosphate and also sucrose as carbon source [102]. Maximum color removal of 82% and 36% removal of N-NH4 was obtained on using low sucrose concentration and KH2PO4 as the only nutrient. Some studies have identified the lignin degradation related enzymes participating in the melanoidin decolorization. Intracellular H2O2 producing sugar oxidases have been isolated from *Coriolus* strains. Also, *C. hirsustus* have been reported to produce enzymes that catalyze melanoidin decolorization directly without additions of sugar and O2. Miyata *et al.* [26] used *C. hirsutus* pellets to decolorise a melanoidin-containing medium. It was elucidated that extracellular H2O2 and two extracellular peroxidases, a manganese-independent peroxidase (MIP) and manganese peroxidase (MnP) were involved in decolorization activity. Lee *et al.* [134] inves‐ tigated the dye-decolorizing peroxidase by cultivating *Geotrichum candidum* Dec1 using molasses as a carbon source. Components in the molasses medium stimulated the production of decolorizing peroxidase but inhibited the decolorizing activity of the purified enzyme. It was found that the inhibitory effect of molasses can be eliminated at dilution ratios of more than 25. Recently D'souza *et al.* [135] reported 100% decolorization of 10% spent wash by a marine fungal isolate whose laccase production increased several folds in the presence of phenolic and non-phenolic inducers. A combined treatment technique consisting of enzyme catalyzed in situ transformation of pollutants followed by aerobic biological oxidation was investigated by Sangave and Pandit [121] for the treatment of alcohol distillery spent wash. It was suggested that enzymatic pretreatment of the distillery effluent leads to *in situ* formation of the hydrolysis products, which have different physical properties and are easier to assimilate than the parent pollutant molecules by the microorganisms, leading to faster initial rates of aerobic oxidation even at lower biomass levels. In another study, Sangave and Pandit [136] used irradiation and ultrasound combined with the use of an enzyme as pretreatment technique for treatment of distillery wastewater. The combination of the ultrasound and enzyme yielded the best COD removal efficiencies as compared to the processes when they were used as stand-alone treatment techniques. Enzymatic decolorization of molasses medium has also been tried using *P. chrysosporium* [111]. Under stationary cultivation conditions, none of the strains could decolorise molasses nor produce enzymes lignin peroxidase, manganese peroxidase and laccase. All of them could produce lignin peroxidase and manganese peroxi‐ dase when cultivated in flat bottom glass bottles under stationary cultivation conditions.

The white-rot basidiomycetous fungus *F. flavus* produces extracelluar enzymes, manganesedependent peroxidase (MNP), lignin peroxidase (LIP) and laccase which decolorizes a number of synthetic dyes like Poly R and remazol brilliant blue R etc. [138]. However, a negative correlation was found between MnP concentrations and decolorization of molasses spent wash (MSW) [139]. The levels of glucose oxidase activity also have correlation with decolorization of MSW. Raghukumar *et al.* [108] proposes that hydrogen peroxide produced as a result of glucose oxidase activity acts as a bleaching agent on MSW. Which has been further confirmed that in the absence of glucose the decolorization of MSW is lesser. With the addition of the starch to the medium, amylase activity stimulates and releasing reducing sugars. Which are oxidized by glucose oxidase producing hydrogen peroxide as a byproduct results in greater

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

93

In the few last decades, interest has been developed in the field of bioremediation by using microbes. Several microorganisms such as bacteria and fungi, show a good ability to decolorize the effluent of the melanoidin based distillery industries. Thus, a better understanding of the microbial activities responsible for the degradation of melanoidins would contribute to enhancing the efficiency of the overall treatment system. In degradation process it would also be necessary to know the end product of melanoidins. Genetic improvement of isolates can be explored in future for improving their decolorization efficiency. Thus, it can be suggested that microbial decolorization holds promise and can be exploited to develop a cost effective, ecofriendly biotechnology package for the treatment of distillery effluent. More technically advanced research efforts are required for searching, exploiting new bacterial species and improvement of practical application to propagate the use of bacteria for bioremediation of industrial effluents. Enzymatic studies would be employed to understand the mechanism of degradation of melanoidins in future prospectives. Such organisms could be used in bioreac‐ tors for treatment of wastewaters or scaling up for enzyme productions. Broader validation of these new technologies and integration of different methods in the current treatment schemes will most likely in the near future, render these both efficient and economically viable. Finally, microbial bioremediation involves a combination of microbiologists, biotechnologists, chemists and engineers and is ideal to plug the ever widening gap between the different

decolorization of MSW [108].

**6. Conclusion**

disciplines.

**Author details**

Anita Rani Santal1

and Nater Pal Singh2

1 Department of Microbiology, M. D. University, Rohtak, Haryana, India

2 Centre for Biotechnology, M. D. University, Rohtak, Haryana, India

The MnP and laccase activity in the culture supernatant decreases with the increase in the time duration and further bacterial incubation showed gradual decrease of both enzyme activities. The enzyme activity has direct co-relation with the melanoidin decolorization. The initiation of MnP activity in culture supernatant starts at 48 h of bacterial growth and remains active up to 192 h, whereas, the laccase induction starts at 96 h and its activity decreased after 192 h. This resulted in the initial role of MnP in melanoidin degradation [70]. The melanoidin decolori‐ zation has also been reported by MnP activity in fungus [8, 108]. The enzyme laccase also helps in decolorizing melanoidin [72, 137].

The white-rot basidiomycetous fungus *F. flavus* produces extracelluar enzymes, manganesedependent peroxidase (MNP), lignin peroxidase (LIP) and laccase which decolorizes a number of synthetic dyes like Poly R and remazol brilliant blue R etc. [138]. However, a negative correlation was found between MnP concentrations and decolorization of molasses spent wash (MSW) [139]. The levels of glucose oxidase activity also have correlation with decolorization of MSW. Raghukumar *et al.* [108] proposes that hydrogen peroxide produced as a result of glucose oxidase activity acts as a bleaching agent on MSW. Which has been further confirmed that in the absence of glucose the decolorization of MSW is lesser. With the addition of the starch to the medium, amylase activity stimulates and releasing reducing sugars. Which are oxidized by glucose oxidase producing hydrogen peroxide as a byproduct results in greater decolorization of MSW [108].

#### **6. Conclusion**

protein corresponding to the major mineralizing enzyme system from *T. versicolor* was isolated by Dehorter and Blondeau [133]. This Mn2+ dependent enzyme system required oxygen and was described to be as peroxidase. Uniform, small and spongy pellets of the fungus *T. versicolor* were used as inoculum for color removal using different nutrients such as ammonium nitrate, manganese phosphate, magnesium sulphate and potassium phosphate and also sucrose as carbon source [102]. Maximum color removal of 82% and 36% removal of N-NH4 was obtained on using low sucrose concentration and KH2PO4 as the only nutrient. Some studies have identified the lignin degradation related enzymes participating in the melanoidin decolorization. Intracellular H2O2 producing sugar oxidases have been isolated from *Coriolus* strains. Also, *C. hirsustus* have been reported to produce enzymes that catalyze melanoidin decolorization directly without additions of sugar and O2. Miyata *et al.* [26] used *C. hirsutus* pellets to decolorise a melanoidin-containing medium. It was elucidated that extracellular H2O2 and two extracellular peroxidases, a manganese-independent peroxidase (MIP) and manganese peroxidase (MnP) were involved in decolorization activity. Lee *et al.* [134] inves‐ tigated the dye-decolorizing peroxidase by cultivating *Geotrichum candidum* Dec1 using molasses as a carbon source. Components in the molasses medium stimulated the production of decolorizing peroxidase but inhibited the decolorizing activity of the purified enzyme. It was found that the inhibitory effect of molasses can be eliminated at dilution ratios of more than 25. Recently D'souza *et al.* [135] reported 100% decolorization of 10% spent wash by a marine fungal isolate whose laccase production increased several folds in the presence of phenolic and non-phenolic inducers. A combined treatment technique consisting of enzyme catalyzed in situ transformation of pollutants followed by aerobic biological oxidation was investigated by Sangave and Pandit [121] for the treatment of alcohol distillery spent wash. It was suggested that enzymatic pretreatment of the distillery effluent leads to *in situ* formation of the hydrolysis products, which have different physical properties and are easier to assimilate than the parent pollutant molecules by the microorganisms, leading to faster initial rates of aerobic oxidation even at lower biomass levels. In another study, Sangave and Pandit [136] used irradiation and ultrasound combined with the use of an enzyme as pretreatment technique for treatment of distillery wastewater. The combination of the ultrasound and enzyme yielded the best COD removal efficiencies as compared to the processes when they were used as stand-alone treatment techniques. Enzymatic decolorization of molasses medium has also been tried using *P. chrysosporium* [111]. Under stationary cultivation conditions, none of the strains could decolorise molasses nor produce enzymes lignin peroxidase, manganese peroxidase and laccase. All of them could produce lignin peroxidase and manganese peroxi‐ dase when cultivated in flat bottom glass bottles under stationary cultivation conditions.

The MnP and laccase activity in the culture supernatant decreases with the increase in the time duration and further bacterial incubation showed gradual decrease of both enzyme activities. The enzyme activity has direct co-relation with the melanoidin decolorization. The initiation of MnP activity in culture supernatant starts at 48 h of bacterial growth and remains active up to 192 h, whereas, the laccase induction starts at 96 h and its activity decreased after 192 h. This resulted in the initial role of MnP in melanoidin degradation [70]. The melanoidin decolori‐ zation has also been reported by MnP activity in fungus [8, 108]. The enzyme laccase also helps

in decolorizing melanoidin [72, 137].

92 Biodegradation of Hazardous and Special Products

In the few last decades, interest has been developed in the field of bioremediation by using microbes. Several microorganisms such as bacteria and fungi, show a good ability to decolorize the effluent of the melanoidin based distillery industries. Thus, a better understanding of the microbial activities responsible for the degradation of melanoidins would contribute to enhancing the efficiency of the overall treatment system. In degradation process it would also be necessary to know the end product of melanoidins. Genetic improvement of isolates can be explored in future for improving their decolorization efficiency. Thus, it can be suggested that microbial decolorization holds promise and can be exploited to develop a cost effective, ecofriendly biotechnology package for the treatment of distillery effluent. More technically advanced research efforts are required for searching, exploiting new bacterial species and improvement of practical application to propagate the use of bacteria for bioremediation of industrial effluents. Enzymatic studies would be employed to understand the mechanism of degradation of melanoidins in future prospectives. Such organisms could be used in bioreac‐ tors for treatment of wastewaters or scaling up for enzyme productions. Broader validation of these new technologies and integration of different methods in the current treatment schemes will most likely in the near future, render these both efficient and economically viable. Finally, microbial bioremediation involves a combination of microbiologists, biotechnologists, chemists and engineers and is ideal to plug the ever widening gap between the different disciplines.

#### **Author details**

Anita Rani Santal1 and Nater Pal Singh2

1 Department of Microbiology, M. D. University, Rohtak, Haryana, India

2 Centre for Biotechnology, M. D. University, Rohtak, Haryana, India

#### **References**

[1] Shweta, K. Current Trends in Bioremediation and Biodegradation. Journal of Biore‐ mediation and Biodegradation (2012). e114.

[14] Chandra, R, Bharagava, R. N, & Rai, V. Melanoidins as Major Colorant in Sugarcane Molasses Based Distillery Effluent and its Degradation. Bioresource Technology

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

95

[15] Mohana, S, Desai, C, & Madamwar, D. Biodegradation and Decolorization of Anae‐ robically Treated Distillery Spent Wash by a Novel Bacterial Consortium. Biore‐

[16] Kumar, P, & Chandra, P. Decolorization and Detoxification of Synthetic Molasses Melanoidins by Individual and Mixed Cultures of Bacillus spp. Bioresource Technol‐

[17] Wedzicha, B. L, & Kaputo, M. T. Melanoidins from Glucose and Glycine: Composi‐ tion, Characteristics and Reactivity towards Sulphite Ion. Food Chemistry (1992). ,

[18] Reynolds, T. M. Chemistry of Non-Enzymatic Browning of the Reaction Between

[19] Kumar, V, & Wati, L. FitzGibbon FJ, Nigam P, Banat IM, Singh D, Marchant R. Biore‐ mediation and Decolorization of Anaerobically Digested Distillery Spent Wash. Bio‐

[20] Kumar, V, Wati, L, Nigam, P, & Banat, I. M. MacMullan G, Singh D, Marchant R. Mi‐ crobial Decolorization and Bioremediation of Anaerobically Digested Molasses Spent

[21] Kim, S. B, Hayase, F, & Kato, H. Decolorization and Degradation Products of Mela‐ noidins on Ozonolysis. Agricultural and Biological Chemistry (1985). , 49(3), 785-792.

[22] Ohmomo, S, Kaneko, Y, Sirianuntapiboon, S, Somachi, P, Atthasumpunna, P, & Na‐ kamura, I. Decolorization of Molasses Wastewater by a Thermophilic Strain Asper‐

[23] Satyawali, Y, & Balakrishnan, M. Removal of Color from Biomethanated Distillery Spentwash by Treatment with Activated Carbons. Bioresource Technology (2007). ,

[24] Ivarson, K. C, & Benzing-purdie, L. M. Degradation of Melanoidins by Soil Microor‐ ganisms Under Laboratory Conditions. Canadian Journal of Soil Science (1987). ,

[25] Kitts, D. D, Wu, C. H, Stich, H. F, & Powrie, W. D. Effect of Glucose-Lysine Maillard Reaction Products on Bacterial and Mammalian Cell Mutagenesis. Journal of Agri‐

[26] Miyata, N, Iwahori, K, & Fujita, M. Manganese-Independent and-Dependent Decol‐ orization of Melanoidin by Extracellular Hydrogen Peroxide and Peroxidases from

culture and Food Chemistry (1993). , 41(12), 2353-2358.

Wash Effluent by Aerobic Bacterial Culture. Microbios (1997b). , 89-81.

gillus fumigatus G- Agricultural and Biological Chemistry (1987). , 2-6.

Aldoses and Amines. Advances in Food Research (1968). , 12-1.

(2008). , 99-4648.

ogy (2006). , 97-2096.

43-359.

98-2629.

67-409.

source Technology (2007). , 98(2), 333-339.

technology Letters (1997a). , 19(4), 311-313.


[14] Chandra, R, Bharagava, R. N, & Rai, V. Melanoidins as Major Colorant in Sugarcane Molasses Based Distillery Effluent and its Degradation. Bioresource Technology (2008). , 99-4648.

**References**

[1] Shweta, K. Current Trends in Bioremediation and Biodegradation. Journal of Biore‐

[2] Asamudo, N. U, Daba, A. S, & Ezeronye, O. U. Bioremediation of Textile Effluent Us‐ ing Phanerochaete chrysosporium. African Journal of Food, Agriculture, Nutrition

[3] Tyagi, M. da Fonseca MMR, de Carvalho CC. Bioaugmentation and Biostimulation Strategies to Improve the Effectiveness of Bioremediation Processes. Biodegradation

[4] Madhavi, A, & Rao, A. P. Effect of Industrial Effluent on Properties of Groundwater.

[5] Saravanamoorthy, M. D. Kumari BDR. Effect of Textile Wastewater on Morpho‐ physiology and Yield on two Varieties of Peanut (Arachis hypogaea L.). Journal of

[6] Naik, N. M, Jagdeesh, K. S, & Alagawadi, A. R. Microbial Decolorization of Spent‐

[7] Joshi, H. C. Bio-Energy Potential of Distillery Effluents. BioEnergy News (1999). ,

[8] Pant, D, & Adholeya, A. Identification Ligninolytic Enzyme Activity and Decoloriza‐ tion Potential of two Fungi Isolated from a Distillery Effluent Contaminated Site. Wa‐

[9] Chandra, R. Development of Microorganism for Removal of Color from Anaerobical‐ ly Treated Distillery Effluent. Final technical report submitted to Department of Bio‐

[10] Chandra, R, Kumar, K, & Singh, J. Impact of Anaerobically Treated and Untreated (Raw) Distillery Effluent Irrigation on Soil Microflora Growth Total Chlorophyll and Protein Contents of Phaseolus aureus L. Journal of Environmental Biology (2004). ,

[11] Pazouki, M, Shayegan, J, & Afshari, A. Screening of Microorganisms for Decoloriza‐ tion of Treated Distillery Wastewater. Iranian Journal of Science and Technology

[12] FitzGibbon FSingh D, McMullan G, Marchant R. The Effect of Phenolic Acids and Molasses Spent Wash Concentration on Distillery Wastewater Remediation by Fungi.

[13] Pant, D, & Adholeya, A. Biological Approaches for Treatment of Distillery Wastewa‐

mediation and Biodegradation (2012). e114.

Journal of Environmental Biology (2003). , 24(2), 187-192.

wash: A Review. Indian Journal of Microbiology (2008). , 48-41.

Agricultural Technology (2007). , 3(2), 335-343.

ter, Air and Soil Pollution (2007b). , 183-165.

Process Biochemistry (1998). , 33(8), 799-803.

ter: A Review. Bioresource Technology (2007a). , 98-2321.

technology New Delhi; (2003).

25(4), 381-385.

(2008). , 32-53.

and Development (2011).

94 Biodegradation of Hazardous and Special Products

(2011). , 22(2), 231-241.

3-10.


Coriolus hirsutus pellets. Journal of Fermentation and Bioengineering (1998). , 85(5), 550-553.

[40] Satyawali, Y, & Balakrishnan, M. Wastewater Treatment in Molasses-Based Alcohol Distilleries for Cod and Color Removal: A Review. Journal of Environmental Man‐

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

97

[41] Bitton, G, Jung, K, & Koopman, B. Evaluation of a Microplate Assay Specific for Heavy Metal Toxicity. Archives of Environmental Contamination and Toxicology

[42] Pandey, R. A, Malhotra, S, Tankhiwale, A, Pande, S, Pathe, P. P, & Kaul, S. N. Treat‐ ment of Biologically Treated Distillery Effluent- A Case Study. International Journal

[43] Migo, V. P, Matsumura, M, Delrosario, E. J, & Kataoka, H. Decolorization of Molass‐ es Wastewater Using an Inorganic Flocculent. Journal of Fermentation and Bioengin‐

[44] Coca, M, Pena, M, & Gonzalez, G. Chemical Oxidation Processes for Decolorization of Brown Colored Molasses Wastewater. Ozone: Science and Engineering (2005). ,

[45] Kumaresan, T. Sheriffa Begum KMM, Sivashanmugam P, Anantharaman N, Sundar‐ am S. Experimental Studies on Treatment of Distillery Effluent by Liquid Membrane

[46] De Wilde FGNDemineralization of a Molasses Distillery Wastewater. Desalination

[47] Bories, A, Sire, Y, & Colin, T. Odorous Compounds Treatment of Winery and Distill‐ ery Effluents During Natural Evaporation in Ponds. Water Science and Technology

[48] Bhandari, H. C, Mitra, A. K, & Kumar, S. Crest's Integrated System: Reduction and Recycling of Effluents in Distilleries. In: Tewari P.K. (ed.) Liquid Asset Proceedings of Indo-EU Workshop on Promoting Efficient Water Use in Agro-based Industries.

[49] Gulati, N. Conservation of Resources Using Evaporation and Spray Drying Technol‐ ogy for Distillery and Paper Industries. In: Tewari P.K. (ed.) Liquid Asset Proceed‐ ings of Indo-EU Workshop on Promoting Efficient Water Use in Agro-Based

[50] Cortez LABPerez LEB. Experiences on Vinasse Disposalpart III: Combustion of Vi‐ nasse Fuel Oil Emulsions. Brazilian Journal of Chemical Engineering (1997). , 14(1),

[51] Joshi, D. L. High Rate Anaerobic Treatment of Industrial Wastewater in Tropics. Thammasat International Journal of Science and Technology (1998). , 3(1), 1-7.

agement (2008). , 86-481.

(1994). , 27(1), 25-28.

eering (1993). , 75-438.

27-365.

9-18.

(1987). , 67-481.

(2005). , 51(1), 129-136.

of Environmental Studies (2003). , 60(3), 263-275.

Extraction. Chemical Engineering Journal (2003).

New Delhi, India: TERI Press; , 167-169.

Industries. New Delhi, India: TERI Press; (2004). , 163-166.


[40] Satyawali, Y, & Balakrishnan, M. Wastewater Treatment in Molasses-Based Alcohol Distilleries for Cod and Color Removal: A Review. Journal of Environmental Man‐ agement (2008). , 86-481.

Coriolus hirsutus pellets. Journal of Fermentation and Bioengineering (1998). , 85(5),

[27] Silvan, J. M. Lagemaat JVD, Olano A, Castillo MDD. Analysis and Biological Proper‐ ties of Amino Acid Derivates Formed by Maillard Reaction in Foods. Journal of Phar‐

[28] Morales, F, & Jimnez-perez, S. Free Radical Scavenging Capacity of Maillard Reac‐ tion Products as Related to Color and Fluorescence. Food Chemistry (2001). , 72-119.

[29] Cammerer, B, Jaluschkov, V, & Kroh, L. W. Carbohydrates structures as part of the

[30] Manisankar, P, Rani, C, & Vishwanathan, S. Effect of Halides in the Electrochemical

[31] Sirianuntapiboon, S, Zohsalam, P, & Ohmomo, S. Decolorization of Molasses Waste‐

[32] Migo, V. P. Del Rosario EJ, Matsumura M. Flocculation of Melanoidins Induced by Inorganic Ions. Journal of Fermentation and Bioengineering (1997). , 83(3), 287-291.

[33] Miura, M, & Gomyo, T. Formation of Blue Pigments in the Earlier Stage of Browning in the System Composed of D-Xylose and Glycine. Nippon Nogeikagaku Kaishi

[34] Hayase, F, Takahashi, Y, Tominaga, S, Miura, M, Gomyo, T, & Kato, H. Identification of Blue Pigment Formed in a D-Xylose-Glycine Reaction System. Bioscience, Biotech‐

[35] Hodge, J. E. Chemistry of Browning Reactions in Model Systems. Journal of Agricul‐

[36] Hayashi, T, & Namiki, M. Role of Sugar Fragmentation in an Early Stage Browning of Amino-Carbonyl Reaction of Sugar with Amino Acid. Agricultural and Biological

[37] Sowmeyan, R, & Swaminathan, G. Effluent Treatment Process in Molasses Based Distillery Industries: A Review. Journal of Hazardous Materials (2008). , 152-453. [38] Venkat, S. M, Krishna, S. M, & Karthikeyan, J. Adsorption Mechanism of Acid-Azo Dye from Aqueous Solution on to Coal/Coal Based Sorbents and Activated Carbon: A Mechanist Study. In: Jayarama Reddy S. (ed.) Analytical Techniques in Monitoring

the Environment. Tirupathi, India: Student offset printers; (2000). , 97-103.

[39] Bernardo, E. C, Egashira, R, & Kawasaki, J. Decolorization of Molasses Wastewater Using Activated Carbon Prepared from Cane Bagasses. Carbon (1997). , 35(9),

melanoidins skeleton. International Congress Series (2002). , 1245-269.

Treatment of Distillery Effluent. Chemosphere (2004). , 57-961.

nology, and Biochemistry (1999). , 63(8), 1512-1514.

ture and Food Chemistry (1953). , 1-928.

Chemistry (1986). , 50(8), 1965-1970.

water by Citeromyces spWR- Process Biochemistry (2004a). , 43-6.

maceutical and Biomedical Analysis (2006). , 41-1543.

550-553.

96 Biodegradation of Hazardous and Special Products

(1982). , 56-417.

1217-1221.


[52] Nandy, T, Shastry, S, & Kaul, S. N. Wastewater Management in a Cane Molasses Dis‐ tillery Involving Bioresource Recovery. Journal of Environmental Management (2002). , 65-25.

[65] Bhatia, D, Vieth, W. R, & Vekatasubramanian, K. Steady State and Transit Behaviour in Microbial Methanification: II. Mathematical Modeling and Verification. Biotech‐

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

99

[66] Parkin, G. F, & Speece, R. E. Yang CHJ, Kocher WM. Response of Methane Formation Systems to Industrial Toxicants. Journal of the Water Pollution Control Federation

[67] Karhadkar, P. P, Handa, B. K, & Khanna, P. Pilot-Scale Distillery Spent Wash Biome‐ thanation. Journal of Environmental Engineering (1990). , 116(6), 1029-1045.

[68] Krzywonos, M, & Seruga, P. Decolorization of Sugar Beet Molasses Vinasse a High Strength Distillery Wastewater by Lactic Acid Bacteria. Polish Journal of Environ‐

[69] Krzywonos, M. Decolorization of Sugar Beet Distillery Effluent Using Mixed Cul‐ tures of Bacteria of the Genus Bacillus. African Journal of Biotechnology (2012). ,

[70] Yadav, S, & Chandra, R. Biodegradation of Organic Compounds of Molasses Mela‐ noidin (MM) from Biomethanated Distillery Spent Wash (BMDS) during the Decolor‐

[71] Santal, A. R, Singh, N. P, & Saharan, B. S. Biodegradation and Detoxification of Mela‐ noidin from Distillery Effluent Using an Aerobic Bacterial Strain SAG5 of Alcaligenes

[72] Bharagava, R. N, Chandra, R, & Rai, V. Isolation and Characterization of Aerobic Bacteria Capable of the Degradation of Synthetic and Natural Melanoidins from Dis‐ tillery Effluent. World Journal of Microbiology and Biotechnology (2009). , 25-737. [73] Tondee, T, & Sirianuntapiboon, S. Decolorization of Molasses Wastewater by Lacto‐

[74] Jiranuntipon, S, Chareonpornwattana, S, Damronglerd, S, Albasi, C, & Delia, M. L. Decolorization of Synthetic Melanoidins-Containing Wastewater by a Bacterial Con‐ sortium. Journal of Industrial Microbiology and Biotechnology (2008). , 35-1313. [75] Adikane, H. V, Dange, M. N, & Selvakumari, K. Optimization of Anaerobically Di‐ gested Distillery Molasses Spent Wash Decolorization Using Soil as Inoculum in the Absence of Additional Carbon and Nitrogen Source. Bioresourse Technology (2006). ,

[76] Sivakumar, N, Saravanan, V, & Balakumar, S. Bacterial Decolorization and Bioreme‐ diation of an Anaerobically Digested Molasses Spent Wash. Asian Journal of Micro‐

biology, Biotechnology and Environment Science (2006). , 8(2), 291-295.

ization by a Potential Bacterial Consortium. Biodegradation (2012). , 23-609.

faecalis. Journal of Hazardous Materials (2011). , 193-319.

bacillus plantarum NoPV Bioresource Technology (2008). , 71-1861.

nology and Bioengineering (1985). , 27(8), 1192-1198.

mental Studies (2012). , 21(4), 943-948.

(1983). , 55-44.

11(14), 3464-3475.

97-2131.


[65] Bhatia, D, Vieth, W. R, & Vekatasubramanian, K. Steady State and Transit Behaviour in Microbial Methanification: II. Mathematical Modeling and Verification. Biotech‐ nology and Bioengineering (1985). , 27(8), 1192-1198.

[52] Nandy, T, Shastry, S, & Kaul, S. N. Wastewater Management in a Cane Molasses Dis‐ tillery Involving Bioresource Recovery. Journal of Environmental Management

[53] Bhoi ZMPMehrotra I, Shrivastava AK. Treatability Studies of Black Liquor by Up‐ flow Anaerobic Sludge Blanket Reactor. Journal of Environmental Engineering and

[54] Farhadian, M, Borghei, M, & Umrania, V. V. Treatment of Beet Sugar Wastewater by

[55] Travieso, L, Sánchez, E, Borja, R, Benítez, F, Raposo, F, Rincón, B, & Jiménez, A. M. Evaluation of a Laboratory-Scale Stabilization Pond for Tertiary Treatment of Distill‐ ery Waste Previously Treated by a Combined Anaerobic Filter-Aerobic Trickling Sys‐

[56] Sharma, J, & Singh, R. Effect of Nutrients Supplementation on Anaerobic Sludge De‐ velopment and Activity for Treating Distillery Effluent. Bioresource Technology

[57] Driessen WJBMTielbaard MH, Vereijken TLFM. Experience on Anaerobic Treatment of Distillery Effluent with the UASB Process. Water Science and Technology (1994). ,

[58] Blonskaja, V, Menert, A, & Vilu, R. Use of Two-Stage Anaerobic Treatment for Dis‐ tillery Waste. Advances in Environmental Research (2003). , 7(3), 671-678.

[59] Beltran, F. J, Garcia-araya, J. F, & Alvarez, P. M. Wine Distillery Wastewater Degra‐ dation Improvement of Aerobic Biodegradation by Means of an Integrated Chemical (Ozone)-Biological Treatment. Journal of Agricultural and Food Chemistry (1999). ,

[60] Kitamura, Y, Maekawa, T, Tagawa, A, Hayashi, H, & Farrell-poe, K. L. Treatment of Strong Organic Nitrogenous Wastewater by an Anaerobic Contact Process Incorpo‐

[61] Yeoh, B. G. Two-Phase Anaerobic Treatment of Cane-Molasses Alcohol Stillage. Wa‐

[62] Harada, H, Uemura, S, Chen, A. C, & Jayadevan, J. Anaerobic Treatment of a Recalci‐ trant Distillery Wastewater by a Thermophilic UASB Reactor. Bioresource Technolo‐

[63] Bhatti, Z. I, Furukawa, K, & Fujita, M. Microbial Diversity in UASB Reactors. Pure

[64] Babu, V. L, Mohan, S. V, & Sharma, P. N. Influence of Reactor Configuration on Fer‐ mentative Hydrogen Production During Wastewater Treatment. International Jour‐

rating Ultrafiltration. Applied Engineering in Agriculture (1996). , 12-709.

UAFB Bioprocess. Bioresource Technology (2007). , 98-3080.

(2002). , 65-25.

98 Biodegradation of Hazardous and Special Products

(2001). , 79-203.

30(12), 193-201.

47-3919.

Science (2003). , 2(4), 307-331.

tem. EcolEngg (2006). , 27(2), 100-108.

ter Science and Technology (1997). , 36-441.

and Applied Chemistry (1997). , 69(11), 2431-2438.

nal of Hydrogen Energy (2009). , 34(8), 3305-3312.

gy (1996). , 55(3), 215-221.


[77] Chavan, M. N, Kulkarni, M. V, Zope, V. P, & Mahulikar, P. P. Microbial Degradation of Melanoidins in Distillery Spent Wash by an Indigenous Isolate. Indian Journal of Biotechnology (2006). , 5-416.

wash Effluent from Biomethanation Plants by White Rot Fungi. Process Biochemistry

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

101

[90] Miranda, P. M, Benito, G. G, Cristobal, N. S, & Nieto, C. H. Color Elimination from Molasses Wastewater by Aspergillus niger. Bioresource Technology (1996). , 57-229.

[91] Angayarkanni, J, Palaniswamy, M, & Sawminathan, K. Biotreatment of Distillery Ef‐ fluent Using Aspergillus niveus. Bulletin of Environmental Contamination and Toxi‐

[92] Shayegan, J, Pazouki, M, & Afshari, A. Continuous Decolorization of Anaerobically

[93] Guimaraes, C. Bento LSM, Mota M. Biodegradation of Colorants in Refinery Efflu‐ ents-Potential Use of the Fungus Phanerochaete chrysosporium. International Sugar

[94] Ohmomo, S, Itoh, N, Watanabe, Y, Kaneko, Y, Tozawa, Y, & Ueda, K. Continuous Decolorization of Molasses Wastewater with Mycelia of Coriolus versicolor Ps4a. Ag‐

[95] Ohmomo, S, Kainuma, M, Kamimura, K, Sirianuntapiboon, S, Oshima, I, & Attha‐ sumpunna, P. Adsorption of Melanoidin to the Mycelia of Aspergillus oryzae Y- Ag‐

[96] Borja, R, Martin, A. M, & Duran, M. M. Enhancement of the Anaerobic Digestion of Wine Distillery Wastewater by the Removal of Phenolics Inhibitors. Bioresource

[97] Sirianuntapiboon, S, Sihanonth, P, Somachai, P, Atthasampunna, P, & Hayashida, S. An Adsorption Mechanism for Melanoidins Decolorization by Rhizoctonia sp. Bio‐

[98] Murata, M, Terasawa, N, & Homma, S. Screening of Microorganisms to Decolorize a Model Melanoidin and the Chemical Properties of a Microbially Treated Melanoidin.

[99] Kida, K, Morimura, S, Abe, N, & Sonoda, Y. Biological Treatment of Shochu Distill‐

[100] Fujita, M, Era, A, Ike, M, Soda, S, Miyata, N, & Hirao, T. Decolorization of Heat-Treatment Liquor of Waste Sludge by a Bioreactor Using Polyurethane Foam-Immo‐ bilized White Rot Fungus Equipped with an Ultramembrane Filtration Unit. Journal

[101] Fumi, M. D, Parodi, G, Parodi, E, & Silva, A. Optimisation of Long Term Activated Sludge Treatment of Winery Wastewater. Bioresource Technology (1995). , 52(1),

Bioscience, Biotechnology, and Biochemistry (1992). , 56(8), 1182-1187.

Digested Distillery Wastewater. Process Biochemistry (2004). , 40-1323.

ricultural and Biological Chemistry (1985b). , 49-2551.

ricultural and Biological Chemistry (1988b). , 2-32.

science, Biotechnology, and Biochemistry (1995). , 59-1185.

ery Wastewater. Process Biochemistry (1995). , 30(2), 125-132.

of Bioscience Bioengineering (2000). , 90(4), 387-94.

(1998). , 33(1), 83-88.

cology (2003). , 70-268.

Journal (1999). , 101-246.

Technology (1993). , 45-99.

45-51.


wash Effluent from Biomethanation Plants by White Rot Fungi. Process Biochemistry (1998). , 33(1), 83-88.

[90] Miranda, P. M, Benito, G. G, Cristobal, N. S, & Nieto, C. H. Color Elimination from Molasses Wastewater by Aspergillus niger. Bioresource Technology (1996). , 57-229.

[77] Chavan, M. N, Kulkarni, M. V, Zope, V. P, & Mahulikar, P. P. Microbial Degradation of Melanoidins in Distillery Spent Wash by an Indigenous Isolate. Indian Journal of

[78] Ghosh, M, Verma, S. C, Mengoni, A, & Tripathi, A. K. Enrichment and Identification of Bacteria Capable of Reducing Chemical Oxygen Demand of Anaerobically Treated

[79] Sirianuntapiboon, S, Phothilangka, P, & Ohmomo, S. Decolorization of Molasses Wastewater by a Strain of Acetogenic Bacteria. Bioresource Technology (2004b).

[80] Cibis, E, Kent, C. A, Krzywonos, M, Garncarek, Z, Garncarek, B, & Miskiewicz, T. Bi‐ odegradation of Potato Slops from a Rural Distillery by Thermophilic Aerobic Bacte‐

[81] Savant, D. V, Shouche, Y. S, Prakash, S, & Ranade, D. R. Methanobrevibacter acidi‐ durans sp. nov a Novel Methanogen from a Sour Anaerobic Digester. International

[82] Ghosh, M, Ganguli, A, & Tripathi, A. K. Treatment of Anaerobically Digested Distill‐ ery Spentwash in a Two-Stage Bioreactor Using Pseudomonas putida and Aeromo‐

[83] Dahiya, J, Singh, D, & Nigam, P. Decolorization of Molasses Wastewater by Cells of Pseudomonas fluorescens Immobilized on Porous Cellulose Carrier. Bioresource

[84] Jain, N, Minocha, A. K, & Verma, C. L. Degradation of Predigested Distillery Effluent by Isolated Bacterial Strains. Indian Journal of Experimental Botany (2002). , 40-101.

[85] Masaharu, S, Hiromi, K, Akihiro, N, Naoshi, F, & Rikiya, T. Screening and Identifica‐ tion of Lactic Acid Bacteria with the Ability to Decolorize Wastewater from a Molass‐ es Alcohol Distillery. Journal of the Brewing Society of Japan (1998). , 93(12), 982-989.

[86] Kambe, T. N, Shimomura, M, Nomura, N, Chanpornpong, T, & Nakahara, T. Decol‐ orization of Molasses Wastewater by Bacillus Spunder Thermophilic and Anaerobic

[87] Sirianuntapiboon, S. Selection of Acid Forming Bacteria Having Decolorization Ac‐ tivity for Removal of Color Substances from Molasses Wastewater. Thammasat Inter‐

[88] Ohmomo, S, Daengsabha, W, Yoshikawa, H, Yui, M, Nozaki, K, Nakajima, T, & Na‐ kamura, I. Screening of Anaerobic Bacteria with the Ability to Decolorize Molasses

[89] Kumar, V, Wati, L, Nigam, P, Banat, I. M, Yadav, B. S, Singh, D, & Marchant, R. De‐ colorization and Biodegradation of Anaerobic Digested Sugarcane Molasses Spent‐

Conditions. Journal of Bioscience and Bioengineering (1999). , 87(1), 119-121.

national Journal of Science and Technology (1999). , 4(2), 1-12.

Melanoidin. Agricultural and Biological Chemistry (1988a). , 57-2429.

Journal of Systematic and Evolutionary Microbiology (2002). , 52-1081.

Molasses Spent Wash. Journal of Applied Microbiology (2004). , 96-1278.

Biotechnology (2006). , 5-416.

100 Biodegradation of Hazardous and Special Products

Technology (2001b). , 78-111.

ria. Bioresource Technology (2002). , 85-57.

nas sp. Process Biochemistry (2002). , 37(8), 857-862.

(BP103), 92-31.


[102] Benito, G. G, Miranda, M. P, & Santos, D. R. Decolorization of Wastewater from an Alcoholic Fermentation Process with Trametes Versicolor. Bioresource Technology (1997). , 61-33.

[115] Valderrama, L. T. Del Campo CM, Rodriguez CM, Bashan LE, Bashan Y. Treatment of Recalcitrant Wastewater from Ethanol and Citric Acid Using the Microalga Chlor‐ ella vulgaris and the Macrophyte Lemna minuscule. Water Research (2002). , 36-4185.

Biodegradation of Melanoidin from Distillery Effluent: Role of Microbes and Their Potential Enzymes

http://dx.doi.org/10.5772/56252

103

[116] Kalavathi, D. F, Uma, L, & Subramanian, G. Degradation and Metabolization of the Pigment-Melanoidin in Distillery Effluent by the Marine Cyanobacterium Oscillato‐

[117] Patel, A, Pawar, P, Mishra, S, & Tewari, A. Exploitation of Marine Cyanobacteria for Removal of Color from Distillery Effluent. Indian Journal of Environmental Protec‐

[118] Moriya, K, Lefuji, H, Shimoi, H, Sato, S, & Tadenuma, M. Treatment of Distillery Wastewater Discharged from Beet Molasses-Spirits Production Using Yeast. Journal

[119] Shojaosadati, S. A, Khalilzadeh, R, Jalilzadeh, A, & Sanaei, H. R. Bioconversion of Molasses Stillage to Protein as an Economic Treatment of this Effluent. Resources,

[120] Karam, J, & Nicell, J. A. Potenital Applications of Enzymes in Waste Treatment. Jour‐

[121] Sangave, P. C, & Pandit, A. B. Enhancement in Biodegradability of Distillery Waste‐ water Using Enzymatic Pretreatment. Journal of Environmental Management

[122] Klibanov, A. M, Alberti, B. N, Morris, E. D, & Felshin, L. M. Enzymatic Removal of Toxic Phenols and Anilines from Wastewaters. Journal of Applied Biochemistry

[123] Klibanov, A. M, & Morris, E. D. Horseradish Peroxidases for the Removal of Carcino‐ genic Aromatic Amines from Water. Enzyme and Microbial Technology (1981). ,

[124] Aitken, M. D, & Irvine, R. L. Stability Testing of Ligninase and Mnperoxidase from Phanerochaete Chrysosporium. Biotechnology and Bioengineering (1989). , 34(10),

[125] Duff, S. J, Moritz, J. W, & Andersen, K. L. Simultaneous Hydrolysis and Fermenta‐ tion of Pulp Mill Primary Clarifier Sludge. Canadian Journal of Chemical Engineer‐

[126] Ferrer, I, Dezotti, M, & Duran, N. Decolorization of Kraft Effluent by Free and Immo‐ bilized Lignin Peroxidases and Horse Radish Peroxidases. Biotechnology Letters

[127] Dec, J, & Bollag, J. M. Use of Plant Material for the Decontamination of Water Pollut‐

ed with Phenols. Biotechnology and Bioengineering (1994). , 44-1132.

nal of Chemical Technology and Biotechnology (1997). , 69-141.

of Fermentation and Bioengineering (1990). , 69-138.

Conservation and Recycling (1999).

ria boryana BDU 92181. Enzyme and Microbial Technology (2001). , 29-246.

tion (2001). , 21-1118.

(2006a). , 78-77.

(1980). , 2-414.

3-119.

1251-1260.

ing (1994). , 72-1013.

(1991). , 13-577.


[115] Valderrama, L. T. Del Campo CM, Rodriguez CM, Bashan LE, Bashan Y. Treatment of Recalcitrant Wastewater from Ethanol and Citric Acid Using the Microalga Chlor‐ ella vulgaris and the Macrophyte Lemna minuscule. Water Research (2002). , 36-4185.

[102] Benito, G. G, Miranda, M. P, & Santos, D. R. Decolorization of Wastewater from an Alcoholic Fermentation Process with Trametes Versicolor. Bioresource Technology

[103] Sirianuntapiboon, S, & Chairattanawan, K. Some Properties of Coriolus sp. No.20 for Removal of Color Substances from Molasses Wastewater. Thammasat International

[104] Miyata, N, Mori, T, Iwahori, K, & Fujita, M. Microbial Decolorization of Melanoidin-Containing Wastewaters: Combined Use of Activated Sludge and the Fungus Corio‐

lus hirsutus. Journal of Bioscience and Bioengineering (2000). , 89(2), 145-150.

[105] Dahiya, J, Singh, D, & Nigam, P. Decolorization of Synthetic and Spentwash Mela‐ noidins Using the White-Rot Fungus Phanerochaete chrysosporium JAG-40. Biore‐

[106] Malandra, L, Wolfaardt, G, Zietsman, A, & Viljoen-bloom, M. Microbiology of a Bio‐ logical Contractor for Winery Wastewater Treatment. Water Research (2003). ,

[107] Kwak, E. J, Lee, Y. S, Murata, M, & Homma, S. Effect of Reaction pH on the Photode‐ gradation of Model Melanoidins. Lebensmittel-Wissenschaft und-Technologie

[108] Raghukumar, C, Mohandass, C, Kamat, S, & Shailaja, M. S. Simultaneous Detoxifica‐ tion and Decolorization of Molasses Spent Wash by the Immobilized White-Rot Fun‐ gus Flavodon flavus Isolated from a Marine Habitat. Enzyme and Microbial

[109] Raghukumar, C, & Rivonkar, G. Decolorization of Molasses Spent Wash by the White-Rot Fungus Flavodon flavus Isolated from a Marine Habitat. Applied Microbi‐

[110] Potentini, M. F, & Rodríguez-malaver, A. J. Vinasse Biodegradation by Phanero‐ chaete chrysosporium. Journal of Environmental Biology (2006). , 27(4), 661-665.

[111] Thakkar, A. P, Dhamankar, V. S, & Kapadnis, B. P. Biocatalytic Decolorization of Mo‐ lasses by Phanerochaete chrysosporium. Bioresource Technology (2006). , 97-1377.

[112] Sirianuntapiboon, S, Somachi, P, & Sihanonth, P. Ohmomo PAS. Microbial Decolori‐ zation of Molasses Wastewater by Mycelia Sterilia D90. Agricultural and Biological

[113] Seyis, I, & Subasioglu, T. Screening of Different Fungi for Decolorization of Molasses.

[114] Kaushik, G, & Thakur, I. S. Isolation of Fungi and Optimization of Process Parame‐ ters for Decolorization of Distillery Mill Effluent. World Journal of Microbiology and

Journal of Science and Technology (1998). , 3(2), 74-79.

source Technology (2001a). , 78-95.

(1997). , 61-33.

102 Biodegradation of Hazardous and Special Products

37-4125.

(2004). , 37-255.

Technology (2004). , 35-197.

ology and Biotechnology (2001). , 55-510.

Chemistry (1988). , 52(2), 393-398.

Biotechnology (2009). , 25-955.

Brazilian Journal of Microbiology (2009). , 40-61.


[128] Watanabe, Y, Sugi, R, & Tanaka, Y. Enzymatic Decolorization of Melanoidin by Cor‐ iolus sp. No 20. Agricultural and Biological Chemistry (1982). , 46(6), 1623-1630.

**Chapter 6**

**Degradability: Enzymatic and in Simulated Compost Soil**

**of PLLA:PCL Blend and on Their Composite with Coconut**

The problem of non-biodegradable plastic waste remains a challenge due to its negative environmental impact. In this sense, poly(L-lactic acid), PLLA, and poly(ε-caprolactone), PCL, have been receiving much attention lately due to their biodegradability in human body as well as in the soil, biocompatibility, environmentally friendly characteristics and non-toxicity [1-5]. PLLA is a poly(α-hydroxy acid) and PCL is a poly(ω-hydroxy acid) [1]. PLLA is a hard, transparent and crystalline polymer. On the other hand, PCL can be used as a polymeric plasticizer because of its ability to lower elastic modulus and to soften other polymers [6]. The original reasons for preparing polymer blends are to reduce costs by combining high-quality polymers with cheaper materials (although this approach is usually accompanied by a drastic worsening of the properties of the polymer) and to create a polymer that has a desired combination of the different properties of its components. However, according to Michler [7] usually different polymers are incompatible. Improved properties can be only realized if the blend exhibits optimum morphology. According to Sawyer et al. [8], in polymer science, the term morphology generally refers to form and organization on a size scale above the atomic arrangement, but smaller than the size and shape of the whole sample. Thus, improving compatibility between the different polymers and optimizing the morphology are the main issues to address when producing polymer blends [3]. Moreover, both polymers PLLA and PCL can be used in biomedical applications, which require a proper sterilization process. Nowadays, the most suitable sterilization method is high energy irradiation. However, it is important to remind that polymeric structural changes are induced by radiation processing of polymers, such as scission and crosslinking [9-12]. According to the principles of radiation chemistry, very reactive intermediate, free radicals, ions and excited states are formed when

> © 2013 Kodama; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

**Fiber**

Yasko Kodama

**1. Introduction**

http://dx.doi.org/10.5772/56231

Additional information is available at the end of the chapter


**Chapter 6**

### **Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with Coconut Fiber**

Yasko Kodama

[128] Watanabe, Y, Sugi, R, & Tanaka, Y. Enzymatic Decolorization of Melanoidin by Cor‐ iolus sp. No 20. Agricultural and Biological Chemistry (1982). , 46(6), 1623-1630. [129] Ohmomo, S, Aoshima, I, Tozawa, Y, Sakurada, N, & Ueda, K. Purification and Some Properties of Melanoidin Decolorizing Enzymes and P-4 from Mycelia of Coriolus

[130] Arora, D. S, Chander, M, & Gill, P. K. Involvement of Lignin Peroxidase, Manganese Peroxidase and Laccase in Degradation and Selective Ligninolysis of Wheat Straw.

[131] Rubia TDLLinares A, Perez J, Dorado JM, Romera J, Martinez J. Characterization of Manganese-Dependent Peroxidase Isoenzymes from the Ligninilytic Fungus Phaner‐

[132] Mansur, M, Suarez, T, & Fernandez-larrea, J. B. Brizuela MAA, Gonzalezi AE. Identi‐ fication of a Laccase Gene Family in the New Lignin-Degrading Basidiomycetes CECT 20197. Applied and Environmental Microbiology (1997). , 63(7), 2637-2646. [133] Dehorter, B, & Blondeau, R. Isolation of an Extracellular Mn-Dependent Enzyme Mineralizing Melanoidins from the White Rot Fungus Trametes versicolor. FEMS

[134] Lee, T. H, Aoki, H, Sugano, Y, & Shoda, M. Effect of Molasses on the Production and Activity of Dye-Decolorizing Peroxidase from Geotrichum candidum. Journal of Bio‐

[135] Souza, D, Tiwari, D. T, Sah, R, & Raghukumar, A. K. C. Enhanced Production of Lac‐ case by a Marine Fungus During Treatment of Colored Effluents and Synthetic Dyes.

[136] Sangave, P. C, & Pandit, A. B. Ultrasound and Enzyme Assisted Biodegradation of Distillery Wastewater. Journal of Environmental Management (2006b). , 80-36. [137] Yadav, S, Chandra, R, & Rai, V. Characterization of Potential MnP Producing Bacte‐ ria and its Metabolic Products During Decolorization of Synthetic Melanoidins due to Biostimulatory Effect of D-Xylose at Stationary Phase. Process Biochemistry

[138] Raghukumar, C, Souza, D, Thorn, T. M, Reddy, R. G, & Lignin-modifying, C. A. En‐ zymes of Flavodon flavus a Basidiomycete isolated from a Coastal Marine Environ‐

[139] Raghukumar, C. Fungi from Marine Habitat: Application in Bioremediation. Myco‐

ment. Applied Environmental Microbiology (1999). , 65-2103.

versicolor Ps4a. Agricultural and Biological Chemistry (1985a). , 3.

International Biodeterioration and Biodegradation (2002). , 50-115.

ochaete flavido-alba. Research in Microbiology (2002). , 153-547.

Microbiology Letters (1993). , 109(1), 117-122.

science and Bioengineering (2000). , 89-545.

(2011). , 86-1774.

104 Biodegradation of Hazardous and Special Products

logical Research (2000). , 104-1222.

Enzyme and Microbial Technology (2006). , 38-504.

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56231

#### **1. Introduction**

The problem of non-biodegradable plastic waste remains a challenge due to its negative environmental impact. In this sense, poly(L-lactic acid), PLLA, and poly(ε-caprolactone), PCL, have been receiving much attention lately due to their biodegradability in human body as well as in the soil, biocompatibility, environmentally friendly characteristics and non-toxicity [1-5]. PLLA is a poly(α-hydroxy acid) and PCL is a poly(ω-hydroxy acid) [1]. PLLA is a hard, transparent and crystalline polymer. On the other hand, PCL can be used as a polymeric plasticizer because of its ability to lower elastic modulus and to soften other polymers [6]. The original reasons for preparing polymer blends are to reduce costs by combining high-quality polymers with cheaper materials (although this approach is usually accompanied by a drastic worsening of the properties of the polymer) and to create a polymer that has a desired combination of the different properties of its components. However, according to Michler [7] usually different polymers are incompatible. Improved properties can be only realized if the blend exhibits optimum morphology. According to Sawyer et al. [8], in polymer science, the term morphology generally refers to form and organization on a size scale above the atomic arrangement, but smaller than the size and shape of the whole sample. Thus, improving compatibility between the different polymers and optimizing the morphology are the main issues to address when producing polymer blends [3]. Moreover, both polymers PLLA and PCL can be used in biomedical applications, which require a proper sterilization process. Nowadays, the most suitable sterilization method is high energy irradiation. However, it is important to remind that polymeric structural changes are induced by radiation processing of polymers, such as scission and crosslinking [9-12]. According to the principles of radiation chemistry, very reactive intermediate, free radicals, ions and excited states are formed when

© 2013 Kodama; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

macromolecules of polymers are submitted to ionizing radiation, where they are then free to react with one another or initiate further reactions among the polymeric chains, thus giving rise to changes in material properties. These intermediates can follow several reactions paths that result in disproportion, hydrogen abstraction, arrangements and/or formation of new bonds. The combination of two radicals leads to cross-linking or recombination in the amor‐ phous and crystalline regions, respectively, whereas chain transfer and the subsequent splitting results in chain scission. Usually both these processes take place simultaneously for many polymers [10,11].

fibers would be obtained from vegetal fibers. So, microorganisms are able to consume completely those materials, eventually releasing carbon dioxide and water as by-products [20]. PCL, PLLA and coconut fiber composites studied in this chapter may be categorized in that

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

107

According to Müller [13], there are different approaches concerning the type of test to be applied to evaluate degradation of polymeric materials in the environment and, also, what conclusion can be obtained from that. As principle, tests can be divided in three categories, field test, simulation and laboratorial tests. Nevertheless field test, for instance, in which samples are buried on the ground, or putting them in a lake or river, or performing general process of composting of polymeric biodegradable material, represent the ideal practical conditions. There are several disadvantages associated to this kind of test. One of the problems would be to control environment conditions like temperature, pH, or humidity. Another point to be considered is to analytically monitor the degradation process, in most cases it would be possible to visually evaluate alterations of the sample, or maybe evaluate the disintegration by measuring weight loss. Most reproducible tests are laboratorial ones, well-defined medium and, inoculated with specific microorganisms to a particular polymer are utilized. In those cases, enzymatic activity is optimized to a particular microorganism and, frequently present more elevated degradation rate than the ones observed in natural conditions. This is consid‐ ered as an advantage to the study of basic mechanism of polymer biodegradation. Although results lead to limited conclusion related to real degradation rate on the natural environment,

Samples of PCL and PLLA homopolymers; PCL:PLLA 20:80 (w:w) blend; and composites of the blend containing 5% and 10% of coconut fiber (chemically untreated and acetylated) were

Coir coconut fibers for composite preparation were kindly provided by Embrapa – Paraipaba

Size reduction of the coconut fibers was carried out using helix mill Marconi – modelo MA 680, from Laboratório de Matéria-prima Particulados e Sólidos Não Metálicos – LMPSol,

The fiber size distribution was measured using sieves of the Tyler series 16, 20, 35 and 48, fiber sizes of 1.0mm, 0.84mm, 0.417mm, and 0.297mm, respectively. The 0.297-0.417mm fibers size was used for the assays. The triturated material was separated using a sieve shaker Produtest,

Departamento de Engenharia de Materiais of Escola Politécnica/USP.

class.

those tests have widely been used.

**2. Material and method**

**2.1. Material**

prepared in triplicate.

**2.2. Coconut fiber**

region, Ceará.

for 1 min.

The morphology of the blends affects the thermo mechanical properties as well as the biode‐ gradation of the polymers. In particular, surface structure and morphology of the biodegrad‐ able polymer blends have a great impact on the enzymatic degradation behavior. The development of polymeric materials susceptible to microbiological degradation and that have similar performance to conventional polymers has been intensely studied. The intention would be that those materials reduce waste volume while suffer degradation in sanitary waste deposit, or they could be treated in composting plants [13]. Enzymatic and non-enzymatic degradations occur easier in the amorphous region [14,15]. Kikkawa et al. [16] cited that one of the approaches used to generate biodegradable materials with a wide range of physical properties is blending, and miscibility of blends is one of the most important factors affecting the final polymer properties.

Nishino et al. [17] cited that cellulose is the most abundant form of biomass and the form most likely to be used as reinforcement fibers, not only because of ecological and economic reasons, but also because of their high mechanical and thermal performance. Thus, incorporating fibers of low cost to the polymeric blend, it is possible to obtain an improvement of the mechanical properties without loss of the original characteristics of polymeric components. Regarding the irradiation effects, vegetable fiber, like as coconut fiber, is composed by cellulose and lignin, which suffer chemical alteration by irradiation such as scission or cross-linking. In the case of natural polymers, such as cellulose, main chain scission occurs predominantly due to irradi‐ ation and as a result molecular weight' decrease [10].

Liu et al. and Lenglet et al. [18,19] cited that biodegradability of PCL and PLLA has also been investigated under environmental conditions. The controlled degradation of polymers is sometimes desired for biomedical applications, besides the environmental purposes [7]. It has been seen that PLLA is bio absorbable, that is, the hydrolytic degradation by-products formed can be fully assimilated by microorganisms such as fungi or bacteria. On the other hand, PCL is promptly biodegraded by environmental microorganisms. So, both PCL and PLLA can be considered as environmentally friendly polymers.

Kolybaba et al. [20] mentioned that biodegradable plastics are those that undergo significant enough modification on their chemical structure under specific environmental condition. Those changes result on mechanical and physical properties losses that are measurable by standard methods of testing. Biodegradable plastics suffer degradation under action of microorganisms that has natural occurrence, for instance, bacteria, fungi and algae. The plastic engineered to be entirely biodegradable is classified within the main classes of polymeric materials. In this category, polymeric matrix can be from natural resources and reinforcement fibers would be obtained from vegetal fibers. So, microorganisms are able to consume completely those materials, eventually releasing carbon dioxide and water as by-products [20]. PCL, PLLA and coconut fiber composites studied in this chapter may be categorized in that class.

According to Müller [13], there are different approaches concerning the type of test to be applied to evaluate degradation of polymeric materials in the environment and, also, what conclusion can be obtained from that. As principle, tests can be divided in three categories, field test, simulation and laboratorial tests. Nevertheless field test, for instance, in which samples are buried on the ground, or putting them in a lake or river, or performing general process of composting of polymeric biodegradable material, represent the ideal practical conditions. There are several disadvantages associated to this kind of test. One of the problems would be to control environment conditions like temperature, pH, or humidity. Another point to be considered is to analytically monitor the degradation process, in most cases it would be possible to visually evaluate alterations of the sample, or maybe evaluate the disintegration by measuring weight loss. Most reproducible tests are laboratorial ones, well-defined medium and, inoculated with specific microorganisms to a particular polymer are utilized. In those cases, enzymatic activity is optimized to a particular microorganism and, frequently present more elevated degradation rate than the ones observed in natural conditions. This is consid‐ ered as an advantage to the study of basic mechanism of polymer biodegradation. Although results lead to limited conclusion related to real degradation rate on the natural environment, those tests have widely been used.

#### **2. Material and method**

#### **2.1. Material**

macromolecules of polymers are submitted to ionizing radiation, where they are then free to react with one another or initiate further reactions among the polymeric chains, thus giving rise to changes in material properties. These intermediates can follow several reactions paths that result in disproportion, hydrogen abstraction, arrangements and/or formation of new bonds. The combination of two radicals leads to cross-linking or recombination in the amor‐ phous and crystalline regions, respectively, whereas chain transfer and the subsequent splitting results in chain scission. Usually both these processes take place simultaneously for

The morphology of the blends affects the thermo mechanical properties as well as the biode‐ gradation of the polymers. In particular, surface structure and morphology of the biodegrad‐ able polymer blends have a great impact on the enzymatic degradation behavior. The development of polymeric materials susceptible to microbiological degradation and that have similar performance to conventional polymers has been intensely studied. The intention would be that those materials reduce waste volume while suffer degradation in sanitary waste deposit, or they could be treated in composting plants [13]. Enzymatic and non-enzymatic degradations occur easier in the amorphous region [14,15]. Kikkawa et al. [16] cited that one of the approaches used to generate biodegradable materials with a wide range of physical properties is blending, and miscibility of blends is one of the most important factors affecting

Nishino et al. [17] cited that cellulose is the most abundant form of biomass and the form most likely to be used as reinforcement fibers, not only because of ecological and economic reasons, but also because of their high mechanical and thermal performance. Thus, incorporating fibers of low cost to the polymeric blend, it is possible to obtain an improvement of the mechanical properties without loss of the original characteristics of polymeric components. Regarding the irradiation effects, vegetable fiber, like as coconut fiber, is composed by cellulose and lignin, which suffer chemical alteration by irradiation such as scission or cross-linking. In the case of natural polymers, such as cellulose, main chain scission occurs predominantly due to irradi‐

Liu et al. and Lenglet et al. [18,19] cited that biodegradability of PCL and PLLA has also been investigated under environmental conditions. The controlled degradation of polymers is sometimes desired for biomedical applications, besides the environmental purposes [7]. It has been seen that PLLA is bio absorbable, that is, the hydrolytic degradation by-products formed can be fully assimilated by microorganisms such as fungi or bacteria. On the other hand, PCL is promptly biodegraded by environmental microorganisms. So, both PCL and PLLA can be

Kolybaba et al. [20] mentioned that biodegradable plastics are those that undergo significant enough modification on their chemical structure under specific environmental condition. Those changes result on mechanical and physical properties losses that are measurable by standard methods of testing. Biodegradable plastics suffer degradation under action of microorganisms that has natural occurrence, for instance, bacteria, fungi and algae. The plastic engineered to be entirely biodegradable is classified within the main classes of polymeric materials. In this category, polymeric matrix can be from natural resources and reinforcement

many polymers [10,11].

106 Biodegradation of Hazardous and Special Products

the final polymer properties.

ation and as a result molecular weight' decrease [10].

considered as environmentally friendly polymers.

Samples of PCL and PLLA homopolymers; PCL:PLLA 20:80 (w:w) blend; and composites of the blend containing 5% and 10% of coconut fiber (chemically untreated and acetylated) were prepared in triplicate.

#### **2.2. Coconut fiber**

Coir coconut fibers for composite preparation were kindly provided by Embrapa – Paraipaba region, Ceará.

Size reduction of the coconut fibers was carried out using helix mill Marconi – modelo MA 680, from Laboratório de Matéria-prima Particulados e Sólidos Não Metálicos – LMPSol, Departamento de Engenharia de Materiais of Escola Politécnica/USP.

The fiber size distribution was measured using sieves of the Tyler series 16, 20, 35 and 48, fiber sizes of 1.0mm, 0.84mm, 0.417mm, and 0.297mm, respectively. The 0.297-0.417mm fibers size was used for the assays. The triturated material was separated using a sieve shaker Produtest, for 1 min.

In order to remove lignin from coconut fiber surface, fibers were soaked with Na2SO3 2% aqueous solution for 2h using ultrasound. Coconut fibers were washed several times with tap water and finally, tree times with deionized water, as described in the literature [21].

**3. Method**

without enzyme.

hours.

**3.1. Enzymatic degradation**

**3.2. Biodegradability in soil**

**4. Results and discussion**

**4.1. Enzymatic degradation**

A buffer solution with phosphate, pH 7, and lipase enzyme obtained from *Pseudomonas cepacia*, of Aldrich, was prepared. Solution concentrations were kept at 35 unities of enzymatic

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

preserved by 7 days (168h). Samples were exposed to enzymatic action for 0, 24, 72, 120 and 168 hours. After enzymatic exposure, samples were washed with water, dried and weighted (mass retention determination). Tests were performed in duplicate, subtracted test control

Samples of approximately 10mm × 10mm were buried in plastic trays containing simulated compost soil previously prepared, with 23 % humus, 23 % organic material (tree leaves, coffee powder, food waste and cattle manure), 23 % sand and distilled water to complete 100 %. Simulated compost was characterized for nitrogen and total carbon content, ABNT 1167 and, pH. Simulated compost soil characterization results: pH 7.8; humidity 30 ± 10 %; total carbon 18.4 %; total nitrogen 0.83 %. Samples were removed from the soil at 30, 60, 90 e 120 days of ageing. After that, they were mechanically cleaned, and dried at room temperature for 24

Enzymatic degradation was performed using lipase enzyme obtained from *Pseudomonas cepacia*. In Fig. 1 it can be observed mass retention variation of non-irradiated and electron beam (EB) irradiated samples through time of degradation of homopolymers, blend and composites. Degradation rate of PCL is higher than PLLA in *Pseudomonas* lipase presence, in agreement with observed by Liu et al. [18] that lipase degrade both crystalline and amorphous PCL. According to Liu et al., enzymatic degradation of PCL has been investigated, mainly in presence of lipase enzyme. It is well known that morphology and its alteration plays an important role on hydrolytic degradability of aliphatic polyesters [18]. When the subject is enzymatic degradation, situation complicates due to specificity of enzymes. *Pseudomonas* lipase is able to break esters linkages in hydrophobic substrates, as it is PCL case. Also, it was described that PCL did not absorb water, by the other side, PLLA absorbed 2% water within 72h. The authors also informed that degradation rate of this polymer is higher in proteinase K than in lipase (8% against 1%). However, in the study of this chapter it was observed that PCL degraded approximately 30%, and PLLA 16%, at the same period of time. According to Tsuji and Ishizaka, no alteration was observed on the molar weight distribution, either mass loss of pure PLLA studied films, indicating that enzymatic hydrolysis effect caused by *Pseudomonas*

C. System buffer-enzyme was

http://dx.doi.org/10.5772/56231

109

activity. Flasks were maintained in hot water bath at 37o

Coconut fiber acetylation was performed as described by d'Almeida et al. [22]. As received fibers from Embrapa were soaked in a solution of acetic anhydrate and acetic acid (1.5:1.0, w:w). It was used as a catalyst, 20 drops of sulfuric acid in 500mL solution. Those groups of sets were submitted to ultrasound for 3h, then for more 24h rest at the same solution. Fibers were washed using tap water and for more 24h rested in deionized water. Fibers were separated from water and washed with acetone, after that, were evaporated at room temper‐ ature.

#### **2.3. Preparation of composite pellets and sheets**

PCL (pellets, *<sup>M</sup>*¯ *<sup>w</sup>* =2.14∙10<sup>5</sup> g∙mol-1; *M*¯ *w* / *M*¯ *n*= 1.423), PLLA (pellets, *<sup>M</sup>*¯ *w*=2.64∙10<sup>5</sup> g∙mol-1 *M*¯ *w* / *M*¯ *<sup>n</sup>*= 1.518 – Gel Permeation Chromatographic values) and dried coconut fiber (from Embrapa – Empresa Brasileira de Pesquisa Agropecuária, Ceará, Brazil) were used to prepare blends and composites. A Labo Plastomil model 50C 150 of Toyoseiki twin screw extruder was used for pellets preparation. Pellets of PCL:PLLA 20:80 (w:w) blend and composites containing 5 and 10% of untreated and chemically treated coconut fiber were prepared at AIST.

Sheets (150mm x 150mm x 0.5mm) of PCL, PLLA, PCL:PLLA 20:80 (w:w) blend and composites containing 5 and 10% untreated and chemically treated coconut fiber were prepared using Ikeda hot press equipment of Japan Atomic Energy Agency, JAEA. Mixed pellets of samples were preheated at 195°C for 3 min and then pressed by under heating at the same temperature for another 3 min under pressure of 150 kgf∙ cm-2. Samples sheets were then cooled in the cold press using water as a coolant for 3 min.

For degradability tests, samples were taken from hot compressed polymeric sheets, cut into 15mm × 15mm pieces. Non-irradiated and, electron beam (EB), irradiated samples with absorbed doses of 50 kGy and 100 kGy were studied.

#### **2.4. Electron beam irradiation**

Irradiation was performed at JAEA using electron beam accelerator (2 MeV; 2 mA), absorbed doses of 50 and 100 kGy, dose rate of 0.6 kGy s-1. The energy and current parameters condition of irradiation were enough to the electron beam goes through the 0.5mm thickness sheets.

Absorbed dose is the amount of energy absorbed per unit mass of irradiated material. The SI unit for absorbed dose is joules per kilogram (J kg-1), which is given the special name gray (symbol, Gy). The absorbed dose rate is the absorbed dose per unit time and has the units gray per unit time, for instance kGy s-1. The absorbed dose is a direct measure of the energy transferred to the irradiated material that is capable of producing chemical or physical change [23].

#### **3. Method**

In order to remove lignin from coconut fiber surface, fibers were soaked with Na2SO3 2% aqueous solution for 2h using ultrasound. Coconut fibers were washed several times with tap

Coconut fiber acetylation was performed as described by d'Almeida et al. [22]. As received fibers from Embrapa were soaked in a solution of acetic anhydrate and acetic acid (1.5:1.0, w:w). It was used as a catalyst, 20 drops of sulfuric acid in 500mL solution. Those groups of sets were submitted to ultrasound for 3h, then for more 24h rest at the same solution. Fibers were washed using tap water and for more 24h rested in deionized water. Fibers were separated from water and washed with acetone, after that, were evaporated at room temper‐

water and finally, tree times with deionized water, as described in the literature [21].

*w* / *M*¯

5 and 10% of untreated and chemically treated coconut fiber were prepared at AIST.

*<sup>n</sup>*= 1.518 – Gel Permeation Chromatographic values) and dried coconut fiber (from

Embrapa – Empresa Brasileira de Pesquisa Agropecuária, Ceará, Brazil) were used to prepare blends and composites. A Labo Plastomil model 50C 150 of Toyoseiki twin screw extruder was used for pellets preparation. Pellets of PCL:PLLA 20:80 (w:w) blend and composites containing

Sheets (150mm x 150mm x 0.5mm) of PCL, PLLA, PCL:PLLA 20:80 (w:w) blend and composites containing 5 and 10% untreated and chemically treated coconut fiber were prepared using Ikeda hot press equipment of Japan Atomic Energy Agency, JAEA. Mixed pellets of samples were preheated at 195°C for 3 min and then pressed by under heating at the same temperature for another 3 min under pressure of 150 kgf∙ cm-2. Samples sheets were then cooled in the cold

For degradability tests, samples were taken from hot compressed polymeric sheets, cut into 15mm × 15mm pieces. Non-irradiated and, electron beam (EB), irradiated samples with

Irradiation was performed at JAEA using electron beam accelerator (2 MeV; 2 mA), absorbed doses of 50 and 100 kGy, dose rate of 0.6 kGy s-1. The energy and current parameters condition of irradiation were enough to the electron beam goes through the 0.5mm thickness sheets.

Absorbed dose is the amount of energy absorbed per unit mass of irradiated material. The SI unit for absorbed dose is joules per kilogram (J kg-1), which is given the special name gray (symbol, Gy). The absorbed dose rate is the absorbed dose per unit time and has the units gray per unit time, for instance kGy s-1. The absorbed dose is a direct measure of the energy transferred to the irradiated material that is capable of producing chemical or physical change

*n*= 1.423), PLLA (pellets, *<sup>M</sup>*¯

*w*=2.64∙10<sup>5</sup> g∙mol-1

ature.

*M*¯ *w* / *M*¯

[23].

PCL (pellets, *<sup>M</sup>*¯

**2.3. Preparation of composite pellets and sheets**

108 Biodegradation of Hazardous and Special Products

press using water as a coolant for 3 min.

**2.4. Electron beam irradiation**

absorbed doses of 50 kGy and 100 kGy were studied.

*<sup>w</sup>* =2.14∙10<sup>5</sup> g∙mol-1; *M*¯

#### **3.1. Enzymatic degradation**

A buffer solution with phosphate, pH 7, and lipase enzyme obtained from *Pseudomonas cepacia*, of Aldrich, was prepared. Solution concentrations were kept at 35 unities of enzymatic activity. Flasks were maintained in hot water bath at 37o C. System buffer-enzyme was preserved by 7 days (168h). Samples were exposed to enzymatic action for 0, 24, 72, 120 and 168 hours. After enzymatic exposure, samples were washed with water, dried and weighted (mass retention determination). Tests were performed in duplicate, subtracted test control without enzyme.

#### **3.2. Biodegradability in soil**

Samples of approximately 10mm × 10mm were buried in plastic trays containing simulated compost soil previously prepared, with 23 % humus, 23 % organic material (tree leaves, coffee powder, food waste and cattle manure), 23 % sand and distilled water to complete 100 %. Simulated compost was characterized for nitrogen and total carbon content, ABNT 1167 and, pH. Simulated compost soil characterization results: pH 7.8; humidity 30 ± 10 %; total carbon 18.4 %; total nitrogen 0.83 %. Samples were removed from the soil at 30, 60, 90 e 120 days of ageing. After that, they were mechanically cleaned, and dried at room temperature for 24 hours.

#### **4. Results and discussion**

#### **4.1. Enzymatic degradation**

Enzymatic degradation was performed using lipase enzyme obtained from *Pseudomonas cepacia*. In Fig. 1 it can be observed mass retention variation of non-irradiated and electron beam (EB) irradiated samples through time of degradation of homopolymers, blend and composites. Degradation rate of PCL is higher than PLLA in *Pseudomonas* lipase presence, in agreement with observed by Liu et al. [18] that lipase degrade both crystalline and amorphous PCL. According to Liu et al., enzymatic degradation of PCL has been investigated, mainly in presence of lipase enzyme. It is well known that morphology and its alteration plays an important role on hydrolytic degradability of aliphatic polyesters [18]. When the subject is enzymatic degradation, situation complicates due to specificity of enzymes. *Pseudomonas* lipase is able to break esters linkages in hydrophobic substrates, as it is PCL case. Also, it was described that PCL did not absorb water, by the other side, PLLA absorbed 2% water within 72h. The authors also informed that degradation rate of this polymer is higher in proteinase K than in lipase (8% against 1%). However, in the study of this chapter it was observed that PCL degraded approximately 30%, and PLLA 16%, at the same period of time. According to Tsuji and Ishizaka, no alteration was observed on the molar weight distribution, either mass loss of pure PLLA studied films, indicating that enzymatic hydrolysis effect caused by *Pseudomonas* lipase on the main chain of PLLA on the bulk was not significant. This confirms that enzymatic degradation occurs preferably on the surface of the sample [15].

of PCL degradation on the blends would be the disturbance caused by superficial adsorption of enzyme molecules on the polymeric films or by slow hydrolytic scission of main chains of

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

111

**Figure 2.** Points of enzyme attack on polyesters by lipase: a) PLA; b) PCL; c) PCL:PLA 14:86 (extracted from Sivalingam

On Fig. 3 it is possible to observe effect of radiation dose on enzymatic degradation of PCL

According to Cottam et al. [26], degradability rate of PCL irradiated with 25 kGy decreased, attributed to irradiation process. Authors cited that lipase catalyzes hydrolysis of carbonyl group linkage and one oxygen atom in the case of fat. It is the same linkage that is broken during PCL hydrolysis. They attributed that degradability rate of PCL was affected by crosslinking occurred due to irradiation. In this study, PCL irradiated with 50 kGy suffered a slight decrease on degradation rate, in agreement with authors' observation. However, PCL samples irradiated with 100 kGy presented a certain increase on degradation rate. This fact probably is related to crystallinity decrease of around 6% observed by Differential Scanning Calorimetry, DSC, of irradiated samples. On Fig. 4 it is possible to observe the effect of radiation

According to Maharana et al. [27], enzymatic degradation occurs only on the surface of a solid substrate by erosion on the surface and by weight loss, because enzymes cannot penetrate a solid polymeric substrate. Enzymes degrade selectively amorphous regions or less ordered that allows them to diffuse through substrate and, subsequently, crystalline regions are eventually degrad‐ ed. In this process, molar weight and molar weight distribution of non-degraded solid sub‐ strate do not change during enzymatic degradation because only the polymer on the surface of substrate is degraded and products of low molar weight from degradation are removed of

dose on enzymatic degradation of PLLA samples irradiated with electron beam.

substrate by solubilization on the surrounding aqueous medium.

et al. [25])

samples irradiated with electron beam.

PCL by molecules enzymes on the presence of PLLA molecules on the blends.

Fig. 2 shows points of lipase enzyme attack on polyesters proposed by [25].

Calil et al. and Sivalingam et al. [24,25] cited that the presence of one polymer affects degrad‐ ability of the other polymer. Lenglet et al. observed that PLLA addition to PCL reduced drastically degradation of PCL of the blends in lipase presence [19]. In the study presented in this chapter, it was possible to observe that the presence of PLLA reduced enzymatic degra‐ dation of PCL of PCL:PLLA 20:80 (w:w) blend, and after 120 hours, mass retention variation moved toward of pure PLLA behavior. Tsuji and Ishizaka [15] studied enzymatic degradation of PCL:PLLA blends using *Rhizopus arrhizus* lipase. They observed that enzymes obtained from fungi cause selective hydrolysis and PCL removal of PCL:PLLA blends without significant PLLA degradation in soil. They also cited that enzymatic degradation of PCL:PLLA in presence of *Pseudomonas* lipase and Proteinase K occurred in the interface of two polymeric phases both on the bulk and on the surface of the sample.

**Figure 1.** Mass retention variation versus degradation time, enzymatic method, of samples: (■) PCL; (●) PLLA; (▴) PCL:PLLA 20:80 (w:w); (□) composite with 5% of untreated fiber; (◇) composite with 10% of untreated fiber; (◁) com‐ posite with 5% of acetylated fiber; and (○) composite with 10% of acetylated fiber, of non-irradiated samples.

In the study of this chapter, composites degraded in a way similar of the blend through time. Mass retention values observed were higher than the blends during the same period of time of test, suggesting that coconut fibers did not significantly enzymatic degraded in this test condition. Furthermore, acetylation did not affect enzymatic degradation of composites significantly.

Tsuji and Ishizaka [15] observed that crystallinity of PCL on blends films did not change with composition variation during degradation, suggesting that this property did not affect enzymatic hydrolysis rate of PCL of the blend as it did not altered during process. Rate of enzymatic hydrolysis of blends was lower than pure PCL, suggesting that PLLA interfered on PCL hydrolysis catalyzed by lipase. One reason postulated by the authors to the deceleration of PCL degradation on the blends would be the disturbance caused by superficial adsorption of enzyme molecules on the polymeric films or by slow hydrolytic scission of main chains of PCL by molecules enzymes on the presence of PLLA molecules on the blends.

Fig. 2 shows points of lipase enzyme attack on polyesters proposed by [25].

lipase on the main chain of PLLA on the bulk was not significant. This confirms that enzymatic

Calil et al. and Sivalingam et al. [24,25] cited that the presence of one polymer affects degrad‐ ability of the other polymer. Lenglet et al. observed that PLLA addition to PCL reduced drastically degradation of PCL of the blends in lipase presence [19]. In the study presented in this chapter, it was possible to observe that the presence of PLLA reduced enzymatic degra‐ dation of PCL of PCL:PLLA 20:80 (w:w) blend, and after 120 hours, mass retention variation moved toward of pure PLLA behavior. Tsuji and Ishizaka [15] studied enzymatic degradation of PCL:PLLA blends using *Rhizopus arrhizus* lipase. They observed that enzymes obtained from fungi cause selective hydrolysis and PCL removal of PCL:PLLA blends without significant PLLA degradation in soil. They also cited that enzymatic degradation of PCL:PLLA in presence of *Pseudomonas* lipase and Proteinase K occurred in the interface of two polymeric phases both

0 20 40 60 80 100 120 140 160 180

Degradation period (hour)

**Figure 1.** Mass retention variation versus degradation time, enzymatic method, of samples: (■) PCL; (●) PLLA; (▴) PCL:PLLA 20:80 (w:w); (□) composite with 5% of untreated fiber; (◇) composite with 10% of untreated fiber; (◁) com‐ posite with 5% of acetylated fiber; and (○) composite with 10% of acetylated fiber, of non-irradiated samples.

In the study of this chapter, composites degraded in a way similar of the blend through time. Mass retention values observed were higher than the blends during the same period of time of test, suggesting that coconut fibers did not significantly enzymatic degraded in this test condition. Furthermore, acetylation did not affect enzymatic degradation of composites

Tsuji and Ishizaka [15] observed that crystallinity of PCL on blends films did not change with composition variation during degradation, suggesting that this property did not affect enzymatic hydrolysis rate of PCL of the blend as it did not altered during process. Rate of enzymatic hydrolysis of blends was lower than pure PCL, suggesting that PLLA interfered on PCL hydrolysis catalyzed by lipase. One reason postulated by the authors to the deceleration

degradation occurs preferably on the surface of the sample [15].

on the bulk and on the surface of the sample.

110 Biodegradation of Hazardous and Special Products

Mass retention (%)

significantly.

**Figure 2.** Points of enzyme attack on polyesters by lipase: a) PLA; b) PCL; c) PCL:PLA 14:86 (extracted from Sivalingam et al. [25])

On Fig. 3 it is possible to observe effect of radiation dose on enzymatic degradation of PCL samples irradiated with electron beam.

According to Cottam et al. [26], degradability rate of PCL irradiated with 25 kGy decreased, attributed to irradiation process. Authors cited that lipase catalyzes hydrolysis of carbonyl group linkage and one oxygen atom in the case of fat. It is the same linkage that is broken during PCL hydrolysis. They attributed that degradability rate of PCL was affected by crosslinking occurred due to irradiation. In this study, PCL irradiated with 50 kGy suffered a slight decrease on degradation rate, in agreement with authors' observation. However, PCL samples irradiated with 100 kGy presented a certain increase on degradation rate. This fact probably is related to crystallinity decrease of around 6% observed by Differential Scanning Calorimetry, DSC, of irradiated samples. On Fig. 4 it is possible to observe the effect of radiation dose on enzymatic degradation of PLLA samples irradiated with electron beam.

According to Maharana et al. [27], enzymatic degradation occurs only on the surface of a solid substrate by erosion on the surface and by weight loss, because enzymes cannot penetrate a solid polymeric substrate. Enzymes degrade selectively amorphous regions or less ordered that allows them to diffuse through substrate and, subsequently, crystalline regions are eventually degrad‐ ed. In this process, molar weight and molar weight distribution of non-degraded solid sub‐ strate do not change during enzymatic degradation because only the polymer on the surface of substrate is degraded and products of low molar weight from degradation are removed of substrate by solubilization on the surrounding aqueous medium.

**Figure 3.** Mass retention variation versus degradation, enzymatic method, PCL of non-irradiated and irradiated sam‐ ples with EB, radiation doses of 50 kGy and 100 kGy

**Figure 5.** Hydrolysis of PLA [27].

PCL:PLLA copolymers with *M*

and then, little degradation is observed.

knowing initial carbon content.

with absorbed doses of 50 kGy and 100 kGy.


On Fig. 6 it can be observed mass retention variation through period of enzymatic degradation of PCL:PLLA 20:80 (w:w) non-irradiated and irradiated with electron beam with absorbed radiation doses of 50 kGy and 100 kGy. Degradation values observed in this study were lower compared to the ones found in the literature, probably due to the fact that studied samples were physical mixtures of polymers. Lenglet et al. [19] studied enzymatic degradation of

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

113

that degradation occurred faster with increasing amount of PCL, attaining approximately 99% for PCL:PLA 75:25 after 72h. They suggested that PCL homopolymer can suffer degradation

In Fig. 6 irradiated blend with absorbed dose of 50 kGy presented slight reduction of degra‐ dation rate compared to non-irradiated blend, similar to the observed for homopolymers. Irradiated sample with 100 kGy showed slight increase of degradation rate after 120 hours,

In Fig. 7 it is shown mass retention variation versus degradation period, by enzymatic method, of composites with 5% of chemically untreated coconut fiber, non-irradiated and EB irradiated

Chemically untreated fiber incorporation caused slight reduction of degradation comparing blend to composite. Probably, it is due to the fact that fibers take more time to degrade. Even though the method used by Salazar and Leão [28] was different from the one used in this study, they observed that fresh coconut fiber degraded 10% in 912 hours (38 days) by immediate degradability test, by measuring carbon dioxide release in open system, in which organic substance is subjected to metabolizing of microorganism mixture culture from environment. This carbon source of the substance can be fully consumed by microorganism metabolism into CO2 and H2O. It is possible to predict theoretically total CO2 production for full biodegradation,

in presence of *Pseudomonas* lipase while PLA did not degrade in the same conditions.

*n* of 29000 to 44000, using *Pseudomonas*lipase. Authors observed

**Figure 4.** Mass retention variation versus degradation period, enzymatic method, PLLA samples non irradiated and irradiated with EB, radiation doses of 50 kGy and 100 kGy

There are two kinds of degradation based on the point of cleavage. Cleavage can occur in random points along polymeric chain (degradation endo-type) or at the end terminal of main chain (degradation exo type). Degradation process of lipases is based on endo type scission, so it does not depend on molar weight and on molar weight distribution. Fig. 5 shows PLA hydrolysis reaction cited by [27].

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with… http://dx.doi.org/10.5772/56231 113

**Figure 5.** Hydrolysis of PLA [27].

0 20 40 60 80 100 120 140 160 180

Degradation period (hour)

**Figure 4.** Mass retention variation versus degradation period, enzymatic method, PLLA samples non irradiated and

There are two kinds of degradation based on the point of cleavage. Cleavage can occur in random points along polymeric chain (degradation endo-type) or at the end terminal of main chain (degradation exo type). Degradation process of lipases is based on endo type scission, so it does not depend on molar weight and on molar weight distribution. Fig. 5 shows PLA

 non irradiated 50 kGy

100 kGy

Degradation period (hour)

**Figure 3.** Mass retention variation versus degradation, enzymatic method, PCL of non-irradiated and irradiated sam‐

 non irradiated 50 kGy 100 kGy

Mass retention (%)

irradiated with EB, radiation doses of 50 kGy and 100 kGy

hydrolysis reaction cited by [27].

Mass retention (%)

112 Biodegradation of Hazardous and Special Products

ples with EB, radiation doses of 50 kGy and 100 kGy

On Fig. 6 it can be observed mass retention variation through period of enzymatic degradation of PCL:PLLA 20:80 (w:w) non-irradiated and irradiated with electron beam with absorbed radiation doses of 50 kGy and 100 kGy. Degradation values observed in this study were lower compared to the ones found in the literature, probably due to the fact that studied samples were physical mixtures of polymers. Lenglet et al. [19] studied enzymatic degradation of

PCL:PLLA copolymers with *M n* of 29000 to 44000, using *Pseudomonas*lipase. Authors observed that degradation occurred faster with increasing amount of PCL, attaining approximately 99% for PCL:PLA 75:25 after 72h. They suggested that PCL homopolymer can suffer degradation in presence of *Pseudomonas* lipase while PLA did not degrade in the same conditions.

In Fig. 6 irradiated blend with absorbed dose of 50 kGy presented slight reduction of degra‐ dation rate compared to non-irradiated blend, similar to the observed for homopolymers. Irradiated sample with 100 kGy showed slight increase of degradation rate after 120 hours, and then, little degradation is observed.

In Fig. 7 it is shown mass retention variation versus degradation period, by enzymatic method, of composites with 5% of chemically untreated coconut fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

Chemically untreated fiber incorporation caused slight reduction of degradation comparing blend to composite. Probably, it is due to the fact that fibers take more time to degrade. Even though the method used by Salazar and Leão [28] was different from the one used in this study, they observed that fresh coconut fiber degraded 10% in 912 hours (38 days) by immediate degradability test, by measuring carbon dioxide release in open system, in which organic substance is subjected to metabolizing of microorganism mixture culture from environment. This carbon source of the substance can be fully consumed by microorganism metabolism into CO2 and H2O. It is possible to predict theoretically total CO2 production for full biodegradation, knowing initial carbon content.

**Figure 6.** Mass retention variation with period of degradation, enzymatic method, of PCL:PLLA 20:80 (w:w) non-irradi‐ ated and EB irradiated samples with 50 kGy and 100 kGy absorbed doses.

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

115

Degradation period (hour)

**Figure 8.** Mass retention variation versus degradation period, enzymatic method, composites samples with 10% of chemically untreated coconut fiber non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

Fig. 9 shows mass retention variation with degradation period increase, enzymatic method, of composites samples with 5% acetylated fibers non-irradiated and EB irradiated with 50 kGy

Acetylation process of coconut fiber did not affect significantly degradation rate of irradiated samples. Irradiated samples suffered slight decrease of degradation rate compared to compo‐

Increase of acetylated fiber content up to 10% did not affect significantly degradation neither degradation rate of EB irradiated composites with 50 kGy and 100 kGy. It was observed slight increaseonmassafter168h,probablydue towater absorptionbyPLLAor coconutfibers,Fig. 10.

0 20 40 60 80 100 120 140 160 180

Degradation period (hour)

**Figure 9.** Mass retention variation versus degradation period, enzymatic method, and composites samples containing

 non-irradiated 50 kGy 100 kGy

5% acetylated fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

 non-irradiated 50 kGy 100 kGy

sites containing 5% of acetylated fibers, non-irradiated.

Mass retention (%)

Mass retention (%)

and 100 kGy.

**Figure 7.** Mass retention variation versus degradation period, enzymatic method, composites with 5% of chemically untreated coconut fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

In Fig. 7 it is observed that degradation rate of composites behaves in a way similar to irradiated blends, this suggests that fiber presence does not affect this parameter. Mass retention values of samples studied in 168 hour probably were affected by water absorption by PLLA and/or coconut fibers.

In Fig. 8 it is observed that when fiber content of chemically untreated coconut fiber increases in the composite, degradation rate suffers slight reduction and degradation decreases with increasing radiation dose.

**Figure 8.** Mass retention variation versus degradation period, enzymatic method, composites samples with 10% of chemically untreated coconut fiber non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

0 20 40 60 80 100 120 140 160 180

Degradation period (hour)

**Figure 7.** Mass retention variation versus degradation period, enzymatic method, composites with 5% of chemically

In Fig. 7 it is observed that degradation rate of composites behaves in a way similar to irradiated blends, this suggests that fiber presence does not affect this parameter. Mass retention values of samples studied in 168 hour probably were affected by water absorption by PLLA and/or

In Fig. 8 it is observed that when fiber content of chemically untreated coconut fiber increases in the composite, degradation rate suffers slight reduction and degradation decreases with

 non-irradiated 50 kGy 100 kGy

untreated coconut fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

Degradation period (hour)

**Figure 6.** Mass retention variation with period of degradation, enzymatic method, of PCL:PLLA 20:80 (w:w) non-irradi‐

 non-irradiated 50 kGy 100 kGy

ated and EB irradiated samples with 50 kGy and 100 kGy absorbed doses.

Mass retention (%)

coconut fibers.

increasing radiation dose.

Mass retention (%)

114 Biodegradation of Hazardous and Special Products

Fig. 9 shows mass retention variation with degradation period increase, enzymatic method, of composites samples with 5% acetylated fibers non-irradiated and EB irradiated with 50 kGy and 100 kGy.

Acetylation process of coconut fiber did not affect significantly degradation rate of irradiated samples. Irradiated samples suffered slight decrease of degradation rate compared to compo‐ sites containing 5% of acetylated fibers, non-irradiated.

Increase of acetylated fiber content up to 10% did not affect significantly degradation neither degradation rate of EB irradiated composites with 50 kGy and 100 kGy. It was observed slight increaseonmassafter168h,probablydue towater absorptionbyPLLAor coconutfibers,Fig. 10.

**Figure 9.** Mass retention variation versus degradation period, enzymatic method, and composites samples containing 5% acetylated fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy.

**Figure 10.** Mass retention variation versus degradation period, enzymatic method, of composites samples containing 10% of acetylated fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy

0 20 40 60 80 100 120

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

Ageing period in simulated compost soil (day)

**Figure 11.** Mass retention variation versus biodegradation period in simulated compost soil, of samples: (■) PCL; (●) PLLA; (▴) PCL:PLLA 20:80 (w:w); (□) composite with 5% chemically untreated coconut fiber; (◇) composite with 10% chemically untreated coconut fiber; (◁) composite with 5% acetylated fiber; and (○) composite with 10% of acetylat‐

Effect of radiation dose on the mass retention of EB irradiated samples with doses of 50 kGy and 100 kGy of PCL, is presented in Fig.12; of PLLA, in Fig.13; of blend PCL:PLLA 20:80 (w:w), in Fig.14; of composite containing 5% of chemically untreated fiber, in Fig.15; of composite with 10% of chemically untreated fiber, in Fig.16; of composite with 5% of acetylated fiber, in

Lotto et al. [30] observed that PCL did not suffer degradation in compost soil at room tem‐

by the authors 36% weight loss of PCL samples in 120 days. This fact was attributed to nonenzymatic hydrolysis of esters bonds due to temperature increase, that condition favored

In this study, it was observed that at room temperature PCL suffered approximately 20% of degradation in simulated compost soil in 120 days. Ionizing radiation induced degradation rate increase with increasing radiation dose in the dose range studied, Fig.12, achieving 55% of degradation in the same degradation period. Probably it was because of aerobic condition

PLLA suffered approximately 35% of degradation in the same period as PCL, Fig.13, and irradiation process promoted degradation rate increase with increasing radiation dose,

C, it was observed

http://dx.doi.org/10.5772/56231

117

Fig.17; and of composite with 10% of acetylated fiber, in Fig.18.

of simulated compost soil test is performed.

achieving 70% of degradation in 120 days.

perature even after 300 days. However, after temperature increase up to 46o

microorganism action that exists in natural soil and uses polymers as nutrient.

Mass retention (%)

ed fiber

#### **4.2. Biodegradation in simulated compost soil**

Mass retention variation versus degradation period in simulated compost soil, of nonirradiated samples PCL, PLLA, PCL:PLLA 20:80 (w:w) blend, composites wth 5% and 10% of chemically untreated fiber and composites containing 5% and 10% of acetylated fiber are shown in Fig. 11. It can be observed that all samples suffer degradation in the period of time studied. Values vary in between of 36% and 10% in 120 days for PLLA and composite with 5% of chemically untreated coconut fiber, respectively.

According to Alauzet et al. [29], PLA ester hydrolysis in abiotic aqueous media depends on autocatalysis via chain end carboxylic groups and diffusion reaction phenomena involving water and oligomer molecules formed by degradation by means of its solubility in aqueous media. When submitted to heat and water, high molecular weight PLLA degrade to oligomer (PLA of low molecular weight), dimer and monomer of lactic acid. That would explain the reason why PLA degrade in humid medium and room temperature, like organic compost or humus.

In the study of this chapter, PLLA biodegradation in simulated compost soil presented degradation rate higher than PCL, different from the behavior observed in enzymatic method, probably due to used enzyme specificity in the assay.

**Figure 11.** Mass retention variation versus biodegradation period in simulated compost soil, of samples: (■) PCL; (●) PLLA; (▴) PCL:PLLA 20:80 (w:w); (□) composite with 5% chemically untreated coconut fiber; (◇) composite with 10% chemically untreated coconut fiber; (◁) composite with 5% acetylated fiber; and (○) composite with 10% of acetylat‐ ed fiber

Degradation period (hour)

**Figure 10.** Mass retention variation versus degradation period, enzymatic method, of composites samples containing

Mass retention variation versus degradation period in simulated compost soil, of nonirradiated samples PCL, PLLA, PCL:PLLA 20:80 (w:w) blend, composites wth 5% and 10% of chemically untreated fiber and composites containing 5% and 10% of acetylated fiber are shown in Fig. 11. It can be observed that all samples suffer degradation in the period of time studied. Values vary in between of 36% and 10% in 120 days for PLLA and composite with 5%

According to Alauzet et al. [29], PLA ester hydrolysis in abiotic aqueous media depends on autocatalysis via chain end carboxylic groups and diffusion reaction phenomena involving water and oligomer molecules formed by degradation by means of its solubility in aqueous media. When submitted to heat and water, high molecular weight PLLA degrade to oligomer (PLA of low molecular weight), dimer and monomer of lactic acid. That would explain the reason why PLA degrade in humid medium and room temperature, like organic compost or

In the study of this chapter, PLLA biodegradation in simulated compost soil presented degradation rate higher than PCL, different from the behavior observed in enzymatic method,

 non-irradiated 50 kGy 100 kGy

10% of acetylated fiber, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy

**4.2. Biodegradation in simulated compost soil**

of chemically untreated coconut fiber, respectively.

probably due to used enzyme specificity in the assay.

humus.

Mass retention (%)

116 Biodegradation of Hazardous and Special Products

Effect of radiation dose on the mass retention of EB irradiated samples with doses of 50 kGy and 100 kGy of PCL, is presented in Fig.12; of PLLA, in Fig.13; of blend PCL:PLLA 20:80 (w:w), in Fig.14; of composite containing 5% of chemically untreated fiber, in Fig.15; of composite with 10% of chemically untreated fiber, in Fig.16; of composite with 5% of acetylated fiber, in Fig.17; and of composite with 10% of acetylated fiber, in Fig.18.

Lotto et al. [30] observed that PCL did not suffer degradation in compost soil at room tem‐ perature even after 300 days. However, after temperature increase up to 46o C, it was observed by the authors 36% weight loss of PCL samples in 120 days. This fact was attributed to nonenzymatic hydrolysis of esters bonds due to temperature increase, that condition favored microorganism action that exists in natural soil and uses polymers as nutrient.

In this study, it was observed that at room temperature PCL suffered approximately 20% of degradation in simulated compost soil in 120 days. Ionizing radiation induced degradation rate increase with increasing radiation dose in the dose range studied, Fig.12, achieving 55% of degradation in the same degradation period. Probably it was because of aerobic condition of simulated compost soil test is performed.

PLLA suffered approximately 35% of degradation in the same period as PCL, Fig.13, and irradiation process promoted degradation rate increase with increasing radiation dose, achieving 70% of degradation in 120 days.

hydrophobicity equilibrium of PLA. Crystallinity degree is one of the main factors that con‐ trols degradation rate of solid polymers. In general, main chain scission occurs at esters bonds sites, leading to oligomer formation, which number after chain scission depends on the quanti‐

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

119

Normally, biodegradation occurs in three steps. In the first step, depolymerization occurs, then, in the second step depolymerized PLA produces lactic acid. Finally, lactic acid is consumed in citric acid cycle where it is transformed into CO2 and H2O in the presence of an enzyme produced by microorganism. PCL:PLLA 20:80 (w:w) blend suffers degradation of approximately 30% in 120 days, PCL slightly affected PLLA degradation in the blend, Fig.14.

Radiation absorbed dose of 50kGy did not affect significant effect of degradation rate, irradiat‐ ed samples with 100kGy suffered few significant increase of degradation rate after 60 days.

0 20 40 60 80 100 120

Ageing period in simulated compost soil (day)

**Figure 14.** Mass retention variation versus biodegradation period in simulated compost soil of PCL:PLLA 20:80 (w:w)

Absorbed radiation dose of 50 kGy did not significantly affect degradation of composite containing 5% of non-chemically treated coconut fiber, neither degradation rate in simulated compost soil, FIG.15. Samples irradiated with 100kGy suffered discrete increase of degradation

Absorbed radiation dose did not affect significantly biodegradation neither degradation rate of studied samples of composites containing 10% of non-chemically treated fibers, Fig.16.

rate after 60 days of test and over 120 days biodegradation tend to stabilize.

 non-irradiated 50 kGy 100 kGy

non-irradiated and EB irradiated with radiation doses of 50 kGy and 100 kGy

Mass retention (%)

ty of ester bonds present on PLA.

**Figure 12.** Mass retention variation versus biodegradation period in simulated compost soil of PCL samples non-irradi‐ ated and EB irradiated with radiation doses of 50 kGy and 100 kGy

**Figure 13.** Mass retention variation versus biodegradation period in simulated compost soil, non-irradiated and EB irradiated PLLA samples with radiation doses of 50 kGy and 100 kGy.

Maharana et al. [27] cited that ionizing radiation does not affect glass transition temperature Tg, melting temperature Tf neither hydrolytic degradation of aliphatic polyesters. However, in our study it was possible to observe slight increase of biodegradation rate in simulated compost soil after60days.Probablythisbehaviorisrelatedtomicroorganismpresenceinsolithatwouldfavor the degradation process by produced oligomer consumption. According to the authors, as radiation induced reactions occur mainly in amorphous regions of polymers, it is important to know their crystallinity degree. Biodegradation is also affected by solid state morphology, primary chemical structure, for instance, functional groups existence and hydrophicity and hydrophobicity equilibrium of PLA. Crystallinity degree is one of the main factors that con‐ trols degradation rate of solid polymers. In general, main chain scission occurs at esters bonds sites, leading to oligomer formation, which number after chain scission depends on the quanti‐ ty of ester bonds present on PLA.

Normally, biodegradation occurs in three steps. In the first step, depolymerization occurs, then, in the second step depolymerized PLA produces lactic acid. Finally, lactic acid is consumed in citric acid cycle where it is transformed into CO2 and H2O in the presence of an enzyme produced by microorganism. PCL:PLLA 20:80 (w:w) blend suffers degradation of approximately 30% in 120 days, PCL slightly affected PLLA degradation in the blend, Fig.14.

Radiation absorbed dose of 50kGy did not affect significant effect of degradation rate, irradiat‐ ed samples with 100kGy suffered few significant increase of degradation rate after 60 days.

0 20 40 60 80 100 120

Ageing period in simulated compost soil (day)

**Figure 12.** Mass retention variation versus biodegradation period in simulated compost soil of PCL samples non-irradi‐

0 20 40 60 80 100 120

Ageing period in simulated compost soil (day)

**Figure 13.** Mass retention variation versus biodegradation period in simulated compost soil, non-irradiated and EB

Maharana et al. [27] cited that ionizing radiation does not affect glass transition temperature Tg,

study it was possible to observe slight increase of biodegradation rate in simulated compost soil after60days.Probablythisbehaviorisrelatedtomicroorganismpresenceinsolithatwouldfavor the degradation process by produced oligomer consumption. According to the authors, as radiation induced reactions occur mainly in amorphous regions of polymers, it is important to know their crystallinity degree. Biodegradation is also affected by solid state morphology, primary chemical structure, for instance, functional groups existence and hydrophicity and

neither hydrolytic degradation of aliphatic polyesters. However, in our

 non-irradiated 50 kGy 100 kGy

 non-irradiated 50 kGy 100 kGy

ated and EB irradiated with radiation doses of 50 kGy and 100 kGy

irradiated PLLA samples with radiation doses of 50 kGy and 100 kGy.

melting temperature Tf

Mass retention (%)

Mass retention (%)

118 Biodegradation of Hazardous and Special Products

**Figure 14.** Mass retention variation versus biodegradation period in simulated compost soil of PCL:PLLA 20:80 (w:w) non-irradiated and EB irradiated with radiation doses of 50 kGy and 100 kGy

Absorbed radiation dose of 50 kGy did not significantly affect degradation of composite containing 5% of non-chemically treated coconut fiber, neither degradation rate in simulated compost soil, FIG.15. Samples irradiated with 100kGy suffered discrete increase of degradation rate after 60 days of test and over 120 days biodegradation tend to stabilize.

Absorbed radiation dose did not affect significantly biodegradation neither degradation rate of studied samples of composites containing 10% of non-chemically treated fibers, Fig.16.

surface and in the bulk. In the case of bulk erosion, fragments of total mass of polymer are lost and its molecular weight is altered because of bond rupture. This rupture is provoked by chemicals (H2O, acids, bases, transition metal and radicals) or by radiation, however not by enzymes. They are very big to penetrate through bulk structure. Whereas in the case of surface erosion, matter is lost, though molecular weight of polymeric matrix does not alter. If chemical substances diffusion through the material is faster than bond scission of polymer, polymer suffers erosion. If the opposite occurs, process occurs mainly on the surface of polymeric

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

121

Radiation dose did not affect significantly biodegradation of composites containing 5% of acetylated fibers up to 60 days of test. Irradiated samples with 100kGy presented slight increase

On Fig. 18 it is possible to observe that composites samples containing 10% of acetylated fiber

0 20 40 60 80 100 120

Ageing period in simulated compost soil (day)

**Figure 17.** Mass retention variation versus period of biodegradation in simulated compost soil of composites samples containing 5% of acetylated fibers, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy

Higher degradation values found of those samples compared to non-chemically treated fibers could be related to the effect of fissures observed on the polymeric matrix during test that

Mass retention results deviations were in average 7%, probably due to weight variation of

 non-irradiated 50 kGy 100 kGy

did not suffer significant alteration of degradation rate with radiation dose increase.

matrix.

on the degradation rate after 90 days, Fig. 17.

Mass retention (%)

probably favored microbiological attack.

sample to sample.

**Figure 15.** Mass retention variation versus biodegradation period in simulated compost soil of composites samples containing 5% of non-chemically treated coconut fibers, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy

**Figure 16.** Mass retention variation versus biodegradation period in simulated compost soil of composites samples containing 10% of non-chemically treated coconut fibers, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy

It was observed during preparation of composites with acetylated coconut fibers that some kind of chemical reaction occurred during extrusion in some few events. Probably some vestiges of chemicals used for acetylation had remained on the coconut fibers. This fact could have affected degradation test of some samples of composites containing acetylated coconut fibers that started to present fissures favoring degradation on these points. Lucas et al. [31] cited that bio deterioration of thermoplastics occurs via two different mechanisms, erosion at surface and in the bulk. In the case of bulk erosion, fragments of total mass of polymer are lost and its molecular weight is altered because of bond rupture. This rupture is provoked by chemicals (H2O, acids, bases, transition metal and radicals) or by radiation, however not by enzymes. They are very big to penetrate through bulk structure. Whereas in the case of surface erosion, matter is lost, though molecular weight of polymeric matrix does not alter. If chemical substances diffusion through the material is faster than bond scission of polymer, polymer suffers erosion. If the opposite occurs, process occurs mainly on the surface of polymeric matrix.

Radiation dose did not affect significantly biodegradation of composites containing 5% of acetylated fibers up to 60 days of test. Irradiated samples with 100kGy presented slight increase on the degradation rate after 90 days, Fig. 17.

On Fig. 18 it is possible to observe that composites samples containing 10% of acetylated fiber did not suffer significant alteration of degradation rate with radiation dose increase.

0 20 40 60 80 100 120

Ageing period in simulated compost soil (day)

**Figure 15.** Mass retention variation versus biodegradation period in simulated compost soil of composites samples containing 5% of non-chemically treated coconut fibers, non-irradiated and EB irradiated with absorbed doses of 50

0 20 40 60 80 100 120

Ageing period in simulated compost soil (days)

**Figure 16.** Mass retention variation versus biodegradation period in simulated compost soil of composites samples containing 10% of non-chemically treated coconut fibers, non-irradiated and EB irradiated with absorbed doses of 50

It was observed during preparation of composites with acetylated coconut fibers that some kind of chemical reaction occurred during extrusion in some few events. Probably some vestiges of chemicals used for acetylation had remained on the coconut fibers. This fact could have affected degradation test of some samples of composites containing acetylated coconut fibers that started to present fissures favoring degradation on these points. Lucas et al. [31] cited that bio deterioration of thermoplastics occurs via two different mechanisms, erosion at

 non-irradiated 50 kGy 100 kGy

 non-irradiated 50 kGy 100 kGy

Mass retention (%)

Mass retention (%)

120 Biodegradation of Hazardous and Special Products

kGy and 100 kGy

kGy and 100 kGy

**Figure 17.** Mass retention variation versus period of biodegradation in simulated compost soil of composites samples containing 5% of acetylated fibers, non-irradiated and EB irradiated with absorbed doses of 50 kGy and 100 kGy

Higher degradation values found of those samples compared to non-chemically treated fibers could be related to the effect of fissures observed on the polymeric matrix during test that probably favored microbiological attack.

Mass retention results deviations were in average 7%, probably due to weight variation of sample to sample.

enzymatic degradation occurs in two steps: adhesion of enzyme on the surface of sample followed by scission of polymeric chain catalyzed by enzyme that generally results in small alterations of properties of polymeric matrix. According those authors, highly crystalline PCL can be fully degraded in a couple of days in presence of *Pseudomonas* lipase, while hydrolytic

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

al. [32] have cited that *Pseudomonas cepacia* lipase accelerates significantly PCL degradation. Interface activation of enzymes lipase type results mainly in conformational alteration of enzymes. Reaching substrate surface, they expose their active site and provide hydrophobic surface to the interaction with substrate molecular chains. The authors cited that several publications deal with the fundamentals of the theory of hydrolytic degradation and erosion of solid polymers. The basic modes, the surface erosion and the bulk degradation, depend on the relation between the rate of water/enzyme diffusion into the polymer, the rate of chain cleavage by water ions/enzymes, and the rate of transportation of scission products out of the solid. The rate of water diffusion into a polymer solid is strongly influenced by a number of structural parameters, its porosity, the crystallinity, the surface roughness, the hydrophobicity and the size of the sample. Most authors treat the enzymatic degradation of polymer solids exclusively as surface process. For hydrophilic enzymes it is usually considered to be difficult

C (average temperature of human body). Kulkarni et

http://dx.doi.org/10.5772/56231

123

degradation can take several years in 37o

to penetrate into a hydrophobic polymer.

**Figure 19.** General mechanism of biodegradation of polymeric materials [13].

Loo et al. [11] cited that the rate of hydrolytic degradation for biopolymers like PGLA and PLLA is controlled by altering their physical properties; such as their molecular weights, degree of crystallinity and glass transition temperature (Tg). As mentioned previously, radiation has been known to alter the physical properties of polymers through main-chain scission and cross-linking. Semi-crystalline polymers, such as PLLA, are nonhomogeneous with a two-phase system consisting of amorphous and crystalline regions. During irradiation, energy is deposited uniformly and radicals are formed throughout the polymer in both the amorphous and crystalline regions. However, crystalline regions consist of chains that were more oriented and closely packed compared to the more open amorphous regions. As a result, oxygen, stabilizers and specific active radical species are excluded from the crystalline phase,

**Figure 18.** Mass retention variation versus period of biodegradation in simulated compost soil of composites samples containing 10% of acetylated fibers, non-irradiated and EB irradiated with radiation doses of 50 kGy and 100 kGy

#### **4.3. Hydrolytic degradation**

This section will present some aspects of hydrolytic degradation because PCL and PLLA homopolymers studied here are biomaterials.

Biodegradability of polymeric materials occurs in several steps. Initially, digestible macromo‐ lecules, that form polymeric chain, suffer enzymatic scission. This is followed by metabolism of scission parts, leading to progressive enzymatic degradation of macromolecules from chain ends. Instead, macromolecular oxidative cleavage occurs, inducing fragments metabolism. Anyway, chain fragments become small enough to be converted by microorganisms [13,20]. Enzymes are catalytic proteins that decrease activation energy of molecules favoring chemical reactions. Those proteins have large diversity and marked specificity, but are easily denatured by heating, radiation, surfactants, among others [31]. In Fig. 19 general mechanism of biode‐ gradation of polymeric materials is presented.

According to Liu et al. [18], hydrolytic degradation of PCL and PLLA has been studied extensively. PLLA artifacts degradation is faster in the inner part than in the surface due to autocatalytic effect of carboxyl end groups. In the case of PCL, hydrolytic degradation is very low because of hydrophobicity and crystallinity. Authors reported that, in presence of proteinase K, PLLA degraded preferably at L-lactil units. Furthermore, enzymatic degradation occurred preferably on amorphous region of semi-crystalline PLLA polymers [11,18,19].

According to Lenglet et al. [19], hydrolytic degradation is a mass phenomenon and polyesters degradation with high size is auto catalyzed by carboxyl end groups initially present, or generated by ester bond cleavage. The three most important discoveries about polyester degradation performed in the last decade were about faster degradation in the inner portion of the sample and that degradation induces morphology and composition alteration. On the other hand, enzymes are macromolecules and cannot penetrate in a solid material. Then, enzymatic degradation occurs in two steps: adhesion of enzyme on the surface of sample followed by scission of polymeric chain catalyzed by enzyme that generally results in small alterations of properties of polymeric matrix. According those authors, highly crystalline PCL can be fully degraded in a couple of days in presence of *Pseudomonas* lipase, while hydrolytic degradation can take several years in 37o C (average temperature of human body). Kulkarni et al. [32] have cited that *Pseudomonas cepacia* lipase accelerates significantly PCL degradation. Interface activation of enzymes lipase type results mainly in conformational alteration of enzymes. Reaching substrate surface, they expose their active site and provide hydrophobic surface to the interaction with substrate molecular chains. The authors cited that several publications deal with the fundamentals of the theory of hydrolytic degradation and erosion of solid polymers. The basic modes, the surface erosion and the bulk degradation, depend on the relation between the rate of water/enzyme diffusion into the polymer, the rate of chain cleavage by water ions/enzymes, and the rate of transportation of scission products out of the solid. The rate of water diffusion into a polymer solid is strongly influenced by a number of structural parameters, its porosity, the crystallinity, the surface roughness, the hydrophobicity and the size of the sample. Most authors treat the enzymatic degradation of polymer solids exclusively as surface process. For hydrophilic enzymes it is usually considered to be difficult to penetrate into a hydrophobic polymer.

**Figure 19.** General mechanism of biodegradation of polymeric materials [13].

0 20 40 60 80 100 120

Ageing period in simulated compost soil (day)

**Figure 18.** Mass retention variation versus period of biodegradation in simulated compost soil of composites samples containing 10% of acetylated fibers, non-irradiated and EB irradiated with radiation doses of 50 kGy and 100 kGy

This section will present some aspects of hydrolytic degradation because PCL and PLLA

Biodegradability of polymeric materials occurs in several steps. Initially, digestible macromo‐ lecules, that form polymeric chain, suffer enzymatic scission. This is followed by metabolism of scission parts, leading to progressive enzymatic degradation of macromolecules from chain ends. Instead, macromolecular oxidative cleavage occurs, inducing fragments metabolism. Anyway, chain fragments become small enough to be converted by microorganisms [13,20]. Enzymes are catalytic proteins that decrease activation energy of molecules favoring chemical reactions. Those proteins have large diversity and marked specificity, but are easily denatured by heating, radiation, surfactants, among others [31]. In Fig. 19 general mechanism of biode‐

According to Liu et al. [18], hydrolytic degradation of PCL and PLLA has been studied extensively. PLLA artifacts degradation is faster in the inner part than in the surface due to autocatalytic effect of carboxyl end groups. In the case of PCL, hydrolytic degradation is very low because of hydrophobicity and crystallinity. Authors reported that, in presence of proteinase K, PLLA degraded preferably at L-lactil units. Furthermore, enzymatic degradation occurred preferably on amorphous region of semi-crystalline PLLA polymers [11,18,19].

According to Lenglet et al. [19], hydrolytic degradation is a mass phenomenon and polyesters degradation with high size is auto catalyzed by carboxyl end groups initially present, or generated by ester bond cleavage. The three most important discoveries about polyester degradation performed in the last decade were about faster degradation in the inner portion of the sample and that degradation induces morphology and composition alteration. On the other hand, enzymes are macromolecules and cannot penetrate in a solid material. Then,

 non-irradiated 50 kGy 100 kGy

Mass retention (%)

122 Biodegradation of Hazardous and Special Products

homopolymers studied here are biomaterials.

gradation of polymeric materials is presented.

**4.3. Hydrolytic degradation**

Loo et al. [11] cited that the rate of hydrolytic degradation for biopolymers like PGLA and PLLA is controlled by altering their physical properties; such as their molecular weights, degree of crystallinity and glass transition temperature (Tg). As mentioned previously, radiation has been known to alter the physical properties of polymers through main-chain scission and cross-linking. Semi-crystalline polymers, such as PLLA, are nonhomogeneous with a two-phase system consisting of amorphous and crystalline regions. During irradiation, energy is deposited uniformly and radicals are formed throughout the polymer in both the amorphous and crystalline regions. However, crystalline regions consist of chains that were more oriented and closely packed compared to the more open amorphous regions. As a result, oxygen, stabilizers and specific active radical species are excluded from the crystalline phase, and the irradiation chemical reaction paths in the amorphous and crystalline phases will therefore be different. According to Loo et al. [11], due to the close packing of the crystalline structure, the poor diffusion of oxygen into the crystalline region limits the formation of peroxyl free radicals and thus, the extent of chain scission. The ''cage effect'' also encourages the recombination of free radicals in the crystalline region. These factors play an important role in reducing the extent of e-beam degradation in PLLA.

**References**

[1] Tsuji, H, & Ikada, Y. Blends of aliphatic polyesters. I. Physical properties and mor‐ phologies of solution-cast blends from poly (DL-lactide) and poly(ε-caprolactone).

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

125

[2] Kammer, H. W, & Kummerlowe, C. (1994). Poly (ε-caprolactone) Comprising Blends-Phase Behavior and Thermal Properties, *in* Finlayson, K. (ed.) *Advances in Polymer*

[3] Dell'Erba RGroeninckx G., Maglio G., Malinconico M., Migliozzi A. Imiscible poly‐ mer blends of semicrystalline biocompatible components: thermal properties and phase morphology analysis of PLLA/PCL blends. Polymer (2001 4). , 2001(42), 1-7831.

[4] Yoshii, F, Darvis, D, Mitomo, H, & Makuuchi, K. Crosslinking of poly (ε-caprolac‐ tone) by radiation technique and its biodegradability. Radiation Physics and Chemis‐

[5] Zhang, J, Duan, Y, Sato, H, Tsuji, H, Noda, I, Yan, S, & Ozaki, Y. Crystal modifica‐ tions and thermal behavior of poly (L-lactic acid) revealed by infrared spectroscopy.

[6] Mochizuki, M, & Hirami, M. Structural effects on the biodegradation of aliphatic pol‐

[8] Sawyer, L. C, Grubb, D. T, & Meyers, G. F. eds) ((2008). Polymer Microscopy 3rd ed,

[9] Azevedo, H. S, & Reis, R. L. (2005). Understanding the enzymatic degradation of bio‐ degradable polymers and strategies to control their degradation rate. In: Reis RL, Ro‐ mán JS (eds). Biodegradable Systems in Tissue Engineering and Regenerative

[10] Chmielewski, A. G. New Trends in radiation processing of polymers, In: Internation‐ al Nuclear Atlantic Conference; Encontro Nacional de Aplicações Nucleares, 7th,

[11] Loo, J. S. C, Ooi, C. P, & Boey, F. Y. C. Degradation of poly(lactide-co-glycolide) (PLGA) and poly(l-lactide) (PLLA) by electron beam radiation. Biomaterials (2005). ,

[12] Kantoglu, Ö, & Güven, O. Radiation induced crystallinity damage in poly(L-lactic acid), Nuclear Instruments and Methods in Physical Research B (2002 1). , 2002(19),

aug. sept. 2, 2005, Santos, SP. Anais... São Paulo: ABEN, (2005). , 28.

yesters. Polymers for AdvancedTechnology (1997 8). , 1997(8), 1-203.

[7] Michler, G. H. ed) ((2008). Electron Microscopy of Polymers, Springer-Verlag.

Journal of Applied Polymer Science (1996). , 60(1), 2367-2375.

*Blends and Alloys Technology*, Technomicv, USA, 5, , 132-160.

try (2000 5). , 2000(57), 1-417.

Springer.

2005(26), 1-1359.

1-259.

Macromolecules (2005 3). , 2005(38), 1-8012.

Medicine. CRC, Boca Raton, , 177-197.

#### **5. Conclusion**

Results of degradability test, enzymatic and in simulated compost soil, indicate that studied materials suffered accentuated degradation in enzymes presence and are not affected by negatively by radiation processing. Even though coconut fibers addition had slightly reduced degradation process, composites keep degrading through time. Artifacts produced utilizing the studied materials can be processed by ionizing radiation up to 100 kGy radiation doses without detriment of their biodegradability.

#### **Acknowledgements**

We are grateful to the financial support from JICA and IAEA. Additionally, to Dr. Akihiro Oishi and Dr. Kazuo Nakayama, from National Institute of Advanced Industrial Science and Technology – AIST, Japan, for samples preparation and valuable discussion; to Dr. Naotsugu Nagasawa and Dr. Masao Tamada, from Japan Atomic Energy Agency – JAEA, Japan, for samples preparation and irradiation. We also would like to thank Dr. Morsyleide Freitas Rosa from Embrapa for providing coconut fiber; to Prof. Dr. Hélio Wiebeck, and Mr. Wilson Maia from Laboratório de Matérias-Primas Particuladas e Sólidos Não Metálicos – LMPSol, Departamento de Engenharia de Materiais, Escola Politécnica da USP (EPUSP) for coconut fiber size reduction and segregation; also to Eng. Elisabeth S.R. Somessari, Eng. Carlos G. da Silveira, and Mr. Paulo de Souza Santos, from IPEN, for blends and composites irradiation. Furthermore, we would like to acknowledge Prof. Dr. Derval dos Santos Rosa, UFABC, Centro de Engenharia, Modelagem e Ciências Sociais Aplicadas (CECS), for degradability tests and valuable discussion. We would like also to thank Dr. Luci Diva Brocardo Machado and Eng. Marcelo Augusto Gonçalves Bardi for helpful discussion.

#### **Author details**

Yasko Kodama

Address all correspondence to: ykodama@ipen.br

Nuclear and Energy Research Institute- IPEN–CNEN/SP, Radiation Technology Center, São Paulo, Brazil

#### **References**

and the irradiation chemical reaction paths in the amorphous and crystalline phases will therefore be different. According to Loo et al. [11], due to the close packing of the crystalline structure, the poor diffusion of oxygen into the crystalline region limits the formation of peroxyl free radicals and thus, the extent of chain scission. The ''cage effect'' also encourages the recombination of free radicals in the crystalline region. These factors play an important

Results of degradability test, enzymatic and in simulated compost soil, indicate that studied materials suffered accentuated degradation in enzymes presence and are not affected by negatively by radiation processing. Even though coconut fibers addition had slightly reduced degradation process, composites keep degrading through time. Artifacts produced utilizing the studied materials can be processed by ionizing radiation up to 100 kGy radiation doses

We are grateful to the financial support from JICA and IAEA. Additionally, to Dr. Akihiro Oishi and Dr. Kazuo Nakayama, from National Institute of Advanced Industrial Science and Technology – AIST, Japan, for samples preparation and valuable discussion; to Dr. Naotsugu Nagasawa and Dr. Masao Tamada, from Japan Atomic Energy Agency – JAEA, Japan, for samples preparation and irradiation. We also would like to thank Dr. Morsyleide Freitas Rosa from Embrapa for providing coconut fiber; to Prof. Dr. Hélio Wiebeck, and Mr. Wilson Maia from Laboratório de Matérias-Primas Particuladas e Sólidos Não Metálicos – LMPSol, Departamento de Engenharia de Materiais, Escola Politécnica da USP (EPUSP) for coconut fiber size reduction and segregation; also to Eng. Elisabeth S.R. Somessari, Eng. Carlos G. da Silveira, and Mr. Paulo de Souza Santos, from IPEN, for blends and composites irradiation. Furthermore, we would like to acknowledge Prof. Dr. Derval dos Santos Rosa, UFABC, Centro de Engenharia, Modelagem e Ciências Sociais Aplicadas (CECS), for degradability tests and valuable discussion. We would like also to thank Dr. Luci Diva Brocardo Machado and Eng.

Nuclear and Energy Research Institute- IPEN–CNEN/SP, Radiation Technology Center, São

role in reducing the extent of e-beam degradation in PLLA.

without detriment of their biodegradability.

Marcelo Augusto Gonçalves Bardi for helpful discussion.

Address all correspondence to: ykodama@ipen.br

**5. Conclusion**

124 Biodegradation of Hazardous and Special Products

**Acknowledgements**

**Author details**

Yasko Kodama

Paulo, Brazil


[13] Müller, R. J. ((2005). Biodegradability of Polymers: Regulations and Methods for Testing *in*: STEINBÜCHEL, A. (ed.), Biopolymers- General Aspects and Special Ap‐ plications, Wiley-VCH Verlag GmbH & Co. KgaA, n. 19, , 10, 365-374.

[25] Sivalingam, G, Vijayalakshmi, S. P, & Madras, G. Enzymatic and thermal degrada‐ tion of poly(e-caprolactone), poly(D,L-lactide, and their blends. Industrial & Engi‐

Degradability: Enzymatic and in Simulated Compost Soil of PLLA:PCL Blend and on Their Composite with…

http://dx.doi.org/10.5772/56231

127

[26] Cottam, E, Hukins, D. W. L, Lee, K, Hewitt, C, & Jenkins, M. J. Effect of sterilization by gamma irradiation on the ability of polycaprolactone (PCL) to act as a scaffold

[27] Maharana, T, Mohanty, B, & Negi, Y. S. Melt-solid polycondensation of lactic acid and its biodegradability. Progress in Polymer Science (2009 3). , 2009(34), 1-99.

[28] Salazar, V. L. P, & Leão, A. L. Biodegradação das fibras de coco e de sisal aplicadas na indústria automotiva. Revista Energia na Agricultura (2006). , 2006(21), 2-99. [29] Alauzet, N, Roussos, S, Garreau, H, & Vert, M. Microflora dynamics in earthworms casts in an artificial soil (biosynthesol) containing lactic acid oligomers Brazilian ar‐

[30] Lotto, N. T, Calil, M. R, Guedes, C. G. F, & Rosa, D. S. The effect of temperature on the biodegradation test. Materials Science and Engineerin*g* C (2004 2). , 2004(24),

[31] Lucas, N, Bienaime, C, Belloy, C, Queneudec, M, Silvestre, F, & Nava-saucedo, J. E. Polymer biodegradation: mechanisms and estimation techniques. Chemosphere

[32] Kulkarni, A, Reiche, J, Hartmann, J, Kratz, K, & Lendlein, A. Selective enzymatic degradation of poly(e-caprolactone) containing multiblock copolymers. European

Journal of pharmaceuticals and biopharmaceutics (2008 6). , 2008(68), 1-46.

neering Chemistry Research (2004 4). , 2004(43), 1-7702.

material. Medical Engineering&Physics (2009 3). , 2009(31), 1-221.

chives of biology and technology (2001 4). , 2001(44), 2-113.

1-659.

(2008 7). , 2008(73), 1-429.


[25] Sivalingam, G, Vijayalakshmi, S. P, & Madras, G. Enzymatic and thermal degrada‐ tion of poly(e-caprolactone), poly(D,L-lactide, and their blends. Industrial & Engi‐ neering Chemistry Research (2004 4). , 2004(43), 1-7702.

[13] Müller, R. J. ((2005). Biodegradability of Polymers: Regulations and Methods for Testing *in*: STEINBÜCHEL, A. (ed.), Biopolymers- General Aspects and Special Ap‐

[14] Kuo, S-W. Hydrogen-bonding in polymer blends. Journal of Polymer Research (2008

[15] Tsuji, H, & Ishizaka, T. Blends of aliphatic polyesters, VI. Lipase-catalyzed hydrolysis and visualized phase strucuture of biodegradable blends from poly(e-caprolactone) and poly(L-lactide). International Journal of Biological Macromolecules (2001 2). ,

[16] Kikkawa, Y, Suzuki, T, Tsuge, T, Kanesato, M, Doi, Y, & Abe, H. Phase structure and enzymatic degradation of poly(L-lactide)/atactic poly(3-hydroxybutyrate) blends: an

[17] Nishino, T, & Hirao, K. Kotera M. X-ray diffraction studies on stress transfer of kenaf reinforced poly(L-lactic acid) composite. Composites: Part A (2006). , 2006(37),

[18] Liu, L, Li, S, Garreau, H, & Vert, M. Selective Enzymatic Degradations of Poly(L-lac‐ tide) and Poly(ε-caprolactone) Blend Films. Biomacromolecules (2000). , 2000(1),

[19] Lenglet, S, Li, S, & Vert, M. Lipase-catalysed degradation of copolymers prepared from e-caprolactone and DL-lactide. Polymer Degradation and Stability (2009 9). ,

[20] Kolybaba, M, Tabil, L. G, Panigrahi, S, Crerar, W. J, Powell, T, & Wang, B. (2003). Bio‐ degradable polymers: past, present and future. 2003 CSAE/ASAE Annual Intercec‐ tional Meeting Sponsered by the Red Rive Section of ASAE. Fargo, North Dakota,

[21] Calado, V, Barreto, D. W, & Almeida, D. J.R.M. The effect of a chemical treatment on the structure and morphology of coir fibers. Journal of Materials Science Letters

[22] Almeida, D, Calado, A. L. F. S, & Barreto, V. D.W. Acetilação da fibra de bucha (Luf‐

[23] Spinks, J. W. T, & Woods, R. J. An Introduction to Radiation Chemistry. 3rd ed. USA:

[24] Calil, M. R, Gaboardi, F, Bardi, M. A. G, Rezende, M. L, & Rosa, D. S. Enzymatic deg‐ radation of poly(ε-caprolactone) and cellulose acetate blends by lipase and α-ami‐

fa cylindrica) Polímeros: Ciência e Tecnologia (2005 1). , 2005(15), 1-59.

atomic force microscopy study. Biomacromolecules (2006). , 2006(7), 1-1921.

plications, Wiley-VCH Verlag GmbH & Co. KgaA, n. 19, , 10, 365-374.

1). , 2008(15), 1-459.

126 Biodegradation of Hazardous and Special Products

2001(29), 1-83.

1-2269.

1-350.

2009(94), 1-688.

USA, October , 3-4.

(2000). , 2000(19), 1-2151.

John Wiley and Sons; (1990).

lase. Polymer Testing (2007 2). , 2007(26), 1-257.


**Chapter 7**

**Intradiol Dioxygenases —**

Danuta Wojcieszyńska

http://dx.doi.org/10.5772/56205

or carcinogenic activity [5-8].

active in this respect [9-22].

**1. Introduction**

Urszula Guzik, Katarzyna Hupert-Kocurek and

Additional information is available at the end of the chapter

**The Key Enzymes in Xenobiotics Degradation**

Aromatic compounds are derived from both natural and anthropogenic sources. Under natural conditions, arenes are formed as a result of the pyrolysis of organic materials at high temperatures during forest, steppe and peatland fires, and during volcanic eruptions. Biogenic aromatic compounds like aromatic amino acids and lignin, the second most abundant organic compound in the environment, are universally distributed in nature. Many species of plants, especially willow (*Salix*), thyme (*Thymus vulgaris*), camomile (*Chamomilla recutita*), bean (*Phaesoli vulgaris*) or strawberry (*Fregaria ananasa*), water plants as sweet flag (*Acorus calamus*) and many species of alga are known to produce aromatic compounds as secondary metabolites [1-4]. A lot of aromatic compounds are introduced to the environment as contaminating compounds from chemical, pharmaceutical, explosive, dyes, and agrochemicals industry. Chloro-, amino- and nitroaromatic derivatives, biphenyls, polycyclic aromatic hydrocarbons accumulate in the soil and water. They are toxic to living systems including humans, animals, and plants. Moreover, most of them may bioaccumulate in the food chain and have mutagenic

Due to the delocalization of π orbitals aromatic compounds are very stable [3]. For many years it has been searching for microorganism able to breakdown these kind of substrates. Among the bacteria, representatives of the genus *Pseudomonas, Sagittula, Streptomyces, Agrobacterium, Acinetobacter, Arthrobacter, Burkholderia, Bacillus, Stenotrophomonas, Brevibacterium, Sphingomo‐ nas, Geobacillus, Rhodococcus, Nocardia, Corynebacterium, Alcaligenes, Gordonia*, or *Citrobacter* are

> © 2013 Guzik et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use,

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

distribution, and reproduction in any medium, provided the original work is properly cited.

and reproduction in any medium, provided the original work is properly cited.

### **Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation**

Urszula Guzik, Katarzyna Hupert-Kocurek and Danuta Wojcieszyńska

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56205

#### **1. Introduction**

Aromatic compounds are derived from both natural and anthropogenic sources. Under natural conditions, arenes are formed as a result of the pyrolysis of organic materials at high temperatures during forest, steppe and peatland fires, and during volcanic eruptions. Biogenic aromatic compounds like aromatic amino acids and lignin, the second most abundant organic compound in the environment, are universally distributed in nature. Many species of plants, especially willow (*Salix*), thyme (*Thymus vulgaris*), camomile (*Chamomilla recutita*), bean (*Phaesoli vulgaris*) or strawberry (*Fregaria ananasa*), water plants as sweet flag (*Acorus calamus*) and many species of alga are known to produce aromatic compounds as secondary metabolites [1-4]. A lot of aromatic compounds are introduced to the environment as contaminating compounds from chemical, pharmaceutical, explosive, dyes, and agrochemicals industry. Chloro-, amino- and nitroaromatic derivatives, biphenyls, polycyclic aromatic hydrocarbons accumulate in the soil and water. They are toxic to living systems including humans, animals, and plants. Moreover, most of them may bioaccumulate in the food chain and have mutagenic or carcinogenic activity [5-8].

Due to the delocalization of π orbitals aromatic compounds are very stable [3]. For many years it has been searching for microorganism able to breakdown these kind of substrates. Among the bacteria, representatives of the genus *Pseudomonas, Sagittula, Streptomyces, Agrobacterium, Acinetobacter, Arthrobacter, Burkholderia, Bacillus, Stenotrophomonas, Brevibacterium, Sphingomo‐ nas, Geobacillus, Rhodococcus, Nocardia, Corynebacterium, Alcaligenes, Gordonia*, or *Citrobacter* are active in this respect [9-22].

© 2013 Guzik et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

The strategy for degradation of aromatic structure comprises hydroxylation and cleavage of the aromatic ring with the usage of oxygenases [8,23,24]. These reactions are initiated by electrophilic attack of molecular oxygen and they are retarded by the presence of electronwithdrawing substituents. For this reason the velocity of reaction is connected with the kind of substituents in the aromatic ring. Substituents of the first group such as: -NH2, -OH, - OCH3, -CH3, -C6H5 activate aromatic ring for electrophilic attack. In contrary, the substituents of the second group (-NO2, -N(CH3)3 + , -CN, -COOH, -COOR, -SO3H, -CHO, -COR) and the third group (-F, -Cl, -Br, -I) impede the electrophilic attack by dioxygenases [3,25-28].

They could be applicable not only in biodegradation but also in industry processes since products of the *ortho* cleavage are valuable intermediates in chemical synthesis [44,45].

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

131

Because some aromatics have been present in the environment for a very short time, bacteria have not evolved efficient pathways for their degradation and for that reason the construction of such pathways may be required. Therefore, a detailed understanding of the molecular bases

Our chapter is divided into three parts: first we classify bacterial intradiol dioxygenases, key enzymes in aromatic compounds degradation, taking under consideration phylogenetic relationships among dioxygenases from various strains and their substrate specificity. Next, we present current knowledge about molecular structure of these dioxygenases and hypo‐ thetical mechanisms for aromatic ring cleavage. And finally, we describe modifications

Many studies on intradiol dioxygenases concentrate on the understanding of their biochemical and structural properties. Based on the results of these studies and taking into consideration substrate specificity of the *ortho* fission dioxygenases they were divided by Vetting and Ohlendorf [47] into two families: catechol 1,2-dioxygenases and protocatechuate 3,4-dioxyge‐ nases, including hydroxyquinol 1,2-dioxygenases. In turn, Hammer et al. [12] and Contzen and Stolz [48] divided protocatechuate 3,4-dioxygenases into two types. Type I dioxygenases catalyze ring fission of protocatechuic acid and its derivatives while type II enzymes cleave *ortho* diphenols carrying a larger substituent at the 4-position of the aromatic ring [12,48].

Although Murakami et al. [14] observed partial activity of hydroxyquinol 1,2-dioxygenase towards catechol and Pandeeti and Siddavattam [49] isolated catechol 1,2-dioxygenase which was active strictly against catechol and 1,2,3-benzenotriol there is no reports about activity of catechol 1,2-dioxygenases towards hydroxyquinol [18,21,50,51]. Therefore, we propose to divide the intradiol dioxygenases into three classes: catechol 1,2-dioxygenases, protocatechu‐ ate 3,4-dioxygenases, and hydroxyquinol 1,2-dioxygenases. Representatives of these three classes are presented in Table 1. Based on the results of biochemical and genetic comparison of intradiol dioxygenases Perez-Pantoja et al. [52] distinguished hydroxyquinol 1,2-dioxyge‐

The catechol 1,2-dioxygenases family can be divided into several subclasses depending on their substrate specificity. Initially this family was divided into two subclasses: type I dioxy‐ genases with activity against catechol and lack or weak activity against chlorocatechol and type II dioxygenases, which cleave only chlorocatechols [53]. Then, based on their amino acid sequences Liu et al. [54] divided family of catechol 1,2-dioxygenases into three groups: catechol 1,2-dioxygenases from gram-negative bacteria, chlorocatechol 1,2-dioxygenases from gramnegative bacteria, and both catechol- and chlorocatechol 1,2-dioxygenases from gram-positive bacteria. However, both of these classifications do not take into consideration substrate

as well as enzymes of known pathways is essential [46].

increasing biodegradation potential of these enzymes.

**2. Classification of bacterial intradiol dioxygenases**

nases as a separate cluster that may support our suggestions.

Hydroxylation into the dihydroxylated intermediates, the first step in the oxidative degrada‐ tion of aromatic compounds, is catalyzed by monooxygenases or hydroxylating dioxygenases (Fig. 1). Aromatic monooxygenases are divided into two groups: activated- ring monooxyge‐ nases and nonactivated- ring enzymes and usually contain FAD as a prosthetic group. These enzymes posses a dinuclear iron centre with two oxo- bridged iron atoms (Fe-O-Fe) in the active site [36-39]. Aromatic monooxygenases activate molecular oxygen through the forma‐ tion of a reactive flavin (hydro)peroxide which can attack the substrate [31,33-35].

Aromatic ring hydroxylating dioxygenases that belong to the Rieske non-heme iron oxygen‐ ases are multicomponent enzyme systems. The reaction catalyzed by hydroxylating dioxyge‐ nases requires two electrons from a NAD(P)H that are consecutively transferred to the terminal oxygenase component through a electron carries such as ferredoxin and/or a reductase [36].

As a result of hydroxylation key intermediates such as catechol, protocatechuic acid, hydrox‐ yquinol, or gentisic acid are formed. These products are substrates for ring-cleaving dioxyge‐ nases [17,37-39]. The ring-cleaving dioxygenases couple O2 bond cleavage with ring fission of hydroxylated derivatives either between the two hydroxyl group (*ortho* cleavage) or beside one of these (*meta* cleavage) [23,40].

Intradiol dioxygenases catalyze the intradiol cleavage of the aromatic ring at 1,2-position of catechol or its derivatives (protocatechuic acid, hydroxyquinol) with incorporation of two atoms of molecular oxygen into the substrate. It leads to the production of *cis,cis*muconic acid or its derivatives (3-carboxy-*cis,cis* muconic acid, 3-hydroxy-*cis,cis* muconic acid). The *cis,cis*-muconic acid is then subsequently transformed by a muconate cycloiso‐ merase to muconolactone. Muconolactone isomerase shifts the double bond to form 3 oxoadipate-enol-lactone, the first common intermediate of the catechol and protocatechuate or hydroxyquinol branch (Fig. 1) [38,41,42].

Extradiol catechol and protocatechuate dioxygenases catalyze the ring fission between position C2 and C3 of catechol ring and between position C2 and C4 or C4 and C5 of proto‐ catechuate ring respectively. Products of these reactions (2-hydroxymuconic semialdehyde or carboxy-2-hydroxymuconic semialdehyde) are transformed finally to pyruvic acid and acetaldehyde in catechol pathway and pyruvic acid and acetaldehyde or pyruvic acid and oxaloacetic acid in protocatechuate pathway (Fig. 1) [37,43].

Degradation of aromatic compounds by the *meta* pathway often leads to more toxic inter‐ mediates such as acyl chloride that is why in the degradation of some arenes the *ortho* cleavage is preferred [5,6]. Recently intradiol dioxygenases with extremely high activity were isolated. They could be applicable not only in biodegradation but also in industry processes since products of the *ortho* cleavage are valuable intermediates in chemical synthesis [44,45].

Because some aromatics have been present in the environment for a very short time, bacteria have not evolved efficient pathways for their degradation and for that reason the construction of such pathways may be required. Therefore, a detailed understanding of the molecular bases as well as enzymes of known pathways is essential [46].

Our chapter is divided into three parts: first we classify bacterial intradiol dioxygenases, key enzymes in aromatic compounds degradation, taking under consideration phylogenetic relationships among dioxygenases from various strains and their substrate specificity. Next, we present current knowledge about molecular structure of these dioxygenases and hypo‐ thetical mechanisms for aromatic ring cleavage. And finally, we describe modifications increasing biodegradation potential of these enzymes.

#### **2. Classification of bacterial intradiol dioxygenases**

The strategy for degradation of aromatic structure comprises hydroxylation and cleavage of the aromatic ring with the usage of oxygenases [8,23,24]. These reactions are initiated by electrophilic attack of molecular oxygen and they are retarded by the presence of electronwithdrawing substituents. For this reason the velocity of reaction is connected with the kind of substituents in the aromatic ring. Substituents of the first group such as: -NH2, -OH, - OCH3, -CH3, -C6H5 activate aromatic ring for electrophilic attack. In contrary, the substituents

, -CN, -COOH, -COOR, -SO3H, -CHO, -COR) and the

+

third group (-F, -Cl, -Br, -I) impede the electrophilic attack by dioxygenases [3,25-28].

tion of a reactive flavin (hydro)peroxide which can attack the substrate [31,33-35].

Hydroxylation into the dihydroxylated intermediates, the first step in the oxidative degrada‐ tion of aromatic compounds, is catalyzed by monooxygenases or hydroxylating dioxygenases (Fig. 1). Aromatic monooxygenases are divided into two groups: activated- ring monooxyge‐ nases and nonactivated- ring enzymes and usually contain FAD as a prosthetic group. These enzymes posses a dinuclear iron centre with two oxo- bridged iron atoms (Fe-O-Fe) in the active site [36-39]. Aromatic monooxygenases activate molecular oxygen through the forma‐

Aromatic ring hydroxylating dioxygenases that belong to the Rieske non-heme iron oxygen‐ ases are multicomponent enzyme systems. The reaction catalyzed by hydroxylating dioxyge‐ nases requires two electrons from a NAD(P)H that are consecutively transferred to the terminal oxygenase component through a electron carries such as ferredoxin and/or a reductase [36]. As a result of hydroxylation key intermediates such as catechol, protocatechuic acid, hydrox‐ yquinol, or gentisic acid are formed. These products are substrates for ring-cleaving dioxyge‐ nases [17,37-39]. The ring-cleaving dioxygenases couple O2 bond cleavage with ring fission of hydroxylated derivatives either between the two hydroxyl group (*ortho* cleavage) or beside

Intradiol dioxygenases catalyze the intradiol cleavage of the aromatic ring at 1,2-position of catechol or its derivatives (protocatechuic acid, hydroxyquinol) with incorporation of two atoms of molecular oxygen into the substrate. It leads to the production of *cis,cis*muconic acid or its derivatives (3-carboxy-*cis,cis* muconic acid, 3-hydroxy-*cis,cis* muconic acid). The *cis,cis*-muconic acid is then subsequently transformed by a muconate cycloiso‐ merase to muconolactone. Muconolactone isomerase shifts the double bond to form 3 oxoadipate-enol-lactone, the first common intermediate of the catechol and protocatechuate

Extradiol catechol and protocatechuate dioxygenases catalyze the ring fission between position C2 and C3 of catechol ring and between position C2 and C4 or C4 and C5 of proto‐ catechuate ring respectively. Products of these reactions (2-hydroxymuconic semialdehyde or carboxy-2-hydroxymuconic semialdehyde) are transformed finally to pyruvic acid and acetaldehyde in catechol pathway and pyruvic acid and acetaldehyde or pyruvic acid and

Degradation of aromatic compounds by the *meta* pathway often leads to more toxic inter‐ mediates such as acyl chloride that is why in the degradation of some arenes the *ortho* cleavage is preferred [5,6]. Recently intradiol dioxygenases with extremely high activity were isolated.

of the second group (-NO2, -N(CH3)3

130 Biodegradation of Hazardous and Special Products

one of these (*meta* cleavage) [23,40].

or hydroxyquinol branch (Fig. 1) [38,41,42].

oxaloacetic acid in protocatechuate pathway (Fig. 1) [37,43].

Many studies on intradiol dioxygenases concentrate on the understanding of their biochemical and structural properties. Based on the results of these studies and taking into consideration substrate specificity of the *ortho* fission dioxygenases they were divided by Vetting and Ohlendorf [47] into two families: catechol 1,2-dioxygenases and protocatechuate 3,4-dioxyge‐ nases, including hydroxyquinol 1,2-dioxygenases. In turn, Hammer et al. [12] and Contzen and Stolz [48] divided protocatechuate 3,4-dioxygenases into two types. Type I dioxygenases catalyze ring fission of protocatechuic acid and its derivatives while type II enzymes cleave *ortho* diphenols carrying a larger substituent at the 4-position of the aromatic ring [12,48].

Although Murakami et al. [14] observed partial activity of hydroxyquinol 1,2-dioxygenase towards catechol and Pandeeti and Siddavattam [49] isolated catechol 1,2-dioxygenase which was active strictly against catechol and 1,2,3-benzenotriol there is no reports about activity of catechol 1,2-dioxygenases towards hydroxyquinol [18,21,50,51]. Therefore, we propose to divide the intradiol dioxygenases into three classes: catechol 1,2-dioxygenases, protocatechu‐ ate 3,4-dioxygenases, and hydroxyquinol 1,2-dioxygenases. Representatives of these three classes are presented in Table 1. Based on the results of biochemical and genetic comparison of intradiol dioxygenases Perez-Pantoja et al. [52] distinguished hydroxyquinol 1,2-dioxyge‐ nases as a separate cluster that may support our suggestions.

The catechol 1,2-dioxygenases family can be divided into several subclasses depending on their substrate specificity. Initially this family was divided into two subclasses: type I dioxy‐ genases with activity against catechol and lack or weak activity against chlorocatechol and type II dioxygenases, which cleave only chlorocatechols [53]. Then, based on their amino acid sequences Liu et al. [54] divided family of catechol 1,2-dioxygenases into three groups: catechol 1,2-dioxygenases from gram-negative bacteria, chlorocatechol 1,2-dioxygenases from gramnegative bacteria, and both catechol- and chlorocatechol 1,2-dioxygenases from gram-positive bacteria. However, both of these classifications do not take into consideration substrate specificity of these enzymes towards methylcatechols. High substrate specificity against these compounds was observed for intradiol dioxygenase isolated from genus *Rhodococcus* [55-57]. Above studies divided catechol dioxygenases into three subclasses. The first subclass com‐ prises the enzymes with high enzymatic activity towards chlorocatechols (CCs), the second one includes enzymes with moderate activities for catechol only or catechol and 4-methylca‐ techol (4-MC). The third subclass consists of the enzymes with high activity for 3- and 4 methylcatechols (3-MC and 4-MC) [50,55].

**Enzyme Microorganism Gene AC Number Substrates References**

*catAA catAB*

*Arthrobacter* BA-5-17 *catA-II* AB109791.1 catechol

*Burkholderia* sp. TH2 *catA1* AB035483 catechol

*Burkholderia* sp. TH2 *catA2* AB035325 catechol

*Pseudomonas* sp. B13 - - catechol

*-* - catechol

*-* - catechol

AF380158 AF182166


*Citrobacter* sp. SA01 - - catechol [22]

*-* - Catechol

3-isopropylcatechol 4-methylcatechol 3-methylcatechol

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

4-methylcatechol 3-methylcatechol

4-methylcatechol

3-methylcatechol 4-methylcatechol

3-methylcatechol 4-methylcatechol

4-methylcatechol

4-methylactechol pyrogallol

3-chlorocatechol 4-chlorocatechol 3-methylcatechol 4-methylcatechol


*catA* EU000396.1 catechol [38]

*catA* D83237 catechol [55]

catechol 4-methylcatechol

*catA* - catechol [49]

*catA* NC\_007973.1 catechol [61]

*-* - catechol [47]

[58]

133

http://dx.doi.org/10.5772/56205

[59]

[53,60]

[25]

[62]

[63]

[63]

[64]

[25]

Catechol 1,2 dioxygenase

*Acinetobacter calcoaceticus* ADP-96

*Acinetobacter calcoaceticus* ADP1

*Acinetobacter radioresistens*

*Acinetobacter radioresistens* S13

*Acinetobacter* sp.

DS002

*Alcaligenes eutrophus* B.9

*Alcaligenes eutrophus* CH34

*Gordonia*

N6

*polyisoprenivorans*

*Pseudomonas putida*

*Pseudomonas aeruginosa* TKU002

*Rhodococcus erythropolis* AN-13

**Figure 1.** Pathways of aromatic compounds degradation [36,40,43].


specificity of these enzymes towards methylcatechols. High substrate specificity against these compounds was observed for intradiol dioxygenase isolated from genus *Rhodococcus* [55-57]. Above studies divided catechol dioxygenases into three subclasses. The first subclass com‐ prises the enzymes with high enzymatic activity towards chlorocatechols (CCs), the second one includes enzymes with moderate activities for catechol only or catechol and 4-methylca‐ techol (4-MC). The third subclass consists of the enzymes with high activity for 3- and 4-

methylcatechols (3-MC and 4-MC) [50,55].

132 Biodegradation of Hazardous and Special Products

**Figure 1.** Pathways of aromatic compounds degradation [36,40,43].


**Enzyme Microorganism Gene AC Number Substrates References**

*-* - protocatechuate [76]

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

135

*pcaHG* U33634 protocatechuate [77]

*pcaHG* L14836.1 protocatechuate [78-80]

*pcaHG* Y18527 protocatechuate [81]

*pcaHG* AF109386.2 protocatechuate [15]

*tftH* AB016258 hydroxyquinol, catechol [14]

*tftH* U19883.1 hydroxyquinol [84]

*pdcDE* FJ821777.2 hydroxyquinol [8]

*pnpG* EF577044.1 hydroxyquinol [51]

*tcpC* AF498371 6-chlorohydroxyquinol [87]

*graB* AB266211.1 hydroxyquinol [89]

*npcC* AB154422 hydroxyquinol [90]

pyrogallol 3-methylcatechol 6-chlorohydroxyquinol

protocatechuate [68]

[41,83]

[17,86]

[88]

chlorohydroxyquinol

6-chlorohydroxyquinol, 5-chlorohydroxyquinol,

*Rhizobium trifolii -* - protocatechuate [82]

JQ364963 JQ364964

*Burkholderia* sp. AK-5 *-* - hydroxyquinol [85]

*chqB* AY822041.2 hydroxyquinol,

*hadC* D86544 hydroxyquinol

*pcaH pcaG*

*Azotobacter* sp. GP1 - - hydroxyquinol,

*Pseudomonas fluorescens* A.3.12

*Pseudomonas marginata*

*Pseudomonas putida* ATCC23975

*Pseudomonas* sp.

*Stenotrophomonas maltophilia KB2*

*Streptomyces* sp.

*Arthrobacter* sp. BA-5-17

*Burkholderia cepacia*

*Nocardioides simplex*

*Pseudomonas* sp.

*Pseudomonas* sp.

*Ralstonia eutropha*

*Ralstonia pickettii* DTP0602

*Rhizobium* sp. MTP-10005

SAO101

*Rhodococcus opacus*

**Table 1.** Intradiol dioxygenases from various microorganisms

HR199

2065

AC1100

3E

1-7

WBC-3

JMP134

Hydroxyquinol 1,2 dioxygenase


**Table 1.** Intradiol dioxygenases from various microorganisms

**Enzyme Microorganism Gene AC Number Substrates References**

*catA* X99622.1 catechol

3-methylcatechol 4-methylcatechol

*clcA* AF003948.1 4-chlorocatechol [65]

*clcA2* AJ439407.1 3-chlorocatechol [65,66]


*-* - catechol [67]

*catA* EU000397.1 catechol [68]

*pcaHG* - protocatechuate [10,69]

*pcaHG* AY099487 protocatechuate [70]

*pcaHG* NC\_005966.1 protocatechuate [16]

*-* - protocatechuate [71]

protocatechuate protocatechuate 4-sulfocatechol

sulfocatechol

4-sulfocatechol 3,4-dihydroxyphenyl-

3,4-dihydroxy-cinnamate

acetate

*-* - protocatechuate [75]

protocatechuate [19]

[50]

[48]

[12]

[12]

*Rhodococcus opacus*

*Rhodococcus opacus*

*Rhodococcus opacus*

*Rhodococcus* sp. NCIM 2891

*Sphingomonas xenophaga* QYY

*Acinetobacter calcoaceticus*

K24

ADP1

*Stenotrophomonas maltophilia KB2*

*Acinetobacter lwoffii*

*Acinetobacter* sp.

*Agrobacterium radiobacter* S2

*Agrobacterium radiobacter S2*

*Brevibacterium fuscum*

*Burkholderia* sp. NCIMB 10467

*Hydrogenophaga palleronii S1*

*Pseudomonas cepacia*

DBO1

*pcaH1G1 pcaH2G2* AF230649 AF230650

YP552569

*Geobacillus* sp. *-* - protocatechuate [18]

*Moraxella* sp.GU2 - - protocatechuate [11] *Pseudomonas cepacia -* - protocatechuate [72-74]

*-* - protocatechuate

*pcaHG* YP552570

*-* - protocatechuate4-

1CP

134 Biodegradation of Hazardous and Special Products

1CP

1CP

Protocatechuate 3,4-dioxygenase

#### **3. Molecular structure and reaction mechanism of intradiol dioxygenases**

Intradiol dioxygenases belong to metalloenzymes with non-heme iron (III) that plays a crucial role for the binding and activation of dioxygen in the cleavage reaction [55,59]. Metal ion is ligated by two tyrosine and two histidine residues that are highly conserved in all investigated intradiol dioxygenases. The fifth ligand is a hydroxyl group derived from the solvent.

Two structurally different intradiol dioxygenases families can be singled out: catechol/ hydroxyquinol 1,2-dioxygenases and protocatechuate 3,4-dioxygenases. Catechol/hydroxy‐ quinol 1,2-dioxygenases are dimmers (αFe3+)2 with identical or similar subunits while proto‐ catechuate 3,4-dioxygenases are typically composed of two homologous subunits in oligomeric complexes (αβFe3+)2-12 [66] (Fig. 3).

#### **3.1. Catechol 1,2-dioxygenases characterization**

Catechol 1,2-dioxygenases catalyze cleavage of catechol and its derivatives. The tertiary structure of the enzymes from this class is similar [53,59,61,64,66]. They form homodimers with two subunits connected with helical zipper motif. Each subunit has a molecular mass of 29.0-38.6 kDa.

Subunits of catechol 1,2-dioxygenases contain two molecules of phospholipids bound into a hydrophobic cavity at the dimeric interface [38,47,49,50,53,59,61,64,66,67]. On the base of electron density and the stereochemistry these phospholipids were identified as a phosphati‐ dylcholine with two C14-15/C12-C17 hydrophobic tails [66]. It is suggested that the phospho‐ lipid chains may modulate the activity of catechol 1,2-dioxygenases by increasing the local concentration of substrate or the lipids act as an effector molecule. The binding of phospholipid to hydrophobic amino acid of the active site can lead to a conformational change at this place. During catalysis catechol is cleaved to *cis,cis-* muconic acid which is known to be toxic to the cell at low levels. Perhaps the next enzyme of intradiol pathway- *cis-cis-* muconate lactonizing enzyme, interacts with the hydrophobic tunnel and allows the proteins to pass the product from one enzyme to next. Moreover, phospholipids can alter the phospholipid bilayer in response to toxic levels of aromatic compounds [47,66].

active site with ferric ion is located between β sheets sandwich and the linker domain [47]. Matera et. al. [50] compared structure of catechol 1,2-dioxygenase from Gram- positive strain *Rhodococcus opacus* 1CP with the enzyme from *Acinetobacter calcoaceticus* described above and observed that the differences in the structure are localized at the N-terminus. In this terminal chain helix 5 is shorter, strand 1 and helix 7 are missing and a new short helix is present between position 124 and 128. Moreover the C-terminal chain of catechol 1,2-dioxygenase from Grampositive strain is shorter than in the other catechol 1,2-dioxygenases from Gram-negative

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

137

As it was mentioned earlier the metal ion is ligated by four amino acid residues and a hydroxyl group from water in trigonal bipyramidal geometry. One molecule of tyrosine and one histidine are the axial ligands, whereas the second Tyr, His and a solvent molecule make up the equatorial plane. [38,47,50,53]. Since the two Fe3+ ions from catalytic domains are separated by more than 40 Å Vetting and Ohlendorf [47] suggested that they act structurally and catalytically independently. It explains the lack of allosteric effect [25,46,57,59,61,70,80]. However, catechol 1,2-dioxygenase from *Acinetobacter radioresistens* exhibits a sigmoidal

The substrate specificity of catechol 1,2-dioxygenases depends on electronic factors, which can play a significant role in determining the efficiency of conversion of catechol deriva‐ tives. Currently, there is little information about the influence of steric factors on catalyt‐

bacteria.

kinetics indicating cooperation of subunits [59].

**Figure 2.** Characterization of intradiol dioxygenases

Unique helical N-terminal domain was first described by Vetting and Ohlendorf [47] for catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus* ADP1. Sequence aligments indicate that this helical domain is conserved among all members of catechol 1,2-dioxygenase class.

Catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus* ADP1 is an elongated molecule, which is composed of helices and β sheets. The overall shape of this enzyme is similar to boomerang and it is divided into three domains: the two catalytic domains at each end of the molecule, and one central linker domain which is composed of several helices. The catalytic domain consists of two mixed topology β sheets which form a β sheets sandwich [47]. The similar structure is also observed in another catechol 1,2-dioxygenases [38,50].

The linker domain of catechol 1,2-dioxygenases contains 12 α helices. Five of them belong to the N-terminal chain of each monomer and one extending from each catalytic domain. The Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation http://dx.doi.org/10.5772/56205 137

**Figure 2.** Characterization of intradiol dioxygenases

**3. Molecular structure and reaction mechanism of intradiol dioxygenases**

Intradiol dioxygenases belong to metalloenzymes with non-heme iron (III) that plays a crucial role for the binding and activation of dioxygen in the cleavage reaction [55,59]. Metal ion is ligated by two tyrosine and two histidine residues that are highly conserved in all investigated intradiol dioxygenases. The fifth ligand is a hydroxyl group derived from the solvent.

Two structurally different intradiol dioxygenases families can be singled out: catechol/ hydroxyquinol 1,2-dioxygenases and protocatechuate 3,4-dioxygenases. Catechol/hydroxy‐ quinol 1,2-dioxygenases are dimmers (αFe3+)2 with identical or similar subunits while proto‐ catechuate 3,4-dioxygenases are typically composed of two homologous subunits in

Catechol 1,2-dioxygenases catalyze cleavage of catechol and its derivatives. The tertiary structure of the enzymes from this class is similar [53,59,61,64,66]. They form homodimers with two subunits connected with helical zipper motif. Each subunit has a molecular mass of

Subunits of catechol 1,2-dioxygenases contain two molecules of phospholipids bound into a hydrophobic cavity at the dimeric interface [38,47,49,50,53,59,61,64,66,67]. On the base of electron density and the stereochemistry these phospholipids were identified as a phosphati‐ dylcholine with two C14-15/C12-C17 hydrophobic tails [66]. It is suggested that the phospho‐ lipid chains may modulate the activity of catechol 1,2-dioxygenases by increasing the local concentration of substrate or the lipids act as an effector molecule. The binding of phospholipid to hydrophobic amino acid of the active site can lead to a conformational change at this place. During catalysis catechol is cleaved to *cis,cis-* muconic acid which is known to be toxic to the cell at low levels. Perhaps the next enzyme of intradiol pathway- *cis-cis-* muconate lactonizing enzyme, interacts with the hydrophobic tunnel and allows the proteins to pass the product from one enzyme to next. Moreover, phospholipids can alter the phospholipid bilayer in

Unique helical N-terminal domain was first described by Vetting and Ohlendorf [47] for catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus* ADP1. Sequence aligments indicate that this helical domain is conserved among all members of catechol 1,2-dioxygenase class.

Catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus* ADP1 is an elongated molecule, which is composed of helices and β sheets. The overall shape of this enzyme is similar to boomerang and it is divided into three domains: the two catalytic domains at each end of the molecule, and one central linker domain which is composed of several helices. The catalytic domain consists of two mixed topology β sheets which form a β sheets sandwich [47]. The

The linker domain of catechol 1,2-dioxygenases contains 12 α helices. Five of them belong to the N-terminal chain of each monomer and one extending from each catalytic domain. The

similar structure is also observed in another catechol 1,2-dioxygenases [38,50].

oligomeric complexes (αβFe3+)2-12 [66] (Fig. 3).

136 Biodegradation of Hazardous and Special Products

29.0-38.6 kDa.

**3.1. Catechol 1,2-dioxygenases characterization**

response to toxic levels of aromatic compounds [47,66].

active site with ferric ion is located between β sheets sandwich and the linker domain [47]. Matera et. al. [50] compared structure of catechol 1,2-dioxygenase from Gram- positive strain *Rhodococcus opacus* 1CP with the enzyme from *Acinetobacter calcoaceticus* described above and observed that the differences in the structure are localized at the N-terminus. In this terminal chain helix 5 is shorter, strand 1 and helix 7 are missing and a new short helix is present between position 124 and 128. Moreover the C-terminal chain of catechol 1,2-dioxygenase from Grampositive strain is shorter than in the other catechol 1,2-dioxygenases from Gram-negative bacteria.

As it was mentioned earlier the metal ion is ligated by four amino acid residues and a hydroxyl group from water in trigonal bipyramidal geometry. One molecule of tyrosine and one histidine are the axial ligands, whereas the second Tyr, His and a solvent molecule make up the equatorial plane. [38,47,50,53]. Since the two Fe3+ ions from catalytic domains are separated by more than 40 Å Vetting and Ohlendorf [47] suggested that they act structurally and catalytically independently. It explains the lack of allosteric effect [25,46,57,59,61,70,80]. However, catechol 1,2-dioxygenase from *Acinetobacter radioresistens* exhibits a sigmoidal kinetics indicating cooperation of subunits [59].

The substrate specificity of catechol 1,2-dioxygenases depends on electronic factors, which can play a significant role in determining the efficiency of conversion of catechol deriva‐ tives. Currently, there is little information about the influence of steric factors on catalyt‐ ic properties of catechol 1,2-dioxygenases [50]. Sauret-Ignazi et al.[61] revealed that very high inhibition constant is connected with steric hindrance, for example by methoxyl group. However another studies have indicated that a steric hindrances are not determining the rate-limiting step but there is a strong correlation between substituent electronegativity and kcat [25,50,61].

The catalytic cycle of catechol 1,2-dioxygenases involve binding of the catechol or its deriva‐ tives as a dianion with simultaneous dissociation of one tyrosine and water molecule (a-e). One hydroxyl group of substrate binds in an axial position *trans* to His and the other binds in an equatorial position *trans* to Tyr. The ligand position *trans* to equatorial His is unoccupied and is postulated to be the oxygen-binding pocket [28,47,50,61]. The active site geometry is converted into a square-pyramidal geometry [91]. The substrate chelates asymmetrically to the iron(III) center. The long Fe-O bonds is *trans* to a tyrosinate ligand and the short Fe-O bond is *trans* to a neutral histidine ligand [40,91]. Binding of the distal oxygen leads to species with a peroxide bridge between the ferric ion and catechol substrate [92]. At the initial stage dioxygen molecule is one-electron reduced by the catecholate ligand and in this way it formes superoxide radical anion that interacts with iron(III) (f,g). In the next step superoxide and catecholate radicals recombine forming peroxo species (h). The last can easily change its conformation because of the hydroxyl group that becomes a ketone one, which is a much weaker ligand. Arginine 457 is supposed to stabilize the carboanion produced by the C3-O bond ketonization [92]. In consequences a coordination site of the axial tyrosine is open and it allows returning of the tyrosine to the coordination sphere of iron. The tyrosine binds to iron (III) as a phenolate (i), whereas the proton either neutralizes the peroxide ligand or it is released from the active site [40,91,92].

In the first case, proton remains in the active site (i) and the peroxide oxygen atom is protonated. It prevents oxidation of iron to a high-valent Fe(IV)=O state during the O-O bond cleavage. As a result peroxide ligand undergoes a Criegee rearrangement with O-O bond heterolysis and migration of the acyl group to the peroxide oxygen (k). It is also possible that the peroxide ligand converts to muconic anhydrate product via O-O bond cleavage made easier by the equatorial tyrosine, which is oxidized to a tyrozyl radical (j). The reduction of tyrozyl radical to tyrosinate is connected with the oxyl radical attack on the carbonyl carbon [40].

In the second case, proton is released from the active site (n) and peroxide bridge is cleaved to reactive oxoferryl group and an alkoxyl radical (o). Alkoxyl radical attacks the adjacent carbonyl carbon, followed by the ring expansion and protonation of the oxo ligand leads to the cyclic anhydride (k). Hydrolysis of the anhydride leads to the final reaction product – muconic acid (l,m) (Fig. 3) [40,92].

**Figure 3.** The catalytic cycle of the intradiol dioxygenases [92].

substrate to the active site.

Sauret-Ignazi et al. [61] observed that monophenolic compounds are also bound by enzyme and they suggested that only one deprotonated hydroxyl may be sufficient for binding of the

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

139

Matera et al. [50] calculated parameter for nucleophilic reactivity for catechol, 3-chloro-, 4 chloro-, 3-methyl-, 4-methylcatechols, and pyrogallol. They revealed that the interaction between the frontier orbitals, the highest occupied molecular orbital (HOMO) of nucleophiles (catechols) and the lowest unoccupied molecular orbital (LUMO) of electrophile (the active site iron ion), is essential and lowers the activation barrier. Since the nuclephilic attack of the

The electrophilic attack of molecular oxygen on iron (III)-bound catecholate is judged to be an essential step in the catalytic reaction of catechol 1,2-dioxygenase [50]. For this type of chemical reaction the σ-complex is assumed to be next to the transition state. The formation of σ-complex requires the highest free energy of activation and must consequent‐ ly be rate-limiting [25].

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation http://dx.doi.org/10.5772/56205 139

**Figure 3.** The catalytic cycle of the intradiol dioxygenases [92].

ic properties of catechol 1,2-dioxygenases [50]. Sauret-Ignazi et al.[61] revealed that very high inhibition constant is connected with steric hindrance, for example by methoxyl group. However another studies have indicated that a steric hindrances are not determining the rate-limiting step but there is a strong correlation between substituent electronegativity and

The catalytic cycle of catechol 1,2-dioxygenases involve binding of the catechol or its deriva‐ tives as a dianion with simultaneous dissociation of one tyrosine and water molecule (a-e). One hydroxyl group of substrate binds in an axial position *trans* to His and the other binds in an equatorial position *trans* to Tyr. The ligand position *trans* to equatorial His is unoccupied and is postulated to be the oxygen-binding pocket [28,47,50,61]. The active site geometry is converted into a square-pyramidal geometry [91]. The substrate chelates asymmetrically to the iron(III) center. The long Fe-O bonds is *trans* to a tyrosinate ligand and the short Fe-O bond is *trans* to a neutral histidine ligand [40,91]. Binding of the distal oxygen leads to species with a peroxide bridge between the ferric ion and catechol substrate [92]. At the initial stage dioxygen molecule is one-electron reduced by the catecholate ligand and in this way it formes superoxide radical anion that interacts with iron(III) (f,g). In the next step superoxide and catecholate radicals recombine forming peroxo species (h). The last can easily change its conformation because of the hydroxyl group that becomes a ketone one, which is a much weaker ligand. Arginine 457 is supposed to stabilize the carboanion produced by the C3-O bond ketonization [92]. In consequences a coordination site of the axial tyrosine is open and it allows returning of the tyrosine to the coordination sphere of iron. The tyrosine binds to iron (III) as a phenolate (i), whereas the proton either neutralizes the peroxide ligand or it is released

In the first case, proton remains in the active site (i) and the peroxide oxygen atom is protonated. It prevents oxidation of iron to a high-valent Fe(IV)=O state during the O-O bond cleavage. As a result peroxide ligand undergoes a Criegee rearrangement with O-O bond heterolysis and migration of the acyl group to the peroxide oxygen (k). It is also possible that the peroxide ligand converts to muconic anhydrate product via O-O bond cleavage made easier by the equatorial tyrosine, which is oxidized to a tyrozyl radical (j). The reduction of tyrozyl radical to tyrosinate is connected with the oxyl radical attack on

In the second case, proton is released from the active site (n) and peroxide bridge is cleaved to reactive oxoferryl group and an alkoxyl radical (o). Alkoxyl radical attacks the adjacent carbonyl carbon, followed by the ring expansion and protonation of the oxo ligand leads to the cyclic anhydride (k). Hydrolysis of the anhydride leads to the final reaction product –

The electrophilic attack of molecular oxygen on iron (III)-bound catecholate is judged to be an essential step in the catalytic reaction of catechol 1,2-dioxygenase [50]. For this type of chemical reaction the σ-complex is assumed to be next to the transition state. The formation of σ-complex requires the highest free energy of activation and must consequent‐

kcat [25,50,61].

138 Biodegradation of Hazardous and Special Products

from the active site [40,91,92].

the carbonyl carbon [40].

ly be rate-limiting [25].

muconic acid (l,m) (Fig. 3) [40,92].

Sauret-Ignazi et al. [61] observed that monophenolic compounds are also bound by enzyme and they suggested that only one deprotonated hydroxyl may be sufficient for binding of the substrate to the active site.

Matera et al. [50] calculated parameter for nucleophilic reactivity for catechol, 3-chloro-, 4 chloro-, 3-methyl-, 4-methylcatechols, and pyrogallol. They revealed that the interaction between the frontier orbitals, the highest occupied molecular orbital (HOMO) of nucleophiles (catechols) and the lowest unoccupied molecular orbital (LUMO) of electrophile (the active site iron ion), is essential and lowers the activation barrier. Since the nuclephilic attack of the aromatic π electrons of the iron(III)-bound catecholate on the oxygen molecule is an important step in the cleavage reaction, the HOMO of the catecholate is assumed to be the relevant frontier orbital. The natural logarithm of the kcat is proportional to the activation energy of the ratelimiting step and is correlated to the energy of HOMO [50].

yquinol 1,2-dioxygenases responsible for substrate selection were selected: Leu80, Asp83,

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

141

One of the most characteristic feature of hydroxyquinol 1,2-dioxygenases is the extensive opening of the upper part of catalytic cavity which is responsible for favorable binding of hydroxyquinol [17,83]. Zaborina et al. [83] connected this fact with lower activation energy. Catalytic process of *ortho* cleavage proceeds through semiquinone radical. This intermediate can be stabilized by mesomeric delocalization of the unpaired electron in the 2-, 4-, or 6-position of semiquinone ring. With a hydroxyquinol as a substrate additional mesomeric delocalization of the unpaired electron is possible in position 1. The increase in the semiquinone character

Protocatechuate 3,4-dioxygenases catalyze cleavage of protocatechuic acid and its derivatives. The mechanism of this reaction is the same as intradiol fission of catechol ring [13,92]. During substrate binding as a dianion to the active site of the enzyme the axial tyrosine is dissociated from the iron(III). It causes conformational changes that yields the chelated iron(III)-protoca‐ techuate complex. Vetting et al. [16] suggested that the function of the structural rearrange‐ ments would be to facilitate the reaction of oxygen with the substrate. Electron transfer from the substrate to dioxygen aids a binding of dioxygen to the iron(III). The activation of substrate for an electrophilic attack by dioxygen leads to the formation of a peroxo bridge between the iron and C4 of substrate. O-O bond cleavage occurs after acyl migration to the peroxo oxygen. It leads to the cyclic anhydride formation and its hydrolysis to the final acyclic product

Although protocatechuate 3,4-dioxygenases are heterodimers with molecular mass of 97.0-700.0 kDa [12,18,72,75,82] theirs tertiary structure is similar to the enzymes from catechol 1,2-dioxygenase family. Protocatechuate 3,4-dioxygenases belong to the non-heme iron dioxygenases and are composed of equimolar amounts of two different α and β subunits [11, 68,78,82,93]. These enzymes have a tetrahedral shape with α subunit forming the apex of the tetrahedron. The central cavity is accessible to solvent through triangular channels. A number of basic residues (mainly arginine and lysine) line the entrance to the active site of the enzyme producing regions of positive electrostatic potential. It is possible that these residues funnel

α and β subunits have similar folds. The core of each subunit contains two four-strand β-sheets that form a β sandwich. One β sheet consists of antiparallel strands, whereas the other has a mixed topology. The core structure is surrounded by series of small helices and large irregular loops. Two β sheets form a part of the interface between the subunits, with surrounding loops providing the remaining contacts. The amino terminus of the α subunit passes completely through a gap between the core β sandwich, supports structure of the β subunit and makes

Aligments of the structure and sequences of α- and β-chain of protocatechuate 3,4-dioxyge‐ nases from more than 26 bacterial strains revealed that the β-chain show higher sequence

Val107, Phe108, Gly109, Pro110, Phe111, Ile199, Pro200, Arg218, and Val251 [17].

leads to a reduction of activation energy [83].

[16,80,92,93].

**3.3. Protocatechuate 3,4-dioxygenases characterization**

the negatively charged substrate into the active site [16].

up one wall of the active-site cavity [16].

The main interaction of catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus* ADP1 with the substrate involves residues Leu73, Pro76, Ile105, Pro108, Leu109, Arg221, Phe253 and Ala254. Vetting and Ohlendorf [47] postulate that Arg221 may play an important role in proper positioning of the substrate's hydroxyl groups to the iron by van der Waals interactions with the aromatic ring. These interactions prevent ring rotation into two equatorial positions [47]. Matera et al. [50] observed that Arg217, Gln236 which play role in the deprotonation of substrate, together with Gly104, Pro105 and Tyr106 are required for the proper placing of aromatic compounds into the active site of catechol 1,2-dioxygenase from Gram- positive strain 1CP. Moreover residue Tyr106 forms a hydrogen bond with Tyr196 during its dissociation from the iron after the binding of catechol.

Catechol derivatives with the substituents on the C3 position can be bound in two different orientations in the enzyme active site from ADP1 strain. The substituent may be located in a pocket created by residues Ile105, Pro108, His226, Gln240 and Arg221 or its positioning may lead to a steric conflict with Tyr200. However, rearrangement in the active site makes possible the interaction of substituent on C3 with the π electrons of Tyr200 [47]. According to Matera et al. [50] the lower affinity of catechol 1,2-dioxygenase from 1CP strain for 3-substituents results from the lack of Tyr106. Tyr 196 is not connected then and it can cause the more open conformation of the space to the 3-substituent binding. It influences a certain degree of disorder in substrates binding to the iron and/or a not suitable coordination for productive conversion.

Ferraroni et al. [66] showed that preference of chlorocatechol dioxygenases for 3-chlorocate‐ chol is connected with Val53 and Tyr78 residues whereas 4-chlorocatechols are cleaved by dioxygenases with Leu and Ile at 49 and 74 position respectively. Moreover, cysteine residues are important for interacting with chlorine substituents.

Presence of large substituents makes impossible active binding of the aromatic substrates to the active site because they interact unfavorably with Pro76, Leu73 and Ile105. Vetting and Ohlendorf [47] suggest that catechol derivatives are ligated the iron through only one of the hydroxyl group or they are trapped in an initial binding state.

#### **3.2. Hydroxyquinol 1,2-dioxygenases characterization**

Hydroxyquinol 1,2-dioxygenases have very similar structure and reaction mechanism to catechol 1,2-dioxygenases. They form homodimers with molecular weight of 58-85 kDa. The iron (III) is coordinated by two molecules of tyrosine, two molecules of histidine and one molecule of water [17,41,84-86,90].

However, Ferraroni et al. [17] revealed that His42 from H2 helix of the linker domain is connected with the copper ion in an oxidation state of +1. Probably it plays a role in stabilization of the enzyme quaternary structure. Moreover, a few different amino acid residues of hydrox‐ yquinol 1,2-dioxygenases responsible for substrate selection were selected: Leu80, Asp83, Val107, Phe108, Gly109, Pro110, Phe111, Ile199, Pro200, Arg218, and Val251 [17].

One of the most characteristic feature of hydroxyquinol 1,2-dioxygenases is the extensive opening of the upper part of catalytic cavity which is responsible for favorable binding of hydroxyquinol [17,83]. Zaborina et al. [83] connected this fact with lower activation energy. Catalytic process of *ortho* cleavage proceeds through semiquinone radical. This intermediate can be stabilized by mesomeric delocalization of the unpaired electron in the 2-, 4-, or 6-position of semiquinone ring. With a hydroxyquinol as a substrate additional mesomeric delocalization of the unpaired electron is possible in position 1. The increase in the semiquinone character leads to a reduction of activation energy [83].

#### **3.3. Protocatechuate 3,4-dioxygenases characterization**

aromatic π electrons of the iron(III)-bound catecholate on the oxygen molecule is an important step in the cleavage reaction, the HOMO of the catecholate is assumed to be the relevant frontier orbital. The natural logarithm of the kcat is proportional to the activation energy of the rate-

The main interaction of catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus* ADP1 with the substrate involves residues Leu73, Pro76, Ile105, Pro108, Leu109, Arg221, Phe253 and Ala254. Vetting and Ohlendorf [47] postulate that Arg221 may play an important role in proper positioning of the substrate's hydroxyl groups to the iron by van der Waals interactions with the aromatic ring. These interactions prevent ring rotation into two equatorial positions [47]. Matera et al. [50] observed that Arg217, Gln236 which play role in the deprotonation of substrate, together with Gly104, Pro105 and Tyr106 are required for the proper placing of aromatic compounds into the active site of catechol 1,2-dioxygenase from Gram- positive strain 1CP. Moreover residue Tyr106 forms a hydrogen bond with Tyr196 during its dissociation

Catechol derivatives with the substituents on the C3 position can be bound in two different orientations in the enzyme active site from ADP1 strain. The substituent may be located in a pocket created by residues Ile105, Pro108, His226, Gln240 and Arg221 or its positioning may lead to a steric conflict with Tyr200. However, rearrangement in the active site makes possible the interaction of substituent on C3 with the π electrons of Tyr200 [47]. According to Matera et al. [50] the lower affinity of catechol 1,2-dioxygenase from 1CP strain for 3-substituents results from the lack of Tyr106. Tyr 196 is not connected then and it can cause the more open conformation of the space to the 3-substituent binding. It influences a certain degree of disorder in substrates binding to the iron and/or a not suitable coordination for productive conversion.

Ferraroni et al. [66] showed that preference of chlorocatechol dioxygenases for 3-chlorocate‐ chol is connected with Val53 and Tyr78 residues whereas 4-chlorocatechols are cleaved by dioxygenases with Leu and Ile at 49 and 74 position respectively. Moreover, cysteine residues

Presence of large substituents makes impossible active binding of the aromatic substrates to the active site because they interact unfavorably with Pro76, Leu73 and Ile105. Vetting and Ohlendorf [47] suggest that catechol derivatives are ligated the iron through only one of the

Hydroxyquinol 1,2-dioxygenases have very similar structure and reaction mechanism to catechol 1,2-dioxygenases. They form homodimers with molecular weight of 58-85 kDa. The iron (III) is coordinated by two molecules of tyrosine, two molecules of histidine and one

However, Ferraroni et al. [17] revealed that His42 from H2 helix of the linker domain is connected with the copper ion in an oxidation state of +1. Probably it plays a role in stabilization of the enzyme quaternary structure. Moreover, a few different amino acid residues of hydrox‐

limiting step and is correlated to the energy of HOMO [50].

from the iron after the binding of catechol.

140 Biodegradation of Hazardous and Special Products

are important for interacting with chlorine substituents.

**3.2. Hydroxyquinol 1,2-dioxygenases characterization**

molecule of water [17,41,84-86,90].

hydroxyl group or they are trapped in an initial binding state.

Protocatechuate 3,4-dioxygenases catalyze cleavage of protocatechuic acid and its derivatives. The mechanism of this reaction is the same as intradiol fission of catechol ring [13,92]. During substrate binding as a dianion to the active site of the enzyme the axial tyrosine is dissociated from the iron(III). It causes conformational changes that yields the chelated iron(III)-protoca‐ techuate complex. Vetting et al. [16] suggested that the function of the structural rearrange‐ ments would be to facilitate the reaction of oxygen with the substrate. Electron transfer from the substrate to dioxygen aids a binding of dioxygen to the iron(III). The activation of substrate for an electrophilic attack by dioxygen leads to the formation of a peroxo bridge between the iron and C4 of substrate. O-O bond cleavage occurs after acyl migration to the peroxo oxygen. It leads to the cyclic anhydride formation and its hydrolysis to the final acyclic product [16,80,92,93].

Although protocatechuate 3,4-dioxygenases are heterodimers with molecular mass of 97.0-700.0 kDa [12,18,72,75,82] theirs tertiary structure is similar to the enzymes from catechol 1,2-dioxygenase family. Protocatechuate 3,4-dioxygenases belong to the non-heme iron dioxygenases and are composed of equimolar amounts of two different α and β subunits [11, 68,78,82,93]. These enzymes have a tetrahedral shape with α subunit forming the apex of the tetrahedron. The central cavity is accessible to solvent through triangular channels. A number of basic residues (mainly arginine and lysine) line the entrance to the active site of the enzyme producing regions of positive electrostatic potential. It is possible that these residues funnel the negatively charged substrate into the active site [16].

α and β subunits have similar folds. The core of each subunit contains two four-strand β-sheets that form a β sandwich. One β sheet consists of antiparallel strands, whereas the other has a mixed topology. The core structure is surrounded by series of small helices and large irregular loops. Two β sheets form a part of the interface between the subunits, with surrounding loops providing the remaining contacts. The amino terminus of the α subunit passes completely through a gap between the core β sandwich, supports structure of the β subunit and makes up one wall of the active-site cavity [16].

Aligments of the structure and sequences of α- and β-chain of protocatechuate 3,4-dioxyge‐ nases from more than 26 bacterial strains revealed that the β-chain show higher sequence identities than the α-chain. It may be connected with the fact that β-chain provides most of the active site [19]. However, amino acids from α-subunit may interact with substrate and in this way stabilize it in the active site. Harnett et al. [69] observed that Arg-133 from α-subunit participates in forming ionic bond with carboxyl group of protocatechuic acid.

the optimal conditions for the microbial growth are disadvantages of biodegradation proc‐ esses. The direct application of enzymes in the environmental treatment processes has been quite limited due to the loss of enzyme activity [95] and therefore, novel methods of enzyme stabilization are developed. One of them is immobilization which has been used as a tool to improve many of enzyme properties such as operational stability, inhibitor resistance and performance in organic solvents [95,96]. Kalogeris et al. [96] showed that immobilization of catechol 1,2-dioxygenase improved its performance at higher temperatures. It extended catechol 1,2-dioxygenase storage stability and retained its activity after operation cycles properties which are fundamental for continuous bioprocesses during removal of toxic pollutants. Catechol 1,2-dioxygenase from *Acinetobacter radioresistens* S13 after immobilization on nanosponges showed the ability to convert the substrate, after 50 reaction cycles. Moreover,

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

143

Immobilized intradiol dioxygenases can also be used as biosensors because of increased interest in the detection of aromatic compounds. Zucolotto et al. [97] successfully fabricated biosensor for catechol detection. Chlorocatechol 1,2-dioxygenase was immobilized in nano‐ structured films in conjuction with poly(amidoamine) dendrimer in a layer-by-layer fashion. Chlorocatechol 1,2-dioxygenase remained active on these films for longer than three weeks. Moreover, this biosensor was able to detect catechol at concentration even 10-10 M [97].

Polychlorinated phenols, amino- and nitrophenols have become widespread environmental pollutants because of their resistance to microbial degradation and their broad toxicity. During their degradation hydroxyquinol as a key intermediate is frequently formed. Therefore, it is believed that hydroxyquinol-metabolized enzymes play an important role in the microbial degradation of various aromatic xenobiotics [51,85,88]. Immobilization of hydroxyquinol 1,2 dioxygenase from *Arthrobacter chlorophenolicus* A6 onto single-walled carbon nanotubes

The important properties of degradation enzymes is a resistance to inhibitors such as metal ions, alcohols, chelators, hydrocarbons, phenols and other [58,59,61]. Metal ions can lead to the conformational changes in the enzymes such as a reduction in α-helices and β sheets, which can result in the loss of enzymatic activity [98] or replacement of the original metal ion at the active site of the enzymes with other metal. It may modulate their activities in accordance with the Irving-Williams studies on the stability of the metal complexes [99]. Moreover, the thiol groups of enzyme structure may bind transition metals that can deactivate the enzyme.

Gou et al. [67] observed that activity of catechol 1,2-dioxygenase from *Sphingomonas*

Hg2+, Ni2+ and Co2+ slow down the enzyme reaction. Only Cu2+, Zn2+ and Fe2+ ions strongly inhibited this enzyme. Catechol 1,2-dioxygenase from *Pseudomonas aeruginosa* was not sensitive to the presence of Mg2+, Fe2+, and Ca2+. It was inhibited by Mn2+, Cu2+, and Ag+ [64]. Ag+ and Hg2+ showed the highest inhibitory effect on catechol 1,2-dioxygenase isozymes from *Arthrobacter* sp. BA-5-17. Catechol 1,2-dioxygenase from *Rhodococcus* sp. NCIM 2891 was inhibited completely in the presence of Fe3+, Cu2+ and Hg2+ [20,62]. In contrary, catechol 1,2-dioxygenase from *Gordonia polyisoprenivorans* was resistant to these cations [21]. Decreased sensitivity of both catechol 1,2-dioxygenase and protocatechuate

, Fe3+ and Ca2+, whereas the addition of Mn2+,

increased its resistance capacity to the variable environmental factors [95].

this enzyme was more thermostable [60].

*xenophaga* QYY was increased by Pb2+, Mg2+, K+

Protocatechuate3,4-dioxygenases fromdifferentbacterial strainsare consistofvariablenumber of αβ protomers (2 – 12) and each protomer has a cylindrical shape [12,16,78]. The minimal catalytic unit of all protocatechuate 3,4-dioxygenases is an αβFe3+ structure, but (αβ)2Fe3+ structure has been also reported [11,72]. The crystal structure of these enzymes shows that the high-spin iron(III) is bound to the active site in a distorted trigonal bipyramidal coordination geometry with two inequivalenttyrosine ligands,two histidines, andexogenous hydroxide ion [68,69,73,77,79,92,93].Kurahashi et al.[93]proposedthatthe electronic effect ofthewaterligand in the active site of protocatechuate 3,4-dioxygenase controls not only the distorted trigonalbipyramidal structurebutalsoplaysaroleintheoxidationof substrate.Thetwotyrosine ligands are thought to stabilize the ferric ion and give protocatechuate 3,4-dioxygenase its characteris‐ tic burgundy-red color via tyrosinate ligand–to-metal charge transfer [13,79].

Orville et al. [94] demonstrated that at least two iron binding sites can be occupied by exoge‐ nous ligand such as substrates, analog of substrate, and small molecules. It leads to the dynamic changes in the iron coordination sphere or the orientation of the aromatic substrate and influences the catalytic cycle. An external ligand with a higher position in the spectrochemical series (Cl«RO<H2O) induces a larger degree of distortion [80,93].

The protocatechuate 3,4-dioxygenases show a high similarity not only for the ligands of the active site but also for the whole pocket in which the iron complex is located [77].

Orville et al. [94] studied cleavage mechanism in the presence of inhibitors. They proposed three different orientations of inhibitors in the active site. First, in which inhibitor protrudes out of the active site cavity but still coordinates the iron to yield trigonal bipyramidal coordi‐ nation geometry. Second, in which inhibitor binds within the active site that is sufficient to dissociate the active site solvent molecules but without direct coordination to the iron. Third, in which inhibitor is completely bound within the active site, directly coordinates the iron, and yields octahedral geometry.

Many protocatechuate dioxygenases have narrow substrate specificities and regioselectivities [9,10,12]. Hou et al. [10] suggested that free sulfhydryl groups located at or near the active center are responsible for substrate specificity of the enzyme. It was showed that protocate‐ chuic acid derivatives are cleaved much slower than protocatechuic acid. Transient kinetic results suggest that slower fission is probably connected with the reduced rates for substrate association and product release rather than the ring opening reaction [13].

#### **4. Application of intradiol dioxygenases in bioremediation**

Bioremediation is an effective treatment process that use organisms through their enzymatic activities. However, slow growth of microorganisms or difficulties in the control and maintain the optimal conditions for the microbial growth are disadvantages of biodegradation proc‐ esses. The direct application of enzymes in the environmental treatment processes has been quite limited due to the loss of enzyme activity [95] and therefore, novel methods of enzyme stabilization are developed. One of them is immobilization which has been used as a tool to improve many of enzyme properties such as operational stability, inhibitor resistance and performance in organic solvents [95,96]. Kalogeris et al. [96] showed that immobilization of catechol 1,2-dioxygenase improved its performance at higher temperatures. It extended catechol 1,2-dioxygenase storage stability and retained its activity after operation cycles properties which are fundamental for continuous bioprocesses during removal of toxic pollutants. Catechol 1,2-dioxygenase from *Acinetobacter radioresistens* S13 after immobilization on nanosponges showed the ability to convert the substrate, after 50 reaction cycles. Moreover, this enzyme was more thermostable [60].

identities than the α-chain. It may be connected with the fact that β-chain provides most of the active site [19]. However, amino acids from α-subunit may interact with substrate and in this way stabilize it in the active site. Harnett et al. [69] observed that Arg-133 from α-subunit

Protocatechuate3,4-dioxygenases fromdifferentbacterial strainsare consistofvariablenumber of αβ protomers (2 – 12) and each protomer has a cylindrical shape [12,16,78]. The minimal catalytic unit of all protocatechuate 3,4-dioxygenases is an αβFe3+ structure, but (αβ)2Fe3+ structure has been also reported [11,72]. The crystal structure of these enzymes shows that the high-spin iron(III) is bound to the active site in a distorted trigonal bipyramidal coordination geometry with two inequivalenttyrosine ligands,two histidines, andexogenous hydroxide ion [68,69,73,77,79,92,93].Kurahashi et al.[93]proposedthatthe electronic effect ofthewaterligand in the active site of protocatechuate 3,4-dioxygenase controls not only the distorted trigonalbipyramidal structurebutalsoplaysaroleintheoxidationof substrate.Thetwotyrosine ligands are thought to stabilize the ferric ion and give protocatechuate 3,4-dioxygenase its characteris‐

Orville et al. [94] demonstrated that at least two iron binding sites can be occupied by exoge‐ nous ligand such as substrates, analog of substrate, and small molecules. It leads to the dynamic changes in the iron coordination sphere or the orientation of the aromatic substrate and influences the catalytic cycle. An external ligand with a higher position in the spectrochemical

The protocatechuate 3,4-dioxygenases show a high similarity not only for the ligands of the

Orville et al. [94] studied cleavage mechanism in the presence of inhibitors. They proposed three different orientations of inhibitors in the active site. First, in which inhibitor protrudes out of the active site cavity but still coordinates the iron to yield trigonal bipyramidal coordi‐ nation geometry. Second, in which inhibitor binds within the active site that is sufficient to dissociate the active site solvent molecules but without direct coordination to the iron. Third, in which inhibitor is completely bound within the active site, directly coordinates the iron, and

Many protocatechuate dioxygenases have narrow substrate specificities and regioselectivities [9,10,12]. Hou et al. [10] suggested that free sulfhydryl groups located at or near the active center are responsible for substrate specificity of the enzyme. It was showed that protocate‐ chuic acid derivatives are cleaved much slower than protocatechuic acid. Transient kinetic results suggest that slower fission is probably connected with the reduced rates for substrate

Bioremediation is an effective treatment process that use organisms through their enzymatic activities. However, slow growth of microorganisms or difficulties in the control and maintain

active site but also for the whole pocket in which the iron complex is located [77].

association and product release rather than the ring opening reaction [13].

**4. Application of intradiol dioxygenases in bioremediation**

participates in forming ionic bond with carboxyl group of protocatechuic acid.

tic burgundy-red color via tyrosinate ligand–to-metal charge transfer [13,79].

series (Cl«RO<H2O) induces a larger degree of distortion [80,93].

yields octahedral geometry.

142 Biodegradation of Hazardous and Special Products

Immobilized intradiol dioxygenases can also be used as biosensors because of increased interest in the detection of aromatic compounds. Zucolotto et al. [97] successfully fabricated biosensor for catechol detection. Chlorocatechol 1,2-dioxygenase was immobilized in nano‐ structured films in conjuction with poly(amidoamine) dendrimer in a layer-by-layer fashion. Chlorocatechol 1,2-dioxygenase remained active on these films for longer than three weeks. Moreover, this biosensor was able to detect catechol at concentration even 10-10 M [97].

Polychlorinated phenols, amino- and nitrophenols have become widespread environmental pollutants because of their resistance to microbial degradation and their broad toxicity. During their degradation hydroxyquinol as a key intermediate is frequently formed. Therefore, it is believed that hydroxyquinol-metabolized enzymes play an important role in the microbial degradation of various aromatic xenobiotics [51,85,88]. Immobilization of hydroxyquinol 1,2 dioxygenase from *Arthrobacter chlorophenolicus* A6 onto single-walled carbon nanotubes increased its resistance capacity to the variable environmental factors [95].

The important properties of degradation enzymes is a resistance to inhibitors such as metal ions, alcohols, chelators, hydrocarbons, phenols and other [58,59,61]. Metal ions can lead to the conformational changes in the enzymes such as a reduction in α-helices and β sheets, which can result in the loss of enzymatic activity [98] or replacement of the original metal ion at the active site of the enzymes with other metal. It may modulate their activities in accordance with the Irving-Williams studies on the stability of the metal complexes [99]. Moreover, the thiol groups of enzyme structure may bind transition metals that can deactivate the enzyme.

Gou et al. [67] observed that activity of catechol 1,2-dioxygenase from *Sphingomonas xenophaga* QYY was increased by Pb2+, Mg2+, K+ , Fe3+ and Ca2+, whereas the addition of Mn2+, Hg2+, Ni2+ and Co2+ slow down the enzyme reaction. Only Cu2+, Zn2+ and Fe2+ ions strongly inhibited this enzyme. Catechol 1,2-dioxygenase from *Pseudomonas aeruginosa* was not sensitive to the presence of Mg2+, Fe2+, and Ca2+. It was inhibited by Mn2+, Cu2+, and Ag+ [64]. Ag+ and Hg2+ showed the highest inhibitory effect on catechol 1,2-dioxygenase isozymes from *Arthrobacter* sp. BA-5-17. Catechol 1,2-dioxygenase from *Rhodococcus* sp. NCIM 2891 was inhibited completely in the presence of Fe3+, Cu2+ and Hg2+ [20,62]. In contrary, catechol 1,2-dioxygenase from *Gordonia polyisoprenivorans* was resistant to these cations [21]. Decreased sensitivity of both catechol 1,2-dioxygenase and protocatechuate

3,4-dioxygenase from *Stenotrophomonas maltophilia* KB2 to metal ions was observed by Guzik et al. [68]. The presence of Mn2+, Ni2+, Cd2+ caused only 20-40% reduction of protocatechuate 3,4-dioxygenase activity, whereas, catechol 1,2-dioxygenase was even activated by Cd2+, Co2+, Zn2+, Cu2+, and Fe2+ [68]. A high resistance of the intradiol dioxyge‐ nases to the metal ions is especially important because these ions are often found togeth‐ er with aromatic xenobiotics in the environment.

**References**

2009;4(1) 29-38.

Biotechnology 2010;26 289-295.

late\_matter.pdf (accessed 27.11.2012).

Microbiology 1984;47(3) 500-505.

Sciences 2009;3(2) 776-783.

27-37.

340-347.

[1] Malawska M, Ekonomiuk A, Wiłkomirski B. Polycyclic aromatic hydrocarbons in peat cores from southern Poland: distribution in stratigraphic profiles as an indicator

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

145

[2] Li J, Yuan H, Yang J. Bacteria and lignin degradation. Frontiers of Biology in China

[3] Greń I, Wojcieszyńska D, Guzik U, Perkosz M, Hupert-Kocurek K. Enhanced bio‐ transformation of mononitrophenols by *Stenotrophomonas maltophilia* KB2 in the pres‐ ence of aromatic compounds of plant origin. World Journal of Microbiology and

[4] Lee B-K, Vu VT. Sources, distribution and toxicity of polycyclic aromatic hydrocar‐ bons (PAHs) in particulate matter. In: Villanyi V. (ed.) Air Pollution. Rijeka: InTech; 2010. p.99-122. Available from http://cdn.intechopen.com/pdfs/11767/InTech-Sour‐ ces\_distribution\_and\_toxicity\_of\_polyaromatic\_hydrocarbons\_pahs\_in\_particu‐

[5] Bartels I, Knackmuss H-J, Reineke W. Suicide inactivation of catechol 2,3-dioxyge‐ nase from *Pseudomonas putida* mt-2 by 3-halocatechols. Applied and Environmental

[6] El-Sayed WS, Ismaeil M, El-Beih F. Isolation of 4-chlorophenol-degrading bacteria, *Bacillus subtilis* OS1 and *Alcaligenes* sp. OS2 from petroleum oil-conatminated soil and characterization of its catabolic pathway. Australian Journal of Basic and Applied

[7] Li X, Zhang T, Min X, Liu P. Toxicity of aromatic compounds to *Tetrahymena* estimat‐

[8] Zhang S, Sun W, Xu L, Zheng X, Chu X, Tian J, Wu N, Fan Y. Identification of the *para*-nitrophenol catabolic pathway, and characterization of three enzymes involved in the hydroquinone pathway, in *pseudomonas* sp. 1-7. BMC Microbiology 2012;12

[9] Fujisawa H, Hayaishi O. Protocatechuate 3,4-dioxygenase. Crystallization and char‐

[10] Hou ChT, Lillard MO, Schwartz RD. Protocatechuate 3,4-dioxygenase from *Acineto‐*

[11] Sterjiades R, Pelmont J. Occurrence of two different forms of protocatechuate 3,4-di‐ oxygenase in a *Moraxella* sp. Applied and Environmental Microbiology 1989;55(2)

ed by microcalorimetry and QSAR. Aquatic Toxicology 2010;98 322-327.

acterization. The Journal of Biological Chemistry 1968;243(10) 2673-2681.

*bacter calcoaceticus*. Biochemistry 1976;15(3) 582-588.

of PAH sources. Mires and Peat 2006;1 1-14.

Some catechol 1,2-dioxgenases can be inhibited by chelating agent, especially by tiron – ferric ion chelating agent [58]. However, catechol 1,2-dioxygenase from *Arthrobacter* sp. BA-5-17 is not sensitive to chelating and sulfhydryl agents [62].

The wide substrate specificity and inhibitor resistance of some intradiol dioxygenases makes them an attractive target in the design of enzyme systems for bioremediation.

#### **5. Conclusion**

Intradiol dioxygenases are responsible for aromatic ring cleavage and for that reason are directly involved in biogeochemical cycles, and can be very useful in the development of bioremediation technology. These enzymes are the members of the aromatic-ring-cleav‐ age dioxygenase superfamily. Based on their substrate specificity intradiol dioxygenases could be divided into three classes: catechol 1,2-dioxygenases, protocatechuate 3,4-dioxyge‐ nases, and hydroxyquinol 1,2-dioxygenase. The mechanism of intradiol fisson is common for enzymes from all clasess. In this mechanism the high-spin iron(III) is ligated by two tyrosines, two histidines, and a water ligand in trigonal bipyramidal geometry. The catalytic cycle of the intradiol dioxygenases involve binding of the catechol as a dianion and dioxygen to the metal. It leads to the formation of a peroxo bridge between iron and C4 of substrate. In the next step the Criegee rearrangement and O-O bond cleavage occurs, leading to the cyclic anhydride formation. Hydrolysis of the anhydride leads to the formation of the final acyclic product. Structural analysis of many of intradiol dioxygenas‐ es revealed only small differences in their active site structure. However, these differen‐ ces significantly influence the substrate specificity and the resistance of the enzymes to the changing environmental factors.

#### **Author details**

Urszula Guzik\* , Katarzyna Hupert-Kocurek and Danuta Wojcieszyńska

\*Address all correspondence to: urszula.guzik@us.edu.pl

University of Silesia in Katowice, Faculty of Biology and Environmental Protection, Depart‐ ment of Biochemistry, Katowice, Poland

#### **References**

3,4-dioxygenase from *Stenotrophomonas maltophilia* KB2 to metal ions was observed by Guzik et al. [68]. The presence of Mn2+, Ni2+, Cd2+ caused only 20-40% reduction of protocatechuate 3,4-dioxygenase activity, whereas, catechol 1,2-dioxygenase was even activated by Cd2+, Co2+, Zn2+, Cu2+, and Fe2+ [68]. A high resistance of the intradiol dioxyge‐ nases to the metal ions is especially important because these ions are often found togeth‐

Some catechol 1,2-dioxgenases can be inhibited by chelating agent, especially by tiron – ferric ion chelating agent [58]. However, catechol 1,2-dioxygenase from *Arthrobacter* sp. BA-5-17 is

The wide substrate specificity and inhibitor resistance of some intradiol dioxygenases makes

Intradiol dioxygenases are responsible for aromatic ring cleavage and for that reason are directly involved in biogeochemical cycles, and can be very useful in the development of bioremediation technology. These enzymes are the members of the aromatic-ring-cleav‐ age dioxygenase superfamily. Based on their substrate specificity intradiol dioxygenases could be divided into three classes: catechol 1,2-dioxygenases, protocatechuate 3,4-dioxyge‐ nases, and hydroxyquinol 1,2-dioxygenase. The mechanism of intradiol fisson is common for enzymes from all clasess. In this mechanism the high-spin iron(III) is ligated by two tyrosines, two histidines, and a water ligand in trigonal bipyramidal geometry. The catalytic cycle of the intradiol dioxygenases involve binding of the catechol as a dianion and dioxygen to the metal. It leads to the formation of a peroxo bridge between iron and C4 of substrate. In the next step the Criegee rearrangement and O-O bond cleavage occurs, leading to the cyclic anhydride formation. Hydrolysis of the anhydride leads to the formation of the final acyclic product. Structural analysis of many of intradiol dioxygenas‐ es revealed only small differences in their active site structure. However, these differen‐ ces significantly influence the substrate specificity and the resistance of the enzymes to the

, Katarzyna Hupert-Kocurek and Danuta Wojcieszyńska

University of Silesia in Katowice, Faculty of Biology and Environmental Protection, Depart‐

\*Address all correspondence to: urszula.guzik@us.edu.pl

ment of Biochemistry, Katowice, Poland

them an attractive target in the design of enzyme systems for bioremediation.

er with aromatic xenobiotics in the environment.

144 Biodegradation of Hazardous and Special Products

not sensitive to chelating and sulfhydryl agents [62].

**5. Conclusion**

changing environmental factors.

**Author details**

Urszula Guzik\*


[12] Hammer A, Stolz A, Knackmuss H-J. Purification and characterization of a novel type of protocatechuate 3,4-dioxyegnase with the ability to oxidize 4-sulfocatechol. Archives of Microbiology 1996;166 92-100.

[24] Hu X-F, Wu L. Oxidation of 3,5-di-tert-butylcatechol in the presence of V-polyoxo‐

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

147

[25] Dorn E, Knackmuss H-J. Chemical structure and biodegradability of halogenated ar‐

[26] Pitter P. Correlation of microbial degradation rates with the chemical structure. Acta

[27] Kovacs A, Hargittai I. Theoretical investigation of the additivity of structural sub‐ stituent effects in benzene derivatives. Structural Chemistry 2000;11(2-3) 193-201.

[28] Pau MYM, Lipscomb JD, Solomon EI. Substrate activation for O2 reactions by oxi‐ dized metal centers in biology. Proceedings of the National Academy of Sciences

[29] Divari S, Valetti F. Caposito P, Pessione E, Cavaletto M, Griva E, Gribaudo G, Gilardi G, Giunta C. The oxygenase component of phenol hydroxylase from *Acinetobacter ra‐*

[30] Murray LJ, Garcia-Serres R, McCormick MS, Davydov R, Naik SG, Kim S-H, Hoff‐ man BM, Huynh BH, Lippard SJ. Dioxygen activation at non-heme diiron centers: oxidation of proximal residue in the I100W variant of toluene/o-xylene monooxyge‐

[31] Wojcieszyńska D, Greń I, Hupert-Kocurek K, Guzik U. Modulation of FAD-depend‐ ent monooxygenase activity from aromatic compounds-degrading *Stenotrophomonas*

[33] Arora PK, Srivastava A, Singh VP. Application of monooxygenase in dehalogenation, desulphurization, denitrification and hydroxylation of aromatic compounds. Journal

[34] Dresen C, Lin LY-C, D'Angelo I, Tocheva EI, Strynadka N, Eltis LD. A flavin-de‐ pendent monooxygenase from *Mycobacterium tuberculosis* involved in cholesterol ca‐

[35] Plazmiňo DE, Winkler M, Glieder A, Fraaije MW. Monooxygenase as biocatalysts: classification, mechanistic aspects and biotechnological applications. Journal of Bio‐

[36] Parales RE, Resnick SM. Aromatic ring hydroxylating dioxygenases. In: Ramos J-L, Levesque RC. (ed.) Pseudomonas. Netherlands: Springer; 2006. p287-340. Available from http://link.springer.com/chapter/10.1007/0-387-28881-3\_9?null (accessed

[37] Karegoudar TB, Kim Ch-K. Microbial degradation of monohydroxybenzoic acid. The

*dioresistens* S13. European Journal of Biochemistry 2003;270(10): 2244-2253.

metalate. Chemical Papers 2012;66(3) 211-215.

2007;104(47) 18355-18362.

omatic compounds. Biochemical Journal 1978;174 85-94.

nase hydroxylase. Biochemistry 2007;46(51): 14795-14809.

*maltophilia* KB2. Acta Biochimica Polonica 2011;58(3) 421-6.

of Bioremediation & Biodegradation 2010;1 1-112.

technology 2010;146(1-2) 9-24.

Journal of Microbiology 2000;38(2) 53-61.

28.11.2012).

[32] Greń I. Microbial transformation of xenobiotics. Chemik 2012;66 835-842.

tabolism. The Journal of Biological Chemistry 2010;285(29) 2264-2275.

Hydrochimica et Hydrobiologics 1985;13(4) 453-460.


[24] Hu X-F, Wu L. Oxidation of 3,5-di-tert-butylcatechol in the presence of V-polyoxo‐ metalate. Chemical Papers 2012;66(3) 211-215.

[12] Hammer A, Stolz A, Knackmuss H-J. Purification and characterization of a novel type of protocatechuate 3,4-dioxyegnase with the ability to oxidize 4-sulfocatechol.

[13] Elgren TE, Orville AM, Kelly KA, Lipscomb JD, Ohlendorf DH, Que L. Crystal struc‐ ture and resonance Raman studies of protocatechuate 3,4-dioxygenase complexed

[14] Murakami S, Okuno T, Matsumura E, Takenaka S, Shinke R, Aoki K. Cloning of a gene encoding hydroxyquinol 1,2-dioxygenase that catalyzes both intradiol and ex‐ tradiol ring cleavage of catechol. Bioscience, Biotechnology, and Biochemistry

[15] Iwagami SG, Yang K, Davies J. Characterization of the protocatechuic acid catabolic gene cluster from *Streptomyces* sp. strain 2065. Applied and Environmental Microbi‐

[16] Vetting MW, D'Argenio DA, Ornston LN, Ohlendorf DH. Structure of *Acinetobacter* strain ADP1 protocatechuate 3,4-dioxygenase at 2.2 Å resolution: implications for the

[17] Ferraroni M, Seifert J, Travkin VM, Thiel M, Kaschabek S, Scozzafava A, Golovleva L, Schlömann M, Briganti F. Crystal structure of the hydroxyquinol 1,2-dioxygenase from *Nocardioides simplex* 3E, a key enzyme involved in polychlorinated aromatics bi‐

[18] Bubinas A, Giedraityte G, Kalediene L. Protocatechuate 3,4-dioxygeanse from ther‐

[19] Luo S, Zhang J-J, Zhou N-Y. Molecular cloning and biochemical characterization of protocatechuate 3,4-dioxygenase in *Burkholderia* sp. NCIMB 10467. Microbiology

[20] Nadaf NH, Ghosh JS. Purification and characterization of catechol 1,2-dioxygenase from *Rhodococcus* sp. NCIM 2891. Research Journal of Environmental and Earth Sci‐

[21] Camargo FAO, Andreazza R, Baldoni DB, Bento FM. Enzymatic activity of catechol 1,2-dioxygenase and catechol 2,3-dioxygenase produced by *Gordonia polyisoprenivor‐*

[22] Li F, Zhu L. Effect of surfactant-induced cell surface modifications on electron trans‐ port system and catechol 1,2-dioxygenase activities and phenanthrene biodegrada‐

[23] Fetzner S. Ring-cleaving dioxygenases with a cupin fold. Applied and Environmen‐

tion by *Citrobacter* sp. SA01. Bioresource Technology 2012;123 42-48.

mechanism of an intradiol dioxygenase. Biochemistry 2000;39 7943-7955.

odegradation. The Journal of Biological Chemistry 2005;280(2) 21144-21154.

mophilic *Geobacillus* sp. strain. Biologija 2007;18(1) 31-34.

with 3,4-dihydroxyphenylacetate. Biochemistry 1997;36 11504-11513.

Archives of Microbiology 1996;166 92-100.

1999;63(5) 859-865.

146 Biodegradation of Hazardous and Special Products

2008;35(5) 712-719.

ence 2011;3(5) 608-613.

*ans*. Quimica Nova 2012;35(8) 1587-1592.

tal Microbiology 2012;78(8) 2505-2514.

ology 2000;66(4) 1499-1508.


[38] Guzik U, Greń I, Hupert-Kocurek K, Wojcieszyńska D. Catechol 1,2-dioxygenase from the New aromatic compounds-degrading *Pseudomonas putida* strain N6. Interna‐ tional Biodeterioration & Biodegradation 2011;65(3) 504-512.

structure/activity relationship and the crystal structures of native enzyme and cate‐

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

149

[51] Wei M, Zhang J-J, Liu H, Zhou N-Y. *para*-Nitrophenol 4-monooxygenase and hy‐ droxyquinol 1,2-dioxygenase catalyze sequential transformation of 4-nitrocatechol in

[52] Perez-Pantoja D, Donoso R, Junca H, Gonzales B, Pieper DH. Phylogenomics of aero‐ bic bacterial degradation of aromatics. In: Timmis KN (ed.) Handbook of Hydrocar‐ bon and Lipid Microbiology. Heidelberg: Springer Verlag; 2010. p1-40. Available from http://www.bio.puc.cl/caseb/pdf/prog7/Vol%202\_00094.pdf (accessed

[53] Caposio P, Pessione E, Giuffrida G, Conti A, Landolfo S, Giunta C, Gribaudo G. Cloning and characterization of two catechol 1,2-dioxygeanse genes from *Acinetobact‐*

[54] Liu S, Ogawa N, Senda T, Hasebe A, Miyashita K. Amino acids in position 48, 52, and 73 differentiate the substrate specificities of the highly homologous chlorocate‐ chol 1,2-dioxygenase CbnA and TcbC. Journal of Bacteriology 2005; 187(15)

[55] Murakami S, Kodama N, Shinke R, Aoki K. Classification of catechol 1,2-dioxygenase family: sequence analysis of a gene for the catechol 1,2-dioxygenase showing high

[56] Suvorova MM, Solyanikova IP, Golovleva LA. Specificity of catechol *ortho*-cleavage during *para*-toluate degradation by *Rhodococcus opacus* 1 cp. Biochemistry (Moscow)

[57] Solyanikova IP, Konovalova EI, Golovleva LA. Methylcatechol 1,2-dioxygenase of *Rhodococcus opacus* 6a is a new type of the catechol-cleaving enzyme. Biochemistry

[58] Patel RN, Hou CT, Felix A, Lillard MO. Catechol 1,2-dioxygenase from *Acinetobacter calcoaceticus*: purification and properties. Journal of Bacteriology 1976;127(1) 536-544.

[59] Briganti F, Pessione E, Giunta C, Mazzoli R, Scozzafava A. Purification and catalytic properties of two catechol 1,2-dioxygenase isozymes from benzoate-grown cells of

[60] Di Nardo G, Roggero C, Campolongo S, Valetti F, Trotta F, Gilardi G. Catalytic prop‐ erties of catechol 1,2-dioxygenase from *Acinetobacter radioresistens* S13 immobilized

[61] Sauert-Ignazi G, Gagnon J, Beguin C, Barrelle M, Markowicz Y, Pelmont J, Toussaint A. Characterization of a chromosomally encoded catechol 1,2-dioxygenase (E.C.

*Acinetobacter radioresistens*. Journal of Protein Chemistry 2000;19(8) 709-716.

on anosponges. Dalton Transactions 2009;33 6507-6512.

aniline-assimilating *Rhodococcus erythropo‐*

chols adducts. Journal of Structural Biology 2010;170(3) 548-564.

*Pseudomonas* sp. strain WBC-3. Biodegradation 2010;21(6) 915-921.

*er radioresistens* S13. Research in Microbiology 2002;153(2) 69-74.

specificity for methylcatechols from Gram+

*lis* An-13. Gene 1997;185(1) 49-54.

(Moscow) 2009;74(9) 994-1001.

2006;71(12) 1316-1323.

28.11.2012).

5427-5436.


structure/activity relationship and the crystal structures of native enzyme and cate‐ chols adducts. Journal of Structural Biology 2010;170(3) 548-564.

[51] Wei M, Zhang J-J, Liu H, Zhou N-Y. *para*-Nitrophenol 4-monooxygenase and hy‐ droxyquinol 1,2-dioxygenase catalyze sequential transformation of 4-nitrocatechol in *Pseudomonas* sp. strain WBC-3. Biodegradation 2010;21(6) 915-921.

[38] Guzik U, Greń I, Hupert-Kocurek K, Wojcieszyńska D. Catechol 1,2-dioxygenase from the New aromatic compounds-degrading *Pseudomonas putida* strain N6. Interna‐

[39] Wojcieszyńska D, Hupert-Kocurek K, Greń I, Guzik U. High activity catechol 2,3-di‐ oxygenase from the cresols-degrading *Stenotrophomonas maltophilia* strain KB2. Inter‐

[40] Vaillancourt FH, Bolin JT, Eltis LD. The ins and outs of ring-cleaving dioxygenases. Critical Reviews in Biochemistry and Molecular Biology 2006;41(4) 241-267.

[41] Latus M, Seitz H, Eberspacher J, Lingens F. Purification and characterization of hy‐ droxyquinol 1,2-dioxygenase from *Azotobacter* sp. strain GP1. Applied and Environ‐

[42] Costas M, Mehn MP, Jensen MP, Que L. Dioxygen activation at mononuclear non‐ heme iron active sites: enzymes, models, and intermediates. Chemical Reviews

[43] Mahiudddin Md, Fakhruddin ANM, Mahin AA. Degradation of phenol via meta cleavage pathway by *Pseudomonas fluorescens* PU1. International Scholarly Research

[44] Mizuno S, Yoshikawa N, Seki M, Mikawa T, Imada Y. Microbial production of *cis,cis*muconic acid from benzoil acid. Applied Microbiology and Biotechnology 1988;28(1)

[45] Wu Ch-M, Lee T-H, Lee S-N, Lee Y-A, Wu J-Y. Microbial synthesis *cis,cis*-muconic acid by *Sphingobacterium* sp. GCG generated from effluent of a styrene monomer (SM) production plant. Enzyme and Microbial Technology 2004;35(6-7) 589-604.

[46] Gibson DT, Parales RE. Aromatic hydrocarbon dioxygenases in environmental bio‐

[47] Vetting MW, Ohlendorf DH. The 1.8 Å crystal structure of catechol 1,2-dioxygenase reveals a novel hydrophobic helical zipper as a subunit linker. Structure 2000;8(4)

[48] Contzen M, Stolz A. Characterization of the genes form two protocatechuate 3,4-di‐ oxygenases from the 4-sulfocatechol-degrading bacterium *Agrobacterium radiobacter*

[49] Pandeeti EVP, Siddavattam D. Purification and characterization of catechol 1,2-diox‐ ygenase from *Acinetobacter* sp. DS002 and cloning, sequencing of partial *catA* gene.

[50] Matera I, Ferraroni M, Kolomytseva M, Golovleva L, Scozzafava A, Briganti F. Cate‐ chol 1,2-dioxygenase from the Gram-positive *Rhodococcus opacus* 1CP: quantitative

technology. Current Opinion in Biotechnology 2000;11(3) 236-243.

strain S2. Journal of Bacteriology 2000;182(21) 6123-6129.

Indian Journal of Microbiology 2011;51(3) 312-318.

tional Biodeterioration & Biodegradation 2011;65(3) 504-512.

national Biodeterioration & Biodegradation 2011;65(6) 853-858.

mental Microbiology 1995;61(7) 2453-2460.

Network Microbiology 2012; 2012 1-6.

2004;104(2) 939-986.

148 Biodegradation of Hazardous and Special Products

20-25.

429-440.


1.13.11.1) from *Alcaligenes eutrophus* CH34. Archives of Micorbiology 1996;166(1) 42-50.

[72] Bull Ch, Ballou DP. Purification and properties of protocatechuate 3,4-dioxygenase from *Pseudomonas putida*. A new iron to subunit stoichiometry. The Journal of Biolog‐

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

151

[73] Walsh TA, Ballou DP. Halogenated protocatechuates as substrates for protocatechu‐ ate dioxygenase from *Pseudomonas cepacia*. The Journal of Biological Chemistry

[74] Ludwig ML, Weber LD, Ballou DP. Characterization of crystals of protocatechuate 3,4-dioxygenase from *Pseudomonas cepacia*. The Journal of Biological Chemistry

[75] Zylstra GJ, Olsen RH, Ballou DP. Genetic organization and sequence of the *Pseudomo‐ nas cepacia* genes for the alpha and beta subunits of protocatechuate 3,4-dioxygenase.

[76] Stanier RY, Ingraham JL. Protocatechuic acid oxidase. The Journal of Biological

[77] Petersen EI, Zuegg J, Ribbons DW, Schwab H. Molecular cloning and homology modeling of protocatechuate 3,4-dioxygenase from *Pseudomonas marginata*. Microbio‐

[78] Frazee RW, Livingston DM, LaPorte DC, Lipscomb JD. Cloning, sequencing, and ex‐ pression of the *Pseudomonas putida* protocatechuate 3,4-dioxygenase genes. Journal of

[79] Orville AM, Lipscomb JD, Ohlendorf DH. Crystal structures of substrate and sub‐ strate analog complexes of protocatechuate 3,4-dioxygenase: endogenous Fe3+ ligand displacement in response to substrate binding. Biochemistry 1997;36(33) 10052-10066.

[80] Valley MP, Brown CK, Burk DL, Vetting MW, Ohlendorf DH, Lipscomb JD. Roles of the equatorial tyrosyl iron ligand of protocatechuate 3,4-dioxygenase in catalysis. Bi‐

[81] Overhage J, Kresse AU, Priefert H, Sommer H, Krammer G, Rabenhorst J, Steinbü‐ chel A. Molecular characterization of the genes *pcaG* and *pcaH*, encoding protocatech‐ uate 3,4-dioxygenase, which are essential for vanillin catabolism in *Pseudomonas* sp.

[82] Chen YP, Dilworth MJ, Glenn AR. Aromatic metabolism in *Rhizobium trifolii* – proto‐ catechuate 3,4-dioxygenase. Archives of Microbiology 1984;138(3) 187-190.

[83] Zaborina O, Seitz H-J, Sidorov I, Eberspächer J, Alexeeva E, Golovleva L, Lingens F. Inhibition analysis of hydroxyquinol-cleaving dioxygenases from the chlorophenoldegrading *Azotobacter* sp. GP1 and *Streptomyces rochei* 303. Journal of Basic Microbiol‐

[84] Daubaras DL, Saido K, Chakrabarty AM. Purification of hydroxyquinol 1,2-dioxyge‐ nase and maleylacetate reductase: the lower pathway of 2,4,5-trichlorophenoxyacetic

strain HR199. Applied and Environmental Microbiology 1999;65(3) 951-960.

ical Chemistry 1981;256(24) 12673-12680.

Journal of Bacteriology 1989;171(11) 5915-5921.

1983;258(23) 14413-14421.

1984;259(23) 14840-14842.

Chemistry 1954;210 799-808.

logical Research 1996;151(4) 359-370.

Bacteriology 1993;175(19) 6194-6202.

ochemistry 2005;44(33) 11024-11039.

ogy 1999;39(1) 61-73.


[72] Bull Ch, Ballou DP. Purification and properties of protocatechuate 3,4-dioxygenase from *Pseudomonas putida*. A new iron to subunit stoichiometry. The Journal of Biolog‐ ical Chemistry 1981;256(24) 12673-12680.

1.13.11.1) from *Alcaligenes eutrophus* CH34. Archives of Micorbiology 1996;166(1)

[62] Murakami S, Wang CL, Naito A, Shinke R, Aoki K. Purification and characterization of four catechol 1,2-dioxygenase isozymes from the benzamide-assimilating bacteri‐ um *Arthrobacter* species BA-5-17. Microbiological Research 1998;153(2) 163-171. [63] Suzuki K, Ichimura A, Ogawa N, Hasebe A, Miyashita K. Differential expression of two catechol 1,2-dioxygenases in *Burkholderia* sp. strain TH2. Journal of Bacteriology

[64] Wang Ch-L, You S-L, Wang S-L. Purification and characterization of a novel catechol 1,2-dioxygenase from *Pseudomonas aeruginosa* with benzoic acid as a carbon source.

[65] König Ch, Eulberg D, Gröning J, Lakner S, Seibert V, Kaschabke SR, Schlömann M. A linear megaplasmid, p1CP, carrying the genes for chlorocatechol catabolism of *Rho‐*

[66] Ferraroni M, kolomytseva MP, Solyanikova IP, Scozzafava A, Golovleva LA, Briganti F. Crystal structure of 3-chlorocatechol 1,2-dioxygenase key enzyme of a new modi‐ fied *ortho*-pwthway from the Gram-positive *Rhodococcus opacus* 1CP grown on 2-

[67] Gou M, Qu YY, Zhou JT, Li A, Uddin MS. Characterization of catechol 1,2-dioxyge‐ nase from cell extracts of *Sphingomonas xenophaga* QYY. Science in China Series B:

[68] Guzik U, Hupert-Kocurek K, Sałek K, Wojcieszyńska D. Influence of metal ions on bioremediation activity of protocatechuate 3,4-dioxygenase from *Stenotrophomonas maltophilia* KB2. World Journal of Microbiology and Biotechnology 2012; DOI

[69] Hartnett Ch, Neidle EL, Ngai K-L, Ornston N. DNA sequences of genes encoding *Acinetobacter calcoaceticus* protocatechuate 3,4-dioxyegnase: evidence indicating shuf‐ fling of genes and of DNA sequences within genes during their evolutionary diver‐

[70] Kahng Y-Y, Cho K, Song S-Y, Kim S-J, Leem S-H, Kim SI. Enhanced detection and characterization of protocatechuate 3,4-dioxygenase in *Acinetobacter lwoffii* K24 by proteomics using a column separation. Biochemical and Biophysical Research Com‐

[71] Whittaker JW, Lipscomb JD, Kent TA, Munck E. *Brevibacterium fuscum* protocatechu‐ ate 3,4-dioxygenase. Purification, crystallization, and characterization. The Journal of

42-50.

2002;184(20) 5714-5722.

150 Biodegradation of Hazardous and Special Products

Chemistry 2009;52(5) 615-620.

10.1007/s11274-012-1178-z.

munications 2002;295(4) 903-909.

Biological Chemistry 1984;259(7) 4466-4475.

Process Biochemistry 2006; 41(7) 1594-1601.

*dococcus opacus* 1CP. Microbiology 2000;150(Pt 9) 3075-3087.

chlorophenol. Journal of Molecular Biology 2006;360(4) 788-799.

gence. Journal of Bacteriology 1990;172(2) 956-966.


acid metabolism by *Burkholderia cepacia* AC1100. Applied and Environmental Micro‐ biology 1996;61(11) 4276-4279.

ternational Conference on Chemical, Environmental Science and Engineering (ICE‐

Intradiol Dioxygenases — The Key Enzymes in Xenobiotics Degradation

http://dx.doi.org/10.5772/56205

153

[96] Kalogeris E, Sanakis Y, Mamma D, Christakopoulos P, Kekos D, Stamatis H. Proper‐ ties of catechol 1,2-dioxygenase from *Pseudomonas putida* immobilized in calcium al‐

[97] Zucolotto V, Pinto APA, Tumolo T, Moraes ML, Baptista MS, Riul A, Araujo APU, Oliveira ON. Catechol biosensing using a nanostructured layer-by-layer film contain‐ ing Cl-catechol 1,2-dioxygenase. Biosensors and Bioelectronics 2006;21(7) 1320-1326.

[98] Latha R, Mandappa IM, Thakur MS, Manonmani HK. Influence of metal ions on de‐ hydrohalogenase activity. African Journal of Basic & Applied Sciences 2011;3(2)

[99] Gopal B, Madan LL, Betz SF, Kossiakoff AA. The crystal structure of a quercetin 2,3 dioxygenase from *Bacillus subtilis* suggests modulation of enzyme activity by a

change in the metal ion at the active site(s). Biochemistry 2005;44(1) 193-201.

ginate hydrogels. Enzyme and Microbial Technology 2006;39(5) 1113-1121.

EBS'2012), July 28-29, 2012, Pattaya, Thailand.

45-51.


ternational Conference on Chemical, Environmental Science and Engineering (ICE‐ EBS'2012), July 28-29, 2012, Pattaya, Thailand.

[96] Kalogeris E, Sanakis Y, Mamma D, Christakopoulos P, Kekos D, Stamatis H. Proper‐ ties of catechol 1,2-dioxygenase from *Pseudomonas putida* immobilized in calcium al‐ ginate hydrogels. Enzyme and Microbial Technology 2006;39(5) 1113-1121.

acid metabolism by *Burkholderia cepacia* AC1100. Applied and Environmental Micro‐

[85] Takenaka S, Okugawa S, Kadowaki M, Murakami S, Aoki K. The metabolic pathway of 4-aminophenol in *Burkholderia* sp. strain AK-5 differs from that of aniline and ani‐

[86] Travkin VM, Jadan AP, Briganti F, Scozzafava A, Golovleva LA. Characterization of an intradiol dioxygenase involved in the biodegradation of the chlorophenoxy herbi‐

[87] Louie TM, Webster ChM, Xun L. Genetic and biochemical characterization of a 2,4,6 trichlorophenol degradation pathway in *Ralstonia eutropha* JMP134. Journal of Bac‐

[88] Hatta T, Nakano O, Imai N, Takizawa N, Kiyohara H. Cloning and sequence analysis of hydroxyquinol 1,2-dioxygenase gene In 2,4,6-trichlorophenol-degrading *Ralstonia pickettii* DTP0602 and characterization of its product. Journal of Bioscience and Bioen‐

[89] Yoshida M, Oikawa T, Obata H, Abe K, Mihara H, Esaki N. Biochemical and genetic analysis of the γ-resorcylate (2,6-dihydrobenzoate) catabolic pathway in *Rhizobium* sp. strain MTP-10005: identification and functional analysis of its gene cluster. Jour‐

[90] Kitagawa W, Kimura N, Kamagata Y. A novel *p*-nitrophenol degradation gene luster from a gram-positive bacterium, *Rhodococcus opacus* SAO101. Journal of bacteriology

[91] Mayilmurugan R, Sankaralingam M, Suresh E, Palaniandavar M. Novel square pyra‐ midal iron(III) complexes of linear tetradentate bis(phenolate) ligands as structural and reactive model for intradiol-cleaving 3,4-PCD enzymes: quinone formation *vs.*

[92] Borowski T, Siegbahn PEM. Mechanism for catechol ring cleavage by non-heme iron intradiol dioxygenases: a hybrid DFT study. Journal of American Chemical Society

[93] Kurahashi T, Oda K, Sugimoto M, Ogura T, Fujii H. Trigonal-bipyramidal geometry induced by an external water ligand In a sterically hindered iron salen complex, re‐ lated to the active site of protocatechuate 3,4-dioxygenase. Inorganic Chemistry

[94] Orville AM, Elango N, Lipscomb JD, Ohlendorf DH. Structures of competitive inhibi‐ tor complexes of protocatechuate 3,4-dioxygenases: multiple exogenous ligand bind‐

[95] Suma Y, Kim D, Lee JW, Park KY, Kim HS. Degradation of catechol by immobilized hydroxyquinol 1,2-dioxygenase (1,2-HQD) onto single-walled carbon nanotubes. In‐

ing orientations within the active site. Biochemistry 1997;36(33) 10039-10051.

intradiol cleavage. Dalton Transactions 2010;39 9611-9625.

cides 2,4-D and 2,4,5-T. FEBS Letters 1997;407(1) 69-72.

biology 1996;61(11) 4276-4279.

152 Biodegradation of Hazardous and Special Products

line with C-4 substituents.

teriology 2002;184(13) 3492-3500.

gineering 1999;87(3) 267-272.

2004;186(15) 4894-4902.

2006;128(39) 12941-12953.

2006;45(19) 7709-7721.

nal of Bacteriology 2007;189(5) 1573-1581.


**Chapter 8**

**Hydrocarbon Biodegradation Potential of Native and**

**Exogenous Microbial Inocula in Mexican Tropical Soils**

Soil and water contamination by oil is one of the central environmental problems in Mexico. In the southeastern part of the country, especially in Tabasco and Veracruz states there are several oil facilities involved in extraction, transportation, processing and storage of oil and oil products. Historically in this region there have been spills that affect large areas of soil and eventually rivers, streams and lagoons. Pollutants often persist for long periods in the soil, sediments and water due to different factors like the environmental conditions prevailing in the region (heavy rains and extensive wetlands), the nature of the soils (mostly clayed soils adjacent to water bodies), and the recalcitrance of oil components (Adams et al. 2011).

The application of bioremediation methods based on degradation of hydrocarbons is a widely used alternative for the recovery of such contaminated soils (Boonchan et al. 2000, Sayara et al. 2012). However, the success of this strategy depends on the degrading capacity of the native or exogenous microorganisms applied as part of the treatment (Liu and Suflita, 1993, Van Hamme, 2003, Liu et al. 2011). In this sense, a key issue for the restoration of soils is to assess their potential for bioremediation under different treatment strategies and culture conditions. This analysis must consider first the biodegrading capacity of native microbial populations (natural attenuation). Alternatively, the use of inoculants composed by exogenous microor‐ ganisms, previously characterized as degraders (bioaugmentation), can be considered as an

Several authors have reported that specific microorganisms are required for the removal of various petroleum fractions, in particular those having enzymatic capabilities for biotransfor‐

and reproduction in any medium, provided the original work is properly cited.

© 2013 Díaz-Ramírez et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

© 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution,

Ildefonso Díaz-Ramírez, Erika Escalante-Espinosa,

Reyna Fócil-Monterrubio and Hugo Ramírez-Saad

Additional information is available at the end of the chapter

Randy Adams Schroeder ,

http://dx.doi.org/10.5772/56233

active bioremediation treatment.

**1. Introduction**

### **Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils**

Ildefonso Díaz-Ramírez, Erika Escalante-Espinosa, Randy Adams Schroeder , Reyna Fócil-Monterrubio and Hugo Ramírez-Saad

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/56233

#### **1. Introduction**

Soil and water contamination by oil is one of the central environmental problems in Mexico. In the southeastern part of the country, especially in Tabasco and Veracruz states there are several oil facilities involved in extraction, transportation, processing and storage of oil and oil products. Historically in this region there have been spills that affect large areas of soil and eventually rivers, streams and lagoons. Pollutants often persist for long periods in the soil, sediments and water due to different factors like the environmental conditions prevailing in the region (heavy rains and extensive wetlands), the nature of the soils (mostly clayed soils adjacent to water bodies), and the recalcitrance of oil components (Adams et al. 2011).

The application of bioremediation methods based on degradation of hydrocarbons is a widely used alternative for the recovery of such contaminated soils (Boonchan et al. 2000, Sayara et al. 2012). However, the success of this strategy depends on the degrading capacity of the native or exogenous microorganisms applied as part of the treatment (Liu and Suflita, 1993, Van Hamme, 2003, Liu et al. 2011). In this sense, a key issue for the restoration of soils is to assess their potential for bioremediation under different treatment strategies and culture conditions. This analysis must consider first the biodegrading capacity of native microbial populations (natural attenuation). Alternatively, the use of inoculants composed by exogenous microor‐ ganisms, previously characterized as degraders (bioaugmentation), can be considered as an active bioremediation treatment.

Several authors have reported that specific microorganisms are required for the removal of various petroleum fractions, in particular those having enzymatic capabilities for biotransfor‐

© 2013 Díaz-Ramírez et al.; licensee InTech. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited. © 2013 The Author(s). Licensee InTech. This chapter is distributed under the terms of the Creative Commons Attribution License http://creativecommons.org/licenses/by/3.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

mation of these compounds (Van der Meer et al. 1992; Prince, 2003). Although the bacteria can be grown in the laboratory, progress on knowledge of microbial ecology of complex com‐ munities requires more extensive studies, regarding the activity of microorganisms in environmental samples that suffered minimal modifications (Amann and Kühl, 1998). Moreover, knowledge of the mechanisms of degradation of contaminants in the environment has been generated in laboratory studies using degraders. As a result the role of microorgan‐ isms in regulating processes such as mineralization and biodegradation of polluting com‐ pounds is not fully understood (Watanabe and Hamamura, 2003).

considered that exogenous inocula must possess two essential properties: i) to degrade oil faster than native populations, and ii) to degrade a broader spectrum of oil compounds than native populations (Van Hamme et al. 2003, Madueño et al. 2011). Obtaining microorganisms that meet these conditions may be more feasible from sites where the microbial population is exposed for long periods to hydrocarbons pollutants (Liu and

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

http://dx.doi.org/10.5772/56233

157

It is generally considered that mixed cultures have a higher metabolic versatility than pure cultures (axenic). Mixed cultures have been used as inocula for treating waste oil in various systems, applying preadaptation strategies of consortia to different types of hydrocarbons and field application conditions (Van Hamme et al. 2000). Some reports have studied the biode‐ grading abilities of defined mixed cultures where the composition of the consortia is previously known. Ko and Lebeault (1999), reported the co-oxidation of decalin and pristane in the presence of hexadecane, by a defined mixed culture of two bacterial strains. Finding that decalin was degraded to a greater extent by an axenic strain than the mixed culture, while pristane was largely degraded by the mixed culture. Richard and Vogel, (1999) compared the biodegradation of diesel by three bacterial strains and a mixed culture composed of the same, degradation of aliphatics was greater with the mixed culture than with the single strains, these authors suggest that degradation of hydrocarbon mixtures by mixed cultures may involve

Often the success of the addition of exogenous microorganisms depends on their adaptation to the environmental conditions prevailing in the soil and their ability to compete with native microorganisms. However some cases have been documented where native populations are better degraders than exogenous inoculants (Thouand et al. 1999). An alternative is the application of microorganisms native of sites contaminated with the compounds of interest, they can pre-selected based on their metabolic capabilities and subsequently be characterized in the laboratory for possible use (Medina-Moreno et al. 2005; Diaz -Ramirez et al. 2008). The application of such microorganisms is useful when the number of native microorganisms is low and they do not have a proper biodegrading capacity (Van Hamme et al. 2003). In general, bacterial cocktails used as inoculants are undefined mixed cultures, where the composition and metabolic capacities of the microbial members is not known precisely; therefore the role

Another useful alternative for the treatment of oil contaminated soils is the use of native plants together with degrading microorganisms associated in their rhizosphere, known as phytore‐ mediation or phytomitigation strategies. The use of plants for remediation of contaminated soil is well accepted, particularly when they are part of a sequence of treatments. The use of pastures allows overall improvement of soil fertility conditions and biological activity (enzymatic functions and respiratory activity), to reduce toxicity, improve, mineralization and / or integration of polluting compounds via humification (Escalante-Espinosa, 2005;

Suflita, 1993, Greenwood et al. 2009).

*1.2.2. Biodegradation of hydrocarbons by mixed cultures*

different strains that possess complementary metabolic capabilities.

of each strain during biodegradation is not clear.

Adams et al. 2011).

#### **1.1. Adaptation and presence of microorganisms in contaminated sites**

Exposure of native microbial soil populations to hydrocarbon contamination is a key factor that determines the biodegradation rates for overall removal of contaminant. This phenom‐ enon allows the increase in oxidation potential of the microbial community and is called adaptation. According to Leahy and Colwell (1990), this term applies to mixed microbial communities and single microbial species. Van der Meer et al. (1992) proposed some mecha‐ nisms on the molecular events that allow adaptation of microbial communities: i) induction and/or de-repression of specific enzymes, ii) new metabolic capabilities as a result of genetic changes, and iii) enrichment of microbial populations capable of transforming the compound of interest.

Generally, degrading-microorganisms are more frequently isolated from hydrocarbon contaminated soils, where they can reach levels between 1 to 10% of the soil microbial population. While in non polluted environments degraders normally represent 1% of the microbial community (Torsvik and Ovreas, 2002). In some cases, predominant genera of degraders are reported, while in others there is a high microbial diversity, which may be due to the composition of the native microbial community, and the environmental conditions at the affected sites. Microbial ability to grow in the presence of oil-derived compounds as a sole carbon source has been addressed by selective microbial enrichment studies (Leahy and Colwell, 1990, Viñas et al. 2002, Roldán-Carrillo et al. 2012).

#### **1.2. Hydrocarbons biodegradation**

The biodegradation process is defined as the change in the chemical structure of a compound carried out by microorganisms, and is the basis of conventional engineering techniques for the treatment of sewage and contaminated soil bioremediation (Madsen, 1998). The petroleum compounds can selectively inhibit certain microorganisms, causing changes in the number and diversity of microbial species in soil. Thus the rate of biodegradation of hydrocarbon is determined by the presence of such native microorganisms, their physiological capabilities and the environmental conditions (Prince, 2003, Zhang et al. 2012).

#### *1.2.1. Applying degraders*

In some cases, hydrocarbon biodegradation levels of native inocula had been higher than commercial or exogenous inocula (Thouand et al. 1999, Mohammed et al. 2007). It is considered that exogenous inocula must possess two essential properties: i) to degrade oil faster than native populations, and ii) to degrade a broader spectrum of oil compounds than native populations (Van Hamme et al. 2003, Madueño et al. 2011). Obtaining microorganisms that meet these conditions may be more feasible from sites where the microbial population is exposed for long periods to hydrocarbons pollutants (Liu and Suflita, 1993, Greenwood et al. 2009).

#### *1.2.2. Biodegradation of hydrocarbons by mixed cultures*

mation of these compounds (Van der Meer et al. 1992; Prince, 2003). Although the bacteria can be grown in the laboratory, progress on knowledge of microbial ecology of complex com‐ munities requires more extensive studies, regarding the activity of microorganisms in environmental samples that suffered minimal modifications (Amann and Kühl, 1998). Moreover, knowledge of the mechanisms of degradation of contaminants in the environment has been generated in laboratory studies using degraders. As a result the role of microorgan‐ isms in regulating processes such as mineralization and biodegradation of polluting com‐

Exposure of native microbial soil populations to hydrocarbon contamination is a key factor that determines the biodegradation rates for overall removal of contaminant. This phenom‐ enon allows the increase in oxidation potential of the microbial community and is called adaptation. According to Leahy and Colwell (1990), this term applies to mixed microbial communities and single microbial species. Van der Meer et al. (1992) proposed some mecha‐ nisms on the molecular events that allow adaptation of microbial communities: i) induction and/or de-repression of specific enzymes, ii) new metabolic capabilities as a result of genetic changes, and iii) enrichment of microbial populations capable of transforming the compound

Generally, degrading-microorganisms are more frequently isolated from hydrocarbon contaminated soils, where they can reach levels between 1 to 10% of the soil microbial population. While in non polluted environments degraders normally represent 1% of the microbial community (Torsvik and Ovreas, 2002). In some cases, predominant genera of degraders are reported, while in others there is a high microbial diversity, which may be due to the composition of the native microbial community, and the environmental conditions at the affected sites. Microbial ability to grow in the presence of oil-derived compounds as a sole carbon source has been addressed by selective microbial enrichment studies (Leahy and

The biodegradation process is defined as the change in the chemical structure of a compound carried out by microorganisms, and is the basis of conventional engineering techniques for the treatment of sewage and contaminated soil bioremediation (Madsen, 1998). The petroleum compounds can selectively inhibit certain microorganisms, causing changes in the number and diversity of microbial species in soil. Thus the rate of biodegradation of hydrocarbon is determined by the presence of such native microorganisms, their physiological capabilities

In some cases, hydrocarbon biodegradation levels of native inocula had been higher than commercial or exogenous inocula (Thouand et al. 1999, Mohammed et al. 2007). It is

pounds is not fully understood (Watanabe and Hamamura, 2003).

Colwell, 1990, Viñas et al. 2002, Roldán-Carrillo et al. 2012).

and the environmental conditions (Prince, 2003, Zhang et al. 2012).

**1.2. Hydrocarbons biodegradation**

156 Biodegradation of Hazardous and Special Products

*1.2.1. Applying degraders*

of interest.

**1.1. Adaptation and presence of microorganisms in contaminated sites**

It is generally considered that mixed cultures have a higher metabolic versatility than pure cultures (axenic). Mixed cultures have been used as inocula for treating waste oil in various systems, applying preadaptation strategies of consortia to different types of hydrocarbons and field application conditions (Van Hamme et al. 2000). Some reports have studied the biode‐ grading abilities of defined mixed cultures where the composition of the consortia is previously known. Ko and Lebeault (1999), reported the co-oxidation of decalin and pristane in the presence of hexadecane, by a defined mixed culture of two bacterial strains. Finding that decalin was degraded to a greater extent by an axenic strain than the mixed culture, while pristane was largely degraded by the mixed culture. Richard and Vogel, (1999) compared the biodegradation of diesel by three bacterial strains and a mixed culture composed of the same, degradation of aliphatics was greater with the mixed culture than with the single strains, these authors suggest that degradation of hydrocarbon mixtures by mixed cultures may involve different strains that possess complementary metabolic capabilities.

Often the success of the addition of exogenous microorganisms depends on their adaptation to the environmental conditions prevailing in the soil and their ability to compete with native microorganisms. However some cases have been documented where native populations are better degraders than exogenous inoculants (Thouand et al. 1999). An alternative is the application of microorganisms native of sites contaminated with the compounds of interest, they can pre-selected based on their metabolic capabilities and subsequently be characterized in the laboratory for possible use (Medina-Moreno et al. 2005; Diaz -Ramirez et al. 2008). The application of such microorganisms is useful when the number of native microorganisms is low and they do not have a proper biodegrading capacity (Van Hamme et al. 2003). In general, bacterial cocktails used as inoculants are undefined mixed cultures, where the composition and metabolic capacities of the microbial members is not known precisely; therefore the role of each strain during biodegradation is not clear.

Another useful alternative for the treatment of oil contaminated soils is the use of native plants together with degrading microorganisms associated in their rhizosphere, known as phytore‐ mediation or phytomitigation strategies. The use of plants for remediation of contaminated soil is well accepted, particularly when they are part of a sequence of treatments. The use of pastures allows overall improvement of soil fertility conditions and biological activity (enzymatic functions and respiratory activity), to reduce toxicity, improve, mineralization and / or integration of polluting compounds via humification (Escalante-Espinosa, 2005; Adams et al. 2011).

Diaz-Ramirez et al. (2003) reported a strategy involving bacterial strains obtained from the rhizosphere of a plant native to a contaminated site, that were used to prepare defined mixed cultures for hydrocarbon biodegradation and mineralization in liquid cultures. Ten bacterial strains were isolated and identified as members of the genera: *Bacillus, Gordonia, Kocuria, Pseudomonas, Arthrobacter* and *Micrococcus.* Then assessed for biodegradation of specific hydrocarbon fractions in liquid medium, as both; single strains and defined mixed cultures (CMD) prepared with standardized mixtures. Additionally predominant strains were selected and their population dynamics during biodegradation was analyzed, in order to design specific inocula for different types of hydrocarbons (Diaz-Ramirez et al. 2008). Under this approach consumption levels of 40% of total hydrocarbons were reached in 11 days of culture, degradation value for aromatics was 31%, and 20% for polar hydrocarbons. The results pointed that co-oxidation of aromatic and polar compounds was possible in the presence of aliphatic fraction, when using a defined mixed culture consisting of 3 *Bacillus* strains, 2 *Pseudomonas* strains and one *Gordonia* isolate (Diaz-Ramirez et al. 2008).

Further biodegradation experiments in soil contaminated with crude Olmeca oil, were carried

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

http://dx.doi.org/10.5772/56233

159

The second experimental approach involved the monitoring of natural attenuation (MNA) as a result of a gasoil spill occurred in an area of ecological value; a riparian environment consisting of two streams and two major rivers located in the southeast of Mexico. We identified five points affected by gasoil on areas where the stream presented meanders and points of plant material accumulation. The initial TPH concentrations ranged from 8 500 to 60 000 mg kg-1 soil at different points, total hydrocarbon content was determined periodically in

Experiments were carried out with tropical soils, artificially contaminated with either diesel or Olmeca crude oil. In both experimental designs total hydrocarbon content was determined at different sampling times (biodegradation kinetics), the overall biodegradation rate, the total number of microorganisms and maximum number of degraders were calculated. The com‐ position of the residual diesel was assessed by gas chromatography and for Olmeca crude oil by column chromatography. The half-life of hydrocarbons in soil and the biodegradation

In addition, another study was conducted in order to assess the natural attenuation potential of soil and sediments of a stream system affected by a gasoil spill. Periodic sampling was carried out and the analysis of kinetic parameters suggested alternatives that would allow

Soil was collected close but outside of an area highly impacted by human activities (truck parking lot), with high deposition of plant material and therefore high microbial activity potential. The soil samples showed dark color with fine texture (silt-clay) and pH 7. Soil humidity at the time of collection was 30% with a water retention capacity of 36% (w / w). Proportions of clay, silt and sand were 40%, 45% and 15%, respectively. Soil organic matter content was 2.88%, corresponding to a soil with medium organic matter content (NOM-021- SEMARNAT-2000); these characteristics are shared by many soils of the study area, considered

Biodegradation experiments of diesel (10,000 ppm) were performed in shaken flasks with liquid medium (adjusting the ratio C/ N ~ 20). In such assays, we tested the degrading activity of two defined mixed cultures, the first one (B-DMC) consisted of six bacterial strains; *Bacillus cereus, Gordonia rubripertincta, Kocuria rosea, Bacillus subtilis* strains 7A and 9A, and a nonidentified strain UAM10AP. The second defined mixed culture was formed by two fungal

constant using a first order model were calculated for both biodegradation assays.

out in the presence and absence of co-culture.

order to follow the natural attenuation process.

**2. Material and methods**

restoration of different affected areas.

representative of tropical soils of southeastern Mexico.

**2.1. Soil samples**

**2.2. Diesel biodegradation**

#### **1.3. Monitored natural attenuation**

Conventional bioremediation techniques generally allow the recovery of soils affected by organic pollutants. Natural attenuation has been considered an important remediation strategy mainly when soil, groundwater or surface water is involved, in places where biogeo‐ chemical conditions favor natural processes that degrade or immobilize harmful contaminants. Natural attenuation includes processes such as biodegradation, dispersion, sorption and volatilization of contaminants (Boettcher and Nyer, 2001). Monitored Natural Attenuation (MNA) is a viable alternative for reduction of toxicity, mobility or concentration of the contaminant compounds in soil and/or water (EPA, 1999). Some authors have noted that natural attenuation processes can be selected alone or in conjunction with more active alternatives, when achieving the objectives of remediation, in reasonable time periods, compared with other alternatives that can be costly or with high impact to ecosystems. Among the parameters that should be determined in order to assess the biodegradation potential of the microbial community are: the number and type of predominant microorganisms, degra‐ dation kinetics (measured in field or lab), enzyme activities (dehydrogenases, lipases, oxy‐ genases), as well as the half-life of contaminants, calculated from experimental data based on zero- or first order kinetics (Boettcher and Nyer 2001, Margesin et al. 2005, Chang et al. 2010).

In this chapter, we present two experimental approaches developed to assess the ability of native and exogenous microbial populations to biodegrade different types of hydrocarbons, under lab and field conditions.

In the first experimental approach, the degrading activity of native and exogenous microor‐ ganisms on diesel and Olmeca crude oil was assessed, under different culture conditions at laboratory scale. All exogenous microorganisms were previously isolated from the rhizo‐ sphere of *Cyperus laxus* Lam., a plant native of a highly contaminated tropical swamp (Diaz-Ramirez et al., 2003, Escalante-Espinosa, et al. 2005). The biodegradation ability of these microorganisms was tested in soil artificially contaminated with diesel and biostimulated. Further biodegradation experiments in soil contaminated with crude Olmeca oil, were carried out in the presence and absence of co-culture.

The second experimental approach involved the monitoring of natural attenuation (MNA) as a result of a gasoil spill occurred in an area of ecological value; a riparian environment consisting of two streams and two major rivers located in the southeast of Mexico. We identified five points affected by gasoil on areas where the stream presented meanders and points of plant material accumulation. The initial TPH concentrations ranged from 8 500 to 60 000 mg kg-1 soil at different points, total hydrocarbon content was determined periodically in order to follow the natural attenuation process.

### **2. Material and methods**

Diaz-Ramirez et al. (2003) reported a strategy involving bacterial strains obtained from the rhizosphere of a plant native to a contaminated site, that were used to prepare defined mixed cultures for hydrocarbon biodegradation and mineralization in liquid cultures. Ten bacterial strains were isolated and identified as members of the genera: *Bacillus, Gordonia, Kocuria, Pseudomonas, Arthrobacter* and *Micrococcus.* Then assessed for biodegradation of specific hydrocarbon fractions in liquid medium, as both; single strains and defined mixed cultures (CMD) prepared with standardized mixtures. Additionally predominant strains were selected and their population dynamics during biodegradation was analyzed, in order to design specific inocula for different types of hydrocarbons (Diaz-Ramirez et al. 2008). Under this approach consumption levels of 40% of total hydrocarbons were reached in 11 days of culture, degradation value for aromatics was 31%, and 20% for polar hydrocarbons. The results pointed that co-oxidation of aromatic and polar compounds was possible in the presence of aliphatic fraction, when using a defined mixed culture consisting of 3 *Bacillus* strains, 2 *Pseudomonas*

Conventional bioremediation techniques generally allow the recovery of soils affected by organic pollutants. Natural attenuation has been considered an important remediation strategy mainly when soil, groundwater or surface water is involved, in places where biogeo‐ chemical conditions favor natural processes that degrade or immobilize harmful contaminants. Natural attenuation includes processes such as biodegradation, dispersion, sorption and volatilization of contaminants (Boettcher and Nyer, 2001). Monitored Natural Attenuation (MNA) is a viable alternative for reduction of toxicity, mobility or concentration of the contaminant compounds in soil and/or water (EPA, 1999). Some authors have noted that natural attenuation processes can be selected alone or in conjunction with more active alternatives, when achieving the objectives of remediation, in reasonable time periods, compared with other alternatives that can be costly or with high impact to ecosystems. Among the parameters that should be determined in order to assess the biodegradation potential of the microbial community are: the number and type of predominant microorganisms, degra‐ dation kinetics (measured in field or lab), enzyme activities (dehydrogenases, lipases, oxy‐ genases), as well as the half-life of contaminants, calculated from experimental data based on zero- or first order kinetics (Boettcher and Nyer 2001, Margesin et al. 2005, Chang et al. 2010).

In this chapter, we present two experimental approaches developed to assess the ability of native and exogenous microbial populations to biodegrade different types of hydrocarbons,

In the first experimental approach, the degrading activity of native and exogenous microor‐ ganisms on diesel and Olmeca crude oil was assessed, under different culture conditions at laboratory scale. All exogenous microorganisms were previously isolated from the rhizo‐ sphere of *Cyperus laxus* Lam., a plant native of a highly contaminated tropical swamp (Diaz-Ramirez et al., 2003, Escalante-Espinosa, et al. 2005). The biodegradation ability of these microorganisms was tested in soil artificially contaminated with diesel and biostimulated.

strains and one *Gordonia* isolate (Diaz-Ramirez et al. 2008).

**1.3. Monitored natural attenuation**

158 Biodegradation of Hazardous and Special Products

under lab and field conditions.

Experiments were carried out with tropical soils, artificially contaminated with either diesel or Olmeca crude oil. In both experimental designs total hydrocarbon content was determined at different sampling times (biodegradation kinetics), the overall biodegradation rate, the total number of microorganisms and maximum number of degraders were calculated. The com‐ position of the residual diesel was assessed by gas chromatography and for Olmeca crude oil by column chromatography. The half-life of hydrocarbons in soil and the biodegradation constant using a first order model were calculated for both biodegradation assays.

In addition, another study was conducted in order to assess the natural attenuation potential of soil and sediments of a stream system affected by a gasoil spill. Periodic sampling was carried out and the analysis of kinetic parameters suggested alternatives that would allow restoration of different affected areas.

#### **2.1. Soil samples**

Soil was collected close but outside of an area highly impacted by human activities (truck parking lot), with high deposition of plant material and therefore high microbial activity potential. The soil samples showed dark color with fine texture (silt-clay) and pH 7. Soil humidity at the time of collection was 30% with a water retention capacity of 36% (w / w). Proportions of clay, silt and sand were 40%, 45% and 15%, respectively. Soil organic matter content was 2.88%, corresponding to a soil with medium organic matter content (NOM-021- SEMARNAT-2000); these characteristics are shared by many soils of the study area, considered representative of tropical soils of southeastern Mexico.

#### **2.2. Diesel biodegradation**

Biodegradation experiments of diesel (10,000 ppm) were performed in shaken flasks with liquid medium (adjusting the ratio C/ N ~ 20). In such assays, we tested the degrading activity of two defined mixed cultures, the first one (B-DMC) consisted of six bacterial strains; *Bacillus cereus, Gordonia rubripertincta, Kocuria rosea, Bacillus subtilis* strains 7A and 9A, and a nonidentified strain UAM10AP. The second defined mixed culture was formed by two fungal strains (F-DMC); *Aspergillus terreus* and *Aspergillus carneus.* Finally, a co-culture (CC) com‐ posed of all of the above strains was also tested. The degrading microorganisms were isolated and characterized as previously described (Diaz-Ramirez et al, 2003). In parallel, non-inocu‐ lated controls were prepared to quantify the hydrocarbons abiotic loss.

geomorphology and direction of the stream meanders; several points with presence of hydrocarbons were identified and selected for monitoring. The residual gasoil content (medium fraction C10-C28) was used as a measure of passive biodegradation or Monitored

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

http://dx.doi.org/10.5772/56233

161

The residual gasoil content (medium fraction C10-C28) was used as a measure of passive biodegradation or Monitored Natural Attenuation. Soil hydrocarbon analyses were performed by Gas Chromatography accordingly to the Mexican regulation (NOM-138-SEMARNAT/ SS-2003), which is based on the EPA 8015b standard procedure. From these values, biodegra‐ dation extent and half life of the contaminant in soil were calculated. In addition, the amounts

To assess the natural attenuation potential of contaminated soils, as well as to develop inocula for bioremediation of such sites, it is necessary to know the degrading abilities of microbial native populations. Where applicable, the persistence and activity of the introduced microorganisms, without bioavailability limitations attributable to the soil, have to be addressed. Contrast the biodegradation process under this scheme and in field is one of the main challenges for the definition of the auto recovery potential of contaminated soils, as well as for the establishment of bioremediation strategies that are economic and

Higher levels of biodegradation were obtained with the CC and F -DMC that behave very similar (Table 1). Statistical analysis did not show significant differences at 95% confidence among these two treatments. While the consumption rate data also showed similar values for these treatments. Similar results were obtained with the previously reported CMD (six bacteria) using aliphatics as carbon source (10 000 mg L-1) obtained from Maya crude oil, consumption rates of 282 mg L-1 d-1 in eleven days of culture were reported (Diaz-Ramirez et

The number of viable microorganisms was higher in cultures in liquid medium with the

These results indicated that the application of the standardized mixture of microorganisms (bacteria - fungal strains) was favorable in terms of overall growth. Considering the high biodegradation results, probably there was a synergistic effect between the populations present in the CC and F-DMC, as it has been previously reported with similarly defined mixed

UFC ml-1)

standardized mixture of bacterial strains and co-culture (0.36 - 8.4 ± 1.1 X108

CFU ml-1).

of oil-degrading and heterotrophic microorganisms were determined by

Natural Attenuation.

**3. Results and discussion**

technically feasible.

al. 2008).

**3.1. Biodegradation of diesel in liquid cultures**

compared to the initial values (<1x107

culture in liquid medium (Diaz-Ramirez et al. 2003).

In order to evaluate the biodegradability of diesel by exogenous and native hydrocarbondegrading microorganisms, an experimental trial was performed using artificially contami‐ nated soil (~ 120,000 ppm of diesel), which has been biostimulated by addition of a commercial fertilizer (Triple 17), in order to get an initial C/N ratio of 20. Two treatments were established for this assay; a) bioaugmented soil (inoculated with exogenous microorganisms), and b) nonbioaugmented soil (only with the native microbiota).

#### *2.2.1. Gas chromatography analysis*

Analytical procedure to determine residual hydrocarbons was based on the gas chromatog‐ raphy standard method EPA8015B. A gas chromatograph (Agilent Technologies Mod 6850 – II) equipped with FID and a column Agilent HP-1. Running conditions were Initial Temp: 70°C, Initial Hold 4.00 min, Ramp 1: 10°C/min to 70°C. Ramp 2: 40°C/min to 310°C, hold for 9.00 min. Injector temp: 250°C. Detector temp: 300°C. Helium was the carrying gas at: 10.00 psi. For calibration, standards were prepared with hydrocarbons known to be present in the used diesel, obtaining a standard curve with 8 quantification points.

#### **2.3. Olmeca crude oil biodegradation**

Assays were performed in trays containing 400 g of artificially contaminated soil (10,000 mg TPH kg-1 soil), amended (biostimulated) with 2% w/w sugar cane bagasse used as organic conditioner, and the commercial fertilizer Gro-green 20:30:10 (Campbell, Mexico) to obtain a C / N initial ratio of 6. Soil humidity was maintained at 40% of the water retention capacity of soil. Four treatments were established as follow: a) Bioaugmented - sterile soil (B-SS), b) Non bioaugmented - sterile soil (NB-SS), c) Bioaugmented - non-sterile soil (B-NSS), and d) Non bioaugmented - non-sterile soil (NB-NSS).

The following variables were determined: TPH content at different sampling times (EPA 3540 method), the number of hydrocarbon-degrading bacteria was assessed by the most probable number assay (Wrenn and Venosa, 1996), while total heterothophic microorganisms were determined after Lorch et al (1995). With the data regarding residual TPH content, the biodegradation extent was calculated, as well as the global and maximum consumption rates. Additionally, kinetic parameters were calculated as the adjustment of the consumption data to the first order model (as described above in the section 3.2). Biodegradation kinetics of diesel in soil), the biodegradation constants and half-life under different treatments were also obtained.

#### **2.4. Monitoring of natural attenuation**

A periodic sampling protocol was designed to monitoring the natural attenuation of a soil highly contaminated with gasoil (medium fraction petroleum). Based on criteria as site geomorphology and direction of the stream meanders; several points with presence of hydrocarbons were identified and selected for monitoring. The residual gasoil content (medium fraction C10-C28) was used as a measure of passive biodegradation or Monitored Natural Attenuation.

The residual gasoil content (medium fraction C10-C28) was used as a measure of passive biodegradation or Monitored Natural Attenuation. Soil hydrocarbon analyses were performed by Gas Chromatography accordingly to the Mexican regulation (NOM-138-SEMARNAT/ SS-2003), which is based on the EPA 8015b standard procedure. From these values, biodegra‐ dation extent and half life of the contaminant in soil were calculated. In addition, the amounts of oil-degrading and heterotrophic microorganisms were determined by

### **3. Results and discussion**

strains (F-DMC); *Aspergillus terreus* and *Aspergillus carneus.* Finally, a co-culture (CC) com‐ posed of all of the above strains was also tested. The degrading microorganisms were isolated and characterized as previously described (Diaz-Ramirez et al, 2003). In parallel, non-inocu‐

In order to evaluate the biodegradability of diesel by exogenous and native hydrocarbondegrading microorganisms, an experimental trial was performed using artificially contami‐ nated soil (~ 120,000 ppm of diesel), which has been biostimulated by addition of a commercial fertilizer (Triple 17), in order to get an initial C/N ratio of 20. Two treatments were established for this assay; a) bioaugmented soil (inoculated with exogenous microorganisms), and b) non-

Analytical procedure to determine residual hydrocarbons was based on the gas chromatog‐ raphy standard method EPA8015B. A gas chromatograph (Agilent Technologies Mod 6850 – II) equipped with FID and a column Agilent HP-1. Running conditions were Initial Temp: 70°C, Initial Hold 4.00 min, Ramp 1: 10°C/min to 70°C. Ramp 2: 40°C/min to 310°C, hold for 9.00 min. Injector temp: 250°C. Detector temp: 300°C. Helium was the carrying gas at: 10.00 psi. For calibration, standards were prepared with hydrocarbons known to be present in the

Assays were performed in trays containing 400 g of artificially contaminated soil (10,000 mg TPH kg-1 soil), amended (biostimulated) with 2% w/w sugar cane bagasse used as organic conditioner, and the commercial fertilizer Gro-green 20:30:10 (Campbell, Mexico) to obtain a C / N initial ratio of 6. Soil humidity was maintained at 40% of the water retention capacity of soil. Four treatments were established as follow: a) Bioaugmented - sterile soil (B-SS), b) Non bioaugmented - sterile soil (NB-SS), c) Bioaugmented - non-sterile soil (B-NSS), and d) Non

The following variables were determined: TPH content at different sampling times (EPA 3540 method), the number of hydrocarbon-degrading bacteria was assessed by the most probable number assay (Wrenn and Venosa, 1996), while total heterothophic microorganisms were determined after Lorch et al (1995). With the data regarding residual TPH content, the biodegradation extent was calculated, as well as the global and maximum consumption rates. Additionally, kinetic parameters were calculated as the adjustment of the consumption data to the first order model (as described above in the section 3.2). Biodegradation kinetics of diesel in soil), the biodegradation constants and half-life under different treatments were also

A periodic sampling protocol was designed to monitoring the natural attenuation of a soil highly contaminated with gasoil (medium fraction petroleum). Based on criteria as site

lated controls were prepared to quantify the hydrocarbons abiotic loss.

used diesel, obtaining a standard curve with 8 quantification points.

bioaugmented soil (only with the native microbiota).

*2.2.1. Gas chromatography analysis*

160 Biodegradation of Hazardous and Special Products

**2.3. Olmeca crude oil biodegradation**

bioaugmented - non-sterile soil (NB-NSS).

**2.4. Monitoring of natural attenuation**

obtained.

To assess the natural attenuation potential of contaminated soils, as well as to develop inocula for bioremediation of such sites, it is necessary to know the degrading abilities of microbial native populations. Where applicable, the persistence and activity of the introduced microorganisms, without bioavailability limitations attributable to the soil, have to be addressed. Contrast the biodegradation process under this scheme and in field is one of the main challenges for the definition of the auto recovery potential of contaminated soils, as well as for the establishment of bioremediation strategies that are economic and technically feasible.

#### **3.1. Biodegradation of diesel in liquid cultures**

Higher levels of biodegradation were obtained with the CC and F -DMC that behave very similar (Table 1). Statistical analysis did not show significant differences at 95% confidence among these two treatments. While the consumption rate data also showed similar values for these treatments. Similar results were obtained with the previously reported CMD (six bacteria) using aliphatics as carbon source (10 000 mg L-1) obtained from Maya crude oil, consumption rates of 282 mg L-1 d-1 in eleven days of culture were reported (Diaz-Ramirez et al. 2008).

The number of viable microorganisms was higher in cultures in liquid medium with the standardized mixture of bacterial strains and co-culture (0.36 - 8.4 ± 1.1 X108 UFC ml-1) compared to the initial values (<1x107 CFU ml-1).

These results indicated that the application of the standardized mixture of microorganisms (bacteria - fungal strains) was favorable in terms of overall growth. Considering the high biodegradation results, probably there was a synergistic effect between the populations present in the CC and F-DMC, as it has been previously reported with similarly defined mixed culture in liquid medium (Diaz-Ramirez et al. 2003).


These results indicated that the application of the standardized mixture of microorganisms (bacteria - fungal strains) was favorable in terms of overall growth. Considering the high biodegradation results, probably there was a synergistic effect between the

The higher degradation rates were observed during the first 30 days of culture, being 37.8% and 45.3% for the bioaugmented and non-bioaugmented soils, respectively. This rapid initial degradation is likely due to the consumption of the easily degradable compounds, for example C10 - C18 aliphatic hydrocarbons, and some low molecular weight aromatic compounds (Vallejo et al. 2005). Subsequently, between 30 and 90 days of cultivation in soil biodegradation and non-bio-enhanced bioaugmented and similarly inoculated was not held constant until the end of the culture (Figure 1), reaching ultimate biodegradation of 52% and 54% for bioaugmented and non-bioaugmented soil, respectively. No significant differences were found between the means of the two

Further analysis of these results showed that in the first 30 days of treatment there was a rapid decline in the concentration of hydrocarbons (diesel) in both; the bioaugmented and the non-bioaugmented soils, thereafter the biodegradation rate decreased. High activity of soil microbial populations has been frequently reported for biostimulated soils (Sabaté et al, 2004; Margesin, 2005, Chemlal et al. 2012). Based on colony-morphology observations, the microorganisms inoculated in the treated soil were prevailing at the end of the assay. The predominance of these hydrocarbon-degrading strains in liquid culture and solid media has been previously reported using molecular tools (Escalante-Espinosa, 2005; Diaz-Ramirez et al, 2008), confirming that inoculated strains

Based on a first order model degradation constant (-k) and half-life (t1 / 2) were determined (Eweis et al, 1998). Table 2 presents the

assayed conditions.

c

Where:

hydrocarbons from the assayed treatments.

**Treatment Biodegradation**

Biodegradation constant was calculated as:

*C*= Total petroleum hydrocarbon concentration at "*t*" time

*k* = Biodegradation constant represented as the slope of the *Ln C/C0 vs time* (days)*.*

**Table 2.** Kinetic parameters for the Diesel biodegradation assay in soil.

*C*0= Initial concentration of petroleum hydrocarbon

**(% )**

The higher degradation rates were observed during the first 30 days of culture, being 37.8% and 45.3% for the bioaugmented and non-bioaugmented soils, respectively. This rapid initial degradation is likely due to the consumption of the easily degradable compounds, for example C10 - C18 aliphatic hydrocarbons, and some low molecular weight aromatic compounds (Vallejo et al. 2005). Subsequently, between 30 and 90 days of culture, diesel biodegradation in bioaugmented and non- bioaugmented treatments was constant until the end of the culture (Figure 1), reaching ultimate biodegradation of 52% and 54% for bioaugmented and nonbioaugmented soil, respectively. No significant differences were found between the means of

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

Further analysis of these results showed that in the first 30 days of treatment there was a rapid decline in the concentration of hydrocarbons (diesel) in both; the bioaugmented and the nonbioaugmented soils, thereafter the biodegradation rate decreased. High activity of soil microbial populations has been frequently reported for biostimulated soils (Sabaté et al, 2004; Margesin, 2005, Chemlal et al. 2012). Based on colony-morphology observations, the microorganisms inoculated in the treated soil were prevailing at the end of the assay. The predominance of these hydrocarbon-degrading strains in liquid culture and solid media has been previously reported using molecular tools (Escalante-Espinosa, 2005; Diaz-Ramirez et al, 2008), confirming that inoculated strains have good capacity to remain in soils under the

Based on a first order model degradation constant (-k) and half-life (t1 / 2) were determined (Eweis et al, 1998). Table 2 presents the kinetic parameters calculated for the removal of diesel

> **GCRa (mg kg-1 d-1)**

Bioaugmented 52.0 ± 5.1 684.4 ± 67.6 2 598 ± 555 -0.0093 74.5 Non-bioaugmented 53.7 ± 5.4 707.7 ± 71.6 2 657 ±119 -0.0106 65.4

<sup>a</sup> Global Consumption Rate (GCR) was calculated considering the initial and residual hydrocarbon content after 90 days.

b Maximum Consumption Rate (MCR) was calculated as described in the kinetic parameter section below.

**MCR b (mg kg-1 d-1)**

*C* =*C*0*e*-*kt* (1)

**-k c (d-1)**

http://dx.doi.org/10.5772/56233

163

**Half life time (d)**

the two treatments for biodegradation of diesel oil (t test, P> 0.05).

**Table 1.** Diesel biodegradation by the different microbial inocula assayed during 16 days of incubation. Biodegradation extent was calculated considering an initial TPH concentration of 0.5 g/flask and corrected considering the diesel abiotic loss in the control. Values followed by the same letter did not show significant differences. considering an initial TPH concentration of 0.5 g/flask and corrected considering the diesel abiotic loss in the control. Values followed by the same letter did not show significant differences.

medium (Diaz-Ramirez et al. 2003).

#### **3.2. Biodegradation kinetics of diesel in soil 3.2. Biodegradation kinetics of diesel in soil**

In order to determine the biodegradation level of diesel in soil and the activity of the exogenous and native microbial populations, the residual oil content was determined during the biode‐ gradation assays, allowing to perform a kinetic study. Additionally, we quantified the number of heterotrophic microorganisms and soil toxicity at the end of treatment. In order to determine the biodegradation level of diesel in soil and the activity of the exogenous and native microbial populations, the residual oil content was determined during the biodegradation assays, allowing to perform a kinetic study. Additionally, we quantified the number of heterotrophic microorganisms and soil toxicity at the end of treatment.

Figure 1 shows the degradation kinetics of diesel in artificially contaminated soil (120,000 mg kg-1 soil), determined as diesel residual hydrocarbons. Figure 1 shows the degradation kinetics of diesel in artificially contaminated soil (120,000 mg kg-1 soil), determined as diesel residual hydrocarbons.

Figure 1. Soil hydrocarbon content throughout 90 day-assay, initial concentration was 120,000 mg Kg-1 for the bioaugmented () and non-**Figure 1.** Soil hydrocarbon content throughout 90 day-assay, initial concentration was 120,000 mg Kg-1 for the bio‐ augmented (■) and non-bioaugmented soil (◆).

treatments for biodegradation of diesel oil (t test, P> 0.05).

have good capacity to remain in soils under the assayed conditions.

kinetic parameters calculated for the removal of diesel hydrocarbons from the assayed treatments.

bioaugmented soil ().

**Consumption rate (mg L-1d-1)**  B-DMC 36.5 ± 6.8 a 156.63 ± 48.6 F-DMC 49.1 ± 2.8 b 204.88 ± 12.0 CC 48.3 ± 5.2 b 201.33 ± 22.0 Table 1. Diesel biodegradation by the different microbial inocula assayed during 16 days of incubation. Biodegradation extent was calculated considering an initial TPH concentration of 0.5 g/flask and corrected considering the diesel abiotic loss in the control. Values followed by the same The higher degradation rates were observed during the first 30 days of culture, being 37.8% and 45.3% for the bioaugmented and non-bioaugmented soils, respectively. This rapid initial degradation is likely due to the consumption of the easily degradable compounds, for example C10 - C18 aliphatic hydrocarbons, and some low molecular weight aromatic compounds (Vallejo et al. 2005). Subsequently, between 30 and 90 days of culture, diesel biodegradation in bioaugmented and non- bioaugmented treatments was constant until the end of the culture (Figure 1), reaching ultimate biodegradation of 52% and 54% for bioaugmented and nonbioaugmented soil, respectively. No significant differences were found between the means of the two treatments for biodegradation of diesel oil (t test, P> 0.05).

In order to determine the biodegradation level of diesel in soil and the activity of the exogenous and native microbial populations, the residual oil content was determined during the biodegradation assays, allowing to perform a kinetic study. Additionally, we quantified the number of heterotrophic microorganisms and soil toxicity at the end of treatment. Figure 1 shows the degradation kinetics of diesel in artificially contaminated soil (120,000 mg kg-1 soil), determined as diesel Further analysis of these results showed that in the first 30 days of treatment there was a rapid decline in the concentration of hydrocarbons (diesel) in both; the bioaugmented and the nonbioaugmented soils, thereafter the biodegradation rate decreased. High activity of soil microbial populations has been frequently reported for biostimulated soils (Sabaté et al, 2004; Margesin, 2005, Chemlal et al. 2012). Based on colony-morphology observations, the microorganisms inoculated in the treated soil were prevailing at the end of the assay. The predominance of these hydrocarbon-degrading strains in liquid culture and solid media has been previously reported using molecular tools (Escalante-Espinosa, 2005; Diaz-Ramirez et al, 2008), confirming that inoculated strains have good capacity to remain in soils under the assayed conditions.

> Based on a first order model degradation constant (-k) and half-life (t1 / 2) were determined (Eweis et al, 1998). Table 2 presents the kinetic parameters calculated for the removal of diesel hydrocarbons from the assayed treatments.


<sup>a</sup> Global Consumption Rate (GCR) was calculated considering the initial and residual hydrocarbon content after 90 days.

b Maximum Consumption Rate (MCR) was calculated as described in the kinetic parameter section below.

c Biodegradation constant was calculated as:

$$\mathbf{C} = \mathbf{C}\_{\mathbf{O}} \mathbf{e}^{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\cdot}}}}}}}}}}}}}}}}}}}}}}}}} \tag{}} \tag{} } \tag{} } \mathbf{O}} \tag{} } \tag{1} \text{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\mathrm{\cdot}}}}}}}}}}}}}} }} } } } } } } } } } $$
} } } } } 

Where:

**Microbial inocula Biodegradation extent**

medium (Diaz-Ramirez et al. 2003).

**3.2. Biodegradation kinetics of diesel in soil**

162 Biodegradation of Hazardous and Special Products

kg-1 soil), determined as diesel residual hydrocarbons.

bioaugmented soil ().

0

20,000

40,000

60,000

**mg diesel kg-1**

augmented (■) and non-bioaugmented soil (◆).

 **soil** 

80,000

100,000

120,000

140,000

residual hydrocarbons.

differences.

**(%)**

B-DMC 36.5 ± 6.8 a 156.63 ± 48.6

F-DMC 49.1 ± 2.8 b 204.88 ± 12.0

CC 48.3 ± 5.2 b 201.33 ± 22.0

In order to determine the biodegradation level of diesel in soil and the activity of the exogenous and native microbial populations, the residual oil content was determined during the biode‐ gradation assays, allowing to perform a kinetic study. Additionally, we quantified the number

**3.2. Biodegradation kinetics of diesel in soil** 

Figure 1 shows the degradation kinetics of diesel in artificially contaminated soil (120,000 mg

treatments for biodegradation of diesel oil (t test, P> 0.05).

0 20 40 60 80 100

**Time (days)**

**Figure 1.** Soil hydrocarbon content throughout 90 day-assay, initial concentration was 120,000 mg Kg-1 for the bio‐

have good capacity to remain in soils under the assayed conditions.

kinetic parameters calculated for the removal of diesel hydrocarbons from the assayed treatments.

**Table 1.** Diesel biodegradation by the different microbial inocula assayed during 16 days of incubation. Biodegradation extent was calculated considering an initial TPH concentration of 0.5 g/flask and corrected considering the diesel abiotic loss in the control. Values followed by the same letter did not show significant

letter did not show significant differences.

of heterotrophic microorganisms and soil toxicity at the end of treatment.

**Consumption rate (mg L-1d-1)**

**Microbial inocula Biodegradation extent** 

These results indicated that the application of the standardized mixture of microorganisms (bacteria - fungal strains) was favorable in terms of overall growth. Considering the high biodegradation results, probably there was a synergistic effect between the

**(%)** 

The higher degradation rates were observed during the first 30 days of culture, being 37.8% and 45.3% for the bioaugmented and non-bioaugmented soils, respectively. This rapid initial degradation is likely due to the consumption of the easily degradable compounds, for example C10 - C18 aliphatic hydrocarbons, and some low molecular weight aromatic compounds (Vallejo et al. 2005). Subsequently, between 30 and 90 days of cultivation in soil biodegradation and non-bio-enhanced bioaugmented and similarly inoculated was not held constant until the end of the culture (Figure 1), reaching ultimate biodegradation of 52% and 54% for bioaugmented and non-bioaugmented soil, respectively. No significant differences were found between the means of the two

Further analysis of these results showed that in the first 30 days of treatment there was a rapid decline in the concentration of hydrocarbons (diesel) in both; the bioaugmented and the non-bioaugmented soils, thereafter the biodegradation rate decreased. High activity of soil microbial populations has been frequently reported for biostimulated soils (Sabaté et al, 2004; Margesin, 2005, Chemlal et al. 2012). Based on colony-morphology observations, the microorganisms inoculated in the treated soil were prevailing at the end of the assay. The predominance of these hydrocarbon-degrading strains in liquid culture and solid media has been previously reported using molecular tools (Escalante-Espinosa, 2005; Diaz-Ramirez et al, 2008), confirming that inoculated strains

Based on a first order model degradation constant (-k) and half-life (t1 / 2) were determined (Eweis et al, 1998). Table 2 presents the

*C*= Total petroleum hydrocarbon concentration at "*t*" time

*C*0= Initial concentration of petroleum hydrocarbon

*k* = Biodegradation constant represented as the slope of the *Ln C/C0 vs time* (days)*.*

Figure 1. Soil hydrocarbon content throughout 90 day-assay, initial concentration was 120,000 mg Kg-1 for the bioaugmented () and non-**Table 2.** Kinetic parameters for the Diesel biodegradation assay in soil. According to the analysis the estimated biodegradation constant was similar for both treat‐ ments. These values are comparable to those reported by Cardona and Iturbe (2003) for the biodegradation of diesel in an agricultural soil, under biostimulated conditions, adding alternate sources of oxygen and nitrogen to the soil. The values for the biodegradation constants were 0.044 d-1 – 0.011 d-1, with concentrations ranging from 28 000 to 40 000 mg diesel kg-1 soil. The half-life time of spilled diesel in this soil was calculated, indicating that from 4.4 to 5 months were required to achieve a complete removal of hydrocarbons.

aliphatics (C10 - C24). Moreover, Marquez-Rocha et al. (2001), reported the degradation of medium sized hydrocarbons (> C12) contained in the diesel, together with the short-chain

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

This<\$%&?>chromatographic<\$%&?>analysis<\$%&?>is<\$%&?>consistent<\$%&?>with<\$%&?>the<\$%&?>results<\$%&?>obtained<\$ %&?>by<\$%&?>Hye<\$%&?>et<\$%&?>al.<\$%&?>(2005),<\$%&?>which<\$%&?>shows<\$%&?>a<\$%&?>comparison<\$%&?>of<\$%&?> diesel<\$%&?>before<\$%&?>and<\$%&?>after<\$%&?>culturing<\$%&?>with<\$%&?>a<\$%&?>*P.<\$%&?>aeruginosa*<\$%&?>strain<\$% &?>IU5,<\$%&?>suggesting<\$%&?>that<\$%&?>such<\$%&?>a<\$%&?>decrease<\$%&?>is<\$%&?>mainly<\$%&?>due<\$%&?>to<\$%&? >the<\$%&?>degradation<\$%&?>of<\$%&?>short<\$%&?>to<\$%&?>medium<\$%&?>chain<\$%&?>aliphatics<\$%&?>(C10<\$%&?>-

Rocha<\$%&?>et<\$%&?>al.<\$%&?>(2001),<\$%&?>reported<\$%&?>the<\$%&?>degradation<\$%&?>of<\$%&?>medium<\$%&?>sized<\$ %&?>hydrocarbons<\$%&?>(><\$%&?>C12)<\$%&?>contained<\$%&?>in<\$%&?>the<\$%&?>diesel,<\$%&?>together<\$%&?>with<\$%&

> (a) Initial Diesel composition

http://dx.doi.org/10.5772/56233

165

(b) Residual Diesel after 90 days Bioaugmented soil

(c) Residual Diesel after

Non bioaugmented soil

90 days

Figure 2. Gas<\$%&?>chromatography<\$%&?>of<\$%&?>Diesel<\$%&?>used<\$%&?>as<\$%&?>carbon<\$%&?>source<\$%&?>after<\$%&?>90<\$%&?>

bioaugmented<\$%&?>artificially<\$%&?>contaminated<\$%&?>soil.<\$%&?>Chromatography<\$%&?>analysis<\$%&?>conditions<\$%&?>were<\$%&?>

2003)<\$%&?>and<\$%&?>the<\$%&?>EPA<\$%&?>method<\$%&?>(EPA<\$%&?>8015b).<\$%&?>Initial<\$%&?>and<\$%&?>final<\$%&?>composition<\$

degrading<\$%&?>bacteria<\$%&?>was<\$%&?>assessed<\$%&?>by<\$%&?>the<\$%&?>most<\$%&?>probable<\$%&?>number<\$%&?> assay<\$%&?>(Wrenn<\$%&?>and<\$%&?>Venosa,<\$%&?>1996).<\$%&?>In<\$%&?>bioaugmented<\$%&?>soil<\$%&?>values<\$%&?> were<\$%&?>between<\$%&?>11.0<\$%&?>to<\$%&?>110<\$%&?>X<\$%&?>105<\$%&?>degrading<\$%&?>microorganisms<\$%&?>g-1<\$%&?>dry<\$%&?>soil,<\$%&?>and<\$%&?>for<\$%&?>the<\$%&?>non<\$%&?>bioaugmented<\$%&?>treatment<\$%&?>values<\$%&

**3.2.2.<\$%&?>Assessment<\$%&?>of<\$%&?>degrading<\$%&?>microorganisms<\$%&?>** 

days<\$%&?>of<\$%&?>culture<\$%&?>on<\$%&?>bioaugmented<\$%&?>and<\$%&?>non-

The<\$%&?>number<\$%&?>of<\$%&?>hydrocarbon-

treatments is shown for comparative analysis.

made<\$%&?>based<\$%&?>on<\$%&?>Mexican<\$%&?>normativity<\$%&?>(NOM-138-SEMARNAT/SS-

%&?>for<\$%&?>both<\$%&?>treatments<\$%&?>is<\$%&?>shown<\$%&?>for<\$%&?>comparative<\$%&?>analysis.

**Figure 2.** Gas chromatography of diesel used as carbon source after 90 days of culture on bioaugmented and nonbioaugmented artificially contaminated soil. Chromatography analysis conditions were made based on Mexican nor‐ mativity (NOM-138-SEMARNAT/SS-2003) and the EPA method (EPA 8015b). Initial and final composition for both

?>ranged<\$%&?>from<\$%&?>0.21<\$%&?>to<\$%&?>2.1<\$%&?>x<\$%&?>105<\$%&?>degraders<\$%&?>g-

compounds.

<\$%&?>C24).<\$%&?>Moreover,<\$%&?>Marquez-

?>the<\$%&?>short-chain<\$%&?>compounds.

In similar biodegradation assays, Hye Hong et al (2005), reported 60% degradation of diesel in 13 days using a *Pseudomonas aeruginosa* (IU5 strain) isolated from an oil-contaminated soil. Marquez-Rocha et al (2001), reported rates of biodegradation (GCR) of diesel (118 g kg-1 dry soil) of about 2 130 to 2 780 mg kg-1 diesel dry soil d-1, with a bacterial consortium previously acclimated to diesel. Diaz-Ramirez (2000) evaluated the biodegradation of hydrocarbons (medium fraction) using a bacterial consortium obtained from the rhizosphere of *Cyperus laxus* Lam., finding a 62% biodegradation within 30 days, the level of biodegradation was similar to the one achieved in this work; 37.8% after 30 days of treatment in the bioaugmented soil.

#### *3.2.1. Analysis of residual diesel in soil after 90 days*

Although the level of biodegradation of diesel in the bioaugmented and non-bioaugmented treatments was similar at the end of culture (Table 2), the composition profiles of the residual hydrocarbons were different in relation to the initial composition of diesel, as shown in the comparative chromatographic analysis (Figure 2).

The commercial diesel fuel used in the biodegradation assay is constituted by a mixture of paraffinic and olefinic hydrocarbons ranging from C10 to C28, plus 30% aromatics. The maximum total sulfur content is <0.05% (PEMEX, 2008). Under Mexican standards this type of diesel fraction is regarded as medium-fraction hydrocarbons, whose presence is limited to 1 200 ppm for agricultural soil, and 5 000 ppm in industrial soil (NOM-138-SEMARNAT/ SS-2003).

Figure 2a shows the initial composition of diesel fuel used in both treatments; the most prominent peaks corresponded to the alkanes present in larger proportion in the mixture. In bioaugmented soil (Figure 2b), there was a marked reduction in the number of peaks and the area under the curve after 90 days, whereas in the bioaugmented soil (Figure 2c) an increase in the area under the curve was observed. This later change indicated that the residual hydrocarbons were composed of a more complex mixture (unresolved complex mixture), characteristic of a soil with residual compounds of higher molecular weight and chemical complexity. In the bioaugmented treatment the main biotransformation of hydrocarbons was probably due to degradation of short chain compounds (low molecular weight) and mediumsized alkanes (Figure 2b).

This chromatographic analysis is consistent with the results obtained by Hye et al. (2005), which shows a comparison of diesel before and after culturing with a *P. aeruginosa* strain IU5, suggesting that such a decrease is mainly due to the degradation of short to medium chain This<\$%&?>chromatographic<\$%&?>analysis<\$%&?>is<\$%&?>consistent<\$%&?>with<\$%&?>the<\$%&?>results<\$%&?>obtained<\$

Rocha<\$%&?>et<\$%&?>al.<\$%&?>(2001),<\$%&?>reported<\$%&?>the<\$%&?>degradation<\$%&?>of<\$%&?>medium<\$%&?>sized<\$ %&?>hydrocarbons<\$%&?>(><\$%&?>C12)<\$%&?>contained<\$%&?>in<\$%&?>the<\$%&?>diesel,<\$%&?>together<\$%&?>with<\$%&

aliphatics (C10 - C24). Moreover, Marquez-Rocha et al. (2001), reported the degradation of medium sized hydrocarbons (> C12) contained in the diesel, together with the short-chain compounds. %&?>by<\$%&?>Hye<\$%&?>et<\$%&?>al.<\$%&?>(2005),<\$%&?>which<\$%&?>shows<\$%&?>a<\$%&?>comparison<\$%&?>of<\$%&?> diesel<\$%&?>before<\$%&?>and<\$%&?>after<\$%&?>culturing<\$%&?>with<\$%&?>a<\$%&?>*P.<\$%&?>aeruginosa*<\$%&?>strain<\$% &?>IU5,<\$%&?>suggesting<\$%&?>that<\$%&?>such<\$%&?>a<\$%&?>decrease<\$%&?>is<\$%&?>mainly<\$%&?>due<\$%&?>to<\$%&? >the<\$%&?>degradation<\$%&?>of<\$%&?>short<\$%&?>to<\$%&?>medium<\$%&?>chain<\$%&?>aliphatics<\$%&?>(C10<\$%&?>- <\$%&?>C24).<\$%&?>Moreover,<\$%&?>Marquez-

?>the<\$%&?>short-chain<\$%&?>compounds.

According to the analysis the estimated biodegradation constant was similar for both treat‐ ments. These values are comparable to those reported by Cardona and Iturbe (2003) for the biodegradation of diesel in an agricultural soil, under biostimulated conditions, adding alternate sources of oxygen and nitrogen to the soil. The values for the biodegradation constants were 0.044 d-1 – 0.011 d-1, with concentrations ranging from 28 000 to 40 000 mg diesel kg-1 soil. The half-life time of spilled diesel in this soil was calculated, indicating that from 4.4

In similar biodegradation assays, Hye Hong et al (2005), reported 60% degradation of diesel in 13 days using a *Pseudomonas aeruginosa* (IU5 strain) isolated from an oil-contaminated soil. Marquez-Rocha et al (2001), reported rates of biodegradation (GCR) of diesel (118 g kg-1 dry soil) of about 2 130 to 2 780 mg kg-1 diesel dry soil d-1, with a bacterial consortium previously acclimated to diesel. Diaz-Ramirez (2000) evaluated the biodegradation of hydrocarbons (medium fraction) using a bacterial consortium obtained from the rhizosphere of *Cyperus laxus* Lam., finding a 62% biodegradation within 30 days, the level of biodegradation was similar to the one achieved in this work; 37.8% after 30 days of treatment in the bioaugmented

Although the level of biodegradation of diesel in the bioaugmented and non-bioaugmented treatments was similar at the end of culture (Table 2), the composition profiles of the residual hydrocarbons were different in relation to the initial composition of diesel, as shown in the

The commercial diesel fuel used in the biodegradation assay is constituted by a mixture of paraffinic and olefinic hydrocarbons ranging from C10 to C28, plus 30% aromatics. The maximum total sulfur content is <0.05% (PEMEX, 2008). Under Mexican standards this type of diesel fraction is regarded as medium-fraction hydrocarbons, whose presence is limited to 1 200 ppm for agricultural soil, and 5 000 ppm in industrial soil (NOM-138-SEMARNAT/

Figure 2a shows the initial composition of diesel fuel used in both treatments; the most prominent peaks corresponded to the alkanes present in larger proportion in the mixture. In bioaugmented soil (Figure 2b), there was a marked reduction in the number of peaks and the area under the curve after 90 days, whereas in the bioaugmented soil (Figure 2c) an increase in the area under the curve was observed. This later change indicated that the residual hydrocarbons were composed of a more complex mixture (unresolved complex mixture), characteristic of a soil with residual compounds of higher molecular weight and chemical complexity. In the bioaugmented treatment the main biotransformation of hydrocarbons was probably due to degradation of short chain compounds (low molecular weight) and medium-

This chromatographic analysis is consistent with the results obtained by Hye et al. (2005), which shows a comparison of diesel before and after culturing with a *P. aeruginosa* strain IU5, suggesting that such a decrease is mainly due to the degradation of short to medium chain

to 5 months were required to achieve a complete removal of hydrocarbons.

*3.2.1. Analysis of residual diesel in soil after 90 days*

164 Biodegradation of Hazardous and Special Products

comparative chromatographic analysis (Figure 2).

soil.

SS-2003).

sized alkanes (Figure 2b).

bioaugmented<\$%&?>artificially<\$%&?>contaminated<\$%&?>soil.<\$%&?>Chromatography<\$%&?>analysis<\$%&?>conditions<\$%&?>were<\$%&?> made<\$%&?>based<\$%&?>on<\$%&?>Mexican<\$%&?>normativity<\$%&?>(NOM-138-SEMARNAT/SS-2003)<\$%&?>and<\$%&?>the<\$%&?>EPA<\$%&?>method<\$%&?>(EPA<\$%&?>8015b).<\$%&?>Initial<\$%&?>and<\$%&?>final<\$%&?>composition<\$ %&?>for<\$%&?>both<\$%&?>treatments<\$%&?>is<\$%&?>shown<\$%&?>for<\$%&?>comparative<\$%&?>analysis. **Figure 2.** Gas chromatography of diesel used as carbon source after 90 days of culture on bioaugmented and nonbioaugmented artificially contaminated soil. Chromatography analysis conditions were made based on Mexican nor‐ mativity (NOM-138-SEMARNAT/SS-2003) and the EPA method (EPA 8015b). Initial and final composition for both treatments is shown for comparative analysis.

?>ranged<\$%&?>from<\$%&?>0.21<\$%&?>to<\$%&?>2.1<\$%&?>x<\$%&?>105<\$%&?>degraders<\$%&?>g-

**3.2.2.<\$%&?>Assessment<\$%&?>of<\$%&?>degrading<\$%&?>microorganisms<\$%&?>** 

days<\$%&?>of<\$%&?>culture<\$%&?>on<\$%&?>bioaugmented<\$%&?>and<\$%&?>non-

The<\$%&?>number<\$%&?>of<\$%&?>hydrocarbon-

Figure 2. Gas<\$%&?>chromatography<\$%&?>of<\$%&?>Diesel<\$%&?>used<\$%&?>as<\$%&?>carbon<\$%&?>source<\$%&?>after<\$%&?>90<\$%&?>

degrading<\$%&?>bacteria<\$%&?>was<\$%&?>assessed<\$%&?>by<\$%&?>the<\$%&?>most<\$%&?>probable<\$%&?>number<\$%&?> assay<\$%&?>(Wrenn<\$%&?>and<\$%&?>Venosa,<\$%&?>1996).<\$%&?>In<\$%&?>bioaugmented<\$%&?>soil<\$%&?>values<\$%&?> were<\$%&?>between<\$%&?>11.0<\$%&?>to<\$%&?>110<\$%&?>X<\$%&?>105<\$%&?>degrading<\$%&?>microorganisms<\$%&?>g-1<\$%&?>dry<\$%&?>soil,<\$%&?>and<\$%&?>for<\$%&?>the<\$%&?>non<\$%&?>bioaugmented<\$%&?>treatment<\$%&?>values<\$%&

#### *3.2.2. Assessment of degrading microorganisms*

The number of hydrocarbon-degrading bacteria was assessed by the most probable number assay (Wrenn and Venosa, 1996). In bioaugmented soil values were between 11.0 to 110 X 105 degrading microorganisms g-1 dry soil, and for the non bioaugmented treatment values ranged from 0.21 to 2.1 x 105 degraders g-1 dry soil. Considering that the used soil has no history of contamination, the number of diesel degraders in both treatments is high. Similar level of degraders (2.1x105 g-1 dry soil) was reported by Vallejo et al (2005) in their evaluation of biostimulation on PAH biodegradation of petroleum contaminated soils.

**Treatment Toxicity**

**3.3. Biodegradation assay of Olmeca crude oil in soil**

*3.3.1. Biodegradation kinetics of Olmeca crude oil in soil*

bioaugmented treatments, after 90 days of culture.

\*Toxicity unit = (1/EC50) x106.

biodegradation experiments.

37.6 ± 2%, respectively.

consumption of oil.

treatments that presented similar behavior.

**(TU)\***

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

Bioaugmented 13.7 ± 2.8 75 218 ± 17 679 Non-bioaugmented 17.0 ± 0.2 59 010 ± 762.2

**Table 3.** Toxicity analysis results for the artificially diesel-contaminated soil, under bioaugmented and non-

Among the different types of oil produced in Mexico are the Maya heavy oil with gravity of 22° API (3.3% sulfur by weight); the Istmo light oil with 33.6° API (1.3% of sulfur by weight) and the Olmeca super light oil with API gravity of 39.3 ° (0.8% sulfur by weight). Olmeca and Maya oils are produced in oil fields located in the southeastern region of the country, in the states of Veracruz, Tabasco and Chiapas. There are several sites affected by the accidental discharge of either Maya and Olmeca oils or their refined products. The removal efficiency of the co-culture (CC) of degrading bacteria and fungi was evaluated by addressing residual total petroleum hydrocarbons (TPH) from Olmeca crude oil. The CC was the same used in the diesel

The results corresponding to biodegradation and concentrations of residual and con‐ sumed hydrocarbons during the 90 days assay are shown in Figure 3. Among the four treatments tested, both bioaugmented treatments recorded the highest levels of biodegrada‐ tion (panels a and c), being 58.3 ± 3% and 55.8 ± 2%, respectively. While in the nonbioaugmented treatments (panels b and d) the final biodegradation was about 48 ± 2% and

The period of increased oil consumption corresponded to days 1 to 45 of assay, for the bioaugmented treatments. This consumption was less accelerated for NB-SS, where after 22 days of treatment, oil removal decreased considerably (Figure 3b). Furthermore, in the B-NSS treatment (Figure 3c), there is a marked decrease of the residual hydrocarbon concentration, between days 1 to 60, which is less pronounced in the non bioaugmented treatments (Figure 3b and 3d). These two treatments also showed a marked decrease in the oil removal trend from day 60 to the end of the assay (90 days). In the case of bioaugmented treatments, in this same period (60 to 90 days) was still detected a slight

Values for residual hydrocarbon content at the end of the assay were statistically analyzed (ANOVA, F-test, p<0.05), bioaugmented treatments showed significant differences compared to the non- bioaugmented. However, no differences were found between both bioaugmented

**EC50 (ppm)**

http://dx.doi.org/10.5772/56233

167

#### *3.2.3. Analysis of toxicity*

In order to evaluate the acute toxicity of soil due to the presence of diesel and thus the effect of microbial activity (biodegradation) during the assay, the final acute toxicity of soil samples under both treatments was addressed. The measurements were made using the Microtox ® bioassay under the Mexican regulation procedure (NMX-AA 112-SECOFI, 1995). Soil samples were prepared following the Toxicity Characteristic Leaching Procedure (TCLP) as described in NOM-053-ECOL-1993. Toxicity was calculated based on the Relative Index of Toxicity (RIT) for contaminated soils located in Mexican southeastern. RIT is a relation between the half effective concentration (EC50) and the toxicity level for contaminated soils (Adams and Guzman-Osorio 2008). In tropical soils, hydrocarbons identified as medium fraction (C10 - C28) have a low toxicity in low concentrations (<10 000 ppm), mainly due to the low molecular weight compounds, with short half-life (2 - 4 months) under natural conditions. In contrast, at higher concentrations (> 10,000 ppm), toxic effects are seen together with changes in the physicochemical properties and functions of soil, like; a decrease in nutrient retention capacity and fertility, with further increases in compaction, repellency and salinization (Adams et al. 2008). Many contaminated sites in the southeast of the country have very high hydrocarbon concentrations (> 25,000 ppm) and a high degree of weathering.

The index provides a null level of toxicity for leachates of soils with more than 95 000 ppm; slightly toxic for soil leachates ranging from 84 700 to 58 900 ppm, while levels ranging between 58 900 and 36 000 ppm are regarded as a toxic. This toxicity levels can be transformed into Toxicity Units (TU= (1/EC50) x106 ). The EC50 is the concentration causing a 50% decrease in the bioluminescence level of test organism.

Toxicity results are presented in Table 3, soil samples of both treatments showed a toxicity of about 15 TU. Regarding the EC50 calculated by comparing the results with those described in Table 3, both values lie within the range indicated as slightly toxic for soils in this region.

Results of low or null toxicity level as determined by the Microtox bioassay have been previously reported for hydrocarbon contaminated clayed soils from the same region (Adams et al. 1999), that may be due to a strong degree of sorption of diesel into the soil, in addition to some degree of degradation of the residual compounds. In our case, biodegradation and weathering processes occurring in both treatments probably generated compounds that increased the toxicity of soil leachates, as compared to the initial TU=2.0 value, equivalent to an EC50 of 179,317 ppm.

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils http://dx.doi.org/10.5772/56233 167


**Table 3.** Toxicity analysis results for the artificially diesel-contaminated soil, under bioaugmented and nonbioaugmented treatments, after 90 days of culture.

#### **3.3. Biodegradation assay of Olmeca crude oil in soil**

*3.2.2. Assessment of degrading microorganisms*

166 Biodegradation of Hazardous and Special Products

ranged from 0.21 to 2.1 x 105

*3.2.3. Analysis of toxicity*

Toxicity Units (TU= (1/EC50) x106

an EC50 of 179,317 ppm.

bioluminescence level of test organism.

105

The number of hydrocarbon-degrading bacteria was assessed by the most probable number assay (Wrenn and Venosa, 1996). In bioaugmented soil values were between 11.0 to 110 X

of contamination, the number of diesel degraders in both treatments is high. Similar level of degraders (2.1x105 g-1 dry soil) was reported by Vallejo et al (2005) in their evaluation of

In order to evaluate the acute toxicity of soil due to the presence of diesel and thus the effect of microbial activity (biodegradation) during the assay, the final acute toxicity of soil samples under both treatments was addressed. The measurements were made using the Microtox ® bioassay under the Mexican regulation procedure (NMX-AA 112-SECOFI, 1995). Soil samples were prepared following the Toxicity Characteristic Leaching Procedure (TCLP) as described in NOM-053-ECOL-1993. Toxicity was calculated based on the Relative Index of Toxicity (RIT) for contaminated soils located in Mexican southeastern. RIT is a relation between the half effective concentration (EC50) and the toxicity level for contaminated soils (Adams and Guzman-Osorio 2008). In tropical soils, hydrocarbons identified as medium fraction (C10 - C28) have a low toxicity in low concentrations (<10 000 ppm), mainly due to the low molecular weight compounds, with short half-life (2 - 4 months) under natural conditions. In contrast, at higher concentrations (> 10,000 ppm), toxic effects are seen together with changes in the physicochemical properties and functions of soil, like; a decrease in nutrient retention capacity and fertility, with further increases in compaction, repellency and salinization (Adams et al. 2008). Many contaminated sites in the southeast of the country have very high hydrocarbon

The index provides a null level of toxicity for leachates of soils with more than 95 000 ppm; slightly toxic for soil leachates ranging from 84 700 to 58 900 ppm, while levels ranging between 58 900 and 36 000 ppm are regarded as a toxic. This toxicity levels can be transformed into

Toxicity results are presented in Table 3, soil samples of both treatments showed a toxicity of about 15 TU. Regarding the EC50 calculated by comparing the results with those described in Table 3, both values lie within the range indicated as slightly toxic for soils in this region.

Results of low or null toxicity level as determined by the Microtox bioassay have been previously reported for hydrocarbon contaminated clayed soils from the same region (Adams et al. 1999), that may be due to a strong degree of sorption of diesel into the soil, in addition to some degree of degradation of the residual compounds. In our case, biodegradation and weathering processes occurring in both treatments probably generated compounds that increased the toxicity of soil leachates, as compared to the initial TU=2.0 value, equivalent to

). The EC50 is the concentration causing a 50% decrease in the

biostimulation on PAH biodegradation of petroleum contaminated soils.

concentrations (> 25,000 ppm) and a high degree of weathering.

degrading microorganisms g-1 dry soil, and for the non bioaugmented treatment values

degraders g-1 dry soil. Considering that the used soil has no history

Among the different types of oil produced in Mexico are the Maya heavy oil with gravity of 22° API (3.3% sulfur by weight); the Istmo light oil with 33.6° API (1.3% of sulfur by weight) and the Olmeca super light oil with API gravity of 39.3 ° (0.8% sulfur by weight). Olmeca and Maya oils are produced in oil fields located in the southeastern region of the country, in the states of Veracruz, Tabasco and Chiapas. There are several sites affected by the accidental discharge of either Maya and Olmeca oils or their refined products. The removal efficiency of the co-culture (CC) of degrading bacteria and fungi was evaluated by addressing residual total petroleum hydrocarbons (TPH) from Olmeca crude oil. The CC was the same used in the diesel biodegradation experiments.

#### *3.3.1. Biodegradation kinetics of Olmeca crude oil in soil*

The results corresponding to biodegradation and concentrations of residual and con‐ sumed hydrocarbons during the 90 days assay are shown in Figure 3. Among the four treatments tested, both bioaugmented treatments recorded the highest levels of biodegrada‐ tion (panels a and c), being 58.3 ± 3% and 55.8 ± 2%, respectively. While in the nonbioaugmented treatments (panels b and d) the final biodegradation was about 48 ± 2% and 37.6 ± 2%, respectively.

The period of increased oil consumption corresponded to days 1 to 45 of assay, for the bioaugmented treatments. This consumption was less accelerated for NB-SS, where after 22 days of treatment, oil removal decreased considerably (Figure 3b). Furthermore, in the B-NSS treatment (Figure 3c), there is a marked decrease of the residual hydrocarbon concentration, between days 1 to 60, which is less pronounced in the non bioaugmented treatments (Figure 3b and 3d). These two treatments also showed a marked decrease in the oil removal trend from day 60 to the end of the assay (90 days). In the case of bioaugmented treatments, in this same period (60 to 90 days) was still detected a slight consumption of oil.

Values for residual hydrocarbon content at the end of the assay were statistically analyzed (ANOVA, F-test, p<0.05), bioaugmented treatments showed significant differences compared to the non- bioaugmented. However, no differences were found between both bioaugmented treatments that presented similar behavior.

Regarding the number of microorganisms, all treatments remained at levels higher than 1x106 CFU g-1 dry soil. Maximum values were obtained at 60 days in the bioaugmented trays (ca 1x108 CFU g-1 dry soil). Fertilizer was added to all trays at the start of assay, this fact probably limited the efficiency of biodegradation, as after 60 days of assay there was a decrease in the trend of oil removal (Fig. 3a – 3c).

**Treatment Biodegradation (%)**

Global Consumption Rate (GCR) calculated as:

a

Where:

*TPH final*

*TPH inital*

*t final*

Where:

*tn*= Sampling at "t" time

*tn*-1= Sampling time previous to *tn*

treatments after 90 days of assay.

*t*0= initial time

*GCR:* Global consumption rate

= final experimental time

b Maximum Consumption Rate (MCR) calculated as:

**GCR a (mg kg-1 d-1)**

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

é ù - ë û <sup>=</sup> - (2)

é ù - ë û <sup>=</sup> - (3)

> 0.94), in contrast, residual hydrocarbon

~ 0.7).

B-SS 55.8 ± 2.0 69.1 259.4 NB-SS 37.6 ± 2.3 46.6 210.6 B-NSS 58.3 ± 3.2 72.3 142.6 NB-NSS 48.1 ± 2.3 59.6 150.5

0

*GCR*

= Total petroleum hydrocarbon concentration at the end of the assay

*MCR*

*MCR:* Maximum consumption rate determined between each sampling interval

*TPH <sup>r</sup> <sup>n</sup>*= Residual total petroleum hydrocarbon concentration at *tn*

*TPH <sup>r</sup> <sup>n</sup>*-1= = Residual total petroleum hydrocarbon concentration at *tn*-1

order linear fit performed for the different treatments.

treatments (B-SS, B-NSS) and even for NB-NSS (r2

concentration in NB-SS samples showed a non-linear behavior (r2

= Initial total petroleum hydrocarbon concentration

*TPH TPH*

*t t*

1 [ ] *<sup>r</sup> <sup>n</sup> r n n n TPH TPH*


**Table 4.** Olmeca oil biodegradation, global and maximum consumption rates, calculated for the different soil

In order to calculate quantitative kinetic parameters, the obtained quantitative data were adjusted to a first order model equation (see section 2.2.2 Biodegradation kinetics of diesel in soil). Table 5 presents the biodegradation constant and half life time obtained from the first-

The better fit to first order model was achieved with the data obtained from bioaugmented

*t t*

[ ] *final inital final*

1


**MCR b (mg kg-1 d-1)** 169

http://dx.doi.org/10.5772/56233

14 Book Title

20 21 **Figure 3.** Kinetics of hydrocarbons consumption in soil under different treatments: a) 22 bioaugmented - sterile soil (B-SS), b) non bioaugmented - sterile soil (NB-SS), c) 23 bioaugmented – non sterile soil (B-NSS), d) non bioaugmented – non sterile soil (NB-NSS). 24 Curves represent values of biodegradation (), consumed hydrocarbons () and residual 25 hydrocarbons (). The values are the average of three replicates at each sampling time. **Figure 3.** Kinetics of hydrocarbons consumption in soil under different treatments: a) bioaugmented - sterile soil (B-SS), b) non bioaugmented - sterile soil (NB-SS), c) bioaugmented – non sterile soil (B-NSS), d) non bioaugmented – non sterile soil (NB-NSS). Curves represent values of biodegradation (◆), consumed hydrocarbons (■) and residual hydro‐ carbons (▲). The values are the average of three replicates at each sampling time. The data were corrected consider‐ ing extractable fat content of the soil.

#### 27 *3.3.2. Kinetic parameters for biodegradation of Olmeca crude oil*

trays (ca 1x10<sup>8</sup>

26 The data were corrected considering extractable fat content of the soil.

29 (ANOVA, F-test, p<0.05), bioaugmented treatments showed significant differences 30 compared to the non- bioaugmented. However, no differences were found between both 31 bioaugmented treatments that presented similar behavior. 32 Regarding the number of microorganisms, all treatments remained at levels higher than 1x106 CFU g-1 33 dry soil. Maximum values were obtained at 60 days in the bioaugmented With the hydrocarbons consumption data we calculated the following consumption rates: a) global consumption rate; b) maximum consumption rate. Both parameters are volumetric values representing the biodegradation rate allowing a more representative comparison (Diaz-Ramirez et al. 2008). Data are presented in Table 4.

28 Values for residual hydrocarbon content at the end of the assay were statistically analyzed

 CFU g-1 34 dry soil). Fertilizer was added to all trays at the start of assay, this 35 fact probably limited the efficiency of biodegradation, as after 60 days of assay there was a 36 decrease in the trend of oil removal (Fig. 3a – 3c). Samples of the B-NSS treatment reached maximum consumption rate after 22 days of experi‐ ment (data not shown), while for samples B-SS and NB-SS (sterile soils) the maximum rate was obtained later (between 22 and 45 days).

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils http://dx.doi.org/10.5772/56233 169


a Global Consumption Rate (GCR) calculated as:

$$GCR = \frac{\left[TPH\_{final}\right] - \left[TPH\_{initial}\right]}{t\_0 - t\_{final}}\tag{2}$$

Where:

Regarding the number of microorganisms, all treatments remained at levels higher than

limited the efficiency of biodegradation, as after 60 days of assay there was a decrease in the

**a) b)**

**c) d)**

21 **Figure 3.** Kinetics of hydrocarbons consumption in soil under different treatments: a) 22 bioaugmented - sterile soil (B-SS), b) non bioaugmented - sterile soil (NB-SS), c)

26 The data were corrected considering extractable fat content of the soil.

31 bioaugmented treatments that presented similar behavior.

*3.3.2. Kinetic parameters for biodegradation of Olmeca crude oil*

36 decrease in the trend of oil removal (Fig. 3a – 3c).

Ramirez et al. 2008). Data are presented in Table 4.

23 bioaugmented – non sterile soil (B-NSS), d) non bioaugmented – non sterile soil (NB-NSS). 24 Curves represent values of biodegradation (), consumed hydrocarbons () and residual 25 hydrocarbons (). The values are the average of three replicates at each sampling time.

**Figure 3.** Kinetics of hydrocarbons consumption in soil under different treatments: a) bioaugmented - sterile soil (B-SS), b) non bioaugmented - sterile soil (NB-SS), c) bioaugmented – non sterile soil (B-NSS), d) non bioaugmented – non sterile soil (NB-NSS). Curves represent values of biodegradation (◆), consumed hydrocarbons (■) and residual hydro‐ carbons (▲). The values are the average of three replicates at each sampling time. The data were corrected consider‐

28 Values for residual hydrocarbon content at the end of the assay were statistically analyzed 29 (ANOVA, F-test, p<0.05), bioaugmented treatments showed significant differences 30 compared to the non- bioaugmented. However, no differences were found between both

With the hydrocarbons consumption data we calculated the following consumption rates: a) global consumption rate; b) maximum consumption rate. Both parameters are volumetric values representing the biodegradation rate allowing a more representative comparison (Diaz-

Samples of the B-NSS treatment reached maximum consumption rate after 22 days of experi‐ ment (data not shown), while for samples B-SS and NB-SS (sterile soils) the maximum rate was

32 Regarding the number of microorganisms, all treatments remained at levels higher than

CFU g-1 33 dry soil. Maximum values were obtained at 60 days in the bioaugmented

 CFU g-1 34 dry soil). Fertilizer was added to all trays at the start of assay, this 35 fact probably limited the efficiency of biodegradation, as after 60 days of assay there was a

14 Book Title

CFU g-1 dry soil. Maximum values were obtained at 60 days in the bioaugmented trays

CFU g-1 dry soil). Fertilizer was added to all trays at the start of assay, this fact probably

1x106

(ca 1x108

27

1x106

trays (ca 1x10<sup>8</sup>

obtained later (between 22 and 45 days).

ing extractable fat content of the soil.

trend of oil removal (Fig. 3a – 3c).

168 Biodegradation of Hazardous and Special Products

*GCR:* Global consumption rate

*TPH final* = Total petroleum hydrocarbon concentration at the end of the assay

*TPH inital* = Initial total petroleum hydrocarbon concentration

*t*0= initial time

*t final* = final experimental time

b Maximum Consumption Rate (MCR) calculated as:

$$\text{MCRR} = \frac{\left[TPH\_r\right]\_{\text{fl}} - \left[TPH\_r\right]\_{\text{n-1}}}{t\_{n-1} - t\_n} \tag{3}$$

Where:

*MCR:* Maximum consumption rate determined between each sampling interval

*TPH <sup>r</sup> <sup>n</sup>*= Residual total petroleum hydrocarbon concentration at *tn*

*TPH <sup>r</sup> <sup>n</sup>*-1= = Residual total petroleum hydrocarbon concentration at *tn*-1

*tn*= Sampling at "t" time

*tn*-1= Sampling time previous to *tn*

**Table 4.** Olmeca oil biodegradation, global and maximum consumption rates, calculated for the different soil treatments after 90 days of assay.

In order to calculate quantitative kinetic parameters, the obtained quantitative data were adjusted to a first order model equation (see section 2.2.2 Biodegradation kinetics of diesel in soil). Table 5 presents the biodegradation constant and half life time obtained from the firstorder linear fit performed for the different treatments.

The better fit to first order model was achieved with the data obtained from bioaugmented treatments (B-SS, B-NSS) and even for NB-NSS (r2 > 0.94), in contrast, residual hydrocarbon concentration in NB-SS samples showed a non-linear behavior (r2 ~ 0.7).

Similarly to the biodegradation experiments of diesel in soil, it was determined a degradation constant (-k) and the half life time average data of three replicate trays are presented in Table 5.

of this area is its diverse basins containing most of the country's water resources. The oil spill occurred on a site adjacent to a river system consisting of two streams and a river that flows into the upper Coatzacoalcos River basin. The first of the streams was the most affected due to the increased flow of hydrocarbons. The stream surrounding area has certain degree of conservation and ecological characteristics of a healthy forest area, with silty riverbed and sediments. Based on current Mexican regulations for soils contaminated with hydrocarbons, characterization and site diagnosis was required to propose alternatives for restoration of the affected areas. Particular interest was put in determining gasoil passive biodegradation potential under field conditions, in a series of points located along the stream and close to the spill site. Table 6 shows the results of three points selected for MNA of the gasoil spill occurred

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

**Gasoil concentration (mg Kg-1)a**

P 2 383 1 985 96.6 73.0

P 9 383 3 351 65.2 192.5

P 10 383 6 715 80.6 51.7

Sediment samples and superficial soil (0-10 cm) were collected from sampling points 9 and 10 (P9 and P10). Soil samples from 30 - 45 cm depth were taken from sampling point 2 (P2). Residual hydrocarbons data were well described by the first order model having a high lineal

P 2 0 59 626 0 P 2 36 44 092 26.0 P 2 87 9 670 83.8 P 2 117 ND ND

P 9 87 3 846 60.5 P 9 117 3 583 62.8

P 10 0 34 678 0 P 10 36 14 817 57.2 P 10 87 12 033 65.3 P 10 117 7 486 78.4

Half life time obtained from first order adjustment of the soil residual concentration data.

**Table 6.** MNA parameters of gasoil measured at three sampling points of the spill-impacted area.

 Residual gasoil determined as medium fraction (C10 – C28) EPA 8015b. b Biodegradation calculated considering the initial content of gasoil.

**Biodegradation extent (%)b**

**Half life time (d)c**

http://dx.doi.org/10.5772/56233

171

in this area.

a

c

**Sampling point Sampling time**

**(d)**

P 9 0 9 627 P 9 36 12 682


**Table 5.** Biodegradation constant and half-life time for Olmeca crude oil in soil, after a 90-day assay.

The biodegradation constant values were consistent with the results described, being higher (1.2 to 1.5 times) in the bioaugmented treatments (B-SS, B-NSS). The half life time was over 100 days (3.4 months) to the NB-NSS, while this value was slightly above 60 days (2 months) in the bioaugmented treatments. This indicates that it would take about 4 months to achieve complete biodegradation of the carbon source by applying the tested inoculum, regardless the soil is previously sterilized. For TPH (1,010 – 2,141 mg kg-1), first-order biodegradation rate constants were obtained by Chang et al. (2010) in similar range (0.011 - 0.024 d-1).

It is noteworthy that as in the case of diesel biodegradation (Figure 1), with Olmeca crude oil also was observed a decrease in the consumption rate after 60 days of assay (Figure 3). It is likely due to the reduction of the easily assimilable compounds after this period, staying in the soil compounds more resistant to biodegradation. Another possibility is a reduction in the availability of secondary nutrients and minerals, provided by the sugar cane bagasse and commercial fertilizer, respectively, restricting the removal of oil compounds still present.

The results obtained are comparable to those obtained in similar studies that used Diesel as carbon source and added inocula to promote biodegradation. Margesin et al, (2007) reported similar results, finding that with concentrations of 10,000 mg TPH (diesel) Kg-1 soil, the biodegradation was 31% to 53% in 38 days, assayed soil was biostimulated using inorganic fertilization with NPK nutrients (C/N ratio of 20:1) or oleophilic fertilization with Inipol EAP22. Mishra et al, (2001), tested the inorganic nutrient addition and the bioaugmentation using a defined mixed culture composed by five oily-sludge-degrading indigenous bacterial strains, these strains belong to genus *Acinetobacter (two strains), Burkholderia and Pseudomonas*, one of them was not identified. It inoculum was used for *in situ* bioremediation of hydrocarbons, finding a 48.5% TPH decrease in presence of the defined mixed culture and 17% in the biostimulated (non-inoculated) treatment after 120 days.

#### **3.4. Monitoring of Natural Attenuation (MNA) of contaminated soil and sediments in a riparian environment**

A severe gasoil spill (TPH-medium fraction) affected soils and rivers in a mountainous area southeast Mexico. In this tropical region most soils have silt-clayed texture characteristics, being the main human activities livestock and rain fed agriculture. Another distinctive feature of this area is its diverse basins containing most of the country's water resources. The oil spill occurred on a site adjacent to a river system consisting of two streams and a river that flows into the upper Coatzacoalcos River basin. The first of the streams was the most affected due to the increased flow of hydrocarbons. The stream surrounding area has certain degree of conservation and ecological characteristics of a healthy forest area, with silty riverbed and sediments. Based on current Mexican regulations for soils contaminated with hydrocarbons, characterization and site diagnosis was required to propose alternatives for restoration of the affected areas. Particular interest was put in determining gasoil passive biodegradation potential under field conditions, in a series of points located along the stream and close to the spill site. Table 6 shows the results of three points selected for MNA of the gasoil spill occurred in this area.


a Residual gasoil determined as medium fraction (C10 – C28) EPA 8015b.

b Biodegradation calculated considering the initial content of gasoil.

Similarly to the biodegradation experiments of diesel in soil, it was determined a degradation constant (-k) and the half life time average data of three replicate trays are presented in Table 5.

> **-k (d-1)**

The biodegradation constant values were consistent with the results described, being higher (1.2 to 1.5 times) in the bioaugmented treatments (B-SS, B-NSS). The half life time was over 100 days (3.4 months) to the NB-NSS, while this value was slightly above 60 days (2 months) in the bioaugmented treatments. This indicates that it would take about 4 months to achieve complete biodegradation of the carbon source by applying the tested inoculum, regardless the soil is previously sterilized. For TPH (1,010 – 2,141 mg kg-1), first-order biodegradation rate

It is noteworthy that as in the case of diesel biodegradation (Figure 1), with Olmeca crude oil also was observed a decrease in the consumption rate after 60 days of assay (Figure 3). It is likely due to the reduction of the easily assimilable compounds after this period, staying in the soil compounds more resistant to biodegradation. Another possibility is a reduction in the availability of secondary nutrients and minerals, provided by the sugar cane bagasse and commercial fertilizer, respectively, restricting the removal of oil compounds still present.

The results obtained are comparable to those obtained in similar studies that used Diesel as carbon source and added inocula to promote biodegradation. Margesin et al, (2007) reported similar results, finding that with concentrations of 10,000 mg TPH (diesel) Kg-1 soil, the biodegradation was 31% to 53% in 38 days, assayed soil was biostimulated using inorganic fertilization with NPK nutrients (C/N ratio of 20:1) or oleophilic fertilization with Inipol EAP22. Mishra et al, (2001), tested the inorganic nutrient addition and the bioaugmentation using a defined mixed culture composed by five oily-sludge-degrading indigenous bacterial strains, these strains belong to genus *Acinetobacter (two strains), Burkholderia and Pseudomonas*, one of them was not identified. It inoculum was used for *in situ* bioremediation of hydrocarbons, finding a 48.5% TPH decrease in presence of the defined mixed culture and 17% in the

**3.4. Monitoring of Natural Attenuation (MNA) of contaminated soil and sediments in a**

A severe gasoil spill (TPH-medium fraction) affected soils and rivers in a mountainous area southeast Mexico. In this tropical region most soils have silt-clayed texture characteristics, being the main human activities livestock and rain fed agriculture. Another distinctive feature

B-SS -0.0103 67.5 ± 5.0 NB-SS -0.0067 103.6 ± 5.2 B-NSS -0.0110 63.0 ± 1.0 NB-NSS -0.0088 79.1 ± 1.4

**Table 5.** Biodegradation constant and half-life time for Olmeca crude oil in soil, after a 90-day assay.

constants were obtained by Chang et al. (2010) in similar range (0.011 - 0.024 d-1).

biostimulated (non-inoculated) treatment after 120 days.

**riparian environment**

**Half life time (d)**

**Treatment**

170 Biodegradation of Hazardous and Special Products

c Half life time obtained from first order adjustment of the soil residual concentration data.

**Table 6.** MNA parameters of gasoil measured at three sampling points of the spill-impacted area.

Sediment samples and superficial soil (0-10 cm) were collected from sampling points 9 and 10 (P9 and P10). Soil samples from 30 - 45 cm depth were taken from sampling point 2 (P2). Residual hydrocarbons data were well described by the first order model having a high lineal correlation (r2 = 0.926 to 0.937). Additionally, in P2 hydrocarbons were detected in the stream‐ bed sediments (17 100 mg kg-1), with a later 84% reduction (2 662 mg kg-1) in 81 days. For some of the samples we found heterogeneous results, even finding increases from baseline (P9, times 0 to 36 d), this is explained by considering the inherent field variability on the presence of hydrocarbons and the stream dynamics in the site. However, the general behavior of the residual hydrocarbons concentration was towards an accelerated biodegradation, with reductions between 60 to 80% compared to the initial concentrations. According to the halflife calculated from gasoil biodegradation data, the residual concentrations will reach levels below the permissible limit established in current Mexican regulations (<1 200 mg kg-1) after 6 – 9 months.

isms showed high capacity for diesel biodegradation in liquid medium and in soil (120,000 mg kg-1 soil). Limitations in biodegradation extent probably were due to depletion of nutrients necessary for the growth or activity of the oil-degrading microorganism. Another possible cause was the low bioavailability of diesel (soil weathering), which affected the activity of both native and exogenous degrading microorganisms. Based on the conspicuous colony morphol‐ ogy of the introduced bacterial strains *B. subtilis* 7A and *G. rubripertincta* during biodegradation in soil assays, they prevailed at the end of the experiments, showing a good degrading capacity

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

The high diesel and Olmeca crude oil degrading activities registered in both liquid medium and soil assays are promising alternatives for the application of this type of co-culture as a part of active remediation strategies for contaminated soils based, together with biostimulation and

Finally, MNA schemes like the one here assayed, could be useful for contaminated sites with a good ecological conservation degree, where an active remediation technique may result in more damage and increased costs. Special attention should be put in the physicochemical effects of hydrocarbons on the soil properties (i e. field capacity, water repellency, salinization, lixiviate residual toxicity). These approaches may favour re-establishment of vegetation and

Financial support is acknowledged to the Mexican Ministry for Public Education (Project PROMEP No. 20110123) and Juárez Autonomous University of Tabasco (Project PFICA No. 2012010). Special thanks are given to Cornelio Contreras Juárez, Francisco Guzmán Osorio and

1 Academic Division of Biological Sciences, Juarez Autonomous University of Tabasco, Ta‐

2 Departament of Biological Systems, Metropolitan Autonomous University – Xochimilco,

, Randy Adams Schroeder 1

,

http://dx.doi.org/10.5772/56233

173

, Erika Escalante-Espinosa1

and Hugo Ramírez-Saad2\*

and ability to compete with native soil bacteria.

Jorge Ortiz Maya for experimental and field assistance.

\*Address all correspondence to: hurasa@correo.xoc.uam.mx

bioaugmentation approaches.

fertility of the soil matrix.

**Acknowledgements**

**Author details**

basco, Mexico

Mexico City, Mexico

Ildefonso Díaz-Ramírez1

Reyna Fócil-Monterrubio1

The calculated global consumption rates (GCR) ranged between 51 mg kg-1 soil d-1 for a low contaminated site to 232 mg kg-1 soil d-1, for the most contaminated (P2). While average maximum consumption rates for P10 and P2 scored 551 and 675 mg kg-1 d-1, respectively, these values were achieved between 36 and 87 days of monitoring. Regarding the presence of PAHs, in most of the studied sampling points the levels were below the quantification threshold for CG analysis, except for benzo [a] pyrene that reached 0.884 - 2.76 mg kg-1.

In general, the numbers of diesel degraders and heterotrophic bacteria and fungi were high in the sampled soils. Values for degraders ranged from 104 to 107 , corresponding to 0.1 to 30% of the total microbial population. This range of degraders is even larger than previous reports. Prince, (2003) found that the number of oil-degrading bacteria in a contaminated soil was around 0.1 to 10% of the total microbial population. Our results can be attributed to the favorable environmental conditions detected on the walls of the stream, with abundant vegetation, high soil moisture, rich in organic matter that has been previously deposited by water flow.

Regarding sediment samples, moderate numbers of oil-degrading microorganisms (103 to 106 ) were found, however, a high removal of hydrocarbons was scored. This may be explained by desorption of the gasoil present in sediments, due to washing effect of water flow. Subse‐ quently, biodegradation could take place in the liquid phase.

Accelerated biodegradation rates were observed for most of the monitored points resulting in significant reduction of gasoil concentration after two to three months (short half-life), even at the points of highest content. On view of these results, monitored natural attenuation can be considered as an option for site restoration. Although a site characterization strategy is needed, in order to determine environmental conditions that may influence hydrocarbons disappear‐ ance, in addition to the sole presence of an oil-degrading community. MNA strategies could be applied to other sites of comparable ecological value or similar tropical regions.

#### **4. Concluding remarks**

Diesel biodegradation in liquid media was higher in presence of the co-culture (bacterial and fungal standardized mixture) and the F-DMC. These previously characterized microorgan‐ isms showed high capacity for diesel biodegradation in liquid medium and in soil (120,000 mg kg-1 soil). Limitations in biodegradation extent probably were due to depletion of nutrients necessary for the growth or activity of the oil-degrading microorganism. Another possible cause was the low bioavailability of diesel (soil weathering), which affected the activity of both native and exogenous degrading microorganisms. Based on the conspicuous colony morphol‐ ogy of the introduced bacterial strains *B. subtilis* 7A and *G. rubripertincta* during biodegradation in soil assays, they prevailed at the end of the experiments, showing a good degrading capacity and ability to compete with native soil bacteria.

The high diesel and Olmeca crude oil degrading activities registered in both liquid medium and soil assays are promising alternatives for the application of this type of co-culture as a part of active remediation strategies for contaminated soils based, together with biostimulation and bioaugmentation approaches.

Finally, MNA schemes like the one here assayed, could be useful for contaminated sites with a good ecological conservation degree, where an active remediation technique may result in more damage and increased costs. Special attention should be put in the physicochemical effects of hydrocarbons on the soil properties (i e. field capacity, water repellency, salinization, lixiviate residual toxicity). These approaches may favour re-establishment of vegetation and fertility of the soil matrix.

#### **Acknowledgements**

correlation (r2 = 0.926 to 0.937). Additionally, in P2 hydrocarbons were detected in the stream‐ bed sediments (17 100 mg kg-1), with a later 84% reduction (2 662 mg kg-1) in 81 days. For some of the samples we found heterogeneous results, even finding increases from baseline (P9, times 0 to 36 d), this is explained by considering the inherent field variability on the presence of hydrocarbons and the stream dynamics in the site. However, the general behavior of the residual hydrocarbons concentration was towards an accelerated biodegradation, with reductions between 60 to 80% compared to the initial concentrations. According to the halflife calculated from gasoil biodegradation data, the residual concentrations will reach levels below the permissible limit established in current Mexican regulations (<1 200 mg kg-1) after

The calculated global consumption rates (GCR) ranged between 51 mg kg-1 soil d-1 for a low contaminated site to 232 mg kg-1 soil d-1, for the most contaminated (P2). While average maximum consumption rates for P10 and P2 scored 551 and 675 mg kg-1 d-1, respectively, these values were achieved between 36 and 87 days of monitoring. Regarding the presence of PAHs, in most of the studied sampling points the levels were below the quantification threshold for

In general, the numbers of diesel degraders and heterotrophic bacteria and fungi were high in

the total microbial population. This range of degraders is even larger than previous reports. Prince, (2003) found that the number of oil-degrading bacteria in a contaminated soil was around 0.1 to 10% of the total microbial population. Our results can be attributed to the favorable environmental conditions detected on the walls of the stream, with abundant vegetation, high soil moisture, rich in organic matter that has been previously deposited by

Regarding sediment samples, moderate numbers of oil-degrading microorganisms (103

be applied to other sites of comparable ecological value or similar tropical regions.

were found, however, a high removal of hydrocarbons was scored. This may be explained by desorption of the gasoil present in sediments, due to washing effect of water flow. Subse‐

Accelerated biodegradation rates were observed for most of the monitored points resulting in significant reduction of gasoil concentration after two to three months (short half-life), even at the points of highest content. On view of these results, monitored natural attenuation can be considered as an option for site restoration. Although a site characterization strategy is needed, in order to determine environmental conditions that may influence hydrocarbons disappear‐ ance, in addition to the sole presence of an oil-degrading community. MNA strategies could

Diesel biodegradation in liquid media was higher in presence of the co-culture (bacterial and fungal standardized mixture) and the F-DMC. These previously characterized microorgan‐

to 107

, corresponding to 0.1 to 30% of

 to 106 )

CG analysis, except for benzo [a] pyrene that reached 0.884 - 2.76 mg kg-1.

the sampled soils. Values for degraders ranged from 104

quently, biodegradation could take place in the liquid phase.

6 – 9 months.

172 Biodegradation of Hazardous and Special Products

water flow.

**4. Concluding remarks**

Financial support is acknowledged to the Mexican Ministry for Public Education (Project PROMEP No. 20110123) and Juárez Autonomous University of Tabasco (Project PFICA No. 2012010). Special thanks are given to Cornelio Contreras Juárez, Francisco Guzmán Osorio and Jorge Ortiz Maya for experimental and field assistance.

#### **Author details**

Ildefonso Díaz-Ramírez1 , Erika Escalante-Espinosa1 , Randy Adams Schroeder 1 , Reyna Fócil-Monterrubio1 and Hugo Ramírez-Saad2\*

\*Address all correspondence to: hurasa@correo.xoc.uam.mx

1 Academic Division of Biological Sciences, Juarez Autonomous University of Tabasco, Ta‐ basco, Mexico

2 Departament of Biological Systems, Metropolitan Autonomous University – Xochimilco, Mexico City, Mexico

#### **References**

[1] Adams RH, Castillo-Acosta O, Escalante-Espinosa E, Zavala-Cruz J. 2011. Natural at‐ tenuation and phytoremediation of petroleum hydrocarbon impacted soil in tropical wetland environments, In: Torres LG, Bandala ER. (Eds). Remediation of Soils and Aquifers. pp 1-24. Nova Publishers, New York.

[12] Díaz-Ramírez IJ, Gutiérrez-Rojas M, Ramírez-Saad H, Favela-Torres E. 2003. Biode‐ gradation of Maya crude oil fractions by bacterial strains and a defined mixed cul‐ ture isolated from *Cyperus laxus* rhizosphere soil in a contaminated site. Canadian

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

http://dx.doi.org/10.5772/56233

175

[13] Díaz-Ramírez IJ. 2000. Biodegradación de hidrocarburos por un consorcio microbia‐ no de la rizósfera de una planta nativa de pantano. Tesis de Maestría, Universidad

[14] EPA (USA) 8015, 1996. Non-halogenated organics using GC / FID. SW 846. Test methods for evaluating soil waste, physical/chemical methods. Available at: http://

[15] EPA (USA) 1999. Use of Monitored Natural Attenuation at Superfund, RCRA Correc‐ tive Action, and Underground Storage Tank Sites. OSWER Directive number:

[16] Escalante-Espinoza E. 2005. Estudio de la capacidad fitorremediadora de *Cyperus laxus* Lam. en suelo contaminado con hidrocarburos. Tesis de Doctorado, Universi‐

[17] Eweis JB, Ergas SJ, Chang, DP. Schroeder ED. 1998. Bioremediation Principles. pp. 88

[18] Greenwood PF, Wibrow S, George SJ, Tibbett M. 2009. Hydrocarbon biodegradation and soil microbial community response to repeated oil exposure. Organic Geochem‐

[19] Hye HJ, Kim JK, Choi O, Kyung-Suk C, Hee WR. 2005. Characterization of a dieseldegrading bacterium, *Pseudomonas aeruginosa* IU5, isolated from oil-contaminated

[20] Ko SH. Lebeault JM. 1999. Effect of a mixed culture on co-oxidation during the deg‐ radation of saturated hydrocarbon mixture. Journal of Applied Microbiology 87:

[21] Leahy JG, Colwell RR. 1990. Microbial degradation of hydrocarbons in the environ‐

[22] Liu GPW, Chang TC, Whang LM, Kao CH, Pan PT, Cheng SS. 2011. Bioremediation of petroleum hydrocarbon contaminated soil: Effects of strategies and microbial com‐ munity shift. International Biodeterioration and Biodegradation 65(8): 1119-1127.

[23] Liu S, Suflita JM. 1993. Ecology and evolution of microbial populations for bioreme‐

[24] Lorch HJ, Benckieser G. Ottow JCG. 1995. Basic methods for counting microorgan‐ isms in soil and water. In: Alef K, Nannipieri P (Eds). Methods in Applied Soil Mi‐

crobiology and Biochemistry, pp. 146-161, Academic Press, Great Britain.

soil in Korea. World Journal of Microbiology y Biotechnology 21:381-384.

9200.4-17P. Available at: http://www.epa.gov/oust/directiv/d9200417.pdf

Journal of Microbiology 49: 755-761.

Autónoma Metropolitana – Iztapalapa, México.

www.caslab.com/EPA-Methods/PDF/8015b.pdf

dad Autónoma Metropolitana – Iztapalapa, México.


ment. Microbiological Reviews 54: 305-315.

diation. Trends in Biotechnology 11: 344-352.

istry. 40(3): 293-300.

72-79.


[12] Díaz-Ramírez IJ, Gutiérrez-Rojas M, Ramírez-Saad H, Favela-Torres E. 2003. Biode‐ gradation of Maya crude oil fractions by bacterial strains and a defined mixed cul‐ ture isolated from *Cyperus laxus* rhizosphere soil in a contaminated site. Canadian Journal of Microbiology 49: 755-761.

**References**

17(2): 159-174.

174 Biodegradation of Hazardous and Special Products

terciencia. 33: 483 -489.

[1] Adams RH, Castillo-Acosta O, Escalante-Espinosa E, Zavala-Cruz J. 2011. Natural at‐ tenuation and phytoremediation of petroleum hydrocarbon impacted soil in tropical wetland environments, In: Torres LG, Bandala ER. (Eds). Remediation of Soils and

[2] Adams RH, Domínguez-Rodríguez VI, García Hernández L. 1999. Potencial de la bi‐ orremediación de suelo y agua impactado por petróleo en el trópico Mexicano. Terra

[3] Adams RH, Guzmán-Osorio FJ. 2008. Evaluation of land farming and chemical-bio‐ logical stabilization for treatment of heavily contaminated sediments in a tropical en‐ vironment. International Journal Environmental Science Technology 5(2): 169-178.

[4] Adams RH, Zavala Cruz J, Morales García FA. 2008. Concentración residual de hi‐ drocarburos en suelo del trópico. II: Afectación a la fertilidad y su recuperación. In‐

[5] Amann R, Kühl M. 1998. *In situ* methods for assessment of microorganism and their

[6] Boettcher G, Nyer EK. 2001. *In situ* Bioremediation. In: Nyer EK, Palmer PL, Carman EP, Boettcher G, Bedessman JM, Lenzo F, Crossman T L, Rorech GJ, Kidd DF (Eds). *In situ* Treatment Technology. pp. 261-281, Arcadis, Geraghty & Miller. FL. USA.

[7] Boonchan S, Britz ML, Stanley GA. 2000. Degradation and mineralization of highweight polycyclic aromatic hydrocarbons by defined fungal-bacterial cocultures. Ap‐

[8] Cardona S, Iturbe R. 2003. Degradación de diésel Mexicano por un consorcio de bac‐ terias de un suelo agrícola. Revista Dyna 70(138):13-26. Available at: http://reda‐

[9] Chang W, Dyen M, Spagnuolo L, Simon P, Whyte L, Ghoshal S. 2010. Biodegradation of semi- and non-volatile petroleum hydrocarbons in aged, contaminated soils from a sub-Arctic site: Laboratory pilot-scale experiments at site temperatures. Chemo‐

[10] Chemlal R, Tassist A, Drouiche M, Lounici H, Drouiche N, Mameri N. 2012. Micro‐ biological aspects study of bioremediation of diesel-contaminated soils by biopile

[11] Díaz-Ramírez IJ, Escalante-Espinosa E, Favela-Torres E, Gutiérrez-Rojas M, Ramírez-Saad H. 2008. Design of bacterial defined mixed cultures for biodegradation of spe‐ cific crude oil fractions, using population dynamics analysis by DGGE. International

technique. International Biodeterioration and Biodegradation 75: 201-206.

Aquifers. pp 1-24. Nova Publishers, New York.

activities. Current Opinion in Microbiology 1: 352-358.

plied and Environmental Microbiology 66(3): 1007-1019.

lyc.uaemex.mx/pdf/496/49613802.pdf

Biodeterioration and Biodegradation 62: 21-30.

sphere 80: 319–326.


[25] Madsen EL. 1998. Epistemology of environmental microbiology. Environmental Sci‐ ence Technology.32(4): 429-439.

[37] Petróleos Mexicanos (PEMEX). 2008. Subdirección de auditoría en seguridad indus‐ trial y protección ambiental. Hoja de datos de seguridad de sustancias (Diésel UBA).

Hydrocarbon Biodegradation Potential of Native and Exogenous Microbial Inocula in Mexican Tropical Soils

http://dx.doi.org/10.5772/56233

177

[38] Prince RC. 2003. Biodegradation of petroleum and other hydrocarbons. In: Encyclo‐ pedia of Environmental Microbiology. John Wiley & Sons, Inc. New Jersey. Available

[39] Richard JY, Vogel TM. 1999. Characterization of a soil bacterial consortium capable of degrading diesel fuel. International Biodeterioration and Biodegradation 44: 93-00.

[40] Roldán-Carrillo T, Castorena-Cortés G, Zapata-Peñasco I, Reyes-Avila J, Olguín-Lora P. 2012. Aerobic biodegradation of sludge with high hydrocarbon content generated by a Mexican natural gas processing facility. Journal of Environmental Management

[41] Sabaté J, Viñas M. Solanas AM. 2004. Laboratory-scale bioremediation experiments on hydrocarbon-contaminated soils. International Biodeterioration and Biodegrada‐

[42] Sayara T, Borràs E, Caminal G, Sarrà M, Sánchez A, 2011. Bioremediation of PAHscontaminated soil through composting: Influence of bioaugmentation and biostimu‐ lation on contaminant biodegradation. International Biodeterioration and

[43] Thouand G, Bauda P, Oudot J, Kirsch G, Sutton C, Vidalie JF. 1999. Laboratory evalu‐ ation of crude oil biodegradation with commercial or natural microbial inocula.

[44] Torsvik V, Ovreas L. 2002. Microbial diversity and function in soil: from genes to eco‐

[45] Vallejo V, Salgado L, Roldán F. 2005. Evaluación de la estimulación en la biodegrada‐ ción de HTPs en suelos contaminados con petróleo. Revista Colombiana de Biotecno‐

[46] Van der Meer JR, De Vos WM, Harayama S, Zehnder AJB. 1992. Molecular mecha‐ nisms of genetic adaptation to xenobiotic compounds. Microbiological Reviews 56

[47] Van Hamme JD, Odumeru JA, Ward OP. 2000. Community dynamics of a mixedbacterial culture growing on petroleum hydrocarbons in batch culture. Canadian

[48] Van Hamme JD, Singh A, Ward OP. 2003. Recent advances in petroleum microbiolo‐

[49] Viñas M, Grifoll M, Sabaté J, Solanas AM. 2002. Biodegradation of a crude oil by three microbial consortia of different origins and metabolic capabilities. Journal of In‐

gy. Microbiology and Molecular Biology Reviews 67(4): 503-549.

dustrial Microbiology and Biotechnology 28: 252–260.

at (http://www.mrw.interscience.wiley.com/eem/articles/env118).

95: S93-S98.

tion 54: 19-25.

logía 3(2): 67-68.

(4): 677-694.

Biodegradation 65: 859-865.

Canadian Journal of Microbiology 45: 106-115.

Journal of Microbiology 46: 441-450.

systems. Current Opinion in Microbiology 5: 240-245.


[37] Petróleos Mexicanos (PEMEX). 2008. Subdirección de auditoría en seguridad indus‐ trial y protección ambiental. Hoja de datos de seguridad de sustancias (Diésel UBA).

[25] Madsen EL. 1998. Epistemology of environmental microbiology. Environmental Sci‐

[26] Madueño L, Coppotelli BM, Alvarez HM, Morelli IS. 2011. Isolation and characteriza‐ tion of indigenous soil bacteria for bioaugmentation of PAH contaminated soil of semiarid Patagonia, Argentina. International Biodeterioration and Biodegradation

[27] Margesin R, Hämmerle M, Tscherko D. 2007. Microbial activity and community com‐ position during bioremediation of diesel-oil-contaminated soil: effects of hydrocar‐ bon concentration, fertilizers, and incubation time. Microbial Ecology 53: 259–269.

[28] Margesin R. 2005. Determination of Enzyme Activities in Contaminated Soil. In: Mar‐ gesin R, Schinner F (Eds). Manual for Soil Analysis – Monitoring and Assessing Soil

[29] Marquez-Rocha FJ, Hernández RV, Lamela MT. 2001. Biodegradation of diesel oil in soil by a microbial consortium. Water, Air and Soil Pollution 128: 313–320.

[30] Medina-Moreno SA, Huerta-Ochoa S, Gutiérrez-Rojas M. 2005. Hydrocarbon biode‐ gradation in oxygen limited sequential batch reactors by consortium from weathered

[31] Mishra S, Jyot J, Kuhad RC, Lal B. 2001. Evaluation of inoculum addition to stimulate *in situ* bioremediation of oily-sludge-contaminated soil. Applied and Environmental

[32] Mohammed D, Ramsubhag A, Beckles DM. 2007. An assessment of the biodegrada‐ tion of petroleum hydrocarbons in contaminated soil using non-indigenous, com‐

[33] NMX-AA 112-SECOFI, 1995. Norma Oficial Mexicana. Water and sediment analysis – Acute Toxicity Evaluation with *Photobacterium phsophoreum* – Test method. Availa‐ ble at: http://www.imta.gob.mx/cotennser/images/docs/NOM/NMX-

[34] NOM-021-SEMARNAT-2000. Norma Oficial Mexicana, que establece las especifica‐ ciones de fertilidad, salinidad, y clasificación de suelos, estudio, muestreo y análisis.

[35] NOM-053-ECOL-1993. Norma Oficial Mexicana. Procedimiento para llevar a cabo la prueba de extracción para determinar los constituyentes que hacen a un residuo peli‐ groso por su toxicidad al ambiente. Available at: http://biblioteca.semarnat.gob.mx/

[36] NOM-138-SEMARNAT/SS-2003. Norma Oficial Mexicana. Límites máximos permisi‐ bles de hidrocarburos en suelos y las especificaciones para su caracterización y reme‐

México, D., F., Diario Oficial de la Federación. Martes 23 de Abril del 2003.

Bioremediation. Soil Biology, Vol. 5. pp. 309-319. Springer, Germany.

oil-contaminated soil. Canadian Journal of Microbiology 51: 231-239.

mercial microbes. Water Air and Soil Pollution 182: 349–356.

janium/Documentos/Ciga/agenda/PPD02/DO2283m.pdf

diación. Diario Oficial de la Federación, México, D.F. 29-Mar-2005.

ence Technology.32(4): 429-439.

176 Biodegradation of Hazardous and Special Products

Microbiology 67: 1675–1681.

AA-112-1995.pdf

65: 345-351.


[50] Watanabe K, Hamamura N. 2003. Molecular and physiological approaches to under‐ standing the ecology of pollutant degradation. Current Opinion in Microbiology 14:

[51] Wrenn BA, Venosa AD. 1996. Selective enumeration of aromatic and aliphatic hydro‐ carbon degrading bacteria by a most-probable-number procedure. Canadian Journal

[52] Zhang D, Mörtelmaier C, Margesin R. 2012. Characterization of the bacterial archaeal diversity in hydrocarbon-contaminated soil. Science of the Total Environment 421:

289-295.

178 Biodegradation of Hazardous and Special Products

184–196.

of Microbiology 42: 252-258.

### *Edited by Rolando Chamy and Francisca Rosenkranz*

This book contains a collection of different research activities that include the biodegradation compounds with contaminant characteristics and special products of different interests as an added value product or that allows following up various biological processes. The chapters consider the degradation of contaminant compounds generated by industrial activities, i.e., oil industry by-product compounds and halogen compounds or compound generated by natural phenomena such as tsunamis, which require interventions to recover damaged soils. In addition, the book contains chapters that involve special product degradation processes such as chlorophyll, which corresponds to a biological process indicator as photosynthesis.

Photo by panaramka / iStock

Biodegradation of Hazardous and Special Products

Biodegradation of Hazardous

and Special Products

*Edited by Rolando Chamy and Francisca Rosenkranz*