**Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant Species of** *Aspergillus awamori*

Bruno Alexandre Quistorp Santos, Seteno Karabo Obed Ntwampe and James Hamuel Doughari

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/53349

**1. Introduction**

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[74] van den Eede, N., Dirtu, A. C., Neels, H., & Covaci, A. (2011). Analytical develop‐ ments and preliminary assessment of human exposure to organophosphate flame re‐

[75] van den Wijngaard, A. J., Janssen, D. B., & Witholt, B. (1989). Degradation of epi‐ chlorohydrin and halohydrins by bacterial cultures isolated from freshwater sedi‐

[76] van den Wijngaard, A. J., Reuvekamp, P. T., & Janssen, D. B. (1991). Purification and characterization of haloalcohol dehalogenase from *Arthrobacter* sp. strain AD2. *J. Bac‐*

[77] van der Veen, I. & de Boer, J. (2012). Phosphorus flame retardants: Properties, pro‐ duction, environmental occurrence, toxicity and analysis. *Chemosphere,* 88(10),

[78] Watts, M. J. & Linden, K. G. (2008). Photooxidation and subsequent biodegradability of recalcitrant tri-alkyl phosphates TCEP and TBP in water. *Water Res.,* 42(20),

[79] Westerhoff, P., Yoon, Y., Snyder, S., & Wert, E. (2005). Fate of endocrine-disruptor, pharmaceutical, and personal care product chemicals during simulated drinking wa‐

[80] Worls Health Organization (WHO) (1997). Environmental Health Criteria 192: Flame Retardants: A General Introduction: World Health Organization, Geneva.

[81] Yonetani, R., Ikatsu, H., Miyake-Nakayama., C., Fujiwara, E., Maehara, Y., Miyoshi., S., Matsuoka, H., & Shinoda, S. (2004). Isolation and characterization of a 1,3-di‐

ter treatment processes. *Environ. Sci. Technol.,* 39(17), 6649-6663.

chloro-2-propanol-degrading bacterium. *J. Health Sci.,* 50(6), 605-612.

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*teriol.,* 173(1), 124-129.

1119-1153.

4949-4954.

Various industries release a combination of free cyanide and cyanide complexes into the en‐ vironment via a variety of disposal methods, particularly as wastewater. These industries utilise cyanide based compounds in various operations, including: the beneficiation of met‐ als, electroplating, case hardening, automotive manufacturing, circuitry board manufactur‐ ing, and in chemical industries [23]. Cyanide is often found in organic, hydrocarbon chains or as inorganic, transition, alkali and alkali earth metal complexes [20]. Many cyanide com‐ plexes are highly unstable, thus temperature, pH and light can degrade the components to form free cyanide which is the most toxic form of cyanide [20, 26].

There is an overwhelming popularity in industry for the use of chemical treatment methods for the treatment of free cyanide and cyanide complexes compared to biochemical treatment methods. Chemical remediation methods like alkaline chlorine oxidation are commonly used to treat cyanide contaminated wastewater [23, 24]. Chemical oxidation is particularly ineffective in the treatment of cyanide-metal complexes containing heavy metals, such as copper, nickel and silver, due to the slow reaction rate [23]. The excess quantity of chlorine used in the treatment process increases the chemical oxygen demand (COD) of the wastewa‐ ter thereby rendering the water undesirable for reuse, toxic to aquatic life and may produce organic substances. In order to reduce operational costs, some manufacturers partially treat the wastewater, resulting in untreated and/or partially decomposed cyanide being dis‐ charged. Other methods of treatment include copper catalysed hydrogen peroxide oxida‐

© 2013 Santos 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 Santos et al.; licensee InTech. This is a paper 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.

tion, ozonation and electrolytic decomposition [23]. However, these methods are unpopular due to the high capital costs, specialist equipment and maintenance requirements.

advised to keep cyanide solutions at a high pH to prevent the evolvement of hydrocyanic gas since there is a direct relationship between the dissociation of hydrocyanic molecule and

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125

**3. Differentiating filamentous fungi:** *Aspergillus niger* **and** *Aspergillus*

Black *Aspergilli* (*Aspergillus* section *Nigri*) species, are aerobic filamentous fungi often de‐ rived from soil. They have shown potential in biotechnology, food and medical applications. The trait of these species causing agricultural products to spoil had recently shown to be of benefit. Agricultural waste can be fermented to produce a variety of industrially important extracellular enzymes, such as cellulase, amylase, xylanase, pectinase, elastase, and organic acids, such as citric, galacturonic including gluconic acid. *Aspergillus niger* (*A. niger*) and *As‐ pergillus awamori* (*A. awamori*) are closely related species and *A. awamori* is often misidenti‐ fied as *A. niger*. They share similar morphology and growth rates at various temperatures

A black filamentous mould was isolated from cyanide contaminated municipal wastewater discharge drain located in the Western Cape, South Africa. Swab samples were taken at var‐ ious points along the drain and grown on 1 % (w/v) Citrus Pectin Agar (CPA) plates incu‐ bated at 37 °C for 5 days. After incubation, black mycelia of filamentous mould grew on the

pH (Figure 1) including temperature [20].

**Figure 1.** Relationship between HCN in solution and pH [20]

and produce several common enzymes [34].

**3.1. Isolation and identification:** *Aspergillus awamori*

*awamori*

plate (Figure 2).

Several microorganisms, bacteria such as *Nocardia sp*. and *Rhodococcus sp*., fungi such as *As‐ pergillus sp.* and *Fusarium sp.* and algae such as *Arthrospira maxima* and *Scenedesmus obliquus*, possess enzymatic mechanisms able to bioremediate free cyanide and cyanide complexes [1, 26]. However, limited studies have been conducted using organic waste and fungal strains in cyanide bioremediation. Several studies have been conducted using varying concentra‐ tions of free cyanide, with moderate success being achieved in some cases [1].

Since the early 1970's, progress has been made to develop economically viable continuous remediation processes such as membrane bioreactors (MBRs) [8]. A membrane is generally defined as being a selective barrier. The membrane utilised in a bioreactor can provide ei‐ ther a barrier to limit the transport of certain components, while being permeable to others, thus prevent certain components from contacting a biocatalyst, or contain reactive sites thus being a catalyst itself [5]. The application of MBRs for the production of enzymes has re‐ ceived considerable attention for their diverse industrial use. A number of microorganisms have been studied in MBR applications for wastewater using fungi, such as white-rot fun‐ gus, *Phanerochaete chrysosporium* (*P. chrysosporium*) [8].

Solid waste generation in South Africa is a problem growing at an exponential rate with the majority of landfill sites reaching maximum capacity. Approximately 427x106 tonnes of solid waste is generated in South Africa every year, of which 40% by mass is organic waste [10]. The average amount of waste generated per person in South Africa is 0.7 kg/annum, which is close to that of developed countries such as the United Kingdom (0.723 kg/annum) and Singapore (0.87 kg/annum), than for developing countries, such as Nepal (0.3 kg/annum) [10]. It is sensible to bioaugment biotechnological processes to utilise organic waste materi‐ als, particularly for industries which produce large quantities of it.

### **2. Overview: Free cyanide**

Free cyanide is the simplest form of cyanide and has two forms, namely as a hydrocyanic molecule or hydrogen cyanide (HCN) which dissociates into an anionic cyanide molecule (CN- ) in solution [20]. By definition, free cyanides are forms of molecular and ionic cyanides that are released in aqueous solution by the dissolution and dissociation of cyanide com‐ plexes. Simple and weak acid dissociable cyanides are the most unstable and most likely to form free cyanide in aqueous solution. Simple cyanide compounds are ionically bonded cya‐ nide anions and alkali earth or alkali metals that are neutral, that exist in solid form and dis‐ sociate into alkali earth or alkali metals and free cyanide when in aqueous solutions [20].

Accordingly the environmental risk of cyanide wastewater is not limited to the effluent but also the possibility of emitting hydrocyanic gas. Hydrocyanic gas is toxic, colourless, distinc‐ tive almond smell at low concentrations, slightly soluble in water and readily dissociates in‐ to hydrogen and anionic cyanide at low pH in aqueous solution [20]. For safety reasons, it is advised to keep cyanide solutions at a high pH to prevent the evolvement of hydrocyanic gas since there is a direct relationship between the dissociation of hydrocyanic molecule and pH (Figure 1) including temperature [20].

**Figure 1.** Relationship between HCN in solution and pH [20]

tion, ozonation and electrolytic decomposition [23]. However, these methods are unpopular

Several microorganisms, bacteria such as *Nocardia sp*. and *Rhodococcus sp*., fungi such as *As‐ pergillus sp.* and *Fusarium sp.* and algae such as *Arthrospira maxima* and *Scenedesmus obliquus*, possess enzymatic mechanisms able to bioremediate free cyanide and cyanide complexes [1, 26]. However, limited studies have been conducted using organic waste and fungal strains in cyanide bioremediation. Several studies have been conducted using varying concentra‐

Since the early 1970's, progress has been made to develop economically viable continuous remediation processes such as membrane bioreactors (MBRs) [8]. A membrane is generally defined as being a selective barrier. The membrane utilised in a bioreactor can provide ei‐ ther a barrier to limit the transport of certain components, while being permeable to others, thus prevent certain components from contacting a biocatalyst, or contain reactive sites thus being a catalyst itself [5]. The application of MBRs for the production of enzymes has re‐ ceived considerable attention for their diverse industrial use. A number of microorganisms have been studied in MBR applications for wastewater using fungi, such as white-rot fun‐

Solid waste generation in South Africa is a problem growing at an exponential rate with the

waste is generated in South Africa every year, of which 40% by mass is organic waste [10]. The average amount of waste generated per person in South Africa is 0.7 kg/annum, which is close to that of developed countries such as the United Kingdom (0.723 kg/annum) and Singapore (0.87 kg/annum), than for developing countries, such as Nepal (0.3 kg/annum) [10]. It is sensible to bioaugment biotechnological processes to utilise organic waste materi‐

Free cyanide is the simplest form of cyanide and has two forms, namely as a hydrocyanic molecule or hydrogen cyanide (HCN) which dissociates into an anionic cyanide molecule

Accordingly the environmental risk of cyanide wastewater is not limited to the effluent but also the possibility of emitting hydrocyanic gas. Hydrocyanic gas is toxic, colourless, distinc‐ tive almond smell at low concentrations, slightly soluble in water and readily dissociates in‐ to hydrogen and anionic cyanide at low pH in aqueous solution [20]. For safety reasons, it is

) in solution [20]. By definition, free cyanides are forms of molecular and ionic cyanides that are released in aqueous solution by the dissolution and dissociation of cyanide com‐ plexes. Simple and weak acid dissociable cyanides are the most unstable and most likely to form free cyanide in aqueous solution. Simple cyanide compounds are ionically bonded cya‐ nide anions and alkali earth or alkali metals that are neutral, that exist in solid form and dis‐ sociate into alkali earth or alkali metals and free cyanide when in aqueous solutions [20].

tonnes of solid

due to the high capital costs, specialist equipment and maintenance requirements.

tions of free cyanide, with moderate success being achieved in some cases [1].

majority of landfill sites reaching maximum capacity. Approximately 427x106

als, particularly for industries which produce large quantities of it.

gus, *Phanerochaete chrysosporium* (*P. chrysosporium*) [8].

124 Environmental Biotechnology - New Approaches and Prospective Applications

**2. Overview: Free cyanide**

(CN-

### **3. Differentiating filamentous fungi:** *Aspergillus niger* **and** *Aspergillus awamori*

Black *Aspergilli* (*Aspergillus* section *Nigri*) species, are aerobic filamentous fungi often de‐ rived from soil. They have shown potential in biotechnology, food and medical applications. The trait of these species causing agricultural products to spoil had recently shown to be of benefit. Agricultural waste can be fermented to produce a variety of industrially important extracellular enzymes, such as cellulase, amylase, xylanase, pectinase, elastase, and organic acids, such as citric, galacturonic including gluconic acid. *Aspergillus niger* (*A. niger*) and *As‐ pergillus awamori* (*A. awamori*) are closely related species and *A. awamori* is often misidenti‐ fied as *A. niger*. They share similar morphology and growth rates at various temperatures and produce several common enzymes [34].

### **3.1. Isolation and identification:** *Aspergillus awamori*

A black filamentous mould was isolated from cyanide contaminated municipal wastewater discharge drain located in the Western Cape, South Africa. Swab samples were taken at var‐ ious points along the drain and grown on 1 % (w/v) Citrus Pectin Agar (CPA) plates incu‐ bated at 37 °C for 5 days. After incubation, black mycelia of filamentous mould grew on the plate (Figure 2).

This was transferred to Potato Dextrose Agar (PDA) plates with 0.2% (v/v) Penicillin-Strep‐ tomycin (PEN-STREP; (10000 units/L of Penicillin and 10 mg of Streptomycin/ml) anti-biotic solution. The plates were again incubated at 37 ˚C for 5 days. One millilitre (1 ml) of 0.1% (w/v) Tween 80 solution was added to each plate and a spatula was used to harvest the spores and mycelium from the plate to form a spore-mycelium suspension. The suspension was then filtered through a glass wool using 20 ml syringes to entrap the mycelium onto the glass wool and produce a spore suspension which was stored at 4 ˚C. Afterwards, serial di‐ lutions were made from the spore suspension and the number of spores in each 1 ml, were determined in duplicate using a Marienfeld Neubauer cell-counter and a Nikon Eclipse E2000 at a phase contrast of one and magnification of 100x. The absorbance of the diluted spore suspension was determined at 750 nm using a Jenway 6715 UV/Visible spectropho‐ tometer with distilled water as a blank [31]. A calibration graph for the spore concentration was determined by plotting the absorbance against the spore concentration (spores/ml), to quantify spore concentration in the inoculum.

To observe the morphological characteristics of the fungus, the isolates were inoculated on Malt Extract Agar (MEA) and Czapek Yeast Agar (CYA) and incubated at 26 °C for 7 days. Based on their growth rate, the fungus was presumptively identified as *A. awamori. A. awa‐ mori* was reported to show rapid growth on the CYA compared to *A. niger* which exhibited restricted growth. However, the growth and sporulation on MEA was better than CYA in the case of both *A. niger* and *A. awamori*. The fruiting bodies were mounted in lactic acid be‐ fore they were observed under an oil immersion. The conidial heads for *A. awamori* were not well-defined columns in comparison to conidia heads observed for *A. niger.* The strain showed colony characteristics of both *A. niger* and *A. awamori.* On a general note, the coni‐ diophores and conidia of *A. awamori* and *A. niger* are similar and morphologically indistin‐

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guishable, as shown in Figure 3 [34].

**Figure 3.** Growth of isolate on (a & b) MEA and (c & d) CYA

**Figure 2.** Growth of *A. awamori* on CPA after incubation at 37 ˚C for 5 days

To observe the morphological characteristics of the fungus, the isolates were inoculated on Malt Extract Agar (MEA) and Czapek Yeast Agar (CYA) and incubated at 26 °C for 7 days. Based on their growth rate, the fungus was presumptively identified as *A. awamori. A. awa‐ mori* was reported to show rapid growth on the CYA compared to *A. niger* which exhibited restricted growth. However, the growth and sporulation on MEA was better than CYA in the case of both *A. niger* and *A. awamori*. The fruiting bodies were mounted in lactic acid be‐ fore they were observed under an oil immersion. The conidial heads for *A. awamori* were not well-defined columns in comparison to conidia heads observed for *A. niger.* The strain showed colony characteristics of both *A. niger* and *A. awamori.* On a general note, the coni‐ diophores and conidia of *A. awamori* and *A. niger* are similar and morphologically indistin‐ guishable, as shown in Figure 3 [34].

**Figure 3.** Growth of isolate on (a & b) MEA and (c & d) CYA

This was transferred to Potato Dextrose Agar (PDA) plates with 0.2% (v/v) Penicillin-Strep‐ tomycin (PEN-STREP; (10000 units/L of Penicillin and 10 mg of Streptomycin/ml) anti-biotic solution. The plates were again incubated at 37 ˚C for 5 days. One millilitre (1 ml) of 0.1% (w/v) Tween 80 solution was added to each plate and a spatula was used to harvest the spores and mycelium from the plate to form a spore-mycelium suspension. The suspension was then filtered through a glass wool using 20 ml syringes to entrap the mycelium onto the glass wool and produce a spore suspension which was stored at 4 ˚C. Afterwards, serial di‐ lutions were made from the spore suspension and the number of spores in each 1 ml, were determined in duplicate using a Marienfeld Neubauer cell-counter and a Nikon Eclipse E2000 at a phase contrast of one and magnification of 100x. The absorbance of the diluted spore suspension was determined at 750 nm using a Jenway 6715 UV/Visible spectropho‐ tometer with distilled water as a blank [31]. A calibration graph for the spore concentration was determined by plotting the absorbance against the spore concentration (spores/ml), to

quantify spore concentration in the inoculum.

126 Environmental Biotechnology - New Approaches and Prospective Applications

**Figure 2.** Growth of *A. awamori* on CPA after incubation at 37 ˚C for 5 days

**Figure 4.** (a & b) Typical *Aspergillus* conidiophores with a radial head and (c & d) roughened, round conidia with regu‐ lar low ridges and bars

Molecular characterization was carried out in order to confirm the identity of the fungal iso‐ lates. DNA was extracted from the pure isolates using the ZR Fungal/Bacterial DNA Kit (Zy‐ mo Research, California, USA). The subsequent Polymerase Chain Reaction (PCR) of the ITS1–5.8S–ITS2 rDNA region was prepared with primers ITS1 and ITS4 [36]. The β-tubulin gene was amplified using primers Bt2a and Bt2b [7] and the calmodulin gene with CL1 and CL2A [22], respectively. Sequencing reactions of the PCR products were set-up using a Big Dye terminator cycle sequencing premix kit (Applied Biosystems, CA).

Sequence reactions were analysed with an ABI PRISM 310 genetic analyser. Sequences were compared to those of a recent study by [34]. Datasets were aligned in Se-Al, including a se‐ quence analysis in Se-Al. This was followed by a sequence analysis in PAUP\* v4.0b10, using the BioNJ option for calculating a single tree for each dataset. Confidence in nodes was cal‐ culated using a bootstrap analysis of 1000 replicates. Only bootstrap values above 90% were indicated on the branches and *Aspergillus flavus* was chosen as the outgroup [34]. The isolat‐ ed fungus was denoted as *Aspergillus* (CPUT) in Figure 5 and 6.

**Figure 5.** NJ tree based on the analysis of the (a) ITS, (b) β-tublin and (c) calmodulin gene regions

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Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant... http://dx.doi.org/10.5772/53349 129

**Figure 5.** NJ tree based on the analysis of the (a) ITS, (b) β-tublin and (c) calmodulin gene regions

**Figure 4.** (a & b) Typical *Aspergillus* conidiophores with a radial head and (c & d) roughened, round conidia with regu‐

Molecular characterization was carried out in order to confirm the identity of the fungal iso‐ lates. DNA was extracted from the pure isolates using the ZR Fungal/Bacterial DNA Kit (Zy‐ mo Research, California, USA). The subsequent Polymerase Chain Reaction (PCR) of the ITS1–5.8S–ITS2 rDNA region was prepared with primers ITS1 and ITS4 [36]. The β-tubulin gene was amplified using primers Bt2a and Bt2b [7] and the calmodulin gene with CL1 and CL2A [22], respectively. Sequencing reactions of the PCR products were set-up using a Big

Sequence reactions were analysed with an ABI PRISM 310 genetic analyser. Sequences were compared to those of a recent study by [34]. Datasets were aligned in Se-Al, including a se‐ quence analysis in Se-Al. This was followed by a sequence analysis in PAUP\* v4.0b10, using the BioNJ option for calculating a single tree for each dataset. Confidence in nodes was cal‐ culated using a bootstrap analysis of 1000 replicates. Only bootstrap values above 90% were indicated on the branches and *Aspergillus flavus* was chosen as the outgroup [34]. The isolat‐

Dye terminator cycle sequencing premix kit (Applied Biosystems, CA).

128 Environmental Biotechnology - New Approaches and Prospective Applications

ed fungus was denoted as *Aspergillus* (CPUT) in Figure 5 and 6.

lar low ridges and bars

According to [34], the only way to separate *A. niger* and *A. awamori* is through a multi-gene phylogenetic analysis. This was done by using ITS, β-tubulin and calmodulin gene regions, as shown in Figure 5. The combined gene region analysis (Figure 6) indicates that the *Asper‐ gillus* strain was similar and indeed identical to the sequence of the type strain of *A. awamori,* a fungus with diverse properties in biotechnology applications, including the production of

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Citrus peels are composed of cellulose, pectin, hemi-cellulose, lignin, chlorophyll pigments, low relative-molecular-mass hydrocarbons including lipids, proteins, simple sugars, starch‐ es, water and ash [14, 29]. The major components in citrus peel are cellulose, hemi-cellulose, pectin and lignin which are inter-wound with each other to provide a rigid cell wall struc‐ ture. *Aspergillus awamori* is able to produce enzymes for the breakdown of cellulose, hemi-

Cellulase, xylanase, pectinase are important enzymes in the hydrolysis of cellulose, hemicellulose and pectin into simpler sugars which can be utilised as a carbon source by the fun‐ gus. Hydrolysis of the orange peel has shown to yield significant quantities of neutral sugars: glucose, fructose and sucrose with low yields of xylose, arabinose, galactose and mannose. Hydrolysis of citrus peel also yield uronic acids, with galacturonic acid being the major of uronic acid liberated with trace quantities of other uronic acids. The optimum tem‐ perature and pH for these enzymes are in the range of 45 to 50 ˚C and 4.0 to 5.5, respectively [18, 19, 32]. Furthermore, *A. awamori* can produce nitrilase which hydrolyses the nitrile (cya‐ nide) group (R-CΞN) into the corresponding carboxylic acid and ammonia, as shown in Fig‐

The majority of the citrus peel components are high carbon source materials and the ammo‐ nia produced from the cyanide degradation can be utilised as a nitrogen source by the mi‐ croorganism. Studies have shown that the optimal pH and temperature for nitrilase

The successful treatment of 400 ppm free cyanide supplemented with refined/readily me‐ tabolisable carbon sources has been reported [1]. However, current technology used can

**3.2. Citrus peel supplemented growth medium for cyanide bioremediation using**

cellulose and pectin and leaving lignin as the remaining structural component.

ure 7 [26]. In aqueous solution, ammonia/ammonium equilibrium is observed.

nitrilase-a cyanide degrading enzyme.

**Figure 7.** Nitrilase hydrolysis on cyanide group [26]

production is 8 and 45 ˚C, respectively [12, 35].

*Aspergillus awamori*

**Figure 6.** NJ tree based on the analysis of the combined gene regions

According to [34], the only way to separate *A. niger* and *A. awamori* is through a multi-gene phylogenetic analysis. This was done by using ITS, β-tubulin and calmodulin gene regions, as shown in Figure 5. The combined gene region analysis (Figure 6) indicates that the *Asper‐ gillus* strain was similar and indeed identical to the sequence of the type strain of *A. awamori,* a fungus with diverse properties in biotechnology applications, including the production of nitrilase-a cyanide degrading enzyme.

### **3.2. Citrus peel supplemented growth medium for cyanide bioremediation using** *Aspergillus awamori*

Citrus peels are composed of cellulose, pectin, hemi-cellulose, lignin, chlorophyll pigments, low relative-molecular-mass hydrocarbons including lipids, proteins, simple sugars, starch‐ es, water and ash [14, 29]. The major components in citrus peel are cellulose, hemi-cellulose, pectin and lignin which are inter-wound with each other to provide a rigid cell wall struc‐ ture. *Aspergillus awamori* is able to produce enzymes for the breakdown of cellulose, hemicellulose and pectin and leaving lignin as the remaining structural component.

Cellulase, xylanase, pectinase are important enzymes in the hydrolysis of cellulose, hemicellulose and pectin into simpler sugars which can be utilised as a carbon source by the fun‐ gus. Hydrolysis of the orange peel has shown to yield significant quantities of neutral sugars: glucose, fructose and sucrose with low yields of xylose, arabinose, galactose and mannose. Hydrolysis of citrus peel also yield uronic acids, with galacturonic acid being the major of uronic acid liberated with trace quantities of other uronic acids. The optimum tem‐ perature and pH for these enzymes are in the range of 45 to 50 ˚C and 4.0 to 5.5, respectively [18, 19, 32]. Furthermore, *A. awamori* can produce nitrilase which hydrolyses the nitrile (cya‐ nide) group (R-CΞN) into the corresponding carboxylic acid and ammonia, as shown in Fig‐ ure 7 [26]. In aqueous solution, ammonia/ammonium equilibrium is observed.

**Figure 7.** Nitrilase hydrolysis on cyanide group [26]

**Figure 6.** NJ tree based on the analysis of the combined gene regions

130 Environmental Biotechnology - New Approaches and Prospective Applications

The majority of the citrus peel components are high carbon source materials and the ammo‐ nia produced from the cyanide degradation can be utilised as a nitrogen source by the mi‐ croorganism. Studies have shown that the optimal pH and temperature for nitrilase production is 8 and 45 ˚C, respectively [12, 35].

The successful treatment of 400 ppm free cyanide supplemented with refined/readily me‐ tabolisable carbon sources has been reported [1]. However, current technology used can be capital intensive for large scale operations. Most studies on free cyanide bioremedia‐ tion efficiency measured the free cyanide reduction periodically as opposed to product formation. This may be misleading since free cyanide is very volatile, even at room tem‐ perature, and the decline in free cyanide concentration observed may be a result of volati‐ lisation into the atmosphere rather than actual biological remediation. The cyanide tolerance of the *A. awamori* (CPUT) isolate was initially assessed up to a 500 ppm CN- (1.2515 g KCN/L) in PDA (Figure 8). **10** 7 (1.2515°g°KCN/L) (1.2515 g KCN/L) **10** Figure 8 **Figure 8.** Cyanide tolerance analysis for *A. awamori* (CPUT) isolate. **Figure 8.** Cyanide tolerance analysis for *A. awamori* (CPUT) isolate **<sup>10</sup>**12 The toxicity of cyanide reduces the functionality of the fungus metabolic processes thus its growth. **<sup>10</sup>**16 A media solution of 42.5 ml of 1% (w/v) refined citrus pectin,

anide (CN-

growth.

of a 1 g CN-

0

50

100

**[CN-] (ppm)**

150

200

250

/L

measure free cyanide reduction periodically, as opposed to

1% (w/v) powered orange peel, Czapek yeast medium and

in solution during biodegradation is indicative of cyanide reduction. However, not all the ammonia/ammonium was

product formation.

) (09701) and Merck ammonium (NH4+) (00683) test kits were used to measure the

Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant...

/L free cyanide

133

http://dx.doi.org/10.5772/53349

Refined pectin 1% (w/v) Orange peel 1% (w/v) Czapek medium Water (control)

2

/NH4+. The or‐

/NH4+, had shown

The orange peel medium showed considerably higher cyanide reduction compared to the other nutrient media evaluated (Figure 9). The change in the cyanide concentration in the

The toxicity of cyanide reduces the functionality of the fungus metabolic processes, thus its

Most studies on free cyanide bioremediation efficiency measured the free cyanide reduction

was again added to each flask to evaluate the robustness of the culture, in each growth me‐ dia. The orange peel culture had the fastest recovery, even with a sudden increase in free cyanide concentration. The numerous enzymes released by the fungus, sufficiently hydro‐ lysed the orange peel which resulted in better supplementation and maintenance of the fun‐

Media solutions of 42.5 ml of 1% (w/v) refined citrus pectin, 1 % (w/v) powered orange peel, Czapek yeast medium and sterile distilled water (standard) were added into 250 ml flasks. To each of the flasks, 1 ml of spore suspension (2x106 spores) was added followed by 7.5 ml

free cyanide and ammonia/ammonium (NH3/NH4+) concentrations in solution.

water medium (control) was due to volatilisation. At day 2, 7.5 ml of 1 g CN-

gus compared to the other media for cyanide bioremediation.

0246

**Figure 9.** Cyanide bioremediation by *A. awamori* (CPUT) in batch cultures.

**Time (days)**

NH3/NH4+ can be used as a nitrogen source for most fungi. The presence of quantifiable NH3/NH4+ in solution during biodegradation is indicative of cyanide reduction. However, not all the NH3/NH4+ was consumed by the fungus (Figure 10). Theoretical NH3/NH4+ was calculated based on the quantity of cyanide degraded and the stoichiometry of the cyanide hydrolysis reaction (Figure 7). The experimental [NH3/NH4+] were lower than that of the

NH3/NH4+ can be used as a nitrogen source for most fungi. The presence of quantifiable NH3/NH4+ in solution during biodegradation is indicative of cyanide reduction. However, not all the NH3/NH4+ was consumed by the fungus (Figure 10). Theoretical NH3/NH4+ was calculated based on the quantity of cyanide degraded and the stoichiometry of the cyanide hydrolysis reaction (Figure 7). The experimental [NH3/NH4+] were lower than that of the the‐ oretical since the fungus metabolised some of the NH3/NH4+ as a nitrogen source. However,

ange peel medium showed the highest concentration of the ammonia/ammonium in solu‐ tion compared to the other media which is indicative of cyanide reduction. The rich carbon

However, for an efficient bioremediation process, a continuous process must be developed to assess the applicability of the *A. awamori* (CPUT) isolate for continuous cyanide bioreme‐ diation processes. One of the effective technologies which have been determined to be effec‐ tive on a large scale is the use of immobilised MBRs, for continuous remediation of

theoretical since the fungus metabolised some of the NH3/NH4+ as a nitrogen source.

the hydrolysis of the cyanide does not result in complete metabolism of NH3

sources present in the orange peel and the further supplementation by NH3

to be an added advantage of using waste orange peel as a potential nutrient source.

periodically as opposed to product formation.

/L cyanide solution.

**Figure 8.** Cyanide tolerance analysis for *A. awamori* (CPUT) isolate

There was a clear decline in the growth of the fungus as the free cyanide concentration was increased. Appreciable growth occurred for the strain for free cyanide concentrations up to 200 ppm. A rapid decline in the growth was observed as the free cyanide concentration ex‐ ceeded 300 ppm. The toxicity of cyanide reduces the functionality of the fungus metabolic processes, thus its growth. There was limited growth observed at cyanide concentrations above 430 ppm. Preliminary analysis on the effect of growth media on *A. awamori* (CPUT) for free cyanide bioremediation was performed in batch cultures, shaken at 180 rpm and 30 ˚C in a ZhiCheng (ZHWY-1102) shaking incubator. Media solutions of 42.5 ml of 1% (w/v) refined citrus pectin, 1 % (w/v) powered orange peel, Czapek yeast medium and sterile dis‐ tilled water (standard) were added into 250 ml flasks. To each of the flasks, 1 ml of spore suspension (2x106 spores) was added followed by 7.5 ml of a 1 g CN- /L cyanide solution.. The experiments were run in duplicate in which sampling was every 48 hours. The samples were centrifuged for 13000 rpm for 5 minutes before any analysis was conducted. Merck cy‐ consumed by the fungus (Figure 10). Theoretical NH3/NH4+ was calculated based on the quantity of cyanide degraded and

anide (CN- ) (09701) and Merck ammonium (NH4+) (00683) test kits were used to measure the free cyanide and ammonia/ammonium (NH3/NH4+) concentrations in solution. **10** 7 (1.2515°g°KCN/L) (1.2515 g KCN/L)

The orange peel medium showed considerably higher cyanide reduction compared to the other nutrient media evaluated (Figure 9). The change in the cyanide concentration in the water medium (control) was due to volatilisation. At day 2, 7.5 ml of 1 g CN- /L free cyanide was again added to each flask to evaluate the robustness of the culture, in each growth me‐ dia. The orange peel culture had the fastest recovery, even with a sudden increase in free cyanide concentration. The numerous enzymes released by the fungus, sufficiently hydro‐ lysed the orange peel which resulted in better supplementation and maintenance of the fun‐ gus compared to the other media for cyanide bioremediation. **10** Figure 8 **Figure 8.** Cyanide tolerance analysis for *A. awamori* (CPUT) isolate. **Figure 8.** Cyanide tolerance analysis for *A. awamori* (CPUT) isolate **<sup>10</sup>**12 The toxicity of cyanide reduces the functionality of the fungus The toxicity of cyanide reduces the functionality of the fungus metabolic processes, thus its growth. **<sup>10</sup>**16 A media solution of 42.5 ml of 1% (w/v) refined citrus pectin, 1% (w/v) powered orange peel, Czapek yeast medium and sterile distilled water (standard) were added into 250 ml flasks. To each of the flasks, 1 ml of spore suspension /L Media solutions of 42.5 ml of 1% (w/v) refined citrus pectin, 1 % (w/v) powered orange peel, Czapek yeast medium and sterile distilled water (standard) were added into 250 ml flasks. To each of the flasks, 1 ml of spore suspension (2x106 spores) was added followed by 7.5 ml of a 1 g CN- /L cyanide solution.

**<sup>11</sup>**11 Ammonia/ammonium can be used as a nitrogen source for NH3/NH4+ can be used as a nitrogen source for most fungi. The presence of quantifiable **Figure 9.** Cyanide bioremediation by *A. awamori* (CPUT) in batch cultures.

be capital intensive for large scale operations. Most studies on free cyanide bioremedia‐ tion efficiency measured the free cyanide reduction periodically as opposed to product formation. This may be misleading since free cyanide is very volatile, even at room tem‐ perature, and the decline in free cyanide concentration observed may be a result of volati‐ lisation into the atmosphere rather than actual biological remediation. The cyanide tolerance of the *A. awamori* (CPUT) isolate was initially assessed up to a 500 ppm CN-

**11** Figure 9

There was a clear decline in the growth of the fungus as the free cyanide concentration was increased. Appreciable growth occurred for the strain for free cyanide concentrations up to 200 ppm. A rapid decline in the growth was observed as the free cyanide concentration ex‐ ceeded 300 ppm. The toxicity of cyanide reduces the functionality of the fungus metabolic processes, thus its growth. There was limited growth observed at cyanide concentrations above 430 ppm. Preliminary analysis on the effect of growth media on *A. awamori* (CPUT) for free cyanide bioremediation was performed in batch cultures, shaken at 180 rpm and 30 ˚C in a ZhiCheng (ZHWY-1102) shaking incubator. Media solutions of 42.5 ml of 1% (w/v) refined citrus pectin, 1 % (w/v) powered orange peel, Czapek yeast medium and sterile dis‐ tilled water (standard) were added into 250 ml flasks. To each of the flasks, 1 ml of spore

spores) was added followed by 7.5 ml of a 1 g CN-

The experiments were run in duplicate in which sampling was every 48 hours. The samples were centrifuged for 13000 rpm for 5 minutes before any analysis was conducted. Merck cy‐

/L cyanide solution..

**<sup>10</sup>**2 Most studies on free cyanide bioremediation efficiency,

metabolic processes thus its growth.

product formation.

cyanide solution.

measure free cyanide reduction periodically, as opposed to

(2x106°spores) was added followed by 7.5°ml of a 1 g CN-

(1.2515 g KCN/L) in PDA (Figure 8).

132 Environmental Biotechnology - New Approaches and Prospective Applications

**Figure 8.** Cyanide tolerance analysis for *A. awamori* (CPUT) isolate

suspension (2x106

2 most fungi. The presence of quantifiable ammonia/ammonium in solution during biodegradation is indicative of cyanide reduction. However, not all the ammonia/ammonium was consumed by the fungus (Figure 10). Theoretical NH3/NH4+ was calculated based on the quantity of cyanide degraded and NH3/NH4+ in solution during biodegradation is indicative of cyanide reduction. However, not all the NH3/NH4+ was consumed by the fungus (Figure 10). Theoretical NH3/NH4+ was calculated based on the quantity of cyanide degraded and the stoichiometry of the cyanide hydrolysis reaction (Figure 7). The experimental [NH3/NH4+] were lower than that of the theoretical since the fungus metabolised some of the NH3/NH4+ as a nitrogen source. NH3/NH4+ can be used as a nitrogen source for most fungi. The presence of quantifiable NH3/NH4+ in solution during biodegradation is indicative of cyanide reduction. However, not all the NH3/NH4+ was consumed by the fungus (Figure 10). Theoretical NH3/NH4+ was calculated based on the quantity of cyanide degraded and the stoichiometry of the cyanide hydrolysis reaction (Figure 7). The experimental [NH3/NH4+] were lower than that of the the‐ oretical since the fungus metabolised some of the NH3/NH4+ as a nitrogen source. However, the hydrolysis of the cyanide does not result in complete metabolism of NH3 /NH4+. The or‐ ange peel medium showed the highest concentration of the ammonia/ammonium in solu‐ tion compared to the other media which is indicative of cyanide reduction. The rich carbon sources present in the orange peel and the further supplementation by NH3 /NH4+, had shown to be an added advantage of using waste orange peel as a potential nutrient source.

> However, for an efficient bioremediation process, a continuous process must be developed to assess the applicability of the *A. awamori* (CPUT) isolate for continuous cyanide bioreme‐ diation processes. One of the effective technologies which have been determined to be effec‐ tive on a large scale is the use of immobilised MBRs, for continuous remediation of

NH3/NH4+ as a nitrogen source.

**Figure 10.** Experimental and theoretical ammonium concentration in batch

into five groups, based on the pore size of the membranes

**<sup>12</sup>**15 Membrane separation processes can be broadly categorised

(Figure 11).

**12** Figure 10

**13** Figure 11

contaminants [8]. The advantage of using MBR technology is that the biomass can be re‐ tained for elongated periods while the continuous remediation of the contaminated water is in progress. the stoichiometry of the cyanide hydrolysis reaction (Figure 10). The experimental NH3/NH4+ were lower than that of the theoretical since the fungus metabolised some of the

**Figure 10.** Experimental and theoretical ammonia/ammonium concentration in batch cultures **Figure 10.** Experimental and theoretical ammonia/ammonium concentration in batch cultures

pore size of the membranes (Figure 11).

### **4. Membrane technology for fixed-film immobilisation in continuous remediation processes**

Membrane separation processes can be broadly categorised into four groups, based on the

**Figure 11.** Filtration spectrum [25]

vantageous property when used in MBRs [28].

chemically and steam sterilized [27].

would limit their application on a large scale.

**4.1. Modes of operation and orientation for membrane bioreactors**

Asymmetric membranes have an excellent mass transfer property which has led to their uti‐ lization in numerous industrial applications, especially for MBRs [8, 25]. The most important ultrafiltration (UF) membrane module types are hollow fibre and capillary tube membranes [21, 27]. They are ideal for biofilm growth because of the nutrients permeation gradient along the membrane and provide a shear free environment [8, 15]. The large surface area of these membranes to their small volume allows for high operational capacity which is an ad‐

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135

Filamentous microorganisms are commonly used in immobilised MBR systems [3, 21, 27]. These particular microorganisms are able to penetrate the membrane due to their apical growth resulting in effective immobilisation on the surface compared to non-motile bacte‐ ria and yeast. Although, comparative studies of ceramic and internally skinned polysul‐ phone (PSu) capillary membranes have shown to provide the best attachment and immobilisation of fungal biofilms than other tubular membrane types [21, 27]. Sheldon and Small [27] showed that the ceramic and internally skinned PSu capillary membrane developed thicker biofilms than tubular membranes. Furthermore, the ceramic capillary membrane can resists mechanical stress caused by the increasing immobilised biofilm build-up on the membrane, as ceramic membranes are mechanically stable and can be

The membrane modules' mode of operation and orientation are important in determining the overall performance of the process. However, one of the biggest challenges in membrane operations is the effect of fouling and feeding mode, orientation and other mechanisms that

Materials used in the construction of membranes include organic or non-organic (e.g. metal, ceramic), homogeneous (e.g. polymer, metal) and heterogeneous (e.g. polymer mixes, mixed glass) solids and solutions (mostly polymers) [25]. Polymeric membranes are commonly used because they are well developed, competitive in separation performances and econom‐ ical [25]. Membrane processes are categorised according to the pore size, molecular cut-off and pressure at which they operate. These categories are inter-related, because as the pore size or the molecular cut-off size is decreased the pressure applied to the membrane increas‐ es [33]. Membrane separation processes can be broadly categorised into four groups, based on the pore size of the membranes (Figure 11).

3 **<sup>15</sup>**14 Immobilised MBRs refers to the fixed microbial biofilm formation on a membrane matrix and the film can be fed a Immobilised MBRs refers to the fixed microbial biofilm formation on a membrane matrix with the film being fed a nutrient medium passing through the lumen. Asymmetric membranes have shown to be effective in immobilising biofilms in MBRs since the membranes allow the transport of nutrients to the biomass immobilised on their external surface [27]. Asymmetric refers to the graded porosity of the membrane substructure and in‐ dicates that the membranes have an inside coating, a skin layer, and combines the high se‐ lectivity of a dense membrane with a high permeation rate of a thin membrane. The exposed microvoids on the externally unskinned surface enable a resilient attachment of the microor‐ ganism on the membrane [21]. The production of asymmetric membranes is by manipulat‐ ing manufacturing parameters during the membrane formation process which results in a unique membrane morphology, characteristics and properties of the membrane [16].

Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant... http://dx.doi.org/10.5772/53349 135

**Figure 11.** Filtration spectrum [25]

3

contaminants [8]. The advantage of using MBR technology is that the biomass can be re‐ tained for elongated periods while the continuous remediation of the contaminated water is

> Pectin - Experimental Pectin - Theoretical

Orange peel - Experimental Orange peel - Theoretical Czapek - Experimental Czapek - Theoretical

0246

134 Environmental Biotechnology - New Approaches and Prospective Applications

**Time (days)**

pore size of the membranes (Figure 11).

on the pore size of the membranes (Figure 11).

**remediation processes**

**Figure 10.** Experimental and theoretical ammonia/ammonium concentration in batch cultures

**Figure 10.** Experimental and theoretical ammonia/ammonium concentration in batch cultures

Membrane separation processes can be broadly categorised into four groups, based on the

Materials used in the construction of membranes include organic or non-organic (e.g. metal, ceramic), homogeneous (e.g. polymer, metal) and heterogeneous (e.g. polymer mixes, mixed glass) solids and solutions (mostly polymers) [25]. Polymeric membranes are commonly used because they are well developed, competitive in separation performances and econom‐ ical [25]. Membrane processes are categorised according to the pore size, molecular cut-off and pressure at which they operate. These categories are inter-related, because as the pore size or the molecular cut-off size is decreased the pressure applied to the membrane increas‐ es [33]. Membrane separation processes can be broadly categorised into four groups, based

**4. Membrane technology for fixed-film immobilisation in continuous**

Immobilised MBRs refers to the fixed microbial biofilm formation on a membrane matrix

unique membrane morphology, characteristics and properties of the membrane [16].

Asymmetric membranes have shown to be effective in immobilising biofilms in MBRs since the membranes allow the transport of nutrients to the biomass immobilised on their external surface [27]. Asymmetric refers to the graded porosity of the membrane substructure and in‐ dicates that the membranes have an inside coating, a skin layer, and combines the high se‐ lectivity of a dense membrane with a high permeation rate of a thin membrane. The exposedmicrovoids on the externally unskinned surface enable a resilient attachment of the microor‐ ganism on the membrane [21]. The production of asymmetric membranes is by manipulat‐ ing manufacturing parameters during the membrane formation process which results in a

with the film being fed a nutrient medium passing through the lumen.

in progress.

**[NH3/NH4+](ppm)**

the stoichiometry of the cyanide hydrolysis reaction (Figure 10). The experimental NH3/NH4+ were lower than that of the theoretical since the fungus metabolised some of the

**Figure 10.** Experimental and theoretical ammonium concentration in batch

into five groups, based on the pore size of the membranes

**<sup>12</sup>**15 Membrane separation processes can be broadly categorised

**<sup>15</sup>**14 Immobilised MBRs refers to the fixed microbial biofilm

formation on a membrane matrix and the film can be fed a

(Figure 11).

cultures 0

NH3/NH4+ as a nitrogen source.

**12** Figure 10

**13** Figure 11

Asymmetric membranes have an excellent mass transfer property which has led to their uti‐ lization in numerous industrial applications, especially for MBRs [8, 25]. The most important ultrafiltration (UF) membrane module types are hollow fibre and capillary tube membranes [21, 27]. They are ideal for biofilm growth because of the nutrients permeation gradient along the membrane and provide a shear free environment [8, 15]. The large surface area of these membranes to their small volume allows for high operational capacity which is an ad‐ vantageous property when used in MBRs [28].

Filamentous microorganisms are commonly used in immobilised MBR systems [3, 21, 27]. These particular microorganisms are able to penetrate the membrane due to their apical growth resulting in effective immobilisation on the surface compared to non-motile bacte‐ ria and yeast. Although, comparative studies of ceramic and internally skinned polysul‐ phone (PSu) capillary membranes have shown to provide the best attachment and immobilisation of fungal biofilms than other tubular membrane types [21, 27]. Sheldon and Small [27] showed that the ceramic and internally skinned PSu capillary membrane developed thicker biofilms than tubular membranes. Furthermore, the ceramic capillary membrane can resists mechanical stress caused by the increasing immobilised biofilm build-up on the membrane, as ceramic membranes are mechanically stable and can be chemically and steam sterilized [27].

### **4.1. Modes of operation and orientation for membrane bioreactors**

The membrane modules' mode of operation and orientation are important in determining the overall performance of the process. However, one of the biggest challenges in membrane operations is the effect of fouling and feeding mode, orientation and other mechanisms that would limit their application on a large scale.

**4.2. Membrane bioreactors: Design and application for cyanide remediation**

MBRs are integrated biological and separation units for the production of value added prod‐ ucts or the bioremediation of toxic components [8]. Immobilised MBRs refers to the fixed microbial biofilm formation on a membrane matrix with the film being fed a nutrient medi‐ um passing through the lumen. These MBR's retain the biomass on the membranes in a low shear environment, separate the nutrient supply from the biomass, as well as continuously remove extracellular metabolic products. These immobilised systems ensure that the bio‐ mass can be maintained in a state of low or non-proliferation for extended periods, while still producing the products desired [8]*.* Filamentous microorganisms have had some degree of success on immobilised MBR systems [3, 21]. However, this has not been evaluated for *A. awamori* biomass for cyanide remediation. Four vertically orientated single fibre membrane

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137

bioreactors (SFMBRs) (Figure 13) were constructed as described by Edwards *et a*l. [4].

The glass housing, was produced by Glasschem (South Africa) and the asymmetric alumini‐ um oxide ceramic capillary membranes were produced and supplied by Hyflux CEPAration

> Outer diameter (m) 0.0028 Inner diameter (m) 0.0018 Wall thickness (m) 0.0005 Burst pressure (Pa) 5.0x106 Maximum temperature (˚C) 1000+ Permeability (m/Pa.s) 6.95x10-10

**Figure 13.** Schematic of SFMBR for filamentous microorganism immobilisation

BV (Netherlands).

**Table 1.** Capillary membrane specifications [27]

**Figure 12.** Modes of operation of membrane modules [2]

The effects of membrane modes of operations, as shown in Figure 12, have been studied us‐ ing both experimental [6, 21, 30] and theoretical approaches [2, 8, 9, 13]. The mode of opera‐ tion is concerned with the distribution and flow of the fluid in the system, as shown in Figure 12. In a dead-end filtration module the feed stream enters the lumen, permeates through the membrane and exit the shell in a continuous stream, with no retentate stream. In a continuous open shell mode, the feed stream enters the lumen and a portion of the feed permeates through the membrane and leaves the shell in a continuous stream and the other portion exit the lumen in a continuous stream. The amount of feed that will permeate through the membrane is dependent on several factors such as trans-membrane pressure, membrane permeability and feed velocity [25]. The pressure in the lumen is always greater than the shell side pressure in both dead-end filtration and continuous open shell modes [30]. This ensures that the trans-membrane flux, fluid's flow direction between the lumen and shell regions, is directed toward the shell [2].

In a closed shell mode, the feed stream enters the lumen and a portion of the feed permeates through the membrane with no outlet shell stream, while the other portion exits the lumen as a continuous stream. The trans-membrane flux for the initial portion of the membrane is directed towards the shell, but for the end portion it is directed toward the lumen [2]. This is referred to as convective recirculation which results in undesired non-uniform distribution of the biomass in the membrane system [2, 30].

In a suction of permeate, the feed stream enters the shell for an equal distribution along the membrane and permeates through the membrane and exits in a continuous stream [8]. The pressure in the shell is kept at greater than that in the lumen for a trans-membrane flux to‐ wards the lumen [30]. There is a negligible pressure gradient in the lumen which results in a relatively uniform trans-membrane flux [30]. The orientation of the MBR is based on its ap‐ plication and is generally operated horizontally or vertically [2, 6, 8, 27]. Studies by Garcin [6] and Ntwampe [21] on Lignin peroxidase and Manganese peroxidase production from *P. chrysosporium* BKMF-1767 (ATCC 24725) in a vertically orientated PSu capillary MBR, had shown the production of these extracellular enzymes were higher in the vertically orientated MBR than in the horizontal orientation. Similarly, the biofilm was denser on the vertically orientated capillary membrane than that in horizontal MBRs [6, 21].

### **4.2. Membrane bioreactors: Design and application for cyanide remediation**

MBRs are integrated biological and separation units for the production of value added prod‐ ucts or the bioremediation of toxic components [8]. Immobilised MBRs refers to the fixed microbial biofilm formation on a membrane matrix with the film being fed a nutrient medi‐ um passing through the lumen. These MBR's retain the biomass on the membranes in a low shear environment, separate the nutrient supply from the biomass, as well as continuously remove extracellular metabolic products. These immobilised systems ensure that the bio‐ mass can be maintained in a state of low or non-proliferation for extended periods, while still producing the products desired [8]*.* Filamentous microorganisms have had some degree of success on immobilised MBR systems [3, 21]. However, this has not been evaluated for *A. awamori* biomass for cyanide remediation. Four vertically orientated single fibre membrane bioreactors (SFMBRs) (Figure 13) were constructed as described by Edwards *et a*l. [4].

**Figure 13.** Schematic of SFMBR for filamentous microorganism immobilisation

The glass housing, was produced by Glasschem (South Africa) and the asymmetric alumini‐ um oxide ceramic capillary membranes were produced and supplied by Hyflux CEPAration BV (Netherlands).


**Table 1.** Capillary membrane specifications [27]

**Figure 12.** Modes of operation of membrane modules [2]

136 Environmental Biotechnology - New Approaches and Prospective Applications

and shell regions, is directed toward the shell [2].

of the biomass in the membrane system [2, 30].

orientated capillary membrane than that in horizontal MBRs [6, 21].

The effects of membrane modes of operations, as shown in Figure 12, have been studied us‐ ing both experimental [6, 21, 30] and theoretical approaches [2, 8, 9, 13]. The mode of opera‐ tion is concerned with the distribution and flow of the fluid in the system, as shown in Figure 12. In a dead-end filtration module the feed stream enters the lumen, permeates through the membrane and exit the shell in a continuous stream, with no retentate stream. In a continuous open shell mode, the feed stream enters the lumen and a portion of the feed permeates through the membrane and leaves the shell in a continuous stream and the other portion exit the lumen in a continuous stream. The amount of feed that will permeate through the membrane is dependent on several factors such as trans-membrane pressure, membrane permeability and feed velocity [25]. The pressure in the lumen is always greater than the shell side pressure in both dead-end filtration and continuous open shell modes [30]. This ensures that the trans-membrane flux, fluid's flow direction between the lumen

In a closed shell mode, the feed stream enters the lumen and a portion of the feed permeates through the membrane with no outlet shell stream, while the other portion exits the lumen as a continuous stream. The trans-membrane flux for the initial portion of the membrane is directed towards the shell, but for the end portion it is directed toward the lumen [2]. This is referred to as convective recirculation which results in undesired non-uniform distribution

In a suction of permeate, the feed stream enters the shell for an equal distribution along the membrane and permeates through the membrane and exits in a continuous stream [8]. The pressure in the shell is kept at greater than that in the lumen for a trans-membrane flux to‐ wards the lumen [30]. There is a negligible pressure gradient in the lumen which results in a relatively uniform trans-membrane flux [30]. The orientation of the MBR is based on its ap‐ plication and is generally operated horizontally or vertically [2, 6, 8, 27]. Studies by Garcin [6] and Ntwampe [21] on Lignin peroxidase and Manganese peroxidase production from *P. chrysosporium* BKMF-1767 (ATCC 24725) in a vertically orientated PSu capillary MBR, had shown the production of these extracellular enzymes were higher in the vertically orientated MBR than in the horizontal orientation. Similarly, the biofilm was denser on the vertically

The two SFMBRs were inoculated with 100 ml of *A. awamori* inoculum (10x106 spores) and two other SFMBRs were used as controls. High pressure reverse filtration of the inoculation spore solution on the external surface of the membranes was used to immobilise the spores onto the external surface of the ceramic membranes.

**Figure 15.** TRS versus time for feed, control and MBR

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139

**Figure 16.** Biofilm development on day (a) 0 and (b) 8

**Figure 14.** Schematic diagram of the experimental setup

The constructed system was setup in a Scientific Manufacturing (SMC) (160 L unit) low tem‐ perature incubator set at 30 ˚C. A Watson Marlow (504S) peristaltic pump was used to sup‐ ply the feed solution to the SFMBRs at a flow rate of 3 ml/hr. The SFMBRs were fitted with two way flow 0.2 μm Millipore air filters, to ensure a monoseptic culture in the system and for aeration. The feed solution initially consisted of 1 % (w/v) orange peel solution which was prepared by adding 10 g milled orange peel to a 1 L Schott bottle. The 1 L solution was filtered through a Whatmann No 1 filter paper and the filtrate produced (orange peel ex‐ tract) was fed to the system for 2 days, in a feed batch mode, to initiate spore germination and biofilm development on the membrane. Thereafter, a feed solution consisting of the or‐ ange peel extract and free cyanide solution (280ppm) was fed to the SFMBR.

The total reduced sugars (TRS), such as glucose, fructose, sucrose, were measured using the dinitrosalicylic (DNS) acid colorimetric method [17]. The Merck cyanide (CN- ) (09701) and Merck ammonium (NH4+) (00683) test kits were used to measure the free cyanide and NH3/NH4+ concentrations, since this is a by-product of the cyanide metabolism. The utilisa‐ tion of the TRS increased for the pure orange peel extract feed in comparison to the control MBRs (Figure 15). The initial feed was rich with TRS and therefore, the metabolism of the TRS was observed. The change in the TRS utilisation for the orange peel extract containing cyanide concentration of 1.7%, 15.2%, 23.7% and 28.8% at day 2, 4, 6 and 8, respectively and therefore the rate of metabolism was drastically hindered which resulted in reduced metab‐ olism of the TRS.

Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant... http://dx.doi.org/10.5772/53349 139

**Figure 15.** TRS versus time for feed, control and MBR

The two SFMBRs were inoculated with 100 ml of *A. awamori* inoculum (10x106

onto the external surface of the ceramic membranes.

138 Environmental Biotechnology - New Approaches and Prospective Applications

**Figure 14.** Schematic diagram of the experimental setup

olism of the TRS.

two other SFMBRs were used as controls. High pressure reverse filtration of the inoculation spore solution on the external surface of the membranes was used to immobilise the spores

The constructed system was setup in a Scientific Manufacturing (SMC) (160 L unit) low tem‐ perature incubator set at 30 ˚C. A Watson Marlow (504S) peristaltic pump was used to sup‐ ply the feed solution to the SFMBRs at a flow rate of 3 ml/hr. The SFMBRs were fitted with two way flow 0.2 μm Millipore air filters, to ensure a monoseptic culture in the system and for aeration. The feed solution initially consisted of 1 % (w/v) orange peel solution which was prepared by adding 10 g milled orange peel to a 1 L Schott bottle. The 1 L solution was filtered through a Whatmann No 1 filter paper and the filtrate produced (orange peel ex‐ tract) was fed to the system for 2 days, in a feed batch mode, to initiate spore germination and biofilm development on the membrane. Thereafter, a feed solution consisting of the or‐

The total reduced sugars (TRS), such as glucose, fructose, sucrose, were measured using the

Merck ammonium (NH4+) (00683) test kits were used to measure the free cyanide and NH3/NH4+ concentrations, since this is a by-product of the cyanide metabolism. The utilisa‐ tion of the TRS increased for the pure orange peel extract feed in comparison to the control MBRs (Figure 15). The initial feed was rich with TRS and therefore, the metabolism of the TRS was observed. The change in the TRS utilisation for the orange peel extract containing cyanide concentration of 1.7%, 15.2%, 23.7% and 28.8% at day 2, 4, 6 and 8, respectively and therefore the rate of metabolism was drastically hindered which resulted in reduced metab‐

ange peel extract and free cyanide solution (280ppm) was fed to the SFMBR.

dinitrosalicylic (DNS) acid colorimetric method [17]. The Merck cyanide (CN-

spores) and

) (09701) and

**Figure 16.** Biofilm development on day (a) 0 and (b) 8

The biofilm development (Figure 16) on the membrane was slow and homogenous along the membrane, as opposed to thick biomass at a particular area of the membrane. The formation of the thin biofilm has its benefits; it allows for better oxygen mass transfer into the biomass, which is vital since *A. awamori*, is an aerobic microorganism. Aerobic microorganisms utilise oxygen for cell maintenance, respiratory oxidation for further growth and for the oxidation of substrates into metabolic products.

The air filter prevented contamination and maintained the oxygen concentration in the MBR due to the gaseous venting, releasing by-products, such as carbon dioxide out of the MBR system and reintroducing oxygen back into the system. The residual cyanide concentration in the permeate decreased with time and a drastic decrease was observed from day 4 as the fungus adapted to the feed (Figure 17). There was also a considerable quantity of cyanide volatilisation compared to actual bioremediation. As shown in the shaken cultures, a majori‐ ty of the ammonia, produced from cyanide hydrolysis, was consumed by the fungus as a nitrogen source. This was an indication that there was further utilisation of ammonia/ ammonium as the fungus adjusts to the cyanide containing feed.

**Figure 18.** Experimental and theoretical ammonia/ammonium concentration

not be done in immersed MBRs.

For the development of a sustainable MBR system for cyanide remediation an alternative MBR system should be used which limits cyanide volatilisation. Immersed MBRs are bio‐ reactors in which the enzyme(s) and/or microorganism(s) or antibiotic(s) are immobilised on membrane(s) and biomass is suspended in the solution and compartmentalised in a re‐ action vessel [3]. Sidestream MBRs are when the membrane module and bioreactor are separate from each other. Immersed and sidestream MBRs are used for conventional bio‐ mass rejection thus allowing for continued biomass utilization [3]. Immersed MBRs re‐ quire less energy than sidestream MBRs. Membrane modules in a pumped sidestream system utilises more energy due to the high pressures to sustain high volumetric flow rates [11]. However, immersed MBRs may be difficult to operate and remove fouling dur‐ ing operation. The separated system of bioreactor and membrane unit in a sidestream MBRs makes it easier to specifically optimise certain parameters for each unit which can‐

Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant...

http://dx.doi.org/10.5772/53349

141

**Figure 17.** Discharge cyanide concentration versus time for feed, control and MBR

**Figure 18.** Experimental and theoretical ammonia/ammonium concentration

The biofilm development (Figure 16) on the membrane was slow and homogenous along the membrane, as opposed to thick biomass at a particular area of the membrane. The formation of the thin biofilm has its benefits; it allows for better oxygen mass transfer into the biomass, which is vital since *A. awamori*, is an aerobic microorganism. Aerobic microorganisms utilise oxygen for cell maintenance, respiratory oxidation for further growth and for the oxidation

The air filter prevented contamination and maintained the oxygen concentration in the MBR due to the gaseous venting, releasing by-products, such as carbon dioxide out of the MBR system and reintroducing oxygen back into the system. The residual cyanide concentration in the permeate decreased with time and a drastic decrease was observed from day 4 as the fungus adapted to the feed (Figure 17). There was also a considerable quantity of cyanide volatilisation compared to actual bioremediation. As shown in the shaken cultures, a majori‐ ty of the ammonia, produced from cyanide hydrolysis, was consumed by the fungus as a nitrogen source. This was an indication that there was further utilisation of ammonia/

ammonium as the fungus adjusts to the cyanide containing feed.

**Figure 17.** Discharge cyanide concentration versus time for feed, control and MBR

of substrates into metabolic products.

140 Environmental Biotechnology - New Approaches and Prospective Applications

For the development of a sustainable MBR system for cyanide remediation an alternative MBR system should be used which limits cyanide volatilisation. Immersed MBRs are bio‐ reactors in which the enzyme(s) and/or microorganism(s) or antibiotic(s) are immobilised on membrane(s) and biomass is suspended in the solution and compartmentalised in a re‐ action vessel [3]. Sidestream MBRs are when the membrane module and bioreactor are separate from each other. Immersed and sidestream MBRs are used for conventional bio‐ mass rejection thus allowing for continued biomass utilization [3]. Immersed MBRs re‐ quire less energy than sidestream MBRs. Membrane modules in a pumped sidestream system utilises more energy due to the high pressures to sustain high volumetric flow rates [11]. However, immersed MBRs may be difficult to operate and remove fouling dur‐ ing operation. The separated system of bioreactor and membrane unit in a sidestream MBRs makes it easier to specifically optimise certain parameters for each unit which can‐ not be done in immersed MBRs.

closed sidestream MBR would provide a suitable system for complete hydrolysis of the sug‐ ars from the orange peel. The unhydrolysed sugar components in the peel can be recycled and further hydrolysed depending on the molecular-weight cut-off size of the membranes

Continuous Biotechnological Treatment of Cyanide Contaminated Waters by Using a Cyanide Resistant...

operate reactor unit which is easier to clean with an added advantage of optimising each

, Seteno Karabo Obed Ntwampe2

1 Faculty of Engineering, Department of Chemical Engineering, Cape Peninsula University

2 Faculty of Applied Science, Department of Agriculture and Food Science, Biotechnology

[1] Akcil A., Karahan AG., Ciftci H., Sagdic O. Biological Treatment of Cyanide by Natu‐ ral Isolated Bacteria (*Pseudomonas sp*.). Minerals Engineering 2003; 16(7) 643-649. [2] Bruining WJ. A General Description of Flows and Pressures in Hollow Fibre Mem‐

[3] De Jager D. *Streptomyces coelicolor* Biofilm Growth Kinetics and Oxygen Mass Trans‐ fer within a Membrane Gradostat Bioreactor. Masters of Technology thesis. Cape

[4] Edwards W., Leukes WD., Fraser SJ. High Throughput Bioprocess Apparatus, SA

[5] Fogler HS. Elements of Chemical Reaction Engineering, 4th edition. New Jersey: Per‐

[6] Garcin CJ. Design and Manufacturing of Membrane Bioreactor Cultivation of Fungi.

programme, Cape Peninsula University of Technology, Cape Town, South Africa

brane Modules. Chemical Engineering Science 1988; 44(6) 1441-1447.

Peninsula University of Technology; 2010.

Masters of Science thesis. Rhodes University; 2002.

patent WO 2007/116266 A1 (2007).

son Education International; 2006.

This research was sponsored by a CPUT University Research Fund (URF RK16).

volatilization thus, offer a safe to

http://dx.doi.org/10.5772/53349

143

and

used in the reactor. The sidestream MBR can reduce CN-

unit individually.

**Author details**

**References**

Bruno Alexandre Quistorp Santos1

of Technology, Cape Town, South Africa

James Hamuel Doughari2

**Acknowledgements**

**Figure 19.** Side stream MBR system

For the continuous bioremediation of cyanide, a sidestream MBR (Figure 19) would be ideal since the solid material, yet to be hydrolysed, can be recycled into the continuous stirred tank reactor (CSTR) and the bioremediated wastewater can be collected as the per‐ meate product. The CSTR can be initially loaded with milled orange peel in a water solu‐ tion and inoculated with *A. awamori* thus the cyanide containing wastewater can be continuously fed into the system.

### **5. Conclusion**

Large quantities of organic wastes are generated every year, but it has been shown that many of the waste, particularly from the agricultural sector, can be utilised as nutrient source for microbial systems. The use of orange peel has shown to be a rich source for culti‐ vation and supplementation for *A. awamori* (CPUT) isolate for cyanide remediation. The am‐ monia produced from the cyanide hydrolysis can be used as a nitrogen source by the fungus, although incomplete metabolism of the ammonia was observed. This could be im‐ proved by changing operating conditions so that the degradation of cyanide, thus the re‐ lease and subsequent consumption of the ammonia is improved.

Challenges that are also evident in bioremediation are due to its volatility, especially in open/agitated cultures. The hydrolysis of sugar components from the orange peel by merely boiling it in water is not an effective true solution since this process results in incomplete hydrolysis/liberation of the sugar components and residual solids. This can be improved by hydrolysis using microorganism such as *A. awamori*, as used in this study. The use of an en‐ closed sidestream MBR would provide a suitable system for complete hydrolysis of the sug‐ ars from the orange peel. The unhydrolysed sugar components in the peel can be recycled and further hydrolysed depending on the molecular-weight cut-off size of the membranes used in the reactor. The sidestream MBR can reduce CN volatilization thus, offer a safe to operate reactor unit which is easier to clean with an added advantage of optimising each unit individually.

### **Acknowledgements**

This research was sponsored by a CPUT University Research Fund (URF RK16).

### **Author details**

**Figure 19.** Side stream MBR system

142 Environmental Biotechnology - New Approaches and Prospective Applications

continuously fed into the system.

**5. Conclusion**

For the continuous bioremediation of cyanide, a sidestream MBR (Figure 19) would be ideal since the solid material, yet to be hydrolysed, can be recycled into the continuous stirred tank reactor (CSTR) and the bioremediated wastewater can be collected as the per‐ meate product. The CSTR can be initially loaded with milled orange peel in a water solu‐ tion and inoculated with *A. awamori* thus the cyanide containing wastewater can be

Large quantities of organic wastes are generated every year, but it has been shown that many of the waste, particularly from the agricultural sector, can be utilised as nutrient source for microbial systems. The use of orange peel has shown to be a rich source for culti‐ vation and supplementation for *A. awamori* (CPUT) isolate for cyanide remediation. The am‐ monia produced from the cyanide hydrolysis can be used as a nitrogen source by the fungus, although incomplete metabolism of the ammonia was observed. This could be im‐ proved by changing operating conditions so that the degradation of cyanide, thus the re‐

Challenges that are also evident in bioremediation are due to its volatility, especially in open/agitated cultures. The hydrolysis of sugar components from the orange peel by merely boiling it in water is not an effective true solution since this process results in incomplete hydrolysis/liberation of the sugar components and residual solids. This can be improved by hydrolysis using microorganism such as *A. awamori*, as used in this study. The use of an en‐

lease and subsequent consumption of the ammonia is improved.

Bruno Alexandre Quistorp Santos1 , Seteno Karabo Obed Ntwampe2 and James Hamuel Doughari2

1 Faculty of Engineering, Department of Chemical Engineering, Cape Peninsula University of Technology, Cape Town, South Africa

2 Faculty of Applied Science, Department of Agriculture and Food Science, Biotechnology programme, Cape Peninsula University of Technology, Cape Town, South Africa

### **References**


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[22] O'Donnell K., Nirenberg HI., Aoki T., Cigelnik E. A Multigene Phylogeny of the *Gib‐ berella fujikuroi* Species Complex: Detection of Additional Phylogenetically Distinct

[23] Patil YB., Paknikar KM. Development of a Process Biodetoxification of Metal Cya‐

[24] Patil YB., Paknikar KM. Removal and Recovery of Metal Cyanides using a Combina‐ tion of Biosorption and Biodegradation Processes. Biotechnology Letters 1999; 21(10)

[25] Perry RH., Green DW., Maloney JO. 1988. Perry's Chemical Engineer's Handbook, 7th

[26] Rao MA., Scelza R., Scotti R., Gianfreda L. Role of Enzymes in the Remediation of Polluted Environments. Journal of Soil Science and Plant Nutrition 2010; 10(3)

[27] Sheldon MS., Small HJ. Immobilisation and Biofilm Development of *Phanerochaete chrysosporium* on Polysulphone and Ceramic Membranes. Journal of Membrane Sci‐

[28] Stamatialis DF., Papenburg BJ., Girones M., Saiful S., Bettahalli SNM., Schmitmeier S., Wessling M. 2008. Medical Applications of Membranes: Drug Delivery, Artificial Organs and Tissue Engineering. Journal of Membrane Science 2008; 308(1-2) 1-34.

[29] Sud D., Mahajan G., Kaur MP. Agricultural Waste Material as Potential Adsorbent for Sequestering Heavy Metal Ions from Aqueous Solutions - a Review. BioResource

[30] Thakaran JP., Chau PC. Operation and Pressure Distribution of Immobilised Hollow Fibre Bioreactors. Biotechnology and Bioengineering 1986; 28(7) 1064-1071.

[31] Torrado AM., Cortés S., Salgado JM., Max B., Rodríguez N., Bibbins BP., Converti A., Domínguez JM. Citric Acid Production from Orange Peel Waste by Solid-State Fer‐

[32] Umsza-Guez MA., Díaz AB., de Ory I., Blandino A., Gomes E., Caro I. Xylanase Pro‐ duction by *Aspergillus awamori* Under Solid State Fermentation Conditions on Toma‐

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[34] Varga J., Frisvad JC., Kocsubé S., Brankovics B., Tóth B., Szigeti G., Samson, RA. New and Revisited Species in *Aspergillus* Section *Nigri*. Studies in Mycology 2011; 69(1)

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[15] Leukes WD. Development and Characterisation of a Membrane Gradostat Bioreactor for the Bioremediation of Aromatic Pollutants using White Rot Fungus. PhD thesis.

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**Chapter 7**

**Biodegradation of Cyanobacterial Toxins**

Water is an essential natural resource, necessary for drinking, agriculture and industrial activities, and providing the human population with safe drinking water is one of the most important issues in public health. Cyanobacteria produce toxins that may present a hazard for drinking water safety. These toxins are structurally diverse and their effects range from liver damage, including liver cancer, to neurotoxicity. Toxic cyanobacteria have been reported in lakes and reservoirs around the world. The World Health Organization (WHO) has set a provisional drinking water guideline of 1 μg/L for microcystin-LR, one of the most commonly

The occurrence of cyanobacteria and their toxins in water bodies used for the production of drinking water causes a technical challenge for water treatment and cleaning. Drinking water should be pure enough to be consumed or used with low risk of immediate or long term harm. The presence of toxins in drinking water creates a potential risk of toxin exposure for water consumers. Conventional water treatment procedures are in some cases insufficient in the removal of cyanobacterial toxins. Besides the chemical and physical methods used, biological degradation could be an efficient method of water detoxification. Therefore there is a need for

This review describes problems related to cyanobacterial toxins and safe drinking water, compares already existing methods of water treatment and cyanotoxin-removal and proposes novel methods of water decontamination. The majority of cyanotoxin-biodegradation studies so far have focused on bacteria isolated from water sources exposed to microcystin-containing blooms. The use of probiotic bacteria is proposed and discussed as a new and efficient means of cyanotoxin-degradation. The removal of cyanobacterial toxins and other environmental contaminants from drinking water is of great importance and probiotic bacteria show prom‐ ising results in this respect. There is a high demand for effective and low-cost approaches for

> © 2013 Nybom; 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 Nybom; licensee InTech. This is a paper 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.

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

Additional information is available at the end of the chapter

Sonja Nybom

**1. Introduction**

http://dx.doi.org/10.5772/55511

occurring cyanotoxin worldwide [1].

simple, low-cost and effective water treatment procedures.

[36] White TJ., Bruns T., Lee S., Taylor J. Amplification and Direct Identification of Fungal Ribosomal RNA Genes for Phylogenetics. In: Innis MA., Gelfand DH., Sinsky JJ., White TJ. (ed.) PCR Protocols: a Guide to Methods and Applications. San Diego: Aca‐ demic Press; 1990. p315-322

## **Biodegradation of Cyanobacterial Toxins**

### Sonja Nybom

[35] Vejvoda V., Kubáč D., Davidová A., Kaplan O., Šulc M., Šveda O., Radka Chaloupko‐ vá R., Martínková L. Purification and Characterization of Nitrilase from *Fusarium sol‐*

[36] White TJ., Bruns T., Lee S., Taylor J. Amplification and Direct Identification of Fungal Ribosomal RNA Genes for Phylogenetics. In: Innis MA., Gelfand DH., Sinsky JJ., White TJ. (ed.) PCR Protocols: a Guide to Methods and Applications. San Diego: Aca‐

*ani* IMI196840. Process Biochemistry 2010; 45(7) 1115-1120.

146 Environmental Biotechnology - New Approaches and Prospective Applications

demic Press; 1990. p315-322

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/55511

### **1. Introduction**

Water is an essential natural resource, necessary for drinking, agriculture and industrial activities, and providing the human population with safe drinking water is one of the most important issues in public health. Cyanobacteria produce toxins that may present a hazard for drinking water safety. These toxins are structurally diverse and their effects range from liver damage, including liver cancer, to neurotoxicity. Toxic cyanobacteria have been reported in lakes and reservoirs around the world. The World Health Organization (WHO) has set a provisional drinking water guideline of 1 μg/L for microcystin-LR, one of the most commonly occurring cyanotoxin worldwide [1].

The occurrence of cyanobacteria and their toxins in water bodies used for the production of drinking water causes a technical challenge for water treatment and cleaning. Drinking water should be pure enough to be consumed or used with low risk of immediate or long term harm. The presence of toxins in drinking water creates a potential risk of toxin exposure for water consumers. Conventional water treatment procedures are in some cases insufficient in the removal of cyanobacterial toxins. Besides the chemical and physical methods used, biological degradation could be an efficient method of water detoxification. Therefore there is a need for simple, low-cost and effective water treatment procedures.

This review describes problems related to cyanobacterial toxins and safe drinking water, compares already existing methods of water treatment and cyanotoxin-removal and proposes novel methods of water decontamination. The majority of cyanotoxin-biodegradation studies so far have focused on bacteria isolated from water sources exposed to microcystin-containing blooms. The use of probiotic bacteria is proposed and discussed as a new and efficient means of cyanotoxin-degradation. The removal of cyanobacterial toxins and other environmental contaminants from drinking water is of great importance and probiotic bacteria show prom‐ ising results in this respect. There is a high demand for effective and low-cost approaches for

© 2013 Nybom; 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 Nybom; licensee InTech. This is a paper 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.

removing cyanotoxins from potable water due to the significant health risk and inadequate access to safe drinking water.

Adda is (2S,3S,8S,9S)-3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4(E),6(E)-dienoic acid (Figure 1). The Adda component of microcystins is contributing to their toxicity [4,9]. There are around 100 structural variants of microcystins described in the literature (listed in [3,10,11]). The most widely-distributed [4] and studied microcystin variant is microcystin-LR (MC-LR), with the amino acid residues leucine and arginine in positions 2 and 4, respectively, and a molecular weight of 994. Production of MC-LR is dependent on various factors like strain specificity, genetic differences and metabolic processes required for toxin production [9]. A single bloom can have both toxigenic and non-toxigenic strains within it [7]. The toxins are generally bound to the cell membrane and are released as cells age and die, and under stress.

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511 149

MC-LR is hepatotoxic and a potent tumour promoter. The primary target organ of MC-LR is the liver [12,13] although it also affects the kidney, gastrointestinal tract and colon [14]. Microcystins are potent and specific inhibitors of serine/threonine-specific protein phospha‐ tases 1 and 2A [15]. Microcystins are distributed in waterbodies worldwide, and the toxicity on exposure to microcystins has been reported worldwide in fish, animals and humans (reviewed in [16]). The World Health Organization has set a provisional drinking water

The cyclic pentapeptide nodularin (NOD) is common in brackish water. It occurs in the Baltic Sea as well as in saline lakes and estuaries. In the Baltic Sea, marine blooms of *Nodularia spumigena* are among some of the largest cyanobacterial mass events in the world. Cylindro‐ spermopsin (CYN), originally isolated from the cyanobacterium *Cylindrospermopsis racibor‐ skii*, is an alkaloid cytotoxin with the structure of a tricyclic guanidine moiety attached to a hydroxymethyluracil [18] and a molecular weight of 415. Cylindrospermopsin inhibits protein synthesis and mainly affects the liver [19], but can also affect the kidney, spleen, thymus, and

They can also passively leak out of cells or be released by lytic bacteria [4].

guideline of 1 μg/L for MC-LR [1]; new edition in [17].

**Figure 1.** General structure of the hepatotoxic cyclic peptides, microcystins.

**2.2. Other cyanobacterial toxins**

### **2. Cyanobacterial toxins**

Cyanobacteria have a long evolutionary history and are among the oldest organisms in the world. There is evidence of the organisms even from around 3500 million years ago [2]. Cyanobacteria carry out oxygen-evolving photosynthesis. In eutrophic water, cyanobacteria recurrently form mass occurrences, so-called water blooms. Mass occurrences of cyanobacteria can be toxic. They have caused a number of animal poisonings and may also pose a threat to human health.

Cyanobacteria produce many different classes of biologically active compounds, including hepatotoxic cyclic peptides, microcystins and nodularins, cytotoxic cylindrospermopsins, neurotoxic anatoxin-a and -a(S), saxitoxins, neurotoxic amino acid β-N-methylamino-Lalanine (BMAA) and non-toxic irritating lipopolysaccharides [3]. Although both neurotoxins and hepatotoxins are distributed worldwide [4,5], it appears that hepatotoxic blooms of cyanobacteria are more commonly found than neurotoxic blooms, and neurotoxins are considered to be of lower risk as they are less stable [6]. In contrast, hepatotoxins are highly stable and exposure to these toxins has resulted in significant toxicity to both animals and humans.

Cyanobacteria are ubiquitous in their distribution in both fresh and marine waters. Toxic cyanobacterial blooms have been reported in most parts of the world, reviewed in [7]. Cyanobacterial blooms are a result of the increasing eutrophication in waterbodies [7]. Most of these cyanobacteria are harmful to animals and humans because of their production of toxins. Over the past several centuries, human nutrient over-enrichment in water, particularly nitrogen and phosphorus, associated with urban, agricultural and industrial development, has promoted eutrophication, which favours algal and cyanobacterial bloom formation. Decay of these excessive blooms results in decreased dissolved oxygen and the release of cyanotoxins in the water, which can result in mortality of animals and even humans [7].

### **2.1. Microcystins**

Globally, the most frequently reported cyanobacterial toxins are cyclic heptapeptide hepato‐ toxins, microcystins (MC). These can be found primarily in some species of the freshwater genera *Microcystis*, *Anabaena*, *Planktothrix*, *Nostoc*, and *Anabaenopsis*. Microcystins are named after *Microcystis aeruginosa*, the cyanobacterium in which the toxin was first isolated and described [8].

Microcystins are cyclic heptapeptides with variable amino acids and a general structure of cyclo(‐D‐Ala(1)–L-X(2)–D‐MeAsp(iso-linkage)(3)–L-Z(4)–Adda(5)–D‐Glu(iso-linkage)(6)– Mdha(7), in which amino acid residues at 2 and 4 are variable L-amino acids, D-MeAsp is Derythro-β-methylaspartic acid, and Mdha is N-methyldehydroalanine, while the amino acid Adda is (2S,3S,8S,9S)-3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4(E),6(E)-dienoic acid (Figure 1). The Adda component of microcystins is contributing to their toxicity [4,9]. There are around 100 structural variants of microcystins described in the literature (listed in [3,10,11]). The most widely-distributed [4] and studied microcystin variant is microcystin-LR (MC-LR), with the amino acid residues leucine and arginine in positions 2 and 4, respectively, and a molecular weight of 994. Production of MC-LR is dependent on various factors like strain specificity, genetic differences and metabolic processes required for toxin production [9]. A single bloom can have both toxigenic and non-toxigenic strains within it [7]. The toxins are generally bound to the cell membrane and are released as cells age and die, and under stress. They can also passively leak out of cells or be released by lytic bacteria [4].

MC-LR is hepatotoxic and a potent tumour promoter. The primary target organ of MC-LR is the liver [12,13] although it also affects the kidney, gastrointestinal tract and colon [14]. Microcystins are potent and specific inhibitors of serine/threonine-specific protein phospha‐ tases 1 and 2A [15]. Microcystins are distributed in waterbodies worldwide, and the toxicity on exposure to microcystins has been reported worldwide in fish, animals and humans (reviewed in [16]). The World Health Organization has set a provisional drinking water guideline of 1 μg/L for MC-LR [1]; new edition in [17].

**Figure 1.** General structure of the hepatotoxic cyclic peptides, microcystins.

### **2.2. Other cyanobacterial toxins**

removing cyanotoxins from potable water due to the significant health risk and inadequate

Cyanobacteria have a long evolutionary history and are among the oldest organisms in the world. There is evidence of the organisms even from around 3500 million years ago [2]. Cyanobacteria carry out oxygen-evolving photosynthesis. In eutrophic water, cyanobacteria recurrently form mass occurrences, so-called water blooms. Mass occurrences of cyanobacteria can be toxic. They have caused a number of animal poisonings and may also pose a threat to

Cyanobacteria produce many different classes of biologically active compounds, including hepatotoxic cyclic peptides, microcystins and nodularins, cytotoxic cylindrospermopsins, neurotoxic anatoxin-a and -a(S), saxitoxins, neurotoxic amino acid β-N-methylamino-Lalanine (BMAA) and non-toxic irritating lipopolysaccharides [3]. Although both neurotoxins and hepatotoxins are distributed worldwide [4,5], it appears that hepatotoxic blooms of cyanobacteria are more commonly found than neurotoxic blooms, and neurotoxins are considered to be of lower risk as they are less stable [6]. In contrast, hepatotoxins are highly stable and exposure to these toxins has resulted in significant toxicity to both animals and

Cyanobacteria are ubiquitous in their distribution in both fresh and marine waters. Toxic cyanobacterial blooms have been reported in most parts of the world, reviewed in [7]. Cyanobacterial blooms are a result of the increasing eutrophication in waterbodies [7]. Most of these cyanobacteria are harmful to animals and humans because of their production of toxins. Over the past several centuries, human nutrient over-enrichment in water, particularly nitrogen and phosphorus, associated with urban, agricultural and industrial development, has promoted eutrophication, which favours algal and cyanobacterial bloom formation. Decay of these excessive blooms results in decreased dissolved oxygen and the release of cyanotoxins

Globally, the most frequently reported cyanobacterial toxins are cyclic heptapeptide hepato‐ toxins, microcystins (MC). These can be found primarily in some species of the freshwater genera *Microcystis*, *Anabaena*, *Planktothrix*, *Nostoc*, and *Anabaenopsis*. Microcystins are named after *Microcystis aeruginosa*, the cyanobacterium in which the toxin was first isolated and

Microcystins are cyclic heptapeptides with variable amino acids and a general structure of cyclo(‐D‐Ala(1)–L-X(2)–D‐MeAsp(iso-linkage)(3)–L-Z(4)–Adda(5)–D‐Glu(iso-linkage)(6)– Mdha(7), in which amino acid residues at 2 and 4 are variable L-amino acids, D-MeAsp is Derythro-β-methylaspartic acid, and Mdha is N-methyldehydroalanine, while the amino acid

in the water, which can result in mortality of animals and even humans [7].

access to safe drinking water.

148 Environmental Biotechnology - New Approaches and Prospective Applications

**2. Cyanobacterial toxins**

human health.

humans.

**2.1. Microcystins**

described [8].

The cyclic pentapeptide nodularin (NOD) is common in brackish water. It occurs in the Baltic Sea as well as in saline lakes and estuaries. In the Baltic Sea, marine blooms of *Nodularia spumigena* are among some of the largest cyanobacterial mass events in the world. Cylindro‐ spermopsin (CYN), originally isolated from the cyanobacterium *Cylindrospermopsis racibor‐ skii*, is an alkaloid cytotoxin with the structure of a tricyclic guanidine moiety attached to a hydroxymethyluracil [18] and a molecular weight of 415. Cylindrospermopsin inhibits protein synthesis and mainly affects the liver [19], but can also affect the kidney, spleen, thymus, and heart. It is a cyanotoxin occurring in tropical or subtropical regions that has recently been detected also in temperate regions.

separated by microscopic identification. To confirm that a particular cyanobacterial strain produces toxins, it is important to isolate a culture of that strain, and to detect and quantify

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511 151

Many cyanobacteria produce potent toxins. As reported in literature, problems caused by cyanobacteria are encountered around the world and problems related to safe drinking water production are common (reviewed in e.g. [7]). The human health effects caused by cyanobac‐ terial toxins vary in severity from mild gastroenteritis to severe and sometimes fatal diarrhoea, dysentery and hepatitis. Microcystins, including the most common variant MC-LR, are hepatotoxic and potent tumour promoters. Acute symptoms reported after exposure to microcystin-containing cyanobacteria include gastrointestinal disorders, nausea, vomiting, fever and irritation of the skin, ears, eyes, throat and respiratory tract, abdominal pain, kidney and liver damage. There are several reports of human health effects associated with ingestion of water containing microcystins, with effects ranging from gastroenteritis [28] to liver damage

Humans can be exposed to a range of cyanotoxins contained either in cyanobacterial cells or released into the water. The dissolved toxins are stable against low pH and enzymatic degradation and will therefore remain intact within the digestive tract. As microcystins do not readily penetrate the cell membrane [31], they enter the body from the intestine via the organic anion transporting polypeptides [32]. From the blood microcystins are then concentrated in the liver as a result of active uptake by hepatocytes [33]. The toxins are covalently bound to protein phosphatases in the hepatocyte cytosol [34]. Human health problems are often associated with chronic exposure to low microcystin concentrations in inadequately treated drinking water, contaminated food (such as fish, mussels and prawns) or with the consump‐ tion of algal supplements contaminated with cyanotoxins. Exposure routes include the oral

Poisonings caused by cyanotoxins produced during heavy blooms have affected both humans and wild and domestic animals. Both hepatotoxic and neurotoxic poisonings have been associated with mass occurrences of cyanobacteria [7]. Many reported incidents of human health effects have involved inappropriate treatment of water supplies. The health risk caused by cyanotoxin exposure is difficult to quantify, since the actual exposure and resulting effects have not been conclusively determined. The most likely route for human exposure is the oral

Due to the growing concern about health effects of cyanotoxins especially via drinking water, WHO has adopted a provisional guideline value of 1.0 μg/L for MC-LR in 1998 [1]. The newest 4th edition to the drinking water guideline was published in 2011 [17]. Assessment of different water treatment procedures has shown that many of the treatment methods result in a

route, through inhalation, through dermal exposure or the nasal mucosa [35,36].

route via drinking water [37], and from recreational use of lakes and rivers [36].

**4. Human health effects caused by cyanobacterial toxins**

toxin concentrations in the pure culture.

[12] and even death [29,30].

**4.1. Risk assessment**

Cyanobacterial neurotoxins belong to a diverse group of heterocyclic compounds called alkaloids. Three types of cyanobacterial neurotoxins, anatoxin-a, anatoxin-a(S), and saxitoxins, are known. A mild neurotoxin, BMAA, has been found in a variety of cyanobacteria [20,21]. Anatoxin-a is a small alkaloid with a molecular weight of 165, and it mimics the effect of acetylcholine and causes rapid death by respiratory arrest. Homoanatoxin-a (MW 179) is an anatoxin-a homologue. Anatoxin-a is perhaps the most common cyanobacterial neurotoxin, especially in North America and Europe, and has caused numerous animal poisonings. Anatoxin-a(S) is an irreversible acetylcholine esterase inhibitor and its characteristic signs of poisonings in mice include salivation. Anatoxin-a(S) was first reported in North America where it has caused animal poisonings and later also in Denmark [22].

Saxitoxins, also known as paralytic shellfish poisons (PSP toxins) were originally isolated and characterised from marine dinoflagellates [23]. Saxitoxins are sodium channel blocking agents causing paralysis and have caused human poisonings due to their ability to concentrate in shellfish [23].

Lipopolysaccharide endotoxins are generally found in the outer membrane of the cell wall of Gram-negative bacteria, also in cyanobacteria. Bacterial lipopolysaccharides are pyrogen‐ ic and toxic [24]. It is often the fatty acid component of lipopolysaccharides that elicits an irritant, pyrogenic or allergenic response in humans and mammals. Cyanobacterial lipopolysaccharides may contribute to human health problems via exposure to mass occurrences of cyanobacteria.

### **3. Occurrence and levels of cyanobacteria and hepatotoxins**

Toxic cyanobacteria are found worldwide both in inland and coastal water environments. Cyanobacteria occur in various environments including water, such as fresh and brackish water, oceans, hot springs, moist terrestrial environments such as soil, and in symbioses with plants, lichens and primitive animals. Some environmental conditions, including sunlight, warm weather, low turbulence and high nutrient levels, can promote growth. A high density of suspended cells may lead to the formation of surface scums and high toxin concentrations.

The toxins are not actively secreted to the surrounding water; most of the toxin is intracellular in growing cells. The release of toxin occurs during senescence of the cultures and when cultures shift from growth phase to stationary and death phases. Under field conditions, the majority of microcystin is intracellular during active growth of the cells [25]. There are reports of hepatotoxic blooms from all continents around the world [7]. Some of the highest reported cyanotoxin concentrations in bloom samples (measured by HPLC) have been 7300 μg/g dry weight microcystin in a *Microcystis* bloom from China [26], 18000 μg/g dry weight nodularin in a *Nodularia* bloom from the Baltic sea [27] and 5500 μg/g dry weight cylindrospermopsin from Australia [3]. Toxic and non-toxic strains from the same cyanobacterial species cannot be separated by microscopic identification. To confirm that a particular cyanobacterial strain produces toxins, it is important to isolate a culture of that strain, and to detect and quantify toxin concentrations in the pure culture.

### **4. Human health effects caused by cyanobacterial toxins**

Many cyanobacteria produce potent toxins. As reported in literature, problems caused by cyanobacteria are encountered around the world and problems related to safe drinking water production are common (reviewed in e.g. [7]). The human health effects caused by cyanobac‐ terial toxins vary in severity from mild gastroenteritis to severe and sometimes fatal diarrhoea, dysentery and hepatitis. Microcystins, including the most common variant MC-LR, are hepatotoxic and potent tumour promoters. Acute symptoms reported after exposure to microcystin-containing cyanobacteria include gastrointestinal disorders, nausea, vomiting, fever and irritation of the skin, ears, eyes, throat and respiratory tract, abdominal pain, kidney and liver damage. There are several reports of human health effects associated with ingestion of water containing microcystins, with effects ranging from gastroenteritis [28] to liver damage [12] and even death [29,30].

Humans can be exposed to a range of cyanotoxins contained either in cyanobacterial cells or released into the water. The dissolved toxins are stable against low pH and enzymatic degradation and will therefore remain intact within the digestive tract. As microcystins do not readily penetrate the cell membrane [31], they enter the body from the intestine via the organic anion transporting polypeptides [32]. From the blood microcystins are then concentrated in the liver as a result of active uptake by hepatocytes [33]. The toxins are covalently bound to protein phosphatases in the hepatocyte cytosol [34]. Human health problems are often associated with chronic exposure to low microcystin concentrations in inadequately treated drinking water, contaminated food (such as fish, mussels and prawns) or with the consump‐ tion of algal supplements contaminated with cyanotoxins. Exposure routes include the oral route, through inhalation, through dermal exposure or the nasal mucosa [35,36].

### **4.1. Risk assessment**

heart. It is a cyanotoxin occurring in tropical or subtropical regions that has recently been

Cyanobacterial neurotoxins belong to a diverse group of heterocyclic compounds called alkaloids. Three types of cyanobacterial neurotoxins, anatoxin-a, anatoxin-a(S), and saxitoxins, are known. A mild neurotoxin, BMAA, has been found in a variety of cyanobacteria [20,21]. Anatoxin-a is a small alkaloid with a molecular weight of 165, and it mimics the effect of acetylcholine and causes rapid death by respiratory arrest. Homoanatoxin-a (MW 179) is an anatoxin-a homologue. Anatoxin-a is perhaps the most common cyanobacterial neurotoxin, especially in North America and Europe, and has caused numerous animal poisonings. Anatoxin-a(S) is an irreversible acetylcholine esterase inhibitor and its characteristic signs of poisonings in mice include salivation. Anatoxin-a(S) was first reported in North America

Saxitoxins, also known as paralytic shellfish poisons (PSP toxins) were originally isolated and characterised from marine dinoflagellates [23]. Saxitoxins are sodium channel blocking agents causing paralysis and have caused human poisonings due to their ability to concentrate in

Lipopolysaccharide endotoxins are generally found in the outer membrane of the cell wall of Gram-negative bacteria, also in cyanobacteria. Bacterial lipopolysaccharides are pyrogen‐ ic and toxic [24]. It is often the fatty acid component of lipopolysaccharides that elicits an irritant, pyrogenic or allergenic response in humans and mammals. Cyanobacterial lipopolysaccharides may contribute to human health problems via exposure to mass

Toxic cyanobacteria are found worldwide both in inland and coastal water environments. Cyanobacteria occur in various environments including water, such as fresh and brackish water, oceans, hot springs, moist terrestrial environments such as soil, and in symbioses with plants, lichens and primitive animals. Some environmental conditions, including sunlight, warm weather, low turbulence and high nutrient levels, can promote growth. A high density of suspended cells may lead to the formation of surface scums and high toxin concentrations.

The toxins are not actively secreted to the surrounding water; most of the toxin is intracellular in growing cells. The release of toxin occurs during senescence of the cultures and when cultures shift from growth phase to stationary and death phases. Under field conditions, the majority of microcystin is intracellular during active growth of the cells [25]. There are reports of hepatotoxic blooms from all continents around the world [7]. Some of the highest reported cyanotoxin concentrations in bloom samples (measured by HPLC) have been 7300 μg/g dry weight microcystin in a *Microcystis* bloom from China [26], 18000 μg/g dry weight nodularin in a *Nodularia* bloom from the Baltic sea [27] and 5500 μg/g dry weight cylindrospermopsin from Australia [3]. Toxic and non-toxic strains from the same cyanobacterial species cannot be

where it has caused animal poisonings and later also in Denmark [22].

**3. Occurrence and levels of cyanobacteria and hepatotoxins**

detected also in temperate regions.

150 Environmental Biotechnology - New Approaches and Prospective Applications

shellfish [23].

occurrences of cyanobacteria.

Poisonings caused by cyanotoxins produced during heavy blooms have affected both humans and wild and domestic animals. Both hepatotoxic and neurotoxic poisonings have been associated with mass occurrences of cyanobacteria [7]. Many reported incidents of human health effects have involved inappropriate treatment of water supplies. The health risk caused by cyanotoxin exposure is difficult to quantify, since the actual exposure and resulting effects have not been conclusively determined. The most likely route for human exposure is the oral route via drinking water [37], and from recreational use of lakes and rivers [36].

Due to the growing concern about health effects of cyanotoxins especially via drinking water, WHO has adopted a provisional guideline value of 1.0 μg/L for MC-LR in 1998 [1]. The newest 4th edition to the drinking water guideline was published in 2011 [17]. Assessment of different water treatment procedures has shown that many of the treatment methods result in a reduction of cyanotoxin concentrations to below acutely toxic levels and below the WHO guideline value of 1.0 μg/L MC-LR in drinking water. During a cyanobacterial bloom the treatment procedures may however be insufficient, and also when different water treatment procedures are not used in combination. Therefore it is important to observe the water treatment efficiency during cyanobacterial blooms.

methods are sometimes considered too expensive to exclusively remove a contaminant that is

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511 153

Coagulation or flocculation involves the aggregation of smaller particles into larger particles using chemicals, such as ferric chloride or aluminium sulphate. Coagulation can be an efficient method for eliminating cyanobacterial cells from water, but soluble cyanotoxins are not very efficiently removed by this method [42]. Coagulation may also cause additional problems such as lysis of cyanobacterial cells leading to release of toxins. The activated carbon approach uses either powdered activated carbon, which can be added occasionally when there is a need, or granular activated carbon adsorbers, which are used continuously [43]. Both microcystins and cylindrospermopsin can be absorbed by activated carbon [43]. The disposal of the carbon

containing cyanobacterial toxins may present a challenge for this type of treatment.

Rapid filtration is a method usually used after a coagulation step in conventional water treatment, but does not effectively remove cyanobacterial cells from water. Conventional water treatment requires regular backwashing of the filters, but if the washing process is inade‐ quately performed, lysis of cyanobacterial cells on the filters can lead to release of toxins into the water [7]. Two types of membrane filtration, microfiltration and ultrafiltration, are commonly used to remove contaminants from drinking water. Both microfiltration and ultrafiltration have been shown to be effective in removal of intact cyanobacterial cells [44].

The most common chemical oxidants used in drinking water treatment are ozone, hydroxyl radicals, chlorine, chlorine dioxide, chloramine and permanganate. Chlorination and ozona‐ tion are effective for the removal of microcystins [43]. However, there are concerns regarding the release of toxin when cyanobacteria are chlorinated and with the formation of undesirable chlorination by-products [45]. Ozonation has been shown to be a very effective method for destroying microcystins and nodularins. In recent years, many water treatment plants have

Removal and inactivation of cyanobacteria and intracellular and extracellular cyanotoxins most often requires a combination of treatment processes or a multiple barrier approach. Furthermore, biological treatment of water is a method used for cyanotoxin-removal from drinking water. Biologically active filtration in the form of river bank filtration and both slow and rapid filtration have been reported to remove or to inactivate microcystins in drinking

Biodegradation is a chemical disruption of organic materials by microorganisms or other biological agents. Microbial degradation of chemicals in the environment is an important route for the removal of these compounds. Biodegradation is also one of the essential processes for the reduction of microcystins in natural eutrophic lakes and reservoirs. Cyanotoxin-degrading bacteria are distributed all over the world. Of all the cyanotoxin-biodegradation studies, most have focused on microcystins as a consequence of their biodegradability in drinking water

water (e.g. [47,48]) and are discussed more in detail in the following section.

included a two-stage ozonation treatment [46].

**6. Biodegradation of cyanotoxins**

irregularly occurring.

### **5. Treatment of drinking water containing cyanotoxins**

Water is an essential natural resource, necessary for drinking, agriculture and industrial activities. Contamination of water can therefore influence humans, agricultural livestock and irrigated field crops, as well as wildlife drinking the water or living in the aquatic environment. Drinking water should be pure enough to be consumed or used with low risk of immediate or long term harm. In large parts of the world, the population has inadequate access to safe potable water and use sources contaminated with disease vectors, pathogens or unacceptable levels of toxins and other harmful substances.

Prevention of bloom formation is naturally the most efficient method for avoiding cyanobac‐ terial toxin contamination of drinking water. Cyanotoxins are produced within the cyanobac‐ terial cells and thus toxin removal involves procedures to destroy or avoid the cells. Cyanotoxins are also water soluble and therefore chemical or biological procedures reducing the toxicity or completely removing the toxins from the drinking water are needed. If high extracellular toxin concentrations are present in the raw water, problems will occur for drinking water treatment plants. Under natural circumstances high toxin concentrations appear during the breakdown of a cyanobacterial bloom. Cyanobacterial cells are also lysed in the presence of chemicals, such as potassium permanganate or chlorine [38].

In cyanotoxin-removal from drinking water there is a need for knowledge of the physical and chemical properties of the toxin, such as the hydrophobicity, molecular size, and functional groups, the nature of the toxin, i.e., intracellular or extracellular, cyanobacterial growth and bloom patterns, and effective treatment processes [7]. However, these treatments may not be sufficient during cyanobacterial blooms or when a high organic load is present, and toxin levels should therefore be monitored during all steps of water treatment processes. Some of the existing methods of drinking water treatment are shortly described in the following section.

### **5.1. Water treatment processes**

Most drinking water plants use conventional treatment methods that are unable to yield complete removal of microcystins or are too expensive [39]. Conventional surface drinking water treatment utilises coagulation, flocculation, sedimentation, filtration and disinfection as basic methods. However, conventional treatment may need to be optimised for cyanotoxinremoval, relating to the form of the toxin to be removed (intra- or extracellular), the background water matrix, and possible dissolved toxin release during the treatment process [40]. Alterna‐ tive processes, such as granular activated carbon, powdered activated carbon, and membrane filtration have been proven efficient for the removal of microcystins [41]. However, these methods are sometimes considered too expensive to exclusively remove a contaminant that is irregularly occurring.

Coagulation or flocculation involves the aggregation of smaller particles into larger particles using chemicals, such as ferric chloride or aluminium sulphate. Coagulation can be an efficient method for eliminating cyanobacterial cells from water, but soluble cyanotoxins are not very efficiently removed by this method [42]. Coagulation may also cause additional problems such as lysis of cyanobacterial cells leading to release of toxins. The activated carbon approach uses either powdered activated carbon, which can be added occasionally when there is a need, or granular activated carbon adsorbers, which are used continuously [43]. Both microcystins and cylindrospermopsin can be absorbed by activated carbon [43]. The disposal of the carbon containing cyanobacterial toxins may present a challenge for this type of treatment.

Rapid filtration is a method usually used after a coagulation step in conventional water treatment, but does not effectively remove cyanobacterial cells from water. Conventional water treatment requires regular backwashing of the filters, but if the washing process is inade‐ quately performed, lysis of cyanobacterial cells on the filters can lead to release of toxins into the water [7]. Two types of membrane filtration, microfiltration and ultrafiltration, are commonly used to remove contaminants from drinking water. Both microfiltration and ultrafiltration have been shown to be effective in removal of intact cyanobacterial cells [44].

The most common chemical oxidants used in drinking water treatment are ozone, hydroxyl radicals, chlorine, chlorine dioxide, chloramine and permanganate. Chlorination and ozona‐ tion are effective for the removal of microcystins [43]. However, there are concerns regarding the release of toxin when cyanobacteria are chlorinated and with the formation of undesirable chlorination by-products [45]. Ozonation has been shown to be a very effective method for destroying microcystins and nodularins. In recent years, many water treatment plants have included a two-stage ozonation treatment [46].

Removal and inactivation of cyanobacteria and intracellular and extracellular cyanotoxins most often requires a combination of treatment processes or a multiple barrier approach. Furthermore, biological treatment of water is a method used for cyanotoxin-removal from drinking water. Biologically active filtration in the form of river bank filtration and both slow and rapid filtration have been reported to remove or to inactivate microcystins in drinking water (e.g. [47,48]) and are discussed more in detail in the following section.

### **6. Biodegradation of cyanotoxins**

reduction of cyanotoxin concentrations to below acutely toxic levels and below the WHO guideline value of 1.0 μg/L MC-LR in drinking water. During a cyanobacterial bloom the treatment procedures may however be insufficient, and also when different water treatment procedures are not used in combination. Therefore it is important to observe the water

Water is an essential natural resource, necessary for drinking, agriculture and industrial activities. Contamination of water can therefore influence humans, agricultural livestock and irrigated field crops, as well as wildlife drinking the water or living in the aquatic environment. Drinking water should be pure enough to be consumed or used with low risk of immediate or long term harm. In large parts of the world, the population has inadequate access to safe potable water and use sources contaminated with disease vectors, pathogens or unacceptable

Prevention of bloom formation is naturally the most efficient method for avoiding cyanobac‐ terial toxin contamination of drinking water. Cyanotoxins are produced within the cyanobac‐ terial cells and thus toxin removal involves procedures to destroy or avoid the cells. Cyanotoxins are also water soluble and therefore chemical or biological procedures reducing the toxicity or completely removing the toxins from the drinking water are needed. If high extracellular toxin concentrations are present in the raw water, problems will occur for drinking water treatment plants. Under natural circumstances high toxin concentrations appear during the breakdown of a cyanobacterial bloom. Cyanobacterial cells are also lysed

In cyanotoxin-removal from drinking water there is a need for knowledge of the physical and chemical properties of the toxin, such as the hydrophobicity, molecular size, and functional groups, the nature of the toxin, i.e., intracellular or extracellular, cyanobacterial growth and bloom patterns, and effective treatment processes [7]. However, these treatments may not be sufficient during cyanobacterial blooms or when a high organic load is present, and toxin levels should therefore be monitored during all steps of water treatment processes. Some of the existing methods of drinking water treatment are shortly described in the following section.

Most drinking water plants use conventional treatment methods that are unable to yield complete removal of microcystins or are too expensive [39]. Conventional surface drinking water treatment utilises coagulation, flocculation, sedimentation, filtration and disinfection as basic methods. However, conventional treatment may need to be optimised for cyanotoxinremoval, relating to the form of the toxin to be removed (intra- or extracellular), the background water matrix, and possible dissolved toxin release during the treatment process [40]. Alterna‐ tive processes, such as granular activated carbon, powdered activated carbon, and membrane filtration have been proven efficient for the removal of microcystins [41]. However, these

in the presence of chemicals, such as potassium permanganate or chlorine [38].

treatment efficiency during cyanobacterial blooms.

152 Environmental Biotechnology - New Approaches and Prospective Applications

levels of toxins and other harmful substances.

**5.1. Water treatment processes**

**5. Treatment of drinking water containing cyanotoxins**

Biodegradation is a chemical disruption of organic materials by microorganisms or other biological agents. Microbial degradation of chemicals in the environment is an important route for the removal of these compounds. Biodegradation is also one of the essential processes for the reduction of microcystins in natural eutrophic lakes and reservoirs. Cyanotoxin-degrading bacteria are distributed all over the world. Of all the cyanotoxin-biodegradation studies, most have focused on microcystins as a consequence of their biodegradability in drinking water sources. This section mainly describes biodegradation studies of microcystins, but studies on nodularin, cylindrospermopsin, saxitoxins and anatoxin-a have also been performed to some extent.

gen. nov.,sp. nov.) was proposed. A bacterium isolated from water samples in Brazil showed high homology with the *Burkholderia* genus, belonging to the beta subdivision of proteobacteria [71], which was the first reported bacterium from the genus *Burkholderia* as a cyanobacterial

**Bacterial strain Degradable toxins Reference** *Arthrobacter* sp. MC-LR [72] *Bacillus* sp. strain EMB MC-LR, MC-RR [74] *Brevibacterium* sp. MC-LR [72] *Burkholderia* sp. MC-LR, [D-Leu1]MC-LR [71]

*Methylobacillus* sp. strain J10 MC-LR, MC-RR [75] *Microbacterium* sp. MC-LR [78] *Morganella morganii* MC-LR [77] *Paucibacter toxinivorans* sp. nov. MC-LR, MC-YR, NOD [69] *Poterioochromonas sp.* MC-LR [81] *Pseudomonas aeruginosa* MC-LR [68] *Rhizobium gallicum* MC-LR [78] *Rhodococcus* sp. MC-LR [72] *Sphingomona stygia* MC-LR, MC-RR, MC-YR [65] *Sphingomonas* sp. 7CY MC-LR, MC-RR, MC-LY, MC-LW, MC-LF [66] *Sphingomonas* sp. ACM-3962 MC-LR, MC-RR [25,63,82]

*Sphingomonas* sp. B9 MC-LR, MC-RR, 3-dmMC-LR, dhMC-LR, MC-LR-Cys, NOD

*Sphingomonas* sp. CBA4 MC-RR [57] *Sphingomonas* sp. MD-1 MC-LR, MC-RR, MC-YR [56] *Sphingomonas* sp. MDB2 MCs [70] *Sphingomonas* sp. MDB3 MCs [70] *Sphingomonas* sp. MJ-PV MC-LR [49] *Sphingomonas* sp. Y2 MC-LR, MC-RR, MC-YR, 6(Z)-Adda-MC-LR [64,70,84] *Sphingopyxis* sp. C-1 MC-LR [58] *Sphingopyxis* sp. LH21 MC-LR, MC-LA [52] *Sphingopyxis* sp. USTB-05 MC-RR [85] *Stenotrophomonas* sp. strain EMS MC-LR, MC-RR [76]

MC-LR, MC-RR, MC-YR, MC-LF, MC-LY, MC-LW [79,80]

MCs [47]

[67,83]

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511 155

toxin degrader.

*Lactobacillus rhamnosus* GG and LC-705,

17 different strains (Gram-negative,

**Table 1.** Reported microcystin-degrading bacteria

*Proteobacteria*)

*Bifidobacterium longum* 46

People are frequently exposed to cyanobacterial toxins as well as other microbial contaminants through drinking water. Conventional water treatment procedures discussed in the previous section are in some cases insufficient in the removal of cyanobacterial toxins from drinking water, especially during cyanobacterial blooms. If the cyanobacterial cells are not removed by traditional water treatment methods, the cells and therefore the toxins remain in the drinking water and must be degraded to non-toxic compounds. Since microcystins have been released into the water body, the toxins can persist for weeks [20] before they are adequately degraded by for example bacteria.

### **6.1. Bacterial degradation of microcystins**

Different biological methods have been applied to remove cyanobacteria and their toxins. One type of these methods is the use of microorganisms or biofilms capable of degrading microcys‐ tins. Biological treatment for removal of toxin contaminants is becoming more useful as toxins can be removed without the addition of chemicals that may have the potential to produce undesira‐ ble by-products. Biodegradation of microcystins in water has been proven to be very effective as they can be used a as carbon source by heterotrophic bacteria [25,38,49,50]. Methods utilizing microcystin-degrading microorganisms can be classified into two groups. One is the use of biofilms grown on the surface of substrates within bioreactors, such as biological sand [48,51,52], biofilm-reactors based on immobilised microorganisms [53], biological treatment facilities combined with conventional treatment processes [54], and granular activated carbon filters [55]. The other group depends on specific microorganisms efficient in microcystin-degradation, such as bacteria of the *Sphingomonas* sp. [56,57] and *Sphingopyxis* sp. [58].

Different variants of microcystins have been demonstrated to be degraded after incubation with water from a lake in Japan, which is frequently contaminated with cyanobacteria [59]. A more effective degradation was observed after adding bed sediment or mud from the lake. Christoffers‐ en *et al.* found out that bacteria can efficiently degrade microcystins in natural waters with previous cyanobacterial contamination and that the degradation process is rapid and without lag phase [60].

Many other studies have also reported biological degradation of microcystin in natural waters from lakes and reservoirs, particularly those containing toxic cyanobacterial blooms [25,50,61,62]. Several strains of the genus *Sphingomonas* have been reported to degrade microcystins [49,57,63–67]. Table 1 lists strains reported to degrade different variants of microcystins and nodularin. A part of the recognised microcystin-degraders so far belonging to the family *Sphingomonadaceae* are closely related and possess homologues of the mlrA gene. Seventeen strains of Gram-negative bacteria with the ability to degrade microcystins were isolated by Lahti *et al*. [47]. Other reported microcystin-degrading bacteria include *Pseudomonas aeruginosa* [68], *Paucibacter toxinivorans* [69] and *Sphingosinicella microcystinivorans* [70]. In a study of Rapala *et al*. thirteen bacteria capable of degrading microcystins and nodularin were isolated from lake sediment [61]. Genomic characterisation of these strains indicated that they formed a single microdiverse species and a novel genus and species (*Paucibacter toxinivorans*

gen. nov.,sp. nov.) was proposed. A bacterium isolated from water samples in Brazil showed high homology with the *Burkholderia* genus, belonging to the beta subdivision of proteobacteria [71], which was the first reported bacterium from the genus *Burkholderia* as a cyanobacterial toxin degrader.


**Table 1.** Reported microcystin-degrading bacteria

sources. This section mainly describes biodegradation studies of microcystins, but studies on nodularin, cylindrospermopsin, saxitoxins and anatoxin-a have also been performed to some

People are frequently exposed to cyanobacterial toxins as well as other microbial contaminants through drinking water. Conventional water treatment procedures discussed in the previous section are in some cases insufficient in the removal of cyanobacterial toxins from drinking water, especially during cyanobacterial blooms. If the cyanobacterial cells are not removed by traditional water treatment methods, the cells and therefore the toxins remain in the drinking water and must be degraded to non-toxic compounds. Since microcystins have been released into the water body, the toxins can persist for weeks [20] before they are adequately degraded

Different biological methods have been applied to remove cyanobacteria and their toxins. One type of these methods is the use of microorganisms or biofilms capable of degrading microcys‐ tins. Biological treatment for removal of toxin contaminants is becoming more useful as toxins can be removed without the addition of chemicals that may have the potential to produce undesira‐ ble by-products. Biodegradation of microcystins in water has been proven to be very effective as they can be used a as carbon source by heterotrophic bacteria [25,38,49,50]. Methods utilizing microcystin-degrading microorganisms can be classified into two groups. One is the use of biofilms grown on the surface of substrates within bioreactors, such as biological sand [48,51,52], biofilm-reactors based on immobilised microorganisms [53], biological treatment facilities combined with conventional treatment processes [54], and granular activated carbon filters [55]. The other group depends on specific microorganisms efficient in microcystin-degradation, such

Different variants of microcystins have been demonstrated to be degraded after incubation with water from a lake in Japan, which is frequently contaminated with cyanobacteria [59]. A more effective degradation was observed after adding bed sediment or mud from the lake. Christoffers‐ en *et al.* found out that bacteria can efficiently degrade microcystins in natural waters with previous cyanobacterial contamination and that the degradation process is rapid and without lag phase [60]. Many other studies have also reported biological degradation of microcystin in natural waters from lakes and reservoirs, particularly those containing toxic cyanobacterial blooms [25,50,61,62]. Several strains of the genus *Sphingomonas* have been reported to degrade microcystins [49,57,63–67]. Table 1 lists strains reported to degrade different variants of microcystins and nodularin. A part of the recognised microcystin-degraders so far belonging to the family *Sphingomonadaceae* are closely related and possess homologues of the mlrA gene. Seventeen strains of Gram-negative bacteria with the ability to degrade microcystins were isolated by Lahti *et al*. [47]. Other reported microcystin-degrading bacteria include *Pseudomonas aeruginosa* [68], *Paucibacter toxinivorans* [69] and *Sphingosinicella microcystinivorans* [70]. In a study of Rapala *et al*. thirteen bacteria capable of degrading microcystins and nodularin were isolated from lake sediment [61]. Genomic characterisation of these strains indicated that they formed a single microdiverse species and a novel genus and species (*Paucibacter toxinivorans*

extent.

by for example bacteria.

**6.1. Bacterial degradation of microcystins**

154 Environmental Biotechnology - New Approaches and Prospective Applications

as bacteria of the *Sphingomonas* sp. [56,57] and *Sphingopyxis* sp. [58].

Recently, Gram-positive bacteria isolated from freshwater, belonging to *Actinobacteria* and identified as *Arthrobacter* sp., *Brevibacterium* sp. and *Rhodococcus* sp., were shown to remove MC-LR [72]. The mechanism of MC-LR removal for *Rhodococcus* sp. C1 [73] was shown to be similar to the previously reported degradation pathway for *Sphingomonas* by Bourne *et al*. [63]. A new strain AMRI-03 with close relationship to the genus *Bacillus* was isolated from a Saudi freshwater lake [74]. Another strain J10 isolated from Lake Taihu in China was identified as *Methylobacillus* sp. [75]. An EMS strain similar to *Stenotrophomonas maltophilia* was described by Chen *et al*. and was the first report of microcystin-degrading bacteria carrying the mlrA gene in the genus of the gamma division of proteobacteria [76]. Other reported examples of bacteria with such ability are *Morganella morganii* and *Pseudomonas* sp. [77]. Further recent findings include two isolates from Lake Okeechobee, Florida, capable of microcystin-degra‐ dation and classified as *Rhizobium gallicum* and *Microbacterium* sp. [78].

isolated from Japanese lakes [56]. More recently, Hoefel *et al*. designed and optimised a quantitative real-time polymerase chain reaction assay for the detection of the mlrA gene [88].

Various studies have designed qualitative polymerase chain reaction assays for detection of mlrA [51,52,56]. Saito *et al.* reported gene homologues of mlrA in two microcystin-degrading bacteria, *Sphingomonas* sp. MD-1 and *Sphingomonas* sp. Y2, both of which were previously isolated from Japanese lakes [56]. More recently, Hoefel *et al*. designed and optimised a quantitative real-time

22 **Fig. 2.** MC-LR degradation pathway by *Sphingomonas* sp. ACM-3962; mlrA-C: microcystinases A-

**Figure 2.** MC-LR degradation pathway by *Sphingomonas* sp. ACM-3962; mlrA-C: microcystinases A-C (modified from

TETRAPEPTIDE (MW 614)

**mlrA mlrB mlrC**

Before biological treatment can be considered a feasible option for effective removal of microcystins, there is a need to determine if any toxic biodegradation by-products are generated. Different studies have demonstrated that the biodegradation of microcystins does not yield toxic by-products. Bourne *et al*. [63] and Harada *et al*. [67] identified two intermediate products from the bacterial degradation of MC-LR by *Sphingomonas* sp. ACM-3962 and *Sphingomonas* sp. B9, respectively. Both studies identified linearized microcystin-LR and a tetrapeptide as the intermediate products, and isolating Adda as one of the final degradation products (Figure 2). Both these intermediate products were less active than the parent microcystin-LR. Studies with *Sphingpoyxis* sp. LH21 in treated reservoir water concluded that the decrease in cytotoxicity indicated that no cytotoxic by-products of microcystins

Before biological treatment can be considered a feasible option for effective removal of microcystins, there is a need to determine if any toxic biodegradation by-products are generated. Different studies have demonstrated that the biodegradation of microcystins does not yield toxic by-products. Bourne *et al*. [63] and Harada *et al*. [67] identified two intermediate products from the bacterial degradation of MC-LR by *Sphingomonas* sp. ACM-3962 and *Sphingomonas* sp. B9, respectively. Both studies identified linearized MC-LR and a tetrapeptide as the intermediate products, and isolated Adda as one of the final degradation products (Figure 2). Both these intermediate products were less active than the parent MC-LR. Studies with *Sphingpoyxis* sp. LH21 in treated reservoir water concluded that the decrease in cytotox‐ icity indicated that no cytotoxic by-products of microcystins were being generated [52].

Different factors may influence the biodegradation efficiency, such as water temperature. Published results suggest that the temperature range for the effective biodegradation of microcystins is between 11 and 37 °C, with more rapid degradation at the higher temperatures in most cases [52,64,88]. In addition, the bacterial composition and cell density within the water body also affects degradation;

40 Only few studies with respect to the biodegradation of a range of cyanobacterial metabolites in water 41 bodies have been performed. This is relevant since multiple classes of cyanobacterial metabolites are

Only few studies with respect to the biodegradation of a range of cyanobacterial metabolites in water bodies have been performed. This is relevant since multiple classes of cyanobacterial metabolites are often simultaneously present in water bodies. The following sections regarding the removal of cyanotoxins by probiotic bacteria will assess this issue, and results regarding the removal of a range of cyanotoxins are presented. The results on a range of bacterial species demonstrate the feasibility of biodegradation as a possible removal option for microcystins. The most important practical use of microbial aggregates, such as biological filters and biofilm, is in biological wastewater treatment, and some new technologies already utilize bacterial

Different factors may influence the biodegradation efficiency, such as water temperature. Published results suggest that the temperature range for the effective biodegradation of microcystins is between 11 and 37 °C, with more rapid degradation at the higher temper‐ atures in most cases [52,64,79,88]. In addition, the bacterial composition and cell density within the water body also affects degradation; both the types of organisms present and

products. The genes mlrA, mlrB and mlrC encode a 336-residue metalloendopeptidase (responsible for linearization of microcystins), a serine protease and a metalloprotease, respectively. Further studies have confirmed the existence of the mlr cluster components also in other microcystin-degrading bacteria; Ho *et al*. identified homologues of four mlr genes in *Sphingopyxis* sp. LH21 [52].

However, as has been recently indicated, mlrC acts not only on the tetrapeptide but is also able to hydrolyze linear microcystin without earlier processing by mlrB [86]. Other products of microcystin-degradation have consequently been documented, but the complete fate of microcystin-derivatives is still unknown [83,87]. Additionally, enzymes other than proteases have been suggested to be involved in microcystin-utilization, and besides typical proteolytic activity, also decarboxylation and

5 Similarly, a homologous gene cluster was also detected in *Sphingopyxis* sp. C-1 [58].

11 demethylation have been proposed as alternative mechanisms [87].

16 polymerase chain reaction assay for the detection of the mlrA gene [88].

24 **6.3. Further aspects of microcystin-biodegradation** 

**6.3. Further aspects of microcystin-biodegradation**

LINEARIZED MC-LR (MW 1012)

39 both the types of organisms present and their concentration.

9

157

SMALLER PEPTIDES AND AMINO ACIDS

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511

Biodegradation of cyanobacterial toxins

17

23 C (modified from [63]).

[63]).

MC-LR (MW 994)

34 were being generated [52].

their concentration.

aggregates for degradation [89].

### **6.2. Enzymatic mechanisms of microcystin-biodegradation**

The first proposal of microcystin-biodegradation suggested a proteolytic mechanism [63]. Within the genome of the first isolated microcystin-degrading bacterium, *Sphingomonas* sp. ACM-3962, Bourne *et al*. identified a gene cluster, mlrA, mlrB, mlrC and mlrD, responsible for the degradation of MC-LR [63,82]. Based on MS-analysis a linear MC-LR (protonated molec‐ ular ion at *m/z* 1013) and a tetrapeptide (protonated molecular ion at *m/z* 615) were recognised as the degradation products. The microcystin-degradation pathway was described as a linear, three-step process. It was suggested that the mlrA gene encoded an enzyme responsible for the hydrolytic cleaving of the cyclic structure of MC-LR (ring-opening at the Adda-Arg peptide bond). The resulting linear MC-LR molecule was then sequentially hydrolysed by peptidases encoded by the mlrB and mlrC genes to a tetrapeptide, and further to smaller peptides and amino acids (Figure 2). The final gene, mlrD, encoded for a possible transporter protein that may have allowed for active transport of microcystin or its degradation products. The genes mlrA, mlrB and mlrC encode a 336-residue metalloendopeptidase (responsible for lineariza‐ tion of microcystins), a serine protease and a metalloprotease, respectively. Further studies have confirmed the existence of the mlr cluster components also in other microcystin-degrad‐ ing bacteria; Ho *et al*. identified homologues of four mlr genes in *Sphingopyxis* sp. LH21 [52]. Similarly, a homologous gene cluster was also detected in *Sphingopyxis* sp. C-1 [58].

However, as has been recently indicated, mlrC acts not only on the tetrapeptide but is also able to hydrolyze linear microcystin without earlier processing by mlrB [86]. Other products of microcystin-degradation have consequently been documented, but the complete fate of microcystin-derivatives is still unknown [83,87]. Additionally, enzymes other than proteases have been suggested to be involved in microcystin-utilisation, and besides typical proteolytic activity, also decarboxylation and demethylation have been proposed as alternative mecha‐ nisms [87].

Various studies have designed qualitative polymerase chain reaction assays for detection of mlrA [51,52,56]. Saito *et al.* reported gene homologues of mlrA in two microcystin-degrading bacteria, *Sphingomonas* sp. MD-1 and *Sphingomonas* sp. Y2, both of which were previously

9

isolated from Japanese lakes [56]. More recently, Hoefel *et al*. designed and optimised a quantitative real-time polymerase chain reaction assay for the detection of the mlrA gene [88]. 13 [51,52,56]. Saito *et al.* reported gene homologues of mlrA in two microcystin-degrading bacteria, 14 *Sphingomonas* sp. MD-1 and *Sphingomonas* sp. Y2, both of which were previously isolated from 15 Japanese lakes [56]. More recently, Hoefel *et al*. designed and optimised a quantitative real-time

12 Various studies have designed qualitative polymerase chain reaction assays for detection of mlrA

products. The genes mlrA, mlrB and mlrC encode a 336-residue metalloendopeptidase (responsible for linearization of microcystins), a serine protease and a metalloprotease, respectively. Further studies have confirmed the existence of the mlr cluster components also in other microcystin-degrading bacteria; Ho *et al*. identified homologues of four mlr genes in *Sphingopyxis* sp. LH21 [52].

However, as has been recently indicated, mlrC acts not only on the tetrapeptide but is also able to hydrolyze linear microcystin without earlier processing by mlrB [86]. Other products of microcystin-degradation have consequently been documented, but the complete fate of microcystin-derivatives is still unknown [83,87]. Additionally, enzymes other than proteases have been suggested to be involved

5 Similarly, a homologous gene cluster was also detected in *Sphingopyxis* sp. C-1 [58].

16 polymerase chain reaction assay for the detection of the mlrA gene [88].

23 C (modified from [63]). 24 **6.3. Further aspects of microcystin-biodegradation Figure 2.** MC-LR degradation pathway by *Sphingomonas* sp. ACM-3962; mlrA-C: microcystinases A-C (modified from [63]).

22 **Fig. 2.** MC-LR degradation pathway by *Sphingomonas* sp. ACM-3962; mlrA-C: microcystinases A-

#### 25 Before biological treatment can be considered a feasible option for effective removal of microcystins, 26 there is a need to determine if any toxic biodegradation by-products are generated. Different studies **6.3. Further aspects of microcystin-biodegradation**

Biodegradation of cyanobacterial toxins

17

Recently, Gram-positive bacteria isolated from freshwater, belonging to *Actinobacteria* and identified as *Arthrobacter* sp., *Brevibacterium* sp. and *Rhodococcus* sp., were shown to remove MC-LR [72]. The mechanism of MC-LR removal for *Rhodococcus* sp. C1 [73] was shown to be similar to the previously reported degradation pathway for *Sphingomonas* by Bourne *et al*. [63]. A new strain AMRI-03 with close relationship to the genus *Bacillus* was isolated from a Saudi freshwater lake [74]. Another strain J10 isolated from Lake Taihu in China was identified as *Methylobacillus* sp. [75]. An EMS strain similar to *Stenotrophomonas maltophilia* was described by Chen *et al*. and was the first report of microcystin-degrading bacteria carrying the mlrA gene in the genus of the gamma division of proteobacteria [76]. Other reported examples of bacteria with such ability are *Morganella morganii* and *Pseudomonas* sp. [77]. Further recent findings include two isolates from Lake Okeechobee, Florida, capable of microcystin-degra‐

The first proposal of microcystin-biodegradation suggested a proteolytic mechanism [63]. Within the genome of the first isolated microcystin-degrading bacterium, *Sphingomonas* sp. ACM-3962, Bourne *et al*. identified a gene cluster, mlrA, mlrB, mlrC and mlrD, responsible for the degradation of MC-LR [63,82]. Based on MS-analysis a linear MC-LR (protonated molec‐ ular ion at *m/z* 1013) and a tetrapeptide (protonated molecular ion at *m/z* 615) were recognised as the degradation products. The microcystin-degradation pathway was described as a linear, three-step process. It was suggested that the mlrA gene encoded an enzyme responsible for the hydrolytic cleaving of the cyclic structure of MC-LR (ring-opening at the Adda-Arg peptide bond). The resulting linear MC-LR molecule was then sequentially hydrolysed by peptidases encoded by the mlrB and mlrC genes to a tetrapeptide, and further to smaller peptides and amino acids (Figure 2). The final gene, mlrD, encoded for a possible transporter protein that may have allowed for active transport of microcystin or its degradation products. The genes mlrA, mlrB and mlrC encode a 336-residue metalloendopeptidase (responsible for lineariza‐ tion of microcystins), a serine protease and a metalloprotease, respectively. Further studies have confirmed the existence of the mlr cluster components also in other microcystin-degrad‐ ing bacteria; Ho *et al*. identified homologues of four mlr genes in *Sphingopyxis* sp. LH21 [52].

Similarly, a homologous gene cluster was also detected in *Sphingopyxis* sp. C-1 [58].

However, as has been recently indicated, mlrC acts not only on the tetrapeptide but is also able to hydrolyze linear microcystin without earlier processing by mlrB [86]. Other products of microcystin-degradation have consequently been documented, but the complete fate of microcystin-derivatives is still unknown [83,87]. Additionally, enzymes other than proteases have been suggested to be involved in microcystin-utilisation, and besides typical proteolytic activity, also decarboxylation and demethylation have been proposed as alternative mecha‐

Various studies have designed qualitative polymerase chain reaction assays for detection of mlrA [51,52,56]. Saito *et al.* reported gene homologues of mlrA in two microcystin-degrading bacteria, *Sphingomonas* sp. MD-1 and *Sphingomonas* sp. Y2, both of which were previously

dation and classified as *Rhizobium gallicum* and *Microbacterium* sp. [78].

**6.2. Enzymatic mechanisms of microcystin-biodegradation**

156 Environmental Biotechnology - New Approaches and Prospective Applications

nisms [87].

27 have demonstrated that the biodegradation of microcystins does not yield toxic by-products. Bourne 28 *et al*. [63] and Harada *et al*. [67] identified two intermediate products from the bacterial degradation 29 of MC-LR by *Sphingomonas* sp. ACM-3962 and *Sphingomonas* sp. B9, respectively. Both studies 30 identified linearized microcystin-LR and a tetrapeptide as the intermediate products, and isolating 31 Adda as one of the final degradation products (Figure 2). Both these intermediate products were less 32 active than the parent microcystin-LR. Studies with *Sphingpoyxis* sp. LH21 in treated reservoir water 33 concluded that the decrease in cytotoxicity indicated that no cytotoxic by-products of microcystins 34 were being generated [52]. 35 Different factors may influence the biodegradation efficiency, such as water temperature. Published 36 results suggest that the temperature range for the effective biodegradation of microcystins is between 37 11 and 37 °C, with more rapid degradation at the higher temperatures in most cases [52,64,88]. In Before biological treatment can be considered a feasible option for effective removal of microcystins, there is a need to determine if any toxic biodegradation by-products are generated. Different studies have demonstrated that the biodegradation of microcystins does not yield toxic by-products. Bourne *et al*. [63] and Harada *et al*. [67] identified two intermediate products from the bacterial degradation of MC-LR by *Sphingomonas* sp. ACM-3962 and *Sphingomonas* sp. B9, respectively. Both studies identified linearized MC-LR and a tetrapeptide as the intermediate products, and isolated Adda as one of the final degradation products (Figure 2). Both these intermediate products were less active than the parent MC-LR. Studies with *Sphingpoyxis* sp. LH21 in treated reservoir water concluded that the decrease in cytotox‐ icity indicated that no cytotoxic by-products of microcystins were being generated [52].

38 addition, the bacterial composition and cell density within the water body also affects degradation; 39 both the types of organisms present and their concentration. 40 Only few studies with respect to the biodegradation of a range of cyanobacterial metabolites in water 41 bodies have been performed. This is relevant since multiple classes of cyanobacterial metabolites are Different factors may influence the biodegradation efficiency, such as water temperature. Published results suggest that the temperature range for the effective biodegradation of microcystins is between 11 and 37 °C, with more rapid degradation at the higher temper‐ atures in most cases [52,64,79,88]. In addition, the bacterial composition and cell density within the water body also affects degradation; both the types of organisms present and their concentration.

Only few studies with respect to the biodegradation of a range of cyanobacterial metabolites in water bodies have been performed. This is relevant since multiple classes of cyanobacterial metabolites are often simultaneously present in water bodies. The following sections regarding the removal of cyanotoxins by probiotic bacteria will assess this issue, and results regarding the removal of a range of cyanotoxins are presented. The results on a range of bacterial species demonstrate the feasibility of biodegradation as a possible removal option for microcystins. The most important practical use of microbial aggregates, such as biological filters and biofilm, is in biological wastewater treatment, and some new technologies already utilize bacterial aggregates for degradation [89].

### **7. Probiotic bacteria involved in cyanotoxin-removal**

Probiotics were earlier defined as "live microbial food supplements which beneficially affect the host either directly or indirectly by improving its intestinal microbial balance" [90]. Today, the most commonly accepted definition by WHO states that probiotics are "live microbial food supplements which, when given in adequate amounts have a demonstrated beneficial effect on human health" [91]. In order to be effective the probiotic micro-organisms must be able to survive the digestive conditions, including bile acids, and they must be able to colonise the gastrointestinal tract at least temporarily without any harm to the host [92]. Only certain strains of micro-organisms have these properties. Most probiotic micro-organisms are grouped in two bacterial genera, *Lactobacillus* (*L.*) and *Bifidobacterium* (*B.*).

the effect of pH, temperature, toxin concentration, bacterial cell density and cell viability, had

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511 159

The removal of MC-LR was shown to be temperature dependent, with the highest removal observed at 37 °C for all studied strains. At 4 °C, practically no removal of MC-LR could be observed and the removal percentages increased with increasing temperature [79]. This can be explained by the fact that at 4 °C, the bacterial cells are metabolically inactive, but at 22 and 37 °C, the bacteria become metabolically active, which is required for enzymatic activity. In addition, the role of glucose in activating the metabolism of the probiotic bacteria was assessed [96]. Since it was shown that viability is a requirement for efficient toxin removal, glucose was added as a source of nutrient to the bacterial solutions to enhance the bacterial viability. Glucose addition improved the removal efficiencies of all tested strains by enhancing both the removal rate and the amount of MC-LR removed after 24 hours of incubation [96]. Supple‐ mentation of glucose provides energy to the bacteria, and thereby, the microcystin-removal

To investigate the role of the probiotic bacterial cell density, a range of bacterial cell densities were screened and tested for their microcystin-removal efficiencies [79]. The removal of MC-LR was shown to be dependent on the bacterial cell density, with a minimum of approximately 109 CFU/mL required for significant MC-LR removal [79]. The removal of MC-LR was further enhanced with increasing bacterial cell density. To assess whether a combination of several probiotic strains could enhance their microcystinremoval efficiencies the microcystin-removal of three probiotic strains (*L. rhamnosus* GG, *L. rhamnosus* LC-705 and *B. longum* 46) separately and in combination was studied [80]. With the probiotic mixture, microcystin-removal percentages of up to 90% could be observed and the results showed that the removal efficiency was improved with a mixture of the

In addition to MC-LR, probiotic bacterial strains were also incubated with other microcystins, including MC-RR, -YR, -LY, -LW and -LF. The results of the study show that probiotic strains were effective in the elimination of several different microcystins from solution [79]. Simulta‐ neous removal of several toxins present in cyanobacterial extracts was also investigated. The time course for the removal of microcystins present in the cyanobacterial extracts *Microcystis* NIES-107 and *Microcystis* PCC 7820 by the probiotic strain *L. rhamnosus* GG is shown in Figures 3 a and b, respectively. The removal of all studied microcystins increased over time. The removal of the microcystins present in *Microcystis* NIES-107 after 24 hours of incubation was around 65–85% of total microcystin and for microcystins present in *Microcystis* PCC 7820 around 60–80%. The toxin-removal was thus shown to be efficient also when several different microcystins were present in the solution. This indicates that there is no competition taking place among the toxins during incubation with probiotic bacteria. In addition, the strains were shown to remove the cytotoxin cylindrospermopsin from aqueous solutions; the removal was

Probiotic bacteria have several advantages in comparison with the previously reported microcystin-degrading bacteria, as they have been classified as food grade, safe bacteria by the European Food Safety Authority (EFSA) [95]. Therefore probiotic bacteria can safely be

an effect on the removal efficiency [80].

efficiencies also increase.

strains and compared to the individual strains [80].

somewhat less efficient, around 30% for all tested strains [80].

The main site of action for the health benefits of probiotic bacteria is the gut. The intestinal mucosa forms a barrier between the external and internal environment of the human body. There are several important modes of action for probiotic bacteria, including modification of gut pH, colonisation ability, inhibition of the colonisation, adhesion and invasion of pathogens, direct antimicrobial effect, replacement of already adhered pathogens, competing for available nutrients and growth factors, regulation of the immune system of the host, normalisation of the gut microbiota, and different metabolic effects (reviewed in [93,94]). It is therefore believed that by adding these bacteria as probiotics to the diet, the normal microbiota can be altered. Many probiotic organisms originate in fermented foods, and they have a long history of safe use in human consumption. Lactobacilli and bifidobacteria common in the food industry belong to the European Qualified Presumption of Safety (QPS) status organisms, which can be used in foods and feeds [95].

### **7.1. Efficiency of probiotic strains in microcystin-removal**

Recently published studies have reported efficient cyanotoxin-removal by several strains of probiotic bacteria [79,80,96,97]. The aim of these studies was to characterise the potential of probiotic lactic acid bacteria and bifidobacteria in removal of microcystins and cylindrosper‐ mopsin from aqueous solutions. Different physiological conditions possibly affecting the removal efficiency were studied and the mechanism of toxin removal was investigated.

In an initial screening study, 15 different strains of probiotic lactic acid bacteria and bifido‐ bacteria were tested for their MC-LR removal capacities and evaluated for their potential in water decontamination [79]. The results showed a reproducible reduction of MC-LR in solution by the majority of the tested bacterial strains; the most efficient removal was achieved with *L. rhamnosus* strains GG and LC-705, *B. lactis* strains 420 and Bb12 and *B. longum* 46 [79]. The removal of MC-LR continued during the entire 24-hour incubation, which indicates that the removal process is quite slow. The effect of pH during incubation was also studied. pH was found to have an influence on toxin removal, with a higher removal percentage observed at neutral pH than at pH 3 [79]. It was also shown that viable bacteria were more efficient in microcystin-removal than non-viable bacteria [80]. Further studies showed that several strains were efficient in microcystin-removal and that different physiological conditions, including the effect of pH, temperature, toxin concentration, bacterial cell density and cell viability, had an effect on the removal efficiency [80].

**7. Probiotic bacteria involved in cyanotoxin-removal**

158 Environmental Biotechnology - New Approaches and Prospective Applications

bacterial genera, *Lactobacillus* (*L.*) and *Bifidobacterium* (*B.*).

**7.1. Efficiency of probiotic strains in microcystin-removal**

be used in foods and feeds [95].

Probiotics were earlier defined as "live microbial food supplements which beneficially affect the host either directly or indirectly by improving its intestinal microbial balance" [90]. Today, the most commonly accepted definition by WHO states that probiotics are "live microbial food supplements which, when given in adequate amounts have a demonstrated beneficial effect on human health" [91]. In order to be effective the probiotic micro-organisms must be able to survive the digestive conditions, including bile acids, and they must be able to colonise the gastrointestinal tract at least temporarily without any harm to the host [92]. Only certain strains of micro-organisms have these properties. Most probiotic micro-organisms are grouped in two

The main site of action for the health benefits of probiotic bacteria is the gut. The intestinal mucosa forms a barrier between the external and internal environment of the human body. There are several important modes of action for probiotic bacteria, including modification of gut pH, colonisation ability, inhibition of the colonisation, adhesion and invasion of pathogens, direct antimicrobial effect, replacement of already adhered pathogens, competing for available nutrients and growth factors, regulation of the immune system of the host, normalisation of the gut microbiota, and different metabolic effects (reviewed in [93,94]). It is therefore believed that by adding these bacteria as probiotics to the diet, the normal microbiota can be altered. Many probiotic organisms originate in fermented foods, and they have a long history of safe use in human consumption. Lactobacilli and bifidobacteria common in the food industry belong to the European Qualified Presumption of Safety (QPS) status organisms, which can

Recently published studies have reported efficient cyanotoxin-removal by several strains of probiotic bacteria [79,80,96,97]. The aim of these studies was to characterise the potential of probiotic lactic acid bacteria and bifidobacteria in removal of microcystins and cylindrosper‐ mopsin from aqueous solutions. Different physiological conditions possibly affecting the removal efficiency were studied and the mechanism of toxin removal was investigated.

In an initial screening study, 15 different strains of probiotic lactic acid bacteria and bifido‐ bacteria were tested for their MC-LR removal capacities and evaluated for their potential in water decontamination [79]. The results showed a reproducible reduction of MC-LR in solution by the majority of the tested bacterial strains; the most efficient removal was achieved with *L. rhamnosus* strains GG and LC-705, *B. lactis* strains 420 and Bb12 and *B. longum* 46 [79]. The removal of MC-LR continued during the entire 24-hour incubation, which indicates that the removal process is quite slow. The effect of pH during incubation was also studied. pH was found to have an influence on toxin removal, with a higher removal percentage observed at neutral pH than at pH 3 [79]. It was also shown that viable bacteria were more efficient in microcystin-removal than non-viable bacteria [80]. Further studies showed that several strains were efficient in microcystin-removal and that different physiological conditions, including

The removal of MC-LR was shown to be temperature dependent, with the highest removal observed at 37 °C for all studied strains. At 4 °C, practically no removal of MC-LR could be observed and the removal percentages increased with increasing temperature [79]. This can be explained by the fact that at 4 °C, the bacterial cells are metabolically inactive, but at 22 and 37 °C, the bacteria become metabolically active, which is required for enzymatic activity. In addition, the role of glucose in activating the metabolism of the probiotic bacteria was assessed [96]. Since it was shown that viability is a requirement for efficient toxin removal, glucose was added as a source of nutrient to the bacterial solutions to enhance the bacterial viability. Glucose addition improved the removal efficiencies of all tested strains by enhancing both the removal rate and the amount of MC-LR removed after 24 hours of incubation [96]. Supple‐ mentation of glucose provides energy to the bacteria, and thereby, the microcystin-removal efficiencies also increase.

To investigate the role of the probiotic bacterial cell density, a range of bacterial cell densities were screened and tested for their microcystin-removal efficiencies [79]. The removal of MC-LR was shown to be dependent on the bacterial cell density, with a minimum of approximately 109 CFU/mL required for significant MC-LR removal [79]. The removal of MC-LR was further enhanced with increasing bacterial cell density. To assess whether a combination of several probiotic strains could enhance their microcystinremoval efficiencies the microcystin-removal of three probiotic strains (*L. rhamnosus* GG, *L. rhamnosus* LC-705 and *B. longum* 46) separately and in combination was studied [80]. With the probiotic mixture, microcystin-removal percentages of up to 90% could be observed and the results showed that the removal efficiency was improved with a mixture of the strains and compared to the individual strains [80].

In addition to MC-LR, probiotic bacterial strains were also incubated with other microcystins, including MC-RR, -YR, -LY, -LW and -LF. The results of the study show that probiotic strains were effective in the elimination of several different microcystins from solution [79]. Simulta‐ neous removal of several toxins present in cyanobacterial extracts was also investigated. The time course for the removal of microcystins present in the cyanobacterial extracts *Microcystis* NIES-107 and *Microcystis* PCC 7820 by the probiotic strain *L. rhamnosus* GG is shown in Figures 3 a and b, respectively. The removal of all studied microcystins increased over time. The removal of the microcystins present in *Microcystis* NIES-107 after 24 hours of incubation was around 65–85% of total microcystin and for microcystins present in *Microcystis* PCC 7820 around 60–80%. The toxin-removal was thus shown to be efficient also when several different microcystins were present in the solution. This indicates that there is no competition taking place among the toxins during incubation with probiotic bacteria. In addition, the strains were shown to remove the cytotoxin cylindrospermopsin from aqueous solutions; the removal was somewhat less efficient, around 30% for all tested strains [80].

Probiotic bacteria have several advantages in comparison with the previously reported microcystin-degrading bacteria, as they have been classified as food grade, safe bacteria by the European Food Safety Authority (EFSA) [95]. Therefore probiotic bacteria can safely be 1

2 3

included in both food and water. Previous studies have also shown the effect of probiot‐ ic bacteria in the removal of other environmental contaminants, such as heavy metals [98] and mycotoxins including aflatoxins and ochratoxins [99].

12 Biodegradation of cyanobacterial toxins – a review

The participation of cell-envelope proteinases in microcystin-removal was investigated. Following standard peptidase assay no proteolytic activity was found in the supernatants of the bacterial cell cultures of the investigated strains; enzymatic activity was found only in the cell suspensions. The activity of cell-associated proteinases of probiotic strain *L. rhamnosus* GG was measured after incubation with protease inhibitors. The protein inhibitor EDTA was shown to inhibit MC-LR removal [97]. The results suggest that the main proteolytic activity observed for the strain was due to metallo-enzymes. A possible extracellular enzymatic degradation of microcystins by probiotic bacteria was therefore investigated and it was suggested that extracellularly located cell-envelope proteinases appear to be involved in the decomposition of MC-LR [97]. A correlation between proteinase activity and MC-LR removal was also found when these parameters were simultaneously measured. The correlation between the activity of cell-envelope proteinases and the decrease of MC-LR concentration suggests that enzymes are involved in microcystin-removal [97]. The findings support the theory that enzymatic degradation of microcystins occurs when the toxin is incubated with

Biodegradation of Cyanobacterial Toxins http://dx.doi.org/10.5772/55511 161

Bacterial degradation of microcystins has previously been reported for strains of *Sphingomo‐ nas* and the degradation products and patterns have been determined for strains ACM-3962 and B9 [63,66,67,82]. For possible identification of toxin removal by probiotic bacteria, the removal process of MC-LR for *L. rhamnosus* GG was compared with the two *Sphingomo‐ nas* strains, and the degradation products were identified [97]. Linearized MC-LR and the tetrapeptide were observed for the two *Sphingomonas* strains, but these degradation products were not obtained using the probiotic strain, suggesting that the removal mechanisms between the strains differ [97]. Furthermore, no additional degradation products could be identified from samples incubated with the probiotic strain, which suggested that microcystin is rapidly degraded to smaller peptides and amino acids. Further studies are needed to identify possible degradation products and the precise steps

The majority of cyanotoxin-biodegradation studies have focused on bacteria isolated from water sources exposed to microcystin-containing blooms. As described in this review, it is clear that many of the cyanobacterial metabolites are susceptible to biodegradation in water supplies. Currently an increasing focus on bacterial degradation of hepatotoxic cyanobacte‐ rial peptides is being observed [56,59,64,66,76]. Previous studies have demonstrated that the ability of bacteria to degrade microcystins is related to the presence of the gene mlrA that encodes a hydrolytic enzyme with specificity to the toxins. The potency to utilize these bacteria in microcystin-degradation has also been demonstrated in laboratory scale [51,54, 100]. Recently, a new type of bacteria, specific probiotic bacterial strains, was presented to be efficient in cyanotoxin-removal. Probiotic bacteria have several advantages in compari‐ son with the previously reported microcystin-degrading bacteria, as they have been classified as food grade, safe bacteria by the EFSA [95]. Therefore probiotic bacteria can

probiotic bacteria, but the exact mechanism still remains unidentified.

of the degradation mechanism by probiotic bacteria.

**8. Discussion and conclusions**

**Figure 3.** Removal of microcystins in cyanobacterial extracts by probiotic strain *Lactobacillus rhamnosus* GG (a) Micro‐ *cystis* NIES-107 and (b) *Microcystis* PCC 7820. Initial concentration of microcystins in extracts: 20-100 µg/L, bacterial concentration 1010 CFU/mL, temperature 37 °C, average ± SD, n = 3 (modified from [80]).

### **7.2. Mechanisms of microcystin-degradation by probiotic bacteria**

As specific probiotic bacterial strains were shown to be efficient in microcystin-removal, the subsequent aim was to identify and specify the removal mechanisms. The location and mechanism of microcystin-removal were investigated by studying a possible extracellular enzymatic degradation of microcystins [97]. Furthermore, a comparison of the degrada‐ tion pathways of previously identified microcystin-degrading bacteria with probiotic bacteria was performed.

The participation of cell-envelope proteinases in microcystin-removal was investigated. Following standard peptidase assay no proteolytic activity was found in the supernatants of the bacterial cell cultures of the investigated strains; enzymatic activity was found only in the cell suspensions. The activity of cell-associated proteinases of probiotic strain *L. rhamnosus* GG was measured after incubation with protease inhibitors. The protein inhibitor EDTA was shown to inhibit MC-LR removal [97]. The results suggest that the main proteolytic activity observed for the strain was due to metallo-enzymes. A possible extracellular enzymatic degradation of microcystins by probiotic bacteria was therefore investigated and it was suggested that extracellularly located cell-envelope proteinases appear to be involved in the decomposition of MC-LR [97]. A correlation between proteinase activity and MC-LR removal was also found when these parameters were simultaneously measured. The correlation between the activity of cell-envelope proteinases and the decrease of MC-LR concentration suggests that enzymes are involved in microcystin-removal [97]. The findings support the theory that enzymatic degradation of microcystins occurs when the toxin is incubated with probiotic bacteria, but the exact mechanism still remains unidentified.

Bacterial degradation of microcystins has previously been reported for strains of *Sphingomo‐ nas* and the degradation products and patterns have been determined for strains ACM-3962 and B9 [63,66,67,82]. For possible identification of toxin removal by probiotic bacteria, the removal process of MC-LR for *L. rhamnosus* GG was compared with the two *Sphingomo‐ nas* strains, and the degradation products were identified [97]. Linearized MC-LR and the tetrapeptide were observed for the two *Sphingomonas* strains, but these degradation products were not obtained using the probiotic strain, suggesting that the removal mechanisms between the strains differ [97]. Furthermore, no additional degradation products could be identified from samples incubated with the probiotic strain, which suggested that microcystin is rapidly degraded to smaller peptides and amino acids. Further studies are needed to identify possible degradation products and the precise steps of the degradation mechanism by probiotic bacteria.

### **8. Discussion and conclusions**

included in both food and water. Previous studies have also shown the effect of probiot‐ ic bacteria in the removal of other environmental contaminants, such as heavy metals [98]

0 6 12 18 24

0 6 12 18 24

time (h)

**Figure 3.** Removal of microcystins in cyanobacterial extracts by probiotic strain *Lactobacillus rhamnosus* GG (a) Micro‐ *cystis* NIES-107 and (b) *Microcystis* PCC 7820. Initial concentration of microcystins in extracts: 20-100 µg/L, bacterial

As specific probiotic bacterial strains were shown to be efficient in microcystin-removal, the subsequent aim was to identify and specify the removal mechanisms. The location and mechanism of microcystin-removal were investigated by studying a possible extracellular enzymatic degradation of microcystins [97]. Furthermore, a comparison of the degrada‐ tion pathways of previously identified microcystin-degrading bacteria with probiotic

concentration 1010 CFU/mL, temperature 37 °C, average ± SD, n = 3 (modified from [80]).

**7.2. Mechanisms of microcystin-degradation by probiotic bacteria**

time (h)

12 Biodegradation of cyanobacterial toxins – a review

dm-MC-RR MC-RR MC-YR MC-LR

MC-LR MC-LY MC-LW MC-LF

and mycotoxins including aflatoxins and ochratoxins [99].

160 Environmental Biotechnology - New Approaches and Prospective Applications

**(a)**

1

**(b)**

microcystins removed (%)

microcystins removed (%)

2 3

bacteria was performed.

The majority of cyanotoxin-biodegradation studies have focused on bacteria isolated from water sources exposed to microcystin-containing blooms. As described in this review, it is clear that many of the cyanobacterial metabolites are susceptible to biodegradation in water supplies. Currently an increasing focus on bacterial degradation of hepatotoxic cyanobacte‐ rial peptides is being observed [56,59,64,66,76]. Previous studies have demonstrated that the ability of bacteria to degrade microcystins is related to the presence of the gene mlrA that encodes a hydrolytic enzyme with specificity to the toxins. The potency to utilize these bacteria in microcystin-degradation has also been demonstrated in laboratory scale [51,54, 100]. Recently, a new type of bacteria, specific probiotic bacterial strains, was presented to be efficient in cyanotoxin-removal. Probiotic bacteria have several advantages in compari‐ son with the previously reported microcystin-degrading bacteria, as they have been classified as food grade, safe bacteria by the EFSA [95]. Therefore probiotic bacteria can safely be included in both food and water, and can also safely be used in food technolo‐ gy. Furthermore, the beneficial health effects of probiotic bacteria give them an advant‐ age for the use in different applications. A potential area of use could be probiotic dietary supplements used as a personal defense mechanism against cyanotoxins in the gastrointes‐ tinal tract when ingested through contaminated drinking water and to reduce the health risks caused by microcystins, as well as applications in biological decontamination of microcystin-containing water.

[2] Schopf JW. Microfossils of the Early Archean Apex chert: new evidence of the antiq‐

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[3] Sivonen K., Jones G. Cyanobacterial toxins. In: Chorus I., Bartram J. (eds) Toxic Cya‐ nobacteria in Water: a Guide to Public Health Significance, Monitoring and Manage‐

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Several reports have showed that biological degradation of cyanotoxins may be a feasi‐ ble method of water treatment. The bacterial strains or possible enzymes identified in the removal process could be used in a degradation process to remove toxins from drinking water. Technologies using potential purified enzymes identified in the removal process of bacteria could be a future approach for efficient cyanotoxin-removal. Today, the best way for cyanotoxin biodegradation is the use of biofilters with immobilised micro-organisms, as most water treatment processes already employ a filtration step. Also the removal of other cyanobacterial toxins, such as anatoxins, saxitoxins, and cylindrospermopsin, should be taken into account. In conclusion, the development of new water treatment technolo‐ gies using efficient bacteria that would be able to remove or inactivate cyanotoxins, as well as other types of environmental contaminants, such as heavy metals, viruses and pathogen‐ ic bacteria found in drinking water, is an important aspect to consider in the future.

### **Acknowledgements**

Dr. Jussi Meriluoto and Prof. Seppo Salminen are gratefully acknowledged for excellent supervision and guidance during the research work. Svenska litteratursällskapet i Finland r.f. is acknowledged for financial support.

### **Author details**

Sonja Nybom

Department of biosciences, Biochemistry, Åbo Akademi University, Finland

### **References**

[1] WHO. Cyanobacterial toxins: Microcystin-LR. In: Guidelines for drinking-water quality, Addendum to Volume 2, World Health Organization, Geneva; 1998. p95– 110.


safely be included in both food and water, and can also safely be used in food technolo‐ gy. Furthermore, the beneficial health effects of probiotic bacteria give them an advant‐ age for the use in different applications. A potential area of use could be probiotic dietary supplements used as a personal defense mechanism against cyanotoxins in the gastrointes‐ tinal tract when ingested through contaminated drinking water and to reduce the health risks caused by microcystins, as well as applications in biological decontamination of

Several reports have showed that biological degradation of cyanotoxins may be a feasi‐ ble method of water treatment. The bacterial strains or possible enzymes identified in the removal process could be used in a degradation process to remove toxins from drinking water. Technologies using potential purified enzymes identified in the removal process of bacteria could be a future approach for efficient cyanotoxin-removal. Today, the best way for cyanotoxin biodegradation is the use of biofilters with immobilised micro-organisms, as most water treatment processes already employ a filtration step. Also the removal of other cyanobacterial toxins, such as anatoxins, saxitoxins, and cylindrospermopsin, should be taken into account. In conclusion, the development of new water treatment technolo‐ gies using efficient bacteria that would be able to remove or inactivate cyanotoxins, as well as other types of environmental contaminants, such as heavy metals, viruses and pathogen‐ ic bacteria found in drinking water, is an important aspect to consider in the future.

Dr. Jussi Meriluoto and Prof. Seppo Salminen are gratefully acknowledged for excellent supervision and guidance during the research work. Svenska litteratursällskapet i Finland r.f.

[1] WHO. Cyanobacterial toxins: Microcystin-LR. In: Guidelines for drinking-water quality, Addendum to Volume 2, World Health Organization, Geneva; 1998. p95–

Department of biosciences, Biochemistry, Åbo Akademi University, Finland

microcystin-containing water.

162 Environmental Biotechnology - New Approaches and Prospective Applications

**Acknowledgements**

**Author details**

Sonja Nybom

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High-performance liquid chromatography of microcystins and nodularins, cyanobacterial peptide toxins. PhD thesis. Åbo Akademi University; 2004.


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**Chapter 8**

**Bioavailability of High Molecular Weight Polycyclic**

Olusola Solomon Amodu, Tunde Victor Ojumu and

Additional information is available at the end of the chapter

Seteno Karabo Obed Ntwampe

http://dx.doi.org/10.5772/54727

**1. Introduction**

Agency [1].

**Aromatic Hydrocarbons Using Renewable Resources**

Polycyclic aromatic hydrocarbons (PAHs) are the world's largest class of carcinogens known to date, not only because of their ability to cause gene mutation and cancer, but due to their persistency in the environment. They are particularly recalcitrant due to their molecular weight, hydrophobic nature and thus, accumulate in various matrices in the environment.

PAHs, also known as polyarenes or polynuclear aromatic hydrocarbons, are formed and released into the environment through natural and anthropogenic sources. Natural sources include volcanoes and forest fires while anthropogenic sources include, majorly, incomplete combustion of fossil fuels, wood burning, municipal and industrial waste incineration. PAHs containing two or three fused benzene rings are classified as low molecular weight (LMW) PAHs and are more water soluble while those with four or more benzene rings are referred to as high molecular weight (HMW) PAHs. They tend to adsorb onto soil and sediment thus, making them recalcitrant in the environment. Sixteen of these organic compounds have been identified as priority pollutants due to their hazardous properties, with HMW PAHs being considered as potential human carcinogens, by the United State Environmental Protection

The cost of biodegradation technology and the low bioavailability including mass transfer limitations of PAHs, especially those with high molecular weight, from several matrices into the aqueous phase for effective enzyme-based microbial biodegradation still constitute major challenges. However, current research efforts have focused on the combined use of biosurfac‐ tants and enzymes produced from renewable resources such as agricultural by-products and/ or agro-industrial waste, through assisted biostimulation and bioaugmentation, for biodegra‐

> © 2013 Amodu 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 Amodu et al.; licensee InTech. This is a paper 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.

