**Analytical Methodology for Determination of Trace Cu in Hydrated Alcohol Fuel**

Fabiana Aparecida Lobo, Fernanda Pollo, Ana Cristina Villafranca and Mercedes de Moraes

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52514

#### **1. Introduction**

Many private and governmental initiatives have been established worldwide to identify viable alternatives to petroleum derivatives [1,2].The goals are to reduce dependence on imported en‐ ergy from non-renewable sources, while mitigating environmental problems caused by petro‐ leum products, and to develop national technologies in the alternative energy field.

Ethyl alcohol (ethanol) is considered to be a highly viable alternative fuel. Its production from biomass means that it can provide a source of energy that is both clean and renewable. The in‐ clusion of ethanol as a component of gasoline can help to reduce problems of pollution in many regions, since it eliminates the needto use tetraethyl lead (historically notorious as a highly tox‐ ic trace component of the atmosphere in major cities) as an anti-knock additive.

The quantitative monitoring of metal elements in fuels (including gasoline, alcohol, and die‐ sel) is important from an economic perspective in the fuel industry as well as in the areas of transport and environment. The presence of metalspecies (ions or organometallic com‐ pounds) in automotive fuels can cause engine corrosion, reduce performance, and contrib‐ ute to environmental contamination [2-5].

The low concentrations of metals in fuels typically require the use of sensitive spectrometric an‐ alytical techniques for the purposes of quality control. Atomic absorption spectrometry (AAS) can be applied for the quantitative determination of many elements (metals and semi-metals) in a wide variety of media including fuels, foodstuffs, and biological, environmental, and geo‐ logical materials, amongst others. The principle of the technique is based on measurement of the absorption of optical radiation, emitted from a source, by ground-state atoms in the gas phase. Atomization can be achieved using a flame, electrothermal heating, or specific chemical

© 2013 Lobo 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 Lobo 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.

reaction (such as the generation of Hg cold vapor). Electrothermalatomizers include graphite tubes, tungsten filaments, and quartz tubes (for atomization of hydrides), as well as metal or ce‐ ramic tubes. Flame atomic absorption spectrometry (FAAS) is mostly used for elemental analy‐ sis at higher concentration levels, of the order of mg L-1[3-5]. Table 1 lists some of the published studies concerning the application of AAS for determination of metals in fuels.

**Technique Matrix Sample preparation Reference**

spectrometry Ethyl alcohol and acids Direct determination <sup>19</sup>


emulsion

Treatment with acid under UV irradiation

Biodiesel Emulsion 21

Biodiesel Microwave digestion and

Analytical Methodology for Determination of Trace Cu in Hydrated Alcohol Fuel

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22

alcohol and biodiesel

Flame atomic absorption spectrometry Ethyl alcohol Solid phase extraction 20

The thermospray (TS) technique was originally developed by Vestal et al.in 1978 [23]as an interface between liquid chromatography and mass spectrometry. In atomic absorption spectrometry, the tube was heated electrically in order to maintain a constant temperature, which restricted use of the method to only a few elements. However, Gáspárand Berndt (2000) proposed the TS-FF-AAS procedure, in which a metal tube is positioned above the flame of the atomic absorption spectrometer, as a reactor. The sample solution is transported through a metal capillary, connected to the tube, and heated simultaneously by the flame. On reaching the hot tip of the capillary, the liquid partially vaporizes, forming an aerosol. In turn, the aerosol is vaporized within the tube, producing an atomic cloud that absorbs the

The TS-FF-AAS method was used as an interface between high performance liquid chroma‐

The objective of this work is to describe the analysis of Cu present in hydrated ethyl alcohol fuel (HEAF) using the technique of atomic absorption spectrometry with thermal nebuliza‐ tion in a tube heated in a flame (TS-FF-AAS). The atomizers used were a metal tube (Ni-Cr

The instrumentation consisted of an atomic absorption spectrometer fitted with a flame atomizer (Perkin-Elmer, model AAnalyst 100), a hollow cathode Cu lamp (λ = 324.8 nm, slit width = 0.7 nm,i = 15 mA), with an air/acetylene (4:2 ratio) flame gas mixture, and back‐

Atomic absorption spectrometry Gasoline, diesel, ethyl

generation (VP-FAAS) Ethyl alcohol

**Table 1.** Analytical methods for the determination of inorganic contaminants in fuels.

tography (HPLC) and FAAS, employing a flow injection system [25-60].

Graphite furnace atomic absorption

Graphite furnace atomic absorption spectrometry

Graphite furnace atomic absorption spectrometry

Atomic absorption spectrometry with vapor

radiation emitted by the lamp.

alloy) and a ceramic tube (Al2O3).

**2. Experimental procedures**

**2.1. Instruments and accessories**



**Table 1.** Analytical methods for the determination of inorganic contaminants in fuels.

The thermospray (TS) technique was originally developed by Vestal et al.in 1978 [23]as an interface between liquid chromatography and mass spectrometry. In atomic absorption spectrometry, the tube was heated electrically in order to maintain a constant temperature, which restricted use of the method to only a few elements. However, Gáspárand Berndt (2000) proposed the TS-FF-AAS procedure, in which a metal tube is positioned above the flame of the atomic absorption spectrometer, as a reactor. The sample solution is transported through a metal capillary, connected to the tube, and heated simultaneously by the flame. On reaching the hot tip of the capillary, the liquid partially vaporizes, forming an aerosol. In turn, the aerosol is vaporized within the tube, producing an atomic cloud that absorbs the radiation emitted by the lamp.

The TS-FF-AAS method was used as an interface between high performance liquid chroma‐ tography (HPLC) and FAAS, employing a flow injection system [25-60].

The objective of this work is to describe the analysis of Cu present in hydrated ethyl alcohol fuel (HEAF) using the technique of atomic absorption spectrometry with thermal nebuliza‐ tion in a tube heated in a flame (TS-FF-AAS). The atomizers used were a metal tube (Ni-Cr alloy) and a ceramic tube (Al2O3).

#### **2. Experimental procedures**

#### **2.1. Instruments and accessories**

The instrumentation consisted of an atomic absorption spectrometer fitted with a flame atomizer (Perkin-Elmer, model AAnalyst 100), a hollow cathode Cu lamp (λ = 324.8 nm, slit width = 0.7 nm,i = 15 mA), with an air/acetylene (4:2 ratio) flame gas mixture, and back‐ ground correction using a deuterium lamp. Other equipment comprised an analytical bal‐ ance (Sartorius BL 2105) and a peristaltic pump (Ismatec, model ICP 8).

ple, injected into the flow of air as the carrier, since previous work has shown that injection

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using carrier solutions results in greater sample dilution and dispersion [40,45,54,58].

**Figure 1.** Schematic arrangement of the TS-FF-AAS system (adapted from Davies & Berndt 2003[59].

ume (50, 100, and 200µL) were evaluated using a standard of 200µg Cu L–1.

ployed the height of the transient signal peak.

**2.5. Construction of analytical curve**

**2.4. Optimization of carrier flow rate and sample volume**

from 12 blank readings for each type of tube (metal or ceramic).

The sample was introduced into the system using a manual Rheodyne valve (Figure 1), after which it was transported to the ceramic capillary in the flow of air. Since the capillary was heated simultaneously with the metal or ceramic reactor tube, the liquid was partially va‐ porized, forming a thermospray, and atomization occurred on arrival in the tube, generating a transient signal that was captured and stored by the software. The determination em‐

The influences of the carrier flow rate (in the range 9.0-18.0 mL min-1) and the sample vol‐

After optimization of the system, analytical curves were constructed in the concentration range 0.1-0.4 mg Cu L-1 in 0.14mol L-1 HNO3. Additions of analyte were made to the sample mixed with an equal volume of 0.14 mol L-1 HNO3. The detection limit (DL) was calculated

The TS-FF-AAS assembly employed a Rheodyne RE9725 injection valve, PEEK tubing, and a ceramic thermocouple insulator capillary (OMEGATITE450, OMEGA, USA). The capillary wascomposed of Al2O3(>99.8%), resistant to temperatures up to 1900 °C, with Øext= 1.6 mm and two orifices with Øint = 0.4 mm (this capillary provided better results than a stainless steel HPLC capillary, with less noise in the absorbance signal). The atomization tubes were a metal tube composed of Ni-Cr super-alloy (Inconel, length 100 mm, Øint = 10.0 mm, Øext = 12.0 mm, 6 orifices with Ø = 2.5 mm, perpendicular to an orifice with Ø = 2.0 mm), and a ceramic tube (99.9% Al2O3, length 100 mm, Øint = 10.0 mm, Øext = 12.0 mm, 6 orifices with Ø = 2.5 mm, perpendicular to an orifice with Ø = 2.0 mm).

Data acquisition employed the software MQDOS (Microquímica), and the absorbance val‐ ues were proportional to the height of the transient signals.

The temperature in the interior of the atomization tube was measured in two ways. The first method employed a thermocouple with an earthed connection, positioned adjacent to the metal tube, oriented towards the orifice where the ceramic capillary used to introduce the sample into the atomizer was located. The temperature measured for the metal tube was 983 ± 1°C. Secondly, the thermocouple with connection exposed was positioned adjacent to the ceramic capillary within the tube, where a temperature of between 1030 °C and 1060°C was measured, at which the tube glowed ruby-red above the flame [16,40,45,54].

When 50 µL of HNO3 (~0.1 mol L-1) was injected at a rate of approximately 1.5 mL min-1,there was a temperature reduction of around 50°C, due to cooling of the tube by the solution, followed by a rapid return to the maximum temperature range.

#### **2.2. Reagents, solutions and samples**

Working standard solutions were prepared from a stock 1000 mg L-1 copper standard solu‐ tion (spectroscopic grade), by dilution in 0.14 mol L-1 HNO3 (Synth).

The HEAF samples were prepared by mixing the fuel with an equal volume of 0.14 mol L-1 HNO3,with final volumes of 50 mL [3-5]. Subsequent quantification employed the standard additions procedure.

#### **2.3. Assembly of the TS-FF-AAS system**

A schematic diagram of the TS-FF-AAS system is shown in Figure 1.

It is recommended that the Inconel tube should only be positioned above the burner head after lighting the flame, to avoid the possibility of an explosion within the tube due to gas accumulation. The TS-FF-AAS system was therefore first assembled, after which the spec‐ trometer flame was ignited immediately after opening the gas valves to avoid any explosion risk. This procedure facilitated the positioning of the tube above the burner head, which was performed while the flame was extinguished. All analyses employed a fixed volume of sam‐ ple, injected into the flow of air as the carrier, since previous work has shown that injection using carrier solutions results in greater sample dilution and dispersion [40,45,54,58].

**Figure 1.** Schematic arrangement of the TS-FF-AAS system (adapted from Davies & Berndt 2003[59].

The sample was introduced into the system using a manual Rheodyne valve (Figure 1), after which it was transported to the ceramic capillary in the flow of air. Since the capillary was heated simultaneously with the metal or ceramic reactor tube, the liquid was partially va‐ porized, forming a thermospray, and atomization occurred on arrival in the tube, generating a transient signal that was captured and stored by the software. The determination em‐ ployed the height of the transient signal peak.

#### **2.4. Optimization of carrier flow rate and sample volume**

The influences of the carrier flow rate (in the range 9.0-18.0 mL min-1) and the sample vol‐ ume (50, 100, and 200µL) were evaluated using a standard of 200µg Cu L–1.

#### **2.5. Construction of analytical curve**

After optimization of the system, analytical curves were constructed in the concentration range 0.1-0.4 mg Cu L-1 in 0.14mol L-1 HNO3. Additions of analyte were made to the sample mixed with an equal volume of 0.14 mol L-1 HNO3. The detection limit (DL) was calculated from 12 blank readings for each type of tube (metal or ceramic).

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

#### **3.1. Optimization of carrier flow rate and sample volume**

Good peak reproducibility was achieved when samples were injected into the air flow as carrier. When samples were injected into 0.14 mol L-1 HNO3, used as the carrier, there was a rise in the baseline (as expected, due to increase of the blank), followed by a fall due to cool‐ ing of the metal or ceramic tubes. This cooling was significant, since no transient signals were obtained following injection of standards, indicating that the temperature within the tubes was insufficient to atomize the analyte, which remained dispersed in the carrier solu‐ tion. This confirmed the findings of earlier work that the use of air (or other gas) as the carri‐ er avoids dilution and dispersion of the sample. Here, all analyses were performed using air as the carrier, not only because it was less expensive than use of a solution, and minimized waste generation, but also because it enabled the TS-FF-AAS system to be used to determine copper, which would not have been possible using a solution as the carrier.

This increase proceeded up to a carrier flow rate of 12.0 mL min-1, above which there was no significant variation in absorbance. The highest absorbance value was obtained at a flow rate of 18.0 mL min-1, which was therefore selected as the best flow rate to use with the metal

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When the ceramic tube was used, maximum absorbance was achieved at a carrier flow rate of 9.0 mL min-1. At higher flow rates, the residence time of the liquid in the heated section of the ceramic capillary was considerably diminished, reducing the time available for evapora‐ tion of the liquid, so that the sample was not delivered in the form of vapor/aerosol, but rather as a flow of liquid. The temperature within the tube decreased, and the color of the tube changed from ruby-red to opaque grey. It was also possible to see droplets emerging from the atomizer tube. Hence, the absorbance values did not increase, while greater varia‐ bility in the signal resulted in elevated standard deviation values. A flow rate of 3.0 mL min-1was selected, at which the absorbance signal was maximized, and the standard devia‐

**Figure 3.** Influence of sample volume on the absorbances obtained for a solution of 200 μg CuL-1in0.14mol L-1HNO3,

The sample volume was varied between 50 and 200 µL, using carrier flow rates of 18.0 and 9.0 mL min-1 for the metal and ceramic tubes, respectively. The results (Figure 3) re‐ vealed that for both tubes a sample volume of 50 µL generated the highest absorbance value, with a low standard deviation, reflecting good repeatability in the experimental measurements. When 100 µL of sample was used, there was a slight cooling of the ceram‐ ic capillary, and consequently of the atomization tubes, while there was no increase in the absorbance values. At a sample volume of 200 µL, the ceramic capillary and the tube

using carrier flow rates of 9.0 and 18.0 mL min-1for the ceramic and metal tubes, respectively.

tube.

tion was minimized.

Figure 2 shows the influence of the carrier (air) flow rate, in the range 9.0-18.0 mL min-1, on the absorbance values obtained using 50 µL of a standard of 200 µg Cu L-1in 0.14 mol L-1 HNO3, using both tubes.In the case of the metal tube, lower absorbance values were ob‐ tained at low flow rates, because the sample arrived slowly at the atomizer,increasing the measurement duration and resulting in an unpredictable and erratic vaporization. Hence, as the flow rate was increased, the absorbance also increased due to a more homogeneous va‐ porization of the sample [23,27,58].

**Figure 2.** Influence of carrier (air) flow rate on the absorbance obtained for 50 μLof a solution of200 μg Cu L-1 in 0.14mol L-1HNO3, using the metal and ceramic tubes.

This increase proceeded up to a carrier flow rate of 12.0 mL min-1, above which there was no significant variation in absorbance. The highest absorbance value was obtained at a flow rate of 18.0 mL min-1, which was therefore selected as the best flow rate to use with the metal tube.

When the ceramic tube was used, maximum absorbance was achieved at a carrier flow rate of 9.0 mL min-1. At higher flow rates, the residence time of the liquid in the heated section of the ceramic capillary was considerably diminished, reducing the time available for evapora‐ tion of the liquid, so that the sample was not delivered in the form of vapor/aerosol, but rather as a flow of liquid. The temperature within the tube decreased, and the color of the tube changed from ruby-red to opaque grey. It was also possible to see droplets emerging from the atomizer tube. Hence, the absorbance values did not increase, while greater varia‐ bility in the signal resulted in elevated standard deviation values. A flow rate of 3.0 mL min-1was selected, at which the absorbance signal was maximized, and the standard devia‐ tion was minimized.

**Figure 3.** Influence of sample volume on the absorbances obtained for a solution of 200 μg CuL-1in0.14mol L-1HNO3, using carrier flow rates of 9.0 and 18.0 mL min-1for the ceramic and metal tubes, respectively.

The sample volume was varied between 50 and 200 µL, using carrier flow rates of 18.0 and 9.0 mL min-1 for the metal and ceramic tubes, respectively. The results (Figure 3) re‐ vealed that for both tubes a sample volume of 50 µL generated the highest absorbance value, with a low standard deviation, reflecting good repeatability in the experimental measurements. When 100 µL of sample was used, there was a slight cooling of the ceram‐ ic capillary, and consequently of the atomization tubes, while there was no increase in the absorbance values. At a sample volume of 200 µL, the ceramic capillary and the tube were substantially cooled, and there was no homogeneous thermospray formation, with erratic generation of droplets that acted to disperse the light radiation (probably to a large degree, since the deuterium lamp was unable to fully correct the resulting background signal). The unpredictable atomization resulted in very high standard deviation values. Using air as the carrier, a sample volume of 50 µL was selected for the subsequent meas‐ urements, due to greater atomization homogeneity, satisfactory absorbance for a 30 mg Cu L-1 standard, and a low SD value.

#### **3.2. Construction of analytical curves**

Figure 4 illustrates the results obtained for the analytical curve in the concentration range 0.1-0.4 µg Cu L–1 in 0.14 mol L-1HNO3, using the optimized conditions of the TS-FF-AAS sys‐ tem. The transient signals were repeatable, and (for both tubes) the curve was linear in the concentration range studied. A two-fold greater sensitivity was achieved using the ceramic tube.

0 100 200 300 400 500

**[Cu] /** m**g L-1**

**Figure 5.** Regression lines fitted to the analytical curves of Cu in 1:1 mixtures of fuel samples and standards prepared in 0.14 mol L-1 HNO3, obtained using the ceramic tube (a) and the metal tube (b) Equations of the lines: A = 1.00x10-2 +

**Analytical characteristics Ceramic tube**

Detection limit, DL (μg L-1) 55.6 56.0

Correlation coefficient (r) 0.9930 0.9978 Analytical frequency (h-1) 26 100

Detection limit, DL (μg L-1) 64.5 128 Characteristic concentration, Co (μg L-1) 33.3 30.3 Analytical curve interval (μg L-1) 100 – 400

Correlation coefficient (r) 0.9918 0.9927 Analytical frequency (h-1) 53 82

**Table 2.** Analytical characteristics for determination of Cu using the TS-FF-AAS system with ceramic and metal tubes.

The analytical parameters obtained for the determination of Cu under the optimized condi‐ tions of the TS-FF-AAS system are provided in Table 2. The analytical curves were linear for a concentration range of 100-400 µg Cu L-1 in 0.14 mol L-1 HNO3. The system could be con‐ sidered to be sensitive, with characteristic concentrations of 8 and 15 µg Cu L-1 for the ce‐ ramic and metal tubes, respectively, and analysis frequencies (using HNO3 medium) of 26

Characteristic concentration, Co (μg L-1) 8.35 15.1 Analytical curve interval (μg L-1) 100 – 400

a b

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**Metal Tube**

0.00

HNO3

HEAF

1.32x10-4 (Cu) (ceramic tube); A = 1.16x10-2 + 1.45x10-4 (Cu) (metal tube).

0.05

0.10

0.15

0.20

0.25

0.30

**A**

**Figure 4.** Regression lines fitted to the analytical curves of Cu obtained using the ceramic tube (a) and the metal tube (b) Equations of the lines: A = 1.16x10-2 + 5.27x10-4(Cu) (ceramic tube);A = 1.20x10-3 + 2.91x10-4(Cu) (metal tube).

Figure 5 illustrates the results obtained for the analytical curves constructed using concen‐ trations of Cu in the range 100-400 µg L–1, with additions of analytein 0.14 mol L-1 HNO3 to equal volumes of sample, under the optimized TS-FF-AAS system conditions. The presence of 75.8 µg Cu L-1 in the sample was calculated from curve (a), obtained using the ceramic tube. This value was slightly above the detection limit (Table 1), although below the concen‐ tration of the first point of the analytical curve. In the case of the metal tube (curve (b)), a Cu concentration of 80.0 µg L-1 was below the detection limit for this tube, but was nevertheless in agreement with the result obtained for the ceramic tube.

**Figure 5.** Regression lines fitted to the analytical curves of Cu in 1:1 mixtures of fuel samples and standards prepared in 0.14 mol L-1 HNO3, obtained using the ceramic tube (a) and the metal tube (b) Equations of the lines: A = 1.00x10-2 + 1.32x10-4 (Cu) (ceramic tube); A = 1.16x10-2 + 1.45x10-4 (Cu) (metal tube).


**Table 2.** Analytical characteristics for determination of Cu using the TS-FF-AAS system with ceramic and metal tubes.

The analytical parameters obtained for the determination of Cu under the optimized condi‐ tions of the TS-FF-AAS system are provided in Table 2. The analytical curves were linear for a concentration range of 100-400 µg Cu L-1 in 0.14 mol L-1 HNO3. The system could be con‐ sidered to be sensitive, with characteristic concentrations of 8 and 15 µg Cu L-1 for the ce‐ ramic and metal tubes, respectively, and analysis frequencies (using HNO3 medium) of 26 and 100 determinations per hour, respectively. Better analytical performance of the system was achieved using the ceramic tube, compared to the metal tube. The data showed that the TS-FF-AAS technique was more sensitive than FAAS, with nine-fold (ceramic tube) and five-fold (metal tube) increases in sensitivity, relative to FAAS with pneumatic nebulization, for which the characteristic concentration was 77 µg L-1. The increase in power of detectio‐ nobtained using the ceramic tube was around twice that for the metal tube. The sensitivity for determination of copper using the ceramic tube was therefore two-fold that obtained us‐ ing the metal tube.

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#### **4. Conclusions**

The TS-FF-AAS system can be used to determine copper at low concentrations, using either metal (Inconel) or ceramic (Al2O3) tubes as atomizers. Following optimization considering the most important experimental variables affecting atomization, these systems provided significantly improved detection limits for Cu determination, with nine-fold (ceramic tube) and five-fold (metal tube) increases in sensitivity, compared to traditional FAAS with pneu‐ matic nebulization. The TS-FF-AAS technique is simple, fast, effective, and inexpensive. It requires low volumes of sample (as little as 50 µL) and reagents, and reduces waste genera‐ tion. The method offers a useful new alternative for the determination of copper in alcohol.

#### **Acknowledgments**

The authors thank UFOP and CNPq for financial assistance.

#### **Author details**

Fabiana Aparecida Lobo1 , Fernanda Pollo2 , Ana Cristina Villafranca2 and Mercedes de Moraes2

1 UFOP - Universidade Federal de Ouro Preto, Brazil

2 UNESP – Universidade Estadual Paulista, Brazil

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nal of Analytical Atomic Spectrometry 2002;17:1308-15


**Chapter 5**

**Gas Fermentation for Commercial Biofuels Production**

With diminishing global reserves of crude oil and increasing demand, especially from developing countries, the pressure on oil supply will grow. Although the 2007-2010 fi‐ nancial crisis brought down the price of crude oil (per barrel) from a record peak of US \$145 in July 2008, factors such as recovering global economies and political instability in the Middle East have restored the price of crude oil to the US\$100 mark. At current rate of consumption, the global reserves of petroleum are predicted to be exhausted within 50 years [1, 2]. This, coupled with the deleterious environmental impacts that result from accumulating atmospheric CO2 from the burning of fossil fuels, the development of af‐ fordable, and environmentally sustainable fuels is urgently required. Many countries have responded to this challenge by legislating mandates and introducing policies to stimulate research and development (R&D) and commercialization of technologies that allow the production of low cost, low fossil carbon emitting fuels. For instance, the Euro‐ pean Union (EU) has mandated member countries to a target of deriving 10% of all transportation fuel from renewable sources by 2020 [3]. Between 2005 and 2010, renewa‐ ble energies such as solar, wind, and biofuels have been increasing at an average annual rate of 15-50% [4]. Renewable energy accounted for an estimated 16% of global final en‐

Biofuels have been defined as solid (bio-char), liquid (bioethanol, biobutanol, and biodie‐ sel) and gaseous (biogas, biosyngas, and biohydrogen) fuels that are mainly derived from biomass [5]. Liquid biofuels provided a small but growing contribution towards worldwide fuel usage, accounting for 2.7% of global road transport fuels in 2010 [4]. The world's largest producer of biofuels is the United States (US), followed by Brazil and the EU [4]. In 2009, US and Brazil accounted for approximately 85% of global bioethanol

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

© 2013 Liew 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.

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

Fung Min Liew, Michael Köpke and

Additional information is available at the end of the chapter

Séan Dennis Simpson

http://dx.doi.org/10.5772/52164

ergy consumption in 2009 [4].

**1. Introduction**

### **Gas Fermentation for Commercial Biofuels Production**

Fung Min Liew, Michael Köpke and Séan Dennis Simpson

Additional information is available at the end of the chapter

http://dx.doi.org/10.5772/52164

#### **1. Introduction**

With diminishing global reserves of crude oil and increasing demand, especially from developing countries, the pressure on oil supply will grow. Although the 2007-2010 fi‐ nancial crisis brought down the price of crude oil (per barrel) from a record peak of US \$145 in July 2008, factors such as recovering global economies and political instability in the Middle East have restored the price of crude oil to the US\$100 mark. At current rate of consumption, the global reserves of petroleum are predicted to be exhausted within 50 years [1, 2]. This, coupled with the deleterious environmental impacts that result from accumulating atmospheric CO2 from the burning of fossil fuels, the development of af‐ fordable, and environmentally sustainable fuels is urgently required. Many countries have responded to this challenge by legislating mandates and introducing policies to stimulate research and development (R&D) and commercialization of technologies that allow the production of low cost, low fossil carbon emitting fuels. For instance, the Euro‐ pean Union (EU) has mandated member countries to a target of deriving 10% of all transportation fuel from renewable sources by 2020 [3]. Between 2005 and 2010, renewa‐ ble energies such as solar, wind, and biofuels have been increasing at an average annual rate of 15-50% [4]. Renewable energy accounted for an estimated 16% of global final en‐ ergy consumption in 2009 [4].

Biofuels have been defined as solid (bio-char), liquid (bioethanol, biobutanol, and biodie‐ sel) and gaseous (biogas, biosyngas, and biohydrogen) fuels that are mainly derived from biomass [5]. Liquid biofuels provided a small but growing contribution towards worldwide fuel usage, accounting for 2.7% of global road transport fuels in 2010 [4]. The world's largest producer of biofuels is the United States (US), followed by Brazil and the EU [4]. In 2009, US and Brazil accounted for approximately 85% of global bioethanol

production while Europe generated about 85% of the world's biodiesel [6]. The global market for liquid biofuels (bioethanol and biodiesel) increased dramatically in recent years, reaching US\$83 billion in 2011 and is projected to US\$139 billion by 2021 [7].

**2. Advantages of gas fermentation**

be a major limitation that needs to be overcome.

The production of first generation biofuels relies on food crops such as sugar beet, sugar cane, corn, wheat and cassava as substrates for bioethanol; and vegetable oils and animal fats for bio‐ diesel. Although years of intense R&D have made methods of bioethanol production (typically using the yeast *Saccharomyces cerevisiae*) technologically mature, there remain some serious questions regarding its sustainability. The use of food crops as a source of carbohydrate feed‐ stocks by these processes requires high-quality agricultural land. The inevitable conflict be‐ tween the increasing diversion of crops or land for fuel rather than food production has been highlighted as one of the prime causes of rising global food prices. Furthermore, corn ethanol producers in the US, have historically enjoyed a 45-cent-a-gallon federal tax credit for years (which ended in early 2012), costing the government US\$30.5 billion between 2005 to 2011, rais‐

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127

These arguments have stimulated the search for so-called second generation biofuels, which utilize non-food lignocellulose biomass such as wood, dedicated energy crops, agricultural residues and municipal solid wastes as feedstocks. Biomass consists of cellulose, hemicellulose and lignin, and the latter of which is extremely resistant to degradation. One approach to un‐ locking the potential in this abundant feedstock is to separate the lignin from the carbohydrate fraction of the biomass via extensive pre-treatment of the lignocellulose involving, for exam‐ ple, steam-explosion and/or acid hydrolysis. These pre-treatments are designed to allow the carbohydrate portion of the biomass to be broken down into simple sugars, for example by en‐ zymatic hydrolysis using exogenously added cellulases to release fermentable sugars [12]. Such approaches have been found to be expensive and rate limiting [6, 12, 13]. Alternatively, processes using cellulolytic microorganisms (such as *C. cellulolyticum*, *C. thermocellum*, and *C. phytofermentans*) to carry out both the hydrolysis of lignocelluloses and sugar fermentation in a single step, termed 'Consolidated Bioprocessing Process (CBP)' [12] have been proposed, how‐ ever the development of these is still at an early stage, and again low conversion rates seem to

Microorganisms such as acetogens, carboxytrophs and methanogens are able to utilize the CO2 + H2, and/or CO available in such syngas as their sole source of carbon and energy for growth as well as the production of biofuels and other valuable products. However, only acetogens are described to synthesize metabolic end products that have potentials as liquid transportation fuels. While biological processes are generally considered slower than chemical reactions, the use of these microbes to carry out syngas fermentation offers several key advantages over alter‐ native thermo-chemical approaches such as the Fischer-Tropsch' process (FTP). First, microbi‐ al processes operate at ambient temperatures and low pressures which offer significant energy and cost savings. Second, the ambient conditions and irreversible nature of biological reactions also avoid thermodynamic equilibrium relationships and allow near complete conversion effi‐ ciencies [14, 15]. Third, biological conversions are commonly more specific due to high enzy‐ matic specificities, resulting in higher product yield with the formation of fewer by-products. Fourth, unlike traditional chemical catalysts which require a set feed gas composition to yield desired product ratios or suite, microbial processes have freedom to operate for the production ofthe same suite of products across a wider range of CO:H2 ratios in the feed gas [16]. Fifth, bio‐

ing questions about its economic competitiveness with gasoline [10, 11].

The use and production of biofuels has a long history, starting with the inventors Niko‐ laus August Otto and Rudolph Diesel, who already envisioned the use of biofuels such as ethanol and natural oils when developing the first Otto cycle combustion and diesel engines [6]. While fermentative production of ethanol has been used for thousands of years, mainly for brewing beer starting in Mesopotamia 5000 B.C., fermentative produc‐ tion of another potential biofuel butanol, has only been discovered over the last century, but had significant impact. During the World War 1, Chaim Weizmann successfully ap‐ plied a process called ABE (acetone-butanol-ethanol) fermentation using *Clostridium aceto‐ butylicum* to generate industrial scale acetone (for cordites, the propellant of cartridges and shells) from starchy materials [6, 8]. His contribution was later recognised in the Balfour declaration in 1917 and he became the first President of the newly founded State of Israel [6, 8]. Intriguingly, the enormous potential of butanol produced at that time was not realized and the substance was simply stored in huge containers [6]. ABE fer‐ mentation became the second biggest ever biotechnological process (after the ethanol fer‐ mentation process) ever performed, but the low demand of acetone following the conclusion of the war led to closure of all the plants [8]. Although ABE fermentation briefly made a comeback during the Second World War, increasing substrate costs and increasing stable supply of low cost crude oil from the Middle East rendered the tech‐ nology economically unviable. Recently, a resurgence of the technology is underway as some old plants are reopened and new plants are being built or planned in China, the US, the United Kingdom (UK), Brazil, France and Austria [6, 8].

Traditionally sugar substrates derived from food crops such as sugar cane, corn (maize) and sugar beet have been the preferred feedstocks for the production of biofuels. How‐ ever, world raw sugar prices have witnessed significant volatility over the last decade or so, ranging from US\$216/ton in year 2000 to a 30 year high of US\$795/ton in February 2011 due to global sugar deficits and crop shortfall [9]. This has created uncertainty and raised sustainability issues about its use as a feedstock for large scale biofuel production. This review aims to shed light on the use of syngas and industrial waste gas as feed‐ stocks, and the emerging field of gas fermentation to generate not only biofuels, but also other high-value added products. The advantages of gas fermentation over conventional sugar-based fermentation and thermochemical conversions, and their flexibility in utiliz‐ ing a spectrum of feedstocks to generate syngas will be discussed. The biochemistry, ge‐ netic and energetic background of the microorganisms that perform this bioconversion process will be critically examined, together with recent advances in systems biology and synthetic biology that offer growing opportunities to improve biocatalysts in terms of both the potential products that can be produced and their process performance. The key processes such as gasification, bioreactor designs, media formulation, and product recovery will be analysed. Finally, the state of commercialization of gas fermentation will be highlighted and an outlook will be provided.

#### **2. Advantages of gas fermentation**

The production of first generation biofuels relies on food crops such as sugar beet, sugar cane, corn, wheat and cassava as substrates for bioethanol; and vegetable oils and animal fats for bio‐ diesel. Although years of intense R&D have made methods of bioethanol production (typically using the yeast *Saccharomyces cerevisiae*) technologically mature, there remain some serious questions regarding its sustainability. The use of food crops as a source of carbohydrate feed‐ stocks by these processes requires high-quality agricultural land. The inevitable conflict be‐ tween the increasing diversion of crops or land for fuel rather than food production has been highlighted as one of the prime causes of rising global food prices. Furthermore, corn ethanol producers in the US, have historically enjoyed a 45-cent-a-gallon federal tax credit for years (which ended in early 2012), costing the government US\$30.5 billion between 2005 to 2011, rais‐ ing questions about its economic competitiveness with gasoline [10, 11].

These arguments have stimulated the search for so-called second generation biofuels, which utilize non-food lignocellulose biomass such as wood, dedicated energy crops, agricultural residues and municipal solid wastes as feedstocks. Biomass consists of cellulose, hemicellulose and lignin, and the latter of which is extremely resistant to degradation. One approach to un‐ locking the potential in this abundant feedstock is to separate the lignin from the carbohydrate fraction of the biomass via extensive pre-treatment of the lignocellulose involving, for exam‐ ple, steam-explosion and/or acid hydrolysis. These pre-treatments are designed to allow the carbohydrate portion of the biomass to be broken down into simple sugars, for example by en‐ zymatic hydrolysis using exogenously added cellulases to release fermentable sugars [12]. Such approaches have been found to be expensive and rate limiting [6, 12, 13]. Alternatively, processes using cellulolytic microorganisms (such as *C. cellulolyticum*, *C. thermocellum*, and *C. phytofermentans*) to carry out both the hydrolysis of lignocelluloses and sugar fermentation in a single step, termed 'Consolidated Bioprocessing Process (CBP)' [12] have been proposed, how‐ ever the development of these is still at an early stage, and again low conversion rates seem to be a major limitation that needs to be overcome.

Microorganisms such as acetogens, carboxytrophs and methanogens are able to utilize the CO2 + H2, and/or CO available in such syngas as their sole source of carbon and energy for growth as well as the production of biofuels and other valuable products. However, only acetogens are described to synthesize metabolic end products that have potentials as liquid transportation fuels. While biological processes are generally considered slower than chemical reactions, the use of these microbes to carry out syngas fermentation offers several key advantages over alter‐ native thermo-chemical approaches such as the Fischer-Tropsch' process (FTP). First, microbi‐ al processes operate at ambient temperatures and low pressures which offer significant energy and cost savings. Second, the ambient conditions and irreversible nature of biological reactions also avoid thermodynamic equilibrium relationships and allow near complete conversion effi‐ ciencies [14, 15]. Third, biological conversions are commonly more specific due to high enzy‐ matic specificities, resulting in higher product yield with the formation of fewer by-products. Fourth, unlike traditional chemical catalysts which require a set feed gas composition to yield desired product ratios or suite, microbial processes have freedom to operate for the production ofthe same suite of products across a wider range of CO:H2 ratios in the feed gas [16]. Fifth, bio‐ catalysts exhibit a much higher tolerance to poisoning by tars, sulphur and chlorine than inor‐ ganic catalysts [6, 16]. However, some challenges have been identified for syngas fermentation to be commercialized, including gas mass transfer limitations, long retention times due to slow cell growth, and lower alcohol production rates and broth concentrations. Recent progress and development to remedy these issues will be highlighted in this review.

Prior to gasification, biomass generally needs to go through a pre-treatment process encom‐ passing drying, size reduction (e.g. chipping, grinding and chopping), pyrolysis, fractiona‐ tion and leaching depending on the gasifier configuration [22, 23]. This upstream pretreatment process can incur significant capital expense and add to the overall biomass feedstock cost, ranging from US\$16-70 per dry ton [22]. Gasification is a thermo-chemical process that converts carbonaceous materials to gaseous intermediates at elevated tempera‐

The resulting syngas contains mainly CO, CO2, H2 and N2, with varying amounts of CH4, water vapour and trace amount of impurities such as H2S, COS, NH3, HCl, HCN, NOx, phe‐ nol, light hydrocarbons and tar [17, 22, 24]. The composition and amount of impurities of syngas depends on the feedstock properties (e.g. moisture, dust and particle size), gasifier type and operational conditions (e.g. temperature, pressure, and oxidant) [17, 22]. Table 1 summarizes typical composition of syngas and other potential gas streams derived from

Depending on the direction of the flows of carbonaceous fuel and oxidant (air or steam), fixed bed gasifier can be further categorized into updraft or downdraft reactor. In the up‐ draft (counter-current) version of the fixed bed gasifier, biomass enters from the top while gasifying agent from the bottom. The biomass moves down the reactor through zones of

[18]. Although this mode of gasifier is often associated with high tar content in the exit gas, recent advances in tar cracking demonstrated that very low tar level is achievable [31]. The direct heat exchange of the oxidizing agent with the entering fuel feed results in low gas exit temperature and hence high thermal efficiency [18, 23]. The downdraft (co-current) gasifier has very similar design as the updraft reactor, except the carbonaceous fuel and oxidizing agent flow in the same direction. In comparison to the updraft gasifier, the downdraft reac‐ tor has lower tar content in the exit gas but exhibit lower thermal efficiency [23]. Due to the size limitation in the constriction (where most of the gasification occurs) of the reactor, this

In fluidized bed reactor, the carbonaceous fuel is mixed together with inert bed material (e.g. silica sand) by forcing fluidization medium (e.g. air and/or steam) through the reactor. The inert bed facilitates better heat exchange between the fuel materials, resulting in nearly isothermal operation conditions and high feedstock conversion efficiencies [18, 22]. The

limited by the melting point of the bed material [18]. Furthermore, the geometry of the reac‐ tor and excellent mixing properties also means that fluidized bed reactors are suitable for

maximum operating temperature of the gasifier is typically around 800 - 900o

C) and finally oxidation zone (1400o

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129

C)

C, which is

C), gasification (900o

mode of gasifier is considered unsuitable for large scale operation [18].

C), in the presence of an oxidizing agent such as air, steam or oxygen [16, 22].

ture (600-1000o

various sources.

drying (100o

**4. Fixed bed gasifier**

**5. Fluidized bed reactor**

C), pyrolysis (300o

#### **3. Feedstock and gasification**

Due to the flexibility of the microbes to ferment syngas with diverse composition, virtually any carbonaceous materials can be used as feedstock for gasification. Non-food biomass that can be employed as feedstock for gasification includes agricultural wastes, dedicated energy crops, forest residues, and municipal organic wastes, or even glycerol and feathers [16-20]. Biomass is available on a renewable basis, either through natural processes or anthropogen‐ ic activities (e.g. organic wastes). It has been estimated that out of a global energy potential from modern biomass of 250 EJ per year in 2005, only 9 EJ (3.6%) was used for energy gener‐ ation [18]. The use of existing waste streams such as municipal organic waste also differenti‐ ate itself from other feedstocks such as dedicated energy crops because these wastes are available today at economically attractive prices, and they are often already aggregated and require less indirect land use. Alternatively, gasification of non-biomass sources such as coal, cokes, oil shale, tar sands, sewage sludge and heavy residues from oil refining, as well as reformed natural gas are commonly applied as feedstocks for the FTP and can also be used for syngas fermentation [15, 21]. Furthermore, some industries such as steel manufac‐ turing, oil refining and chemical production generate large volume of CO and/or CO2 rich gas streams as wastes. Tapping into these sources using microbial fermentation process es‐ sentially convert existing toxic waste gas streams into valuable commodities such as bio‐ fuels. The overall process of gas fermentation is outlined in Figure 1.

**Figure 1.** Overview of gas fermentation process

Prior to gasification, biomass generally needs to go through a pre-treatment process encom‐ passing drying, size reduction (e.g. chipping, grinding and chopping), pyrolysis, fractiona‐ tion and leaching depending on the gasifier configuration [22, 23]. This upstream pretreatment process can incur significant capital expense and add to the overall biomass feedstock cost, ranging from US\$16-70 per dry ton [22]. Gasification is a thermo-chemical process that converts carbonaceous materials to gaseous intermediates at elevated tempera‐ ture (600-1000o C), in the presence of an oxidizing agent such as air, steam or oxygen [16, 22]. The resulting syngas contains mainly CO, CO2, H2 and N2, with varying amounts of CH4, water vapour and trace amount of impurities such as H2S, COS, NH3, HCl, HCN, NOx, phe‐ nol, light hydrocarbons and tar [17, 22, 24]. The composition and amount of impurities of syngas depends on the feedstock properties (e.g. moisture, dust and particle size), gasifier type and operational conditions (e.g. temperature, pressure, and oxidant) [17, 22]. Table 1 summarizes typical composition of syngas and other potential gas streams derived from various sources.

#### **4. Fixed bed gasifier**

Depending on the direction of the flows of carbonaceous fuel and oxidant (air or steam), fixed bed gasifier can be further categorized into updraft or downdraft reactor. In the up‐ draft (counter-current) version of the fixed bed gasifier, biomass enters from the top while gasifying agent from the bottom. The biomass moves down the reactor through zones of drying (100o C), pyrolysis (300o C), gasification (900o C) and finally oxidation zone (1400o C) [18]. Although this mode of gasifier is often associated with high tar content in the exit gas, recent advances in tar cracking demonstrated that very low tar level is achievable [31]. The direct heat exchange of the oxidizing agent with the entering fuel feed results in low gas exit temperature and hence high thermal efficiency [18, 23]. The downdraft (co-current) gasifier has very similar design as the updraft reactor, except the carbonaceous fuel and oxidizing agent flow in the same direction. In comparison to the updraft gasifier, the downdraft reac‐ tor has lower tar content in the exit gas but exhibit lower thermal efficiency [23]. Due to the size limitation in the constriction (where most of the gasification occurs) of the reactor, this mode of gasifier is considered unsuitable for large scale operation [18].

#### **5. Fluidized bed reactor**

In fluidized bed reactor, the carbonaceous fuel is mixed together with inert bed material (e.g. silica sand) by forcing fluidization medium (e.g. air and/or steam) through the reactor. The inert bed facilitates better heat exchange between the fuel materials, resulting in nearly isothermal operation conditions and high feedstock conversion efficiencies [18, 22]. The maximum operating temperature of the gasifier is typically around 800 - 900o C, which is limited by the melting point of the bed material [18]. Furthermore, the geometry of the reac‐ tor and excellent mixing properties also means that fluidized bed reactors are suitable for up-scaling [18, 22]. Due to these properties, fluidized bed reactor is currently the most com‐ monly used gasifier for biomass feedstock [32]. However, this mode of gasifier is not suita‐ ble for feedstocks with high levels of ash and alkali metals because the melting of these components causes stickiness and formation of bigger lumps, which ultimately negatively affect the hydrodynamics of the reactor [18].

[18]. At operating temperature of 1200-1500o

C, this method is able to convert tars and meth‐

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131

ane, resulting in better syngas quality [18]. Importantly this technology requires the feed‐ stocks to be pulverised into fine particles of ~50 µm before feeding, which is not a major

Acetogens are defined as obligate anaerobes that utilize the reductive acetyl-CoA pathway for the reduction of CO2 to the acetyl moiety of acetyl-coenzyme A (CoA), for the conserva‐ tion of energy, and for the assimilation of CO2 into cell carbon [33]. In addition to the reduc‐ tive acetyl-CoA pathway, four other biological pathways are known for complete autotrophic CO2 fixation: the Calvin cycle, the reductive tricarboxylic acid (TCA) cycle, the 3-hydroxypropionate/malyl-CoA cycle and the 3-hydroxypropionate/4-hydroxybutyrate cy‐ cle [34]. Since the earlier atmosphere of earth was anoxic and the acetyl-CoA pathway is bio‐ chemically the simplest among the autotrophic pathways (the only linear pathway, whereas the other four pathways are cyclic), it has been postulated to be the first autotrophic process on earth [35, 36]. The reductive acetyl-CoA pathway is also known as the 'Wood-Ljungdahl' pathway, in recognition of the two pioneers, Lars G. Ljungdahl and Harland G. Wood, who elucidated the chemical and enzymology of the pathway using *Moorella thermoacetica* (for‐ merly: *Clostridium thermoaceticum*) [35] or CODH/ACS pathway after the key enzyme of the pathway Carbon Monoxide dehydrogenase/Acetyl-CoA synthase. This ancient pathway is diversely distributed among at least 23 different bacterial genera: *Acetitomaculum*, *Acetoa‐ naerobium*, *Acetobacterium*, *Acetohalobium*, *Acetonema*, *Alkalibaculum,* "*Bryantella"*, "*Butyribacte‐ rium",Caloramator*, *Clostridium*, *Eubacterium*, *Holophaga*, *Moorella*, *Natroniella*, *Natronincola*, *Oxobacter*, *Ruminococcus*, *Sporomusa*, *Syntrophococcus*, *Tindallia*, *Thermoacetogenium*, *Thermoa‐ naerobacter*, and *Treponema* [33]. A selection of mesophilic and thermophilic acetogens are presented in Table 2. Acetogens are able to utilize gases CO2 + H2, and/or CO to produce

223 2 2 4 *CO H CH COOH H O H kJ mol* +Þ + 2 75.3 / D =- (1)

2 2 25 <sup>2</sup> 2 6 *CO H C H OH H O* +Þ + 3 97.3 / D =- *H kJ mol* (2)

23 2 4 2 *CO H O CH COOH CO* +Þ + 2 154.9 / D =- *H kJ mol* (3)

2 25 <sup>2</sup> 6 3 *CO H O C H OH CO* +Þ + 4 217.9 / D =- *H kJ mol* (4)

The Acetyl-CoA pathway is essentially a terminal electron-accepting process that assimilates CO2 into biomass [35]. It constitutes an *Eastern* (or Carbonyl) branch and a *Western* (or Meth‐

issue for coal but very difficult and costly for biomass sources [18, 22].

**7. Microbes and biochemistry of gas fermentation**

acetic acid and ethanol according to the following stoichiometries:


Note: NR, not reported

The factors that determine which type of gasifier to employ are scale of operation, feedstock size and composition, tar yield and sensitivity towards ash [18]. Currently, three main types of gasifier are commercially employed: fixed bed, fluidized bed and entrained flow reactors [18].

**Table 1.** Typical composition of syngas and other potential gas streams from various sources

#### **6. Entrained flow reactor**

Entrained flow reactor is the preferred route for large scale gasification of coal, petcoke and refinery residues because of high carbon conversion efficiencies and low tar production [22]. This mode of gasifier does not require inert bed material but relies on feeding the feedstocks co-currently with oxidizing agent at high velocity to achieve a pneumatic transport regime [18]. At operating temperature of 1200-1500o C, this method is able to convert tars and meth‐ ane, resulting in better syngas quality [18]. Importantly this technology requires the feed‐ stocks to be pulverised into fine particles of ~50 µm before feeding, which is not a major issue for coal but very difficult and costly for biomass sources [18, 22].

#### **7. Microbes and biochemistry of gas fermentation**

Acetogens are defined as obligate anaerobes that utilize the reductive acetyl-CoA pathway for the reduction of CO2 to the acetyl moiety of acetyl-coenzyme A (CoA), for the conserva‐ tion of energy, and for the assimilation of CO2 into cell carbon [33]. In addition to the reduc‐ tive acetyl-CoA pathway, four other biological pathways are known for complete autotrophic CO2 fixation: the Calvin cycle, the reductive tricarboxylic acid (TCA) cycle, the 3-hydroxypropionate/malyl-CoA cycle and the 3-hydroxypropionate/4-hydroxybutyrate cy‐ cle [34]. Since the earlier atmosphere of earth was anoxic and the acetyl-CoA pathway is bio‐ chemically the simplest among the autotrophic pathways (the only linear pathway, whereas the other four pathways are cyclic), it has been postulated to be the first autotrophic process on earth [35, 36]. The reductive acetyl-CoA pathway is also known as the 'Wood-Ljungdahl' pathway, in recognition of the two pioneers, Lars G. Ljungdahl and Harland G. Wood, who elucidated the chemical and enzymology of the pathway using *Moorella thermoacetica* (for‐ merly: *Clostridium thermoaceticum*) [35] or CODH/ACS pathway after the key enzyme of the pathway Carbon Monoxide dehydrogenase/Acetyl-CoA synthase. This ancient pathway is diversely distributed among at least 23 different bacterial genera: *Acetitomaculum*, *Acetoa‐ naerobium*, *Acetobacterium*, *Acetohalobium*, *Acetonema*, *Alkalibaculum,* "*Bryantella"*, "*Butyribacte‐ rium",Caloramator*, *Clostridium*, *Eubacterium*, *Holophaga*, *Moorella*, *Natroniella*, *Natronincola*, *Oxobacter*, *Ruminococcus*, *Sporomusa*, *Syntrophococcus*, *Tindallia*, *Thermoacetogenium*, *Thermoa‐ naerobacter*, and *Treponema* [33]. A selection of mesophilic and thermophilic acetogens are presented in Table 2. Acetogens are able to utilize gases CO2 + H2, and/or CO to produce acetic acid and ethanol according to the following stoichiometries:

$$2\text{ CO}\_2 + 4\text{ H}\_2 \rightleftharpoons \text{CH}\_3\text{COOH} + 2\text{ H}\_2\text{O} \qquad \qquad \Delta H = -75.3 \text{ kJ/mol} \tag{1}$$

$$2\text{ CO}\_2 + 6\text{ H}\_2 \rightleftharpoons \text{C}\_2\text{H}\_5\text{OH} + 3\text{ H}\_2\text{O} \qquad\qquad\qquad \Delta H = -97.3\text{ kJ/mol}\tag{2}$$

$$2\text{ }\text{C}\\
\text{O} + 2\text{ }H\_2\text{O} \Rightarrow \text{CH}\_3\text{COOH} + \text{ }2\text{ }\text{CO}\_2\text{O} \qquad\qquad \qquad \Delta H = -154.9 \text{ kJ/mol}\tag{3}$$

$$6CO + 3H\_2O \rightleftharpoons C\_2H\_5OH + 4\text{ CO}\_2 \tag{4} \qquad \qquad \Delta H = -217.9 \text{ kJ/mol} \tag{4}$$

The Acetyl-CoA pathway is essentially a terminal electron-accepting process that assimilates CO2 into biomass [35]. It constitutes an *Eastern* (or Carbonyl) branch and a *Western* (or Meth‐


yl) branch (Figure 2.). The *Western* branch employs a series of enzymes to carry out a sixelectron reduction of CO2 to the methyl group of acetyl-CoA, starting from the conversion of CO2 to formate by formate dehydrogenase. Formyl-H4folate synthase then condenses for‐ mate with H4folate to form 10-formyl-H4folate, which is then converted to 5,10-methenyl-H4folate by a cyclohydrolase. This is followed by a dehydrogenase that reducesmethenyl- to 5,10-methylene-H4hydrofolate, before (6S)-5-CH3-H4folate is formed by methylene-H4folate reductase [37]. A B12-depedent methyltransferase (MeTr) then transfer the methyl group of (6S)-5-CH3-H4folate to corrinoid iron-sulphur protein (CoFeSP) of the bi-functional carbon monoxide dehydrogenase/acetyl-CoA synthase (CODH/ACS) complex [37]. The bi-function‐ al CODH/ACS enzyme complex is formed by two autonomous proteins, an α2β<sup>2</sup> tetramer (CODH/ACS) and a γδ heterodimer (CoFeSP), and the genes are often arranged in an oper‐ on, together with MeTr [37, 38]. In the *Eastern* branch, the CODH component catalyzes the reduction of CO2 to CO. The central molecule, acetyl-CoA, is finally generated when CO, methyl group (bound to CoFeSP) and CoASH are condensed by ACS. Given the pivotal role of CODH/ACS, it is unsurprising that this complex was found to be the most highly ex‐ pressed transcripts under autotrophic conditions in *C. autoethanogenum* [27], and can repre‐ sent up to 2% of the soluble cell protein of an acetogen [39]. CODH/ACS is not unique to acetogenic bacteria, as it is also present in sulphate-reducing bacteria, desulfitobacteria, and

The reducing equivalents required for fixation of CO2 carbon into acetyl-CoA come from the oxidation of molecular hydrogen under chemolithoautotrophic growth, or NADH and re‐ duced ferredoxin under heterotrophic growth [75]. An extensive review by Calusinska *et al.* (2010) highlighted the diversity of ubiquitous hydrogenases that Clostridia possess although only one acetogen *C. carboxidivorans* was included in this study [76], which catalyze the re‐

The direction of the hydrogenase reaction is directed by the redox potential of the compo‐ nents able to interact with the enzyme. Hydrogen evolution occurs when electron donor is available, whereas the presence of electron acceptor results in hydrogen oxidation [77]. Hy‐ drogenases can be classified into three phylogenetically distinct classes of metalloenzymes: [NiFe]-, [FeFe]-, and [Fe]-hydrogenases [76]. In *Methanosarcina barkeri*, the Ech hydrogenase, a [NiFe]-hydrogenase, was demonstrated to oxidize H2 to reduce ferredoxin [78]. During acetoclastic methanogenesis, Ech hydrogenase oxidize ferredoxin to generate H2 [78]. Al‐ though genome analysis revealed the presence of Ech-like hydrogenase in *C. thermocellum, C. phytofermentans*, *C. papyrosolvens*, and *C. cellulolyticum*, their physiological roles remained unknown [76]. Clostridia harbour multiple distinct [FeFe]-hydrogenases, perhaps reflecting their ability to respond swiftly to changing environmental conditions [76]. The monomeric, soluble [FeFe]-hydrogenase of *C. pasteurianum* is one of the best studied. It transfer electrons from reduced ferredoxins or flavodoxins to protons, forming H2 [79]. A trimeric [FeFe]-hy‐ drogenase found in *C. difficile*, *C. beijerinckii*, and *C. carboxidivorans* were hypothesized to

<sup>2</sup> *H He* 2 2 Û + + - (5)

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Archaea (methanogens and Archaeoglobus) [38, 40].

versible oxidation of hydrogen:

**Table 2.** Acetogens

yl) branch (Figure 2.). The *Western* branch employs a series of enzymes to carry out a sixelectron reduction of CO2 to the methyl group of acetyl-CoA, starting from the conversion of CO2 to formate by formate dehydrogenase. Formyl-H4folate synthase then condenses for‐ mate with H4folate to form 10-formyl-H4folate, which is then converted to 5,10-methenyl-H4folate by a cyclohydrolase. This is followed by a dehydrogenase that reducesmethenyl- to 5,10-methylene-H4hydrofolate, before (6S)-5-CH3-H4folate is formed by methylene-H4folate reductase [37]. A B12-depedent methyltransferase (MeTr) then transfer the methyl group of (6S)-5-CH3-H4folate to corrinoid iron-sulphur protein (CoFeSP) of the bi-functional carbon monoxide dehydrogenase/acetyl-CoA synthase (CODH/ACS) complex [37]. The bi-function‐ al CODH/ACS enzyme complex is formed by two autonomous proteins, an α2β<sup>2</sup> tetramer (CODH/ACS) and a γδ heterodimer (CoFeSP), and the genes are often arranged in an oper‐ on, together with MeTr [37, 38]. In the *Eastern* branch, the CODH component catalyzes the reduction of CO2 to CO. The central molecule, acetyl-CoA, is finally generated when CO, methyl group (bound to CoFeSP) and CoASH are condensed by ACS. Given the pivotal role of CODH/ACS, it is unsurprising that this complex was found to be the most highly ex‐ pressed transcripts under autotrophic conditions in *C. autoethanogenum* [27], and can repre‐ sent up to 2% of the soluble cell protein of an acetogen [39]. CODH/ACS is not unique to acetogenic bacteria, as it is also present in sulphate-reducing bacteria, desulfitobacteria, and Archaea (methanogens and Archaeoglobus) [38, 40].

The reducing equivalents required for fixation of CO2 carbon into acetyl-CoA come from the oxidation of molecular hydrogen under chemolithoautotrophic growth, or NADH and re‐ duced ferredoxin under heterotrophic growth [75]. An extensive review by Calusinska *et al.* (2010) highlighted the diversity of ubiquitous hydrogenases that Clostridia possess although only one acetogen *C. carboxidivorans* was included in this study [76], which catalyze the re‐ versible oxidation of hydrogen:

$$2H\_2 \Leftrightarrow 2\ H^+ + \text{ 2 }e^- \tag{5}$$

The direction of the hydrogenase reaction is directed by the redox potential of the compo‐ nents able to interact with the enzyme. Hydrogen evolution occurs when electron donor is available, whereas the presence of electron acceptor results in hydrogen oxidation [77]. Hy‐ drogenases can be classified into three phylogenetically distinct classes of metalloenzymes: [NiFe]-, [FeFe]-, and [Fe]-hydrogenases [76]. In *Methanosarcina barkeri*, the Ech hydrogenase, a [NiFe]-hydrogenase, was demonstrated to oxidize H2 to reduce ferredoxin [78]. During acetoclastic methanogenesis, Ech hydrogenase oxidize ferredoxin to generate H2 [78]. Al‐ though genome analysis revealed the presence of Ech-like hydrogenase in *C. thermocellum, C. phytofermentans*, *C. papyrosolvens*, and *C. cellulolyticum*, their physiological roles remained unknown [76]. Clostridia harbour multiple distinct [FeFe]-hydrogenases, perhaps reflecting their ability to respond swiftly to changing environmental conditions [76]. The monomeric, soluble [FeFe]-hydrogenase of *C. pasteurianum* is one of the best studied. It transfer electrons from reduced ferredoxins or flavodoxins to protons, forming H2 [79]. A trimeric [FeFe]-hy‐ drogenase found in *C. difficile*, *C. beijerinckii*, and *C. carboxidivorans* were hypothesized to couple formate oxidation to reduce protons into H2 [76]. In *Thermotoga maritima*, an electron bifurcating, trimeric [FeFe]-hydrogenase was identified, that was shown to simultaneously oxidize reduced ferredoxin and NADH to evolve hydrogen under low H2 partial pressure [80]. Under high H2 partial pressure, the authors hypothesized that the NADH is oxidized to produce ethanol. *In silico* analysis revealed homologs of this bifurcating hydrogenase in a few Clostridia including *C. beijerinckii* and *C. thermocellum* [80]. In addition to classical hy‐ drogenases, CODH/ACS and pyruvate:ferredoxin oxidoreductase (PFOR) from *M. thermoa‐ cetica* were shown to have hydrogen evolving capability, possibly as a mean of disposing excess reducing equivalents when electron carriers are limited and/or CO concentration is sufficient to inhibit conventional hydrogenases [81].

> **Figure 3.** The organization of genes involved in acetogenesis and energy conservation from sequenced key aceto‐ gens. (A) Wood-Ljungdahl cluster; (B) carbon monoxide dehydrogenase (CODH) cluster; (C) Rnf complex cluster. *acsA*, CODH subunit; *acsB*, ACS subunit; *acsC*, corrinoid iron-sulfur protein large subunit; *acsD*, corrinoid iron-sulfur protein small subunit; *acsE*, methyltransferase subunit; *cooC*, gene for CODH accessory protein; *cooS*, CODH; *fchA*, formimidotetrahydrofolate cyclodeaminase; *fdx*, ferredoxin; *fhs*, formyl-tetrahydrofolate synthase; *folD*, bifunctional methylenetetrahydrofolate dehydrogenase/formyl-tetrahydrofolate cyclohydrolase; *gcvH*, gene for glycine cleavage system H protein; *hyp*, hypothetical protein; *lpdA*, dihydrolipoamide dehydrogenase; *metF*, methylene-tetrahydrofolate reduc‐ tase; *rnfA, rnfB, rnfC, rnfD, rnfE, rnfG,* electron transport complex protein subunits; *rseC,* sigma E positive regulator. ^,

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Most acetogens are also able to utilize another gas carbon monoxide (CO). In contrast to CO2, CO can serve as both a source of carbon btut also as source of electrons such that hy‐ drogen is not necessarily required. With a CO2/CO reduction potential of -524 to -558mV, CO is approximately 1000-fold more capable of generating extremely low potential electrons than NADH, capable of reducing cellular electron carriers such as ferredoxin and flavodoxin [38, 82]. The reducing equivalents generated from CO oxidation can be coupled to reduction of CO2 into acetate, butyrate and/or methane, evolution of molecular hydrogen from pro‐ tons, reduction of nitrate/nitrite, reduction of sulfur species and reduction of aldehydes into alcohols [35, 83]. However, relatively few microorganisms are able to utilize CO as sole car‐ bon and energy source, probably due to growth inhibition from sensitivity of their metallo‐ proteins and hydrogenases towards CO [38, 83]. During exponential growth of *Pseudomonas carboxydovorans* (an aerobic carboxydotroph), it was demonstrated via immunological locali‐ zation studies that 87% of the key enzyme CODH is associated with the inner cytoplasmic membrane, but this association was lost at the end of the exponential growth phase and a reduction in CO-dependent respiration rate was observed [84, 85]. It should be mentioned that aerobic and anaerobic CODH enzymes are structurally very different. CODH has been reported to be a very rapid and efficient CO oxidizer at rates between 4,000 and 40,000 s−1, and reduces CO2 at 11s-1 [86, 87]. Other electron donors commonly used by acetogens in‐ clude formate, CH3Cl, lactate, pyruvate, alcohols, betaine, carbohydrate, acetoin, oxalate and citrate [88]. CODH is able to split water in a biological water-gas shift reaction into hydro‐

2 2 *CO H O CO H e* 2 2 + - + Þ+ + (6)

truncated *acsA*. #, truncated fdx. \*, lack *rseC*.

gen and electron according to the stoichiometry:

**Figure 2.** Wood-Ljungdahl Pathway. Ack, acetate kinase; ACS, acetyl-CoA synthase; CODH, carbon monoxide dehy‐ drogenase; CoFeSP, corrinoid iron sulfur protein; FDH, formate dehydrogenase; Pta, phosphotransacetylase; THF, tet‐ rahydrofolate.

**Figure 3.** The organization of genes involved in acetogenesis and energy conservation from sequenced key aceto‐ gens. (A) Wood-Ljungdahl cluster; (B) carbon monoxide dehydrogenase (CODH) cluster; (C) Rnf complex cluster. *acsA*, CODH subunit; *acsB*, ACS subunit; *acsC*, corrinoid iron-sulfur protein large subunit; *acsD*, corrinoid iron-sulfur protein small subunit; *acsE*, methyltransferase subunit; *cooC*, gene for CODH accessory protein; *cooS*, CODH; *fchA*, formimidotetrahydrofolate cyclodeaminase; *fdx*, ferredoxin; *fhs*, formyl-tetrahydrofolate synthase; *folD*, bifunctional methylenetetrahydrofolate dehydrogenase/formyl-tetrahydrofolate cyclohydrolase; *gcvH*, gene for glycine cleavage system H protein; *hyp*, hypothetical protein; *lpdA*, dihydrolipoamide dehydrogenase; *metF*, methylene-tetrahydrofolate reduc‐ tase; *rnfA, rnfB, rnfC, rnfD, rnfE, rnfG,* electron transport complex protein subunits; *rseC,* sigma E positive regulator. ^, truncated *acsA*. #, truncated fdx. \*, lack *rseC*.

Most acetogens are also able to utilize another gas carbon monoxide (CO). In contrast to CO2, CO can serve as both a source of carbon btut also as source of electrons such that hy‐ drogen is not necessarily required. With a CO2/CO reduction potential of -524 to -558mV, CO is approximately 1000-fold more capable of generating extremely low potential electrons than NADH, capable of reducing cellular electron carriers such as ferredoxin and flavodoxin [38, 82]. The reducing equivalents generated from CO oxidation can be coupled to reduction of CO2 into acetate, butyrate and/or methane, evolution of molecular hydrogen from pro‐ tons, reduction of nitrate/nitrite, reduction of sulfur species and reduction of aldehydes into alcohols [35, 83]. However, relatively few microorganisms are able to utilize CO as sole car‐ bon and energy source, probably due to growth inhibition from sensitivity of their metallo‐ proteins and hydrogenases towards CO [38, 83]. During exponential growth of *Pseudomonas carboxydovorans* (an aerobic carboxydotroph), it was demonstrated via immunological locali‐ zation studies that 87% of the key enzyme CODH is associated with the inner cytoplasmic membrane, but this association was lost at the end of the exponential growth phase and a reduction in CO-dependent respiration rate was observed [84, 85]. It should be mentioned that aerobic and anaerobic CODH enzymes are structurally very different. CODH has been reported to be a very rapid and efficient CO oxidizer at rates between 4,000 and 40,000 s−1, and reduces CO2 at 11s-1 [86, 87]. Other electron donors commonly used by acetogens in‐ clude formate, CH3Cl, lactate, pyruvate, alcohols, betaine, carbohydrate, acetoin, oxalate and citrate [88]. CODH is able to split water in a biological water-gas shift reaction into hydro‐ gen and electron according to the stoichiometry:

$$\text{CO} + H\_2\text{O} \rightleftharpoons \text{CO}\_2 + 2H^+ + 2e^- \tag{6}$$

The operation of this water gas shift reaction is the biochemical basis for the tremendous flexibility that acetogens have in terms of input gas composition. Via this reaction these or‐ ganisms can flexibly use CO or H2 as a source of electrons.Recently, some acetogens such as *C. ljungdahlii*, *C. aceticum*, *M. thermoacetica*, *Sporomusa ovata*, and *S. sphaeroides* have addition‐ ally been shown to utilize electrons derived from electrodes to reduce CO2 into organic com‐ pounds such as acetate, formate, fumarate, caffeine, and 2-oxo-butyrate [89]. Termed microbial electrosynthesis, this nascent concept offers another route for acetogens to harvest the electrons generated from sustainable sources (e.g. solar and wind) to reduce CO2 into useful multi-carbon products such as biofuels [90].

to be coupled with the Rnf complex for additional energy conservation [62]. However, no

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In an attempt to generate an autotrophic *E. coli*, the genes encoding MeTr, the two subunits of CODH/ACS, and the two subunits of CoFeSP from *M. thermoacetica* were cloned and het‐ erologously expressed in *E. coli* [97]. Although the MeTr was found to be active, the other subunits misassembled hence no active enzymes were found [97]. Autotrophic capability is clearly a very complex process that involves many genes other than the CODH/ACS com‐ plex and tetrahydrofolate pathway, including compatible cofactors, electron carriers, specif‐ ic chaperones and energy conservation mechanisms. For instance, more than 200 genes are predicted to be involved in methanogenesis and energy conservation from CO2 and H2 in methanogens [98]. A recent patent application described the introduction of three Wood-Ljungdahl pathway genes encoding MeTr, CoFeSP subunit α and β from *C. difficile* into *C. acetobutylicum* [99]*.* The recombinant strain was shown to incorporate more CO2 into extrac‐

Acetyl-CoA generated via the Wood-Ljungdahl pathway serves as key intermediate for syn‐ thesis of cell mass as well as products. All acetogens are described to produce acetate, in or‐ der to gain energy via SLP to compensate for the energy invested in activating formate in the *Western* branch of the reductive acetyl-CoA pathway. Acetate and ATP are formed via acetyl-phosphate through the successive actions of Pta and Ack. *pta* and *ack* are arranged in the same operon and they were reported to be constitutively expressed [100]. With CO2 and H2 as substrate, only acetate has been observed as major product [44], with minor amounts of ethanol produced in rare cases with *C. ljungdahlii* [101], *C. autoethanogenum* [53], or "*Moor‐ ella sp.*" [102, 103]. Using the more reduced substrate CO, production of a range of other products have been reported, such as ethanol, butanol, butyrate, 2,3-butanediol [104], and lactate (Figure 4.) [105]. From a biofuel perspective, ethanol and butanol are of particular in‐ terest. Ethanol and butanol have even been described as the main fermentation products over acetate in some acetogens under specific conditions. Ethanol producers include *C. ljungdahlii* [62, 63], *C. autoethanogenum* [53], *"C. ragsdalei"* ("*Clostridium* strain P11") [106, 107], "*Moorella sp.*" [102, 103], *Alkalibaculum bacchii* [44], *C. carboxidivorans* ("*Clostridium* strain P7") [54, 55], and *B. methylotrophicum* [49, 108]. The latter two have also been descri‐

Due to historical roles in ABE fermentation, organisms like *C. acetobutylicum*, *C. beijerinckii*, *C. saccharobutylicum*, and *C. saccharoperbutylacetonicum* have been much more extensively characterized than acetogenic Clostridia [95]. Since *C. acetobutylicum* was the first *Clostridium* to be fully sequenced [109] and it remains the most commonly used species for industrial production of solvents to date [110], it provides a model for study of solventogenesis. Al‐ though sugar- and starch-utilizing ABE Clostridia and acetogens exhibit clear distinctions in substrate utilization and thus metabolism, they share some similarities in the biochemical

experimental proof to support this hypothesis has been published to date.

ellular products than wild-type [99].

**8. Products of gas fermentation**

bed to produce butanol.

Under chemolithoautotrophic conditions, acetogenesis must not only fix carbon but also conserve energy. Approximately 0.1 mol of ATP is required for generation of 1g of dry bio‐ mass in anaerobes [82]. Acetyl-CoA is an energy rich molecule that through the combined actions of Pta (phosphotransacetylase) and Ack (acetate kinase), one ATP can be generated via substrate level phosphorylation (SLP). However, the activation of formate to 10-formyl-H4folate in the methyl-branch of Acetyl-CoA pathway consumes one ATP so no net gain in ATP is achieved via this mechanism [35, 75]. Furthermore, the reduction of CO2 to the car‐ bonyl group also requires energy, estimated at one third of ATP equivalent [35]. Recent ad‐ vances indicated that other modes of energy conservation such as electron transport phosphorylation (ETP) or chemiosmotic processes that are coupled to the translocation of protons or sodium ions are implicated in acetogens. Acetogens such as *M. thermoacetica* har‐ bour membrane-associated electron transport system containing cytochrome, menaqui‐ nones, and oxidoreductases that translocate H+ out of the cell [33]. For acetogens that lack such membranous electron transport system, such as *Acetobacterium woodii* and *C. ljungdahlii*, a membrane-bound corrinoid protein is hypothesized to facilitate extrusion of Na+ or pro‐ tons during the transfer of methyl group from methyl-H4F to CODH/ACS [75]. However, all enzymes involved are predicted to be soluble rather than membrane bound. Recent evi‐ dence suggested coupling to an Rnf complex in *A. woodii*, and *C. ljungdahlii* (Figure 3) which acts as ferredoxin:NAD+ -oxidoreductase [62, 91-93]. The Rnf complex is also found in other Clostridia (but not in ABE model organism *C. acetobutylicum*) and bacteria, and was original‐ ly discovered in *Rhodobacter capsulatus* where it is involved in nitrogen fixation [93]. Using reduced ferredoxin (Fd2-) generated from CO oxidation, carbohydrate utilization and/or hy‐ drogenase reactions, this membrane-bound electron transfer complex is predicted to reduce NAD+ with concomitant translocation of Na+ / H+ . The ion gradient generated from the above processes is harvested by H+ - or Na+ - ATP synthase to generate ATP [33, 93]. The recent ge‐ nome sequencing of *A. woodii* revealed that Rnf complex is likely to be the only ion-pump‐ ing enzyme active during autotrophic growth and the organism's entire catabolic metabolism is optimized to maximize the Fd2-/NAD+ ratio [42]. Recently, a third mechanism of energy conservation which involves bifurcation of electrons by hydrogenases was pro‐ posed for anaerobes [94] and demonstrated for enzymes hydrogenase (see above; [80]), bu‐ tyryl-CoA dehydrogenase [94, 95], or an iron-sulfur flavoprotein Nfn [96]. A similar mechanism has also been proposed for the methylene-THF reductase of the reductive acetyl-CoA pathway, which would enable this highly exergonic reduction step (∆G0′ = −22 kJ/mol) to be coupled with the Rnf complex for additional energy conservation [62]. However, no experimental proof to support this hypothesis has been published to date.

In an attempt to generate an autotrophic *E. coli*, the genes encoding MeTr, the two subunits of CODH/ACS, and the two subunits of CoFeSP from *M. thermoacetica* were cloned and het‐ erologously expressed in *E. coli* [97]. Although the MeTr was found to be active, the other subunits misassembled hence no active enzymes were found [97]. Autotrophic capability is clearly a very complex process that involves many genes other than the CODH/ACS com‐ plex and tetrahydrofolate pathway, including compatible cofactors, electron carriers, specif‐ ic chaperones and energy conservation mechanisms. For instance, more than 200 genes are predicted to be involved in methanogenesis and energy conservation from CO2 and H2 in methanogens [98]. A recent patent application described the introduction of three Wood-Ljungdahl pathway genes encoding MeTr, CoFeSP subunit α and β from *C. difficile* into *C. acetobutylicum* [99]*.* The recombinant strain was shown to incorporate more CO2 into extrac‐ ellular products than wild-type [99].

#### **8. Products of gas fermentation**

Acetyl-CoA generated via the Wood-Ljungdahl pathway serves as key intermediate for syn‐ thesis of cell mass as well as products. All acetogens are described to produce acetate, in or‐ der to gain energy via SLP to compensate for the energy invested in activating formate in the *Western* branch of the reductive acetyl-CoA pathway. Acetate and ATP are formed via acetyl-phosphate through the successive actions of Pta and Ack. *pta* and *ack* are arranged in the same operon and they were reported to be constitutively expressed [100]. With CO2 and H2 as substrate, only acetate has been observed as major product [44], with minor amounts of ethanol produced in rare cases with *C. ljungdahlii* [101], *C. autoethanogenum* [53], or "*Moor‐ ella sp.*" [102, 103]. Using the more reduced substrate CO, production of a range of other products have been reported, such as ethanol, butanol, butyrate, 2,3-butanediol [104], and lactate (Figure 4.) [105]. From a biofuel perspective, ethanol and butanol are of particular in‐ terest. Ethanol and butanol have even been described as the main fermentation products over acetate in some acetogens under specific conditions. Ethanol producers include *C. ljungdahlii* [62, 63], *C. autoethanogenum* [53], *"C. ragsdalei"* ("*Clostridium* strain P11") [106, 107], "*Moorella sp.*" [102, 103], *Alkalibaculum bacchii* [44], *C. carboxidivorans* ("*Clostridium* strain P7") [54, 55], and *B. methylotrophicum* [49, 108]. The latter two have also been descri‐ bed to produce butanol.

Due to historical roles in ABE fermentation, organisms like *C. acetobutylicum*, *C. beijerinckii*, *C. saccharobutylicum*, and *C. saccharoperbutylacetonicum* have been much more extensively characterized than acetogenic Clostridia [95]. Since *C. acetobutylicum* was the first *Clostridium* to be fully sequenced [109] and it remains the most commonly used species for industrial production of solvents to date [110], it provides a model for study of solventogenesis. Al‐ though sugar- and starch-utilizing ABE Clostridia and acetogens exhibit clear distinctions in substrate utilization and thus metabolism, they share some similarities in the biochemical pathway and genetic organization of product synthesis and can be used as model for com‐ parison. Structure of key genes and operons (except for the absence of acetone biosynthetic genes) have been found to be very similar in sequenced acetogen *C. carboxidivorans* [54], and in respect of acetate and ethanol genes to some extent also in *C. ljungdahlii* [62]. For instance, the operon structure of *pta-ack*, *ptb-buk* and the *bcs* cluster of acetogen *C. carboxidivorans* are highly similar to starch-utilizing *C. acetobutylicum* and *C. beijerinckii* [54, 109] (Figure 5). Due to these reasons, solventogenic genes from starch-utilizing Clostridia are ideal targets for heterologous expression in acetogens for improvement of product yield and expansion of product range.

> **Figure 5.** Similarity of acidogenesis and butanol formation gene clusters of acetogens and sugar-utilizing Clostridia. (A) Acetate-forming operon; (B) butyrate-forming operon; (C) butanol-forming operon. *ack*, acetate kinase; *buk,* buty‐ rate kinase; *bcd*, butyryl-CoA dehydrogenase; *crt*, crotonase; *etfA,* electron-transferring flavoprotein subunit A; *etfB*, electron-transferring flavoprotein subunit B; *hbd,* 3-hydroxybutyryl-CoA dehydrogenase; *ptb*, phosphotransbutyrylase;

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Similar to sugar- and starch-utilizing ABE Clostridia, acetogens such as *C. carboxidivorans* [111, 112], *C. ljungdahlii* [113], and *C. autoethanogenum* [27] also typically undergo biphasic fermentation under autotrophic conditions. The first phase involves the production of car‐ boxylic acids (acidogenic), H2 and CO2 during exponential growth. This is followed by the solventogenic phase in which part of the produced acids are reassimilated or reduced in‐ to solvents, which usually occurs during stationary growth phase [114]. This shift from acidogenesis to solventogenesis is of industrial importance and several transcriptional analysis on *C. acetobutylicum* [100, 115], and *C. beijerinckii* [116] have been performed to shed light on this process. In both organisms, the onset of solventogenesis coincides with an increase in expression of master sporulation/solventogenesis regulator gene *spo0A*, sol‐ ventogenic genes such as *ald*, *ctfA-ctfB*, and *adc*, as well as down-regulation of chemotaxis/ motility genes [100, 115, 116]. Physiologically, the signals that induce solventogenesis were hypothesized to involve temperature, low pH, high concentrations of undissociated acetic and butyric acids, limiting concentrations of sulphate or phosphate, ATP/ADP ratio

For Clostridia such as acetogen *C. carboxidivorans* [54], which harbour the genes thiolase (*thlA*), 3-hydroxybutyryl-CoA dehydrogenase (*hbd*), crotonase (*crt*) and butyryl-CoA dehy‐ drogenase (*bcd*), the two carbon acetyl-CoA can be converted to four carbon butyryl-CoA [95]. ThlA compete with the activities of Pta, Ald (aldehyde dehydrogenase), and PFOR to condense two acetyl-CoA into one acetoacetyl-CoA, and plays a key role in regulating the C2:C4 acid ratio [110, 118]. Since the formation of acetate yields twice as much ATP per mole of acetyl-CoA relative to butyrate formation, thiolase activity indirectly affects ATP yield [118]. Under physiological conditions, Crt catalyzes dehydration of β-hydroxybutyryl-CoA to crotonyl-CoA [119]. Bcd was shown to require a pair of electron transfer flavoproteins (Et‐ fA and EtfB) to convert crotonyl-CoA to butyryl-CoA [120]. Furthermore, the Bcd was dem‐

*thlA*, thiolase.

and/or NAD(P)H levels [117].

**Figure 4.** Scheme of metabolite production from gas fermentation using native and genetically modified Clostridia. Black denotes well-characterized pathways in Clostridia. Blue shows demonstrated heterologous pathways that have been engineered into Clostridia*.* Purple designates hypothetical pathways that can be engineered into Clostridia*.* Products are highlighted in boxes. Aad, aldehyde/alcohol dehydrogenase; Ack, acetate kinase; Adc, acetoacetate de‐ carboxylase; Adh, alcohol dehydrogenase; Ald, aldehyde dehydrogenase; Aldc, acetolactate decarboxylase; Aor, alde‐ hyde oxidoreductase; Bcd, butyryl-CoA dehydrogenase; Bk, butyrate kinase; Crt, crotonase; CtfA & CtfB, CoA transferase A & B; Etf, electron-transferring flavoprotein; Hbd, hydroxybutyryl-CoA dehydrogenase; IlvA, threonine de‐ aminase; IlvIHCD, valine and isoleucine biosynthesis; Kdc, 2-ketoacid decarboxylase; Ldh, lactate dehydrogenase; LeuABCD, leucine and norvaline biosynthesis; Pfor, Pyruvate ferredoxin oxidoreductase; Pta, phosphotransacetylase; Ptb, phosphotransbutyrylase; Thl, thiolase; 2,3-Bdh, 2,3-butanediol dehydrogenase.

**Figure 5.** Similarity of acidogenesis and butanol formation gene clusters of acetogens and sugar-utilizing Clostridia. (A) Acetate-forming operon; (B) butyrate-forming operon; (C) butanol-forming operon. *ack*, acetate kinase; *buk,* buty‐ rate kinase; *bcd*, butyryl-CoA dehydrogenase; *crt*, crotonase; *etfA,* electron-transferring flavoprotein subunit A; *etfB*, electron-transferring flavoprotein subunit B; *hbd,* 3-hydroxybutyryl-CoA dehydrogenase; *ptb*, phosphotransbutyrylase; *thlA*, thiolase.

Similar to sugar- and starch-utilizing ABE Clostridia, acetogens such as *C. carboxidivorans* [111, 112], *C. ljungdahlii* [113], and *C. autoethanogenum* [27] also typically undergo biphasic fermentation under autotrophic conditions. The first phase involves the production of car‐ boxylic acids (acidogenic), H2 and CO2 during exponential growth. This is followed by the solventogenic phase in which part of the produced acids are reassimilated or reduced in‐ to solvents, which usually occurs during stationary growth phase [114]. This shift from acidogenesis to solventogenesis is of industrial importance and several transcriptional analysis on *C. acetobutylicum* [100, 115], and *C. beijerinckii* [116] have been performed to shed light on this process. In both organisms, the onset of solventogenesis coincides with an increase in expression of master sporulation/solventogenesis regulator gene *spo0A*, sol‐ ventogenic genes such as *ald*, *ctfA-ctfB*, and *adc*, as well as down-regulation of chemotaxis/ motility genes [100, 115, 116]. Physiologically, the signals that induce solventogenesis were hypothesized to involve temperature, low pH, high concentrations of undissociated acetic and butyric acids, limiting concentrations of sulphate or phosphate, ATP/ADP ratio and/or NAD(P)H levels [117].

For Clostridia such as acetogen *C. carboxidivorans* [54], which harbour the genes thiolase (*thlA*), 3-hydroxybutyryl-CoA dehydrogenase (*hbd*), crotonase (*crt*) and butyryl-CoA dehy‐ drogenase (*bcd*), the two carbon acetyl-CoA can be converted to four carbon butyryl-CoA [95]. ThlA compete with the activities of Pta, Ald (aldehyde dehydrogenase), and PFOR to condense two acetyl-CoA into one acetoacetyl-CoA, and plays a key role in regulating the C2:C4 acid ratio [110, 118]. Since the formation of acetate yields twice as much ATP per mole of acetyl-CoA relative to butyrate formation, thiolase activity indirectly affects ATP yield [118]. Under physiological conditions, Crt catalyzes dehydration of β-hydroxybutyryl-CoA to crotonyl-CoA [119]. Bcd was shown to require a pair of electron transfer flavoproteins (Et‐ fA and EtfB) to convert crotonyl-CoA to butyryl-CoA [120]. Furthermore, the Bcd was dem‐ onstrated to form a stable complex with EtfA and EtfB, and they were shown to couple the reduction of crotonyl-CoA to butyryl-CoA with concomitant generation of reduced ferre‐ doxins, which can be used for energy conservation via Rnf complex [94, 119]. Subsequent actions of phosphotransbutyrylase (*ptb*) and butyrate kinase (*buk*) then generate ATP and butyrate from butyryl-CoA [118].

111]. This is likely the result of gene duplication [62]. qRT-PCR analysis from *C. carboxidivor‐ ans* fed with syngas showed that the two *adhE* showed differential expression, and the more abundant *adhE2* was significantly upregulated over 1000 fold in a time span that coincided

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Pyruvate is a central molecule for anabolism and it is predominantly generated from glycol‐ ysis during heterotrophic growth. But under autotrophic growth, this four carbon molecule can be synthesized by PFOR and potentially also the pyruvate-formate lyase (PFL). Two variants of PFOR were reported in *C. autoethanogenum*, and transcriptional analysis showed that they were differentially expressed when grown using industrial waste gases (containing CO, CO2 and H2) [104]. Unlike PFL from most other microorganisms that only catalyze the lysis of pyruvate into formate and acetyl-CoA, clostridial PFL (*C. kluyveri, C. butylicum*, and *C. butyricum*) were reported to readily catalyze the reverse reaction (i.e. pyruvate formation) [129]. Apart from roles in anabolism, pyruvate is also a precursor to other products such as lactic acid and 2,3-butanediol. Small amounts of lactic acid are converted from pyruvate in acetogens, a reaction which is catalyzed by lactate dehydrogenase (Ldh) [104, 118]. Recently, Köpke *et al*. (2011) reported the production of 2mM 2,3-butanediol from acetogenic bacteria (*C. autoethanogenum*, *C. ljungdahlii*, and *C. ragsdalei*) using industrial waste gases (containing CO, CO2 and H2) as feedstock [104]. Pyruvate is first converted into α-acetolactate by the en‐ zyme acetolactate synthase, followed by acetolactate decarboxylase which split acetolactate into acetoin and CO2, before a final reduction of acetoin into 2,3-butanediol by 2,3-butane‐

The genomes of several solventogenic Clostridia, including gas fermenting species, have been sequenced since 2001 [54, 62, 109, 119, 123, 130], and an array of transcriptomic [100, 116, 121, 131, 132], proteomic [132] and systems analysis [133, 134] are being made increas‐ ingly available. However, the generation of stable recombinant Clostridia has been severely hindered by the difficulties encountered introducing foreign DNA into cells and a lack of established genetic tools for this genera of bacteria. In comparison to starch-utilizing Clostri‐ dia, very little information is available for metabolic engineering of acetogens. Although this section describes recent advances in the development of genetic tools for mostly sugar-uti‐ lizing Clostridia, these techniques are highly relevant and applicable to the closely related

The ideal microbial catalyst for industrial scale gas fermentation might exhibit the following traits: high product yield and selectivity, low product inhibition, no strain degeneration, as‐ porogenous, prolonged cell viability, strong aero-tolerance, high biomass density and effi‐ cient utilization of gas substrates. These can be achieved by directed evolution, random mutagenesis and/or targeted genetic engineering. Traditionally, chemical mutagenesis [135-137] and adaption strategies [138, 139] have been deployed to select for these traits. However, these strategies are limited and often come with the expense of unwanted events.

acetogenic Clostridia for biofuels or chemical production via gas fermentation.

with the greatest rate of butanol production [111].

diol dehydrogenase [104] (Figure 4).

**9. Strain improvement and metabolic engineering**

Under low extracellular pH of 4-4.5, the secreted undissociated acetic acid (p*K*<sup>a</sup> 4.79) and/or butyric acid (p*K*a 4.82) diffuse back into cell cytoplasm and then dissociate into the respec‐ tive salts and protons because of the more alkaline intracellular conditions. Without further interventions, the result of this is abolishment of the proton gradient and inevitable cell death [95]. The conversion of acetate and butyrate into solvents increase the pH, thus pro‐ vide some time for the organism to sporulate and secure long term survival. However, the solvents produced are toxic because they increase membrane fluidity and disrupt critical membrane-associated functions such as ATP synthesis, glucose uptake and other transport processes [114, 121]. In *C. acetobutylicum*, it has been demonstrated that the addition of 7-13 g/l of butanol, or up to 40 g/l of acetone and ethanol resulted in 50% growth inhibition [122]. The bacterium is likely to experience a different cytotoxic effect from endogenously pro‐ duced solvents because the organism has time to adapt to increasing amount of solvents.

The reassimilation of acetate and butyrate into the respective acyl-CoA and acetoacetate is catalyzed by acetoacetyl-CoA:acetate/butyrate CoA transferase (CtfA and CtfB) [110, 117, 118]. Acetoacetate is deconstructed by acetoacetate decarboxylase (Adc) into acetone and CO2. This enzyme is missing in acetogenic *C. carboxidivorans* compared to the ABE strains [54, 123]. Some ABE strains such as *C. beijerinckii* NRRL B593 also possess a primary/secon‐ dary alcohol dehydrogenase that converts acetone to isopropanol [124]. In acetogenic "*C. ragsdalei*", reduction of acetone to isopropanol was also observed although the mechanism of this reduction is as yet unknown [124, 125]. Again, *C. carboxidivorans* lacks this activity [125]. The recycled acetyl-CoA and butyryl-CoA can be converted to ethanol and butanol through the actions of coenzyme A-acylating aldehyde dehydrogenase (Ald) and alcohol de‐ hydrogenase (Adh) [110, 118]. Ald converts acyl-CoA into aldehydes, and the enzyme has been purified from *C. beijerinckii* NRRL B593 and was shown to be NADH-specific, exhibit higher affinity with butyraldehyde than acetaldehyde, but possess no Adh activity [126]. In *C. ljungdahlii*, two variants of aldehyde:ferredoxin oxidoreductases (AOR) are present in the genome, and they are hypothesized to couple reduced ferredoxin from CO oxidation via the CODH (see above) to perform the reversible reduction of acetate into acetaldehyde, which can be further reduced into ethanol [62].

The final step of solventogenesis utilizes Adh to reduce acetaldehyde and butyraldehyde in‐ to ethanol and butanol, respectively. For ethanol synthesis, transposon mutagenesis and en‐ zymatic assay in *C. acetobutylicum* showed the involvement of a specific Ald that does not interact with butyryl-CoA, and a NAD(P)H-dependent Adh [127, 128]. The production of butanol by *C. acetobutylicum* is mainly due to the action of butanol dehydrogenase A and B (BdhA and BdhB), and bifunctional butyraldehyde/butanol dehydrogenase 1 and 2 (AdhE1 and AdhE2) [95]. In *C. carboxidivorans* [54] and *C. ljungdahlii* [62] both *adhE1* and *adhE2* are arranged in tandem and separated by a 200bp gap which contains a putative terminator [62, 111]. This is likely the result of gene duplication [62]. qRT-PCR analysis from *C. carboxidivor‐ ans* fed with syngas showed that the two *adhE* showed differential expression, and the more abundant *adhE2* was significantly upregulated over 1000 fold in a time span that coincided with the greatest rate of butanol production [111].

Pyruvate is a central molecule for anabolism and it is predominantly generated from glycol‐ ysis during heterotrophic growth. But under autotrophic growth, this four carbon molecule can be synthesized by PFOR and potentially also the pyruvate-formate lyase (PFL). Two variants of PFOR were reported in *C. autoethanogenum*, and transcriptional analysis showed that they were differentially expressed when grown using industrial waste gases (containing CO, CO2 and H2) [104]. Unlike PFL from most other microorganisms that only catalyze the lysis of pyruvate into formate and acetyl-CoA, clostridial PFL (*C. kluyveri, C. butylicum*, and *C. butyricum*) were reported to readily catalyze the reverse reaction (i.e. pyruvate formation) [129]. Apart from roles in anabolism, pyruvate is also a precursor to other products such as lactic acid and 2,3-butanediol. Small amounts of lactic acid are converted from pyruvate in acetogens, a reaction which is catalyzed by lactate dehydrogenase (Ldh) [104, 118]. Recently, Köpke *et al*. (2011) reported the production of 2mM 2,3-butanediol from acetogenic bacteria (*C. autoethanogenum*, *C. ljungdahlii*, and *C. ragsdalei*) using industrial waste gases (containing CO, CO2 and H2) as feedstock [104]. Pyruvate is first converted into α-acetolactate by the en‐ zyme acetolactate synthase, followed by acetolactate decarboxylase which split acetolactate into acetoin and CO2, before a final reduction of acetoin into 2,3-butanediol by 2,3-butane‐ diol dehydrogenase [104] (Figure 4).

#### **9. Strain improvement and metabolic engineering**

The genomes of several solventogenic Clostridia, including gas fermenting species, have been sequenced since 2001 [54, 62, 109, 119, 123, 130], and an array of transcriptomic [100, 116, 121, 131, 132], proteomic [132] and systems analysis [133, 134] are being made increas‐ ingly available. However, the generation of stable recombinant Clostridia has been severely hindered by the difficulties encountered introducing foreign DNA into cells and a lack of established genetic tools for this genera of bacteria. In comparison to starch-utilizing Clostri‐ dia, very little information is available for metabolic engineering of acetogens. Although this section describes recent advances in the development of genetic tools for mostly sugar-uti‐ lizing Clostridia, these techniques are highly relevant and applicable to the closely related acetogenic Clostridia for biofuels or chemical production via gas fermentation.

The ideal microbial catalyst for industrial scale gas fermentation might exhibit the following traits: high product yield and selectivity, low product inhibition, no strain degeneration, as‐ porogenous, prolonged cell viability, strong aero-tolerance, high biomass density and effi‐ cient utilization of gas substrates. These can be achieved by directed evolution, random mutagenesis and/or targeted genetic engineering. Traditionally, chemical mutagenesis [135-137] and adaption strategies [138, 139] have been deployed to select for these traits. However, these strategies are limited and often come with the expense of unwanted events. First attempts of targeted genetic modification of Clostridia were made in the early 1990s by the laboratory of Prof. Terry Papoutsakis [140-142]. While these pioneering efforts relied on use of plasmids for (over)expression of genes in *C. acetobutylicum*, more sophisticated tools were later developed for a range of solventogenic and pathogenic Clostridia.

**Organism Genetic modification Phenotypes/Effects Ref**

Produced 2 mM butanol from syngas [62]

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Produced 26 mM butanol using steel

Increased alcohol tolerance [168]

Produced up to 140 µM acetone

Produced same amount of butanol as control but relatively more ethanol, corresponding to a total alcohol tolerance of 21.2 g/l

by more than 97% resulted in 80% less acetate produced but similar

Showed 4 fold increase in ethanol yield (122 mM instead of 28 mM)

Generated 8.5 times higher ethanol yield (56.4 mM) than wild type (6.5

Produced 85 mM isopropanol [176]

Produced 216 mM butanol [174]

diverting C4 products

final solvent amount

mM)

[167]

143

[169, 170]

[171]

[172]

[173]

[156]

[175]

mill gas

using gas

biosynthetic genes from *C. acetobutylicum* (*thlA, crt, hbd, bcd,*

biosynthetic genes from *C. acetobutylicum* (*thlA, crt, hbd, bcd*,

operon from *C. acetobutylicum* (*adc,*

*C. acetobutylicum* Inactivation of *hbd* using ClosTron Produced 716 mM ethanol by

overexpression of *adhE2* from *C.*

homologous recombination

dehydrogenase) using ClosTron

acetoneoperon (*adc, ctfA, ctfB*) and primary/secondary *adh* from *C. beijerinckii* NRRL B593

*acetobutylicum*

*C. acetobutylicum* Inactivation of *ack* using ClosTron Reduction in acetate kinase activity

*adhE* and *bdhA*)

*etfA, & etfB*)

*ctfAB, thlA*)

overexpression of *aad*

*C. ljungdahlii* Plasmid overexpression of butanol

*C. autoethanogenum* Plasmid overexpression of butanol

*C. autoethanogenum* Plasmid expression of native *groES* and *groEL*

*C. aceticum* Plasmid overexpression of acetone

*C. acetobutylicum* Inactivation of *buk* and

*C. tyrobutylicum* Inactivation of *ack* and plasmid

*C. thermocellum* Inactivation of *ldh* and *pta* via

*C. cellulolyticum* Inactivation of*ldh* and *mdh* (malate

*C. acetobutylicum* Plasmid overexpression of a synthetic

**Acidogenesis and Solventogenesis**

**Acetogens**

Antisense RNA (asRNA) has been employed to down-regulate genes. Here, single stranded RNA binds to a complementary target mRNA and prevents translation by hindering ribo‐ some-binding site interactions [143]. For instance, this method has been used to knockdown *ctfB* resulting in production of 30 g/l solvents with significantly suppressed acetone yield in *C. acetobutylicum* ATCC 824 [144, 145].

Several homologous recombination methods have been developed for integration or knock-out of genes in a range of sugar-utilizing Clostridia. In early stage, knockout mu‐ tants were almost exclusively generated from single crossover events that could revert back to wild-type [146-152], with stable double crossovers only observed in rare cases [153, 154]. For *C. acetobutylicum* [155] and cellulolytic *C. thermocellum* [156] counter selecta‐ ble markers have been developed to allow more efficient screening for the rare second re‐ combination event.

ClosTron utilizes the specificity of mobile group II intron Ll.*ltrB* from *Lactoccocus lactis* to propagate into a specified site in the genome via a RNA-mediated, retro-homing mecha‐ nism which can be used to disrupt genes [157]. This technique has initially been devel‐ oped by InGex and Sigma-Aldrich under the name 'TargeTron™' and successfully adapted to a range of solventogenic and pathogenic Clostridia including *C. acetobutyli‐ cum*, *C. difficile*, *C. sporogenes*, *C. perfringens*, and *C. botulinum* [158-160] by the laboratory of Prof. Nigel Minton.

The same laboratory recently also developed another method for integration of DNA into the genome. Termed Allele-Coupled Exchange (ACE), this approach does not employ a counter selective marker to select for the rare second recombination event. Rather, it utilizes the activation or inactivation of gene(s) that result in a selectable phenotype, and asymmetri‐ cal homology arms to direct the order of recombination events [161]. Remarkably, the whole genome of phage lambda (48.5kb minus a 6kb region) was successfully inserted into the ge‐ nome of *C. acetobutylicum* ATCC 824 in three successive steps using this genetic tool. This technique was also demonstrated in *C. difficile* and *C. sporogenes* [161].

For reverse engineering, mainly transposon mutagenesis has been utilized. Earlier efforts of transposon mutagenesis were demonstrated in *C. acetobutylicum* P262 (now: *C. saccharobuty‐ licum* [162]), *C. acetobutylicum* DSM792, *C. acetobutylicum* DSM1732, and *C. beijerinckii* NCIM 8052, but issues with multiple transposon insertions per mutant, and non-random distribu‐ tion of insertion were reported [163, 164]. Recent developments have seen the successful generation of mono-copy random insertion of transposon *Tn1545* into cellulolytic *C. cellulo‐ lyticum* [165] and mariner transposon *Himar1* into pathogenic *C. difficile* [166].

While there is still a lack of some other essential metabolic engineering tools such as efficient inducible promoters, the array of available tools that enabled significant improvements to the ABE process and cellulolytic Clostridia fermentations as summarized in Table 3.



Besides the classical Clostridial butanol pathway (which constitutes genes *thlA, crt, hbd, bcd, etfA and etfB*; see earlier section), a non-fermentative approach has been described and dem‐ onstrated in *E. coli* for branched chain higher alcohol production [182]. This alternative ap‐ proach requires a combination of highly active amino acid biosynthetic pathway and artificial diversion of 2-keto acid intermediates into alcohols by introduction of two addi‐ tional genes: broad substrate range 2-keto-acid decarboxylase (*kdc*) which converts 2-keto acids into aldehydes, followed by *Adh* to form alcohols [182]. Engineered strains of *E. coli* have been shown to produce alcohols such as isobutanol, n-butanol, 2-methyl-1-butanol, 3 methyl-1-butanol and 2-phenylethanol via this strategy [182]. For instance, the overexpres‐ sion of *kivD* (KDC from *Lactococcus lactis*)*, adh2, ilvA,* and *leuABCD* operon, coupled with deletion of *ilvD* gene and supplementation of L-threonine, increased n-butanol yield to 9 mM while producing 10 mM of 1-propanol [182].An even more remarkable yield of 300 mM isobutanol was achieved through introduction of k*ivD, adh2, alsS* (from *B. subtilis*)*,* and *ilvCD* into *E. coli* [182]. Like butanol, isobutanol exhibits superior properties as a transporta‐ tion fuel when compared to ethanol [177]. By applying similar strategy into *C. cellulolyticum,* 8.9 mM isobutanol was produced from cellulose when k*ivD, yqhD, alsS, ilvC,* and *ilvD* were overexpressed [177]. This result suggests that such non-fermentative pathway is suitable tar‐ get for metabolic engineering of acetogens for the biosynthesis of branched chain higher al‐ cohols. Via synthetic biology and metabolic engineering, production of additional potential liquid transportation fuels like farnesese or fatty acid based fuels has successfully been dem‐ onstrated in *E. coli* or yeast from sugar [183, 184]. Given the unsolved energetics in aceto‐ gens, it is unclear if production of such energy dense liquid fuels could be viable via gas

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An optimum gas fermentation system requires efficient mass transfer of gaseous substrates to the culture medium (liquid phase) and microbial catalysts (solid phase). Gas-to-liquid mass transfer has been identified as the rate-limiting step and bottleneck for gas fermenta‐ tion because of the low aqueous solubility of CO and H2, respectively at only 77% and 68%

ficient gas-to-liquid mass transfer in an energy-efficient manner at commercial scale for gas fermentation represents a significant engineering challenge. A brief overview of reactor con‐

In continuous stirred tank reactor (CSTR), gas substrates are continuously fed into the reac‐ tor and mechanically sheared by baffled impellers into smaller bubbles, which has greater interfacial surface area for mass transfer [16]. In addition, finer bubbles have a slower rising velocity and a longer retention time in the aqueous medium, resulting in higher gas-to-liq‐

figurations reported in gas fermentation operations is given below.

C [185]. Hence, a bioreactor design that delivers suf‐

fermentation.

**10.1. Bioreactor design**

**10. Fermentation and product recovery**

of that of oxygen (on molar basis) at 35o

**10.2. Continuous Stirred Tank Reactor (CSTR)**

**Table 3.** Genetically modified solventogenic Clostridia

In contrast, to date only a limited number of acetogenic Clostridia have been successfully modified. Pioneering work in this area has been undertaken in the laboratory of Prof. Peter Dürre. *C. ljungdahlii*, a species that does not naturally produce butanol, was modified with butanol biosynthetic genes (*thlA*, *hbd*, *crt*, *bcd*, *adhE* and *bdhA*) from *C. acetobutylicum* ATCC 824 resulting in production of up to 2 mM of butanol using synthesis gas as sole energy and carbon source [62]. By delivering a plasmid with acetone biosynthesis genes *ctfA*, *ctfB*, *adc*, and *thlA* in *C. aceticum,* production of up to 140 µM acetone was demonstrated from various gas mixes (80% H2/20% CO2 and 67% H2/33% CO2) [169, 170]. Recent patent filings by Lanza‐ Tech describe the production of butanol as main fermentation product and increased alcohol tolerance in genetically engineered acetogens. Up to 26 mM butanol were produced with ge‐ netically modified *C. ljungdahlii* and *C. autoethanogenum* using steel mill gas (composition 44% CO, 32% N2, 22% CO2, and 2% H2) as the only source of carbon and energy when the butanol biosynthetic genes *thlA, hbd, crt, bcd, etfA,* and *etfB* were heterologously expressed [167]. Overexpression of native *groESL* operon in *C. autoethanogenum* resulted in a strain that displayed higher alcohol tolerance relative to wild-type when challenged with ethanol [168].

Besides the classical Clostridial butanol pathway (which constitutes genes *thlA, crt, hbd, bcd, etfA and etfB*; see earlier section), a non-fermentative approach has been described and dem‐ onstrated in *E. coli* for branched chain higher alcohol production [182]. This alternative ap‐ proach requires a combination of highly active amino acid biosynthetic pathway and artificial diversion of 2-keto acid intermediates into alcohols by introduction of two addi‐ tional genes: broad substrate range 2-keto-acid decarboxylase (*kdc*) which converts 2-keto acids into aldehydes, followed by *Adh* to form alcohols [182]. Engineered strains of *E. coli* have been shown to produce alcohols such as isobutanol, n-butanol, 2-methyl-1-butanol, 3 methyl-1-butanol and 2-phenylethanol via this strategy [182]. For instance, the overexpres‐ sion of *kivD* (KDC from *Lactococcus lactis*)*, adh2, ilvA,* and *leuABCD* operon, coupled with deletion of *ilvD* gene and supplementation of L-threonine, increased n-butanol yield to 9 mM while producing 10 mM of 1-propanol [182].An even more remarkable yield of 300 mM isobutanol was achieved through introduction of k*ivD, adh2, alsS* (from *B. subtilis*)*,* and *ilvCD* into *E. coli* [182]. Like butanol, isobutanol exhibits superior properties as a transporta‐ tion fuel when compared to ethanol [177]. By applying similar strategy into *C. cellulolyticum,* 8.9 mM isobutanol was produced from cellulose when k*ivD, yqhD, alsS, ilvC,* and *ilvD* were overexpressed [177]. This result suggests that such non-fermentative pathway is suitable tar‐ get for metabolic engineering of acetogens for the biosynthesis of branched chain higher al‐ cohols. Via synthetic biology and metabolic engineering, production of additional potential liquid transportation fuels like farnesese or fatty acid based fuels has successfully been dem‐ onstrated in *E. coli* or yeast from sugar [183, 184]. Given the unsolved energetics in aceto‐ gens, it is unclear if production of such energy dense liquid fuels could be viable via gas fermentation.

#### **10. Fermentation and product recovery**

#### **10.1. Bioreactor design**

An optimum gas fermentation system requires efficient mass transfer of gaseous substrates to the culture medium (liquid phase) and microbial catalysts (solid phase). Gas-to-liquid mass transfer has been identified as the rate-limiting step and bottleneck for gas fermenta‐ tion because of the low aqueous solubility of CO and H2, respectively at only 77% and 68% of that of oxygen (on molar basis) at 35o C [185]. Hence, a bioreactor design that delivers suf‐ ficient gas-to-liquid mass transfer in an energy-efficient manner at commercial scale for gas fermentation represents a significant engineering challenge. A brief overview of reactor con‐ figurations reported in gas fermentation operations is given below.

#### **10.2. Continuous Stirred Tank Reactor (CSTR)**

In continuous stirred tank reactor (CSTR), gas substrates are continuously fed into the reac‐ tor and mechanically sheared by baffled impellers into smaller bubbles, which has greater interfacial surface area for mass transfer [16]. In addition, finer bubbles have a slower rising velocity and a longer retention time in the aqueous medium, resulting in higher gas-to-liq‐ uid mass transfer [24]. Fermentation reactions using *C. ljungdahlii* have been successfully maintained in a 2 litre CSTR under autotrophic conditions for more than a month, while achieving peak ethanol level of 6.5 g/l and CO conversion rate of 93% [186]. The production of 49 g/l of ethanol from gas substrates using *C. ljungdahlii* was demonstrated using CSTR [113]. In another example, a 100 litre stirred tank reactor was demonstrated to produce up to 24.57 g/l ethanol, 9.25 g/l isopropanol and 0.47 g/l n-butanol during a 59-day semi-batch gas fermentation using "*C. ragsdalei*" strain P11 as biocatalysts [112]. An improved version of CSTR incorporates microbubble sparger to generate finer bubbles to achieve higher mass transfer coefficient [187]. Although CSTR offers complete mixing and uniform distribution of gas substrates to the microbes, the high power per unit volume required to drive the stir‐ rer are thought to make this approach economically unviable for commercial scale gas fer‐ mentation systems [187].

**11. Gas fermentation parameters**

the feed gas readily convert into NH4

**11.2. Substrate pressure**

**11.3. Medium formulation**

acetogens such as *C. ljungdahlii* up to 5.2% (v/v) [193].

hanced acetate productivity of *A. woodii* to 7.4g acetate/ l/day [199].

The gas composition and its impurities can have an impact on the productivity of the gas fer‐ mentation process. Greater molar ratio of H2:CO allows greater efficiency in the conversion of the carbon from CO into products such as ethanol, because reducing equivalents are generat‐ ed from oxidation of H2 (rather than CO). However, CO is also a known inhibitor of hydroge‐ nase which can affect utilization of H2 during fermentation. In *B. methylotrophicum*, H2 utilization was inhibited until CO was exhausted [108]. When CO is consumed, acetogens are able to grow using CO2 and H2. Common impurities from biomass gasification or other waste gases are tar, ash, char, ethane, ethylene, acetylene, H2S, NH3 and NO [17, 22, 24, 188].These have been shown to cause cell dormancy, inhibition of hydrogen uptake, low cell growth and shift between acidogenesis and solventogenesis in acetogens [13, 188]. For instance, NH3 from

to inhibit hydrogenase and cell growth of acetogen "*C. ragsdalei*" [189]. A number of strategies to mitigate the impact of such impurities have been proposed, for example installing 0.025 mm filters, or the use of gas scrubbers or cyclones, and improvement in gasification efficiency and scavenging for contaminants in the gas stream using agents such as potassium permanganate, sodium hydroxide or sodium hypochlorite [24, 190-192]. H2S does not have a negative effect on

The partial pressure of syngas components have a major influence on microbial growth and product profiles because the enzymes involved are sensitive to substrate exposure [194]. Due to the low solubility of CO and H2 in water, the growth of dense bacterial cell cultures can face mass transfer limitations, so increasing the partial pressure of gaseous substrates can help alle‐ viate this problem. For instance, studies in which the CO partial pressure (PCO) increased from 0.35 to 2.0 atm showed that this resulted in a 440% increase in maximum cell density, a signifi‐ cant increase in ethanol productivity and a decrease in acetate production in *C. carboxidivorans* strain P7 [195]. In another study involving *C. ljungdahlii*, the increase of PCO from 0.8 to 1.8 atm had a positive effect on ethanol production, and the microbe did not exhibit any substrate in‐ hibition at high PCO [196].In less CO-tolerant microorganisms, the effect of increasing PCO parti‐ al pressure range from non-appreciable in the case of *Rhodospirillum rubrum* [197], to negative impact on doubling time of *Peptostreptococcus productus* (now: *Blautia product*) [194] and *Eubac‐ terium limosum* [198]. Similar to CO, the increase in partial pressure of H2 (pH2) to 1700 mbar en‐

Although acetogens are able of utilizing CO and CO2/H2 as carbon and energy source, other constituents such as vitamins, trace metal elements, minerals and reducing agents are also required for maintenance of high metabolic activity [16, 113]. Studies indicated that forma‐ tion of ethanol in solventogenic Clostridiais non-growth associated and limitation of growth

in the culture media and these ions were recently shown

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+

**11.1. Gas composition**

#### **10.3. Bubble column reactor**

In contrast to CSTR, gas mixing in bubble column reactor is achievable by gas sparging, without mechanical agitation. This reactor configuration has fewer moving parts, and conse‐ quently has a lower associated capital and operational costs while exhibiting good heat and mass transfer efficiencies, making it a good candidate for large scale gas fermentation [17]. However, excessive level of gas inflow for enhanced mixing have been cited as an issue that leads to heterogeneous flow and back-mixing of the gas substrates [16, 17]. *C. carboxidivorans* strain P7 was cultured in a 4 litre bubble column reactor for 20 days using a combination of producer gas and synthetic syngas, generating a peak ethanol concentration of 6 g/l [13].

#### **10.4. Immobilized cell column reactor**

One of the key challenges of gas fermentation is cell density. Immobilization of microbes through crosslinking or adsorption to insoluble biosupport materials and the subsequent packing within the column offers a range of benefits [14]. These include high cell densities, plug flow operation, high mass transfer rate via direct contact between microbe and gas, re‐ duction of retention time, and operation without mechanical agitation [14, 16]. However, channelling issues may arise when the microbe overgrows and completely fill the interstitial space. Due to limitations in column dimensions and packing, this reactor configuration lacks flexibility to operate in various gas fermentation conditions [14, 16].

#### **10.5. Trickle-bed reactor**

Trickle-bed reactor is a gas- or liquid- continuous reactor consisting of packed bed, which liquid culture trickles down through packing media containing suspended or immobilized cells [16, 24, 187]. The gas substrate is delivered either co-currently or counter-currently to the liquid flow, and no mechanical agitation is required [187].In this reactor format, low gas and liquid flow rates are typically applied, generating relatively low pressure drops [187]. Trickle-bed reactor was found to exhibit excellent gas conversion rates and higher produc‐ tivities than CSTR and bubble column reactor [15].

#### **11. Gas fermentation parameters**

#### **11.1. Gas composition**

The gas composition and its impurities can have an impact on the productivity of the gas fer‐ mentation process. Greater molar ratio of H2:CO allows greater efficiency in the conversion of the carbon from CO into products such as ethanol, because reducing equivalents are generat‐ ed from oxidation of H2 (rather than CO). However, CO is also a known inhibitor of hydroge‐ nase which can affect utilization of H2 during fermentation. In *B. methylotrophicum*, H2 utilization was inhibited until CO was exhausted [108]. When CO is consumed, acetogens are able to grow using CO2 and H2. Common impurities from biomass gasification or other waste gases are tar, ash, char, ethane, ethylene, acetylene, H2S, NH3 and NO [17, 22, 24, 188].These have been shown to cause cell dormancy, inhibition of hydrogen uptake, low cell growth and shift between acidogenesis and solventogenesis in acetogens [13, 188]. For instance, NH3 from the feed gas readily convert into NH4 + in the culture media and these ions were recently shown to inhibit hydrogenase and cell growth of acetogen "*C. ragsdalei*" [189]. A number of strategies to mitigate the impact of such impurities have been proposed, for example installing 0.025 mm filters, or the use of gas scrubbers or cyclones, and improvement in gasification efficiency and scavenging for contaminants in the gas stream using agents such as potassium permanganate, sodium hydroxide or sodium hypochlorite [24, 190-192]. H2S does not have a negative effect on acetogens such as *C. ljungdahlii* up to 5.2% (v/v) [193].

#### **11.2. Substrate pressure**

The partial pressure of syngas components have a major influence on microbial growth and product profiles because the enzymes involved are sensitive to substrate exposure [194]. Due to the low solubility of CO and H2 in water, the growth of dense bacterial cell cultures can face mass transfer limitations, so increasing the partial pressure of gaseous substrates can help alle‐ viate this problem. For instance, studies in which the CO partial pressure (PCO) increased from 0.35 to 2.0 atm showed that this resulted in a 440% increase in maximum cell density, a signifi‐ cant increase in ethanol productivity and a decrease in acetate production in *C. carboxidivorans* strain P7 [195]. In another study involving *C. ljungdahlii*, the increase of PCO from 0.8 to 1.8 atm had a positive effect on ethanol production, and the microbe did not exhibit any substrate in‐ hibition at high PCO [196].In less CO-tolerant microorganisms, the effect of increasing PCO parti‐ al pressure range from non-appreciable in the case of *Rhodospirillum rubrum* [197], to negative impact on doubling time of *Peptostreptococcus productus* (now: *Blautia product*) [194] and *Eubac‐ terium limosum* [198]. Similar to CO, the increase in partial pressure of H2 (pH2) to 1700 mbar en‐ hanced acetate productivity of *A. woodii* to 7.4g acetate/ l/day [199].

#### **11.3. Medium formulation**

Although acetogens are able of utilizing CO and CO2/H2 as carbon and energy source, other constituents such as vitamins, trace metal elements, minerals and reducing agents are also required for maintenance of high metabolic activity [16, 113]. Studies indicated that forma‐ tion of ethanol in solventogenic Clostridiais non-growth associated and limitation of growth by reducing availability of carbon-, nitrogen- and phosphate- nutrients shift the balance from acidogenesis to solventogenesis [113, 200, 201]. Optimization of medium formulation for *C. ljungdahlii* through reduction of B-vitamin concentrations and elimination of yeast ex‐ tract significantly enhanced the final ethanol yield to 48 g/l in a CSTR with cell recycling (23 g/l without cell recycling) [113]. Another study by Klasson *et al.* showed thatthe replacement of yeast extract with cellobiose not only increased maximum cell concentration, but also en‐ hanced ethanol yield by 4-fold [14]. Media formulation for *C. autoethanogenum* was investi‐ gated using Plackett-Burman and central composite designs, but only low ethanol yield was recorded overall [202]. In an attempt to reduce the cost of fermentation medium and im‐ prove process economics, 0.5 g/l of cotton seed extract without other nutrient supplementa‐ tion was shown to be a superior medium for *C. carboxidivorans* strain P7 in producing ethanol from syngas fermentation [203]. A recent study showed that increasing concentra‐ tions of trace metal ions such as Ni2+, Zn2+, SeO4 - , WO4 - , Fe2+ and elimination of Cu2+ from medium improved enzymatic activities (FDH, CODH, and hydrogenase), growth and etha‐ nol production in "*C. ragsdalei*" under autotrophic conditions [107].

substrate utilization, growth rate and membrane lipid composition of the acetogens, but also gas substrate availability because gas solubility increases with decreasing temperature [24,

To retain high cell densities in reactor, microbes can be grown as biofilm attached to carrier ma‐ terial. Planktonic cells can be retained in the fermentation broth by installing solid/liquid sepa‐ rators such as membranous ultra-filtration units, spiral wound filtration systems, hollow fibres, cell-recycling membranes and centrifuges [214-216]. The concentrations of solvents from gas fermentation rarely exceed 6% [w/v] so a cost- and energy- efficient product recovery process is required. Furthermore, acetogens also exhibit low resistance towards solvents like ethanol [217, 218] and butanol [219, 220] so an *in situ*/online product recovery system can en‐ hance solvent productivity by decreasing solvent concentrations (and hence toxicity) in the fer‐ mentation broth. Distillation has been the traditional method of product recovery but the associated high energy costs have led to the development of alternative methods such as liq‐

In liquid-liquid extraction, a water-insoluble organic extractant is mixed with the fermenta‐ tion broth [222]. Because solvents are more soluble in the organic phase than in the aqueous phase, they get selectively concentrated in the extractant.Although this technique does not re‐ move water or nutrients from the fermentation broth, some gaseous substrates might be re‐ moved because CO and H2 have much higher solubility in organic solvents than water [222, 223]. Oleyl alcohol has been the extractant of choice due to its relatively non-toxicity [224].

Liquid-liquid extraction is associated with several problems including toxicity to the mi‐ crobes, formation of emulsion, and the accumulation of microbes at the extractant and fer‐ mentation broth interphase [222]. In an attempt to remediate these problems, perstraction was developed and this technique employs membrane to separate the extractant from the fermentation broth. This physical barrier prevent direct contact between the microbe and the toxicity of extractant, but it can also limit the rate of solvent extraction and is susceptible to

In a product recovery technique termed pervaporation, a membrane that directly comes in contact with fermentation broth is used to selectively remove volatile compounds such as ethanol and butanol [219, 222]. The volatile compounds diffuse through the membrane as

uid-liquid extraction, pervaporation, perstraction, and gas stripping [24, 221].

C than at the optimum

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211]. "*C. ragsdalei*"was reported to produce more ethanol at 32°

C [211].

**12. Cell separation and product recovery**

growth temperature of 37°

**12.1. Liquid-liquid extraction**

**12.2. Perstraction**

fouling [219, 221]

**12.3. Pervaporation**

A low redox potential is necessary for strict anaerobes to grow, hence reducing agents such as sodium thioglycolate, ascorbic acid, methyl viologen, benzyl viologen, titanium (III)–cit‐ rate, potassium ferricyanide, cysteine-HCl and sodium sulfide are commonly added to fer‐ mentation medium [14, 16, 204]. Furthermore, the addition of reducing agent directs the electron and carbon flow towards solventogenesis by enhancing the availability of reducing equivalents to form NADH for alcohol production [16, 205]. Excessive addition of reducing agents can cause slower microbial growth due to reduced ATP formation from acetogenesis so it is important to determine the optimum concentration of reducing agents [14, 16]. The sulfur containing gases (e.g. H2S) present in syngas are toxic to chemical catalysts but can be beneficial for microbial catalysts by reducing medium redox potential, stimulate redox sen‐ sitive enzymes such as CODH, and promote alcohol formation [206, 207].

#### **11.4. Medium pH**

Like other organisms, acetogens have a limited range of pH for optimal growth so the pH of the fermentation medium needs to be closely controlled. The extracellular pH directly influ‐ ences the intracellular pH, membrane potential, proton motive force, and consequently sub‐ strate utilization and product profile [208, 209]. In most studies, lowering pH medium divert carbon and electron flow from cell and acid formation towards alcohol production [113, 209-211]. By applying this knowledge, Gaddy and Clausen performed a two-stage CSTR syngas fermentation systems using *C. ljungdahlii* where they set the first reactor at pH 5 to promote cell growth, and pH 4 - 4.5 in the second reactor to induce ethanol production [212]. One recent study with *C. ljungdahlii* showed conflicting results in which cell density and ethanol production were both higher at pH 6.8 when compared to pH 5.5 [213].

#### **11.5. Temperature**

The optimum temperature for mesophilic acetogens are between 30-40° C, while thermophil‐ ic acetogens grow best between 55 and 58° C. The fermentation temperature not only affects substrate utilization, growth rate and membrane lipid composition of the acetogens, but also gas substrate availability because gas solubility increases with decreasing temperature [24, 211]. "*C. ragsdalei*"was reported to produce more ethanol at 32° C than at the optimum growth temperature of 37° C [211].

#### **12. Cell separation and product recovery**

To retain high cell densities in reactor, microbes can be grown as biofilm attached to carrier ma‐ terial. Planktonic cells can be retained in the fermentation broth by installing solid/liquid sepa‐ rators such as membranous ultra-filtration units, spiral wound filtration systems, hollow fibres, cell-recycling membranes and centrifuges [214-216]. The concentrations of solvents from gas fermentation rarely exceed 6% [w/v] so a cost- and energy- efficient product recovery process is required. Furthermore, acetogens also exhibit low resistance towards solvents like ethanol [217, 218] and butanol [219, 220] so an *in situ*/online product recovery system can en‐ hance solvent productivity by decreasing solvent concentrations (and hence toxicity) in the fer‐ mentation broth. Distillation has been the traditional method of product recovery but the associated high energy costs have led to the development of alternative methods such as liq‐ uid-liquid extraction, pervaporation, perstraction, and gas stripping [24, 221].

#### **12.1. Liquid-liquid extraction**

In liquid-liquid extraction, a water-insoluble organic extractant is mixed with the fermenta‐ tion broth [222]. Because solvents are more soluble in the organic phase than in the aqueous phase, they get selectively concentrated in the extractant.Although this technique does not re‐ move water or nutrients from the fermentation broth, some gaseous substrates might be re‐ moved because CO and H2 have much higher solubility in organic solvents than water [222, 223]. Oleyl alcohol has been the extractant of choice due to its relatively non-toxicity [224].

#### **12.2. Perstraction**

Liquid-liquid extraction is associated with several problems including toxicity to the mi‐ crobes, formation of emulsion, and the accumulation of microbes at the extractant and fer‐ mentation broth interphase [222]. In an attempt to remediate these problems, perstraction was developed and this technique employs membrane to separate the extractant from the fermentation broth. This physical barrier prevent direct contact between the microbe and the toxicity of extractant, but it can also limit the rate of solvent extraction and is susceptible to fouling [219, 221]

#### **12.3. Pervaporation**

In a product recovery technique termed pervaporation, a membrane that directly comes in contact with fermentation broth is used to selectively remove volatile compounds such as ethanol and butanol [219, 222]. The volatile compounds diffuse through the membrane as vapour and are then collected by condensation. To facilitate volatilization of permeates into vapour, a partial pressure difference across the membrane is usually maintained by apply‐ ing a vacuum or inert gas (e.g. N2) across the permeate side of the membrane [219]. Polydi‐ methylsiloxane (PDMS) is the current material of choice for the membrane, but other materials such as poly(1-trimethylsilyl-1-propyne) (PTMSP), hydrophobic zeolite mem‐ branes, and composite membranes have also been investigated [225].

tools for expansion of product range, the industry might witness an increasing emphasis on

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Several companies are actively engaged in the development of the gas fermentation technol‐ ogy and some are approaching commercialization. Bioengineering Resources Inc (BRI) founded by Prof. James Gaddy of University of Arkensas, Fayetteville, an early pioneer in the investigation of gas fermentation at scale, was the first company to explore the potential of gas fermentation for industrial bioethanol production. BRI was acquired by chemical company INEOS and rebranded as INEOS Bio (www.ineosbio.com). A pilot-scale facility in Arkansas has been operated since 2003 using several isolates of *C. ljungdahlii* [231] and is building a US\$130 million commercial facility in Florida with its joint venture partner New Planet Energy Florida [232]. The commercial facility is expected to start operation in the sec‐ ond quarter of 2012 and is aiming to generate 8 million gallon of cellulosic ethanol per an‐ num and 6 MW of power to the local communities [232]. INEOS Bio also announced design

Founded in 2006, Coskata Inc. (www.coskata.com) is a US-based company that has reported achieving ethanol yields of 100 gallons per dry ton of wood biomass in a semi-commercial facility in Pennsylvania [234]. The company licensed several microbial strains from the Uni‐ versity of Oklahoma [235], which has filed patents and journal publications for acetogens such as *"C. ragsdalei*" [211, 236, 237] and *C. carboxidivorans* [55, 112]. A patent documenting a new ethanologenic species, "*C. coskatii*" was also recently filed by Coskata [238]. Backed by a conditional US\$250 million loan guarantee from the US Department of Agriculture (USDA), Coskata has announced that it is planning to build a commercial plant with the capacity to produce 55 million gallon fuel grade ethanol per annum in Alabama [234, 239]. While the initial strategy saw biomass as feedstock, the company recently announced its first commer‐ cial plant will be switched to 100% natural gas as feedstock [240]. A planned IPO with the aim to tap into private investors to finance the plant was put on hold [241]. In 2012, Coskata and INEOS Bio were involved in a trade secret dispute which culminated in a settlement that see INEOS Bio receiving US\$2.5 million cash payment, shares and right to receive 2.5%

LanzaTech is a NZ/US based company that has developed a gas fermentation technology to utilize industrial off-gases from steel making and other sources, as well as syngas pro‐ duced from biomass as feedstocks. The company has reported the development of a pro‐ prietary Clostridial biocatalyst that is able to convert the CO-rich waste gas with minimal gas conditioning into bioethanol and the platform chemical 2,3-butanediol. The use of in‐ dustrial off-gases as feedstock not only helps to reduce the carbon footprint of the steelmaking operations but also allows the production of valuable commodities without the costs associated with feedstock gasification. The company has estimated that up to 30 bil‐ lion gallon of bioethanol per year can be produced from the CO-rich off gases produced through steel manufacturers globally [243]. Founded in 2005, LanzaTech has successfully demonstrated bioethanol production at a pilot plant at BlueScope Steel in Glenbrook, NZ, since 2008 and the company has recently started operating its 100,000 gallon bioethanol per year demonstration facility in Shanghai, China, using waste gas collected from an ad‐

the production of high-value commodities in addition to biofuels.

of a second plant, the Seal Sands Biorefinery in Teeside, UK [233].

of future ethanol royalties from Coskata [242].

#### **12.4. Gas stripping**

Gas stripping is an attractive product recovery method for gas fermentation because the exit gas stream from the bioreactor can be used for *in situ*/online product recovery [219]. Following product recovery via condensation, the effluent and gas can be recycled back into the bioreactor. In sugar-based fermentation using *C. beijerinckii* mutant strain BA101, *in situ* gas stripping was shown to improve ABE productivity by 200%, complete sub‐ strate utilization and also complete acid conversion into solvents, when compared to non-integrated process [226].

#### **13. Commercialization**

The growing commercial interests in using gas fermentation as a platform for biofuels pro‐ duction is evident in the recent spike in patent fillings within the field [105]. A 2009 report compared mass and energy conversion efficiencies from a process engineering standpoint between enzymatic hydrolysis fermentation of lignocellulose, syngas fermentation and FTP [227]. The authors concluded that while syngas fermentation offers a range of advantages such as low pretreatment requirement and low energy requirement for bioconversion, the technology is severely limited by low ethanol productivity [227]. Another report document‐ ed the techno-economic analysis of gas fermentation and concluded that the selling price of ethanol using this technology would still be significantly higher than gasoline in 2009 [228]. In contrast, Griffin and Schultz recently compared the production of ethanol from CO-rich gas using thermo-chemical route and biological gas fermentation route [22]. The authors concluded that gas fermentation offers superior fuel yield per volume of biomass feed, car‐ bon conversion to fuel, energy efficiency and lower carbon emissions relative to the thermochemical approach to bioethanol production.

Ethanol and butanol are the most attractive fuel products from current gas fermentation but other by-products such as 2,3-butanediol, acetic acid and butyric acid are also valuable com‐ modities that have the potential to provide significant additional revenue streams, setting off costs for biofuel production. 2,3-butanediol is a high value commodity which can be used to synthesize chemical products such as 1,3-butanediane, methyl ethyl ketone, and gamma butyrolactone, with a combined potential market value of \$43 billion [104]. Acetic acid is an important precursor for synthesis of polymers while butyric acid can be used as a flavouring agent in the food industry [229, 230]. With the development of advanced genetic tools for expansion of product range, the industry might witness an increasing emphasis on the production of high-value commodities in addition to biofuels.

Several companies are actively engaged in the development of the gas fermentation technol‐ ogy and some are approaching commercialization. Bioengineering Resources Inc (BRI) founded by Prof. James Gaddy of University of Arkensas, Fayetteville, an early pioneer in the investigation of gas fermentation at scale, was the first company to explore the potential of gas fermentation for industrial bioethanol production. BRI was acquired by chemical company INEOS and rebranded as INEOS Bio (www.ineosbio.com). A pilot-scale facility in Arkansas has been operated since 2003 using several isolates of *C. ljungdahlii* [231] and is building a US\$130 million commercial facility in Florida with its joint venture partner New Planet Energy Florida [232]. The commercial facility is expected to start operation in the sec‐ ond quarter of 2012 and is aiming to generate 8 million gallon of cellulosic ethanol per an‐ num and 6 MW of power to the local communities [232]. INEOS Bio also announced design of a second plant, the Seal Sands Biorefinery in Teeside, UK [233].

Founded in 2006, Coskata Inc. (www.coskata.com) is a US-based company that has reported achieving ethanol yields of 100 gallons per dry ton of wood biomass in a semi-commercial facility in Pennsylvania [234]. The company licensed several microbial strains from the Uni‐ versity of Oklahoma [235], which has filed patents and journal publications for acetogens such as *"C. ragsdalei*" [211, 236, 237] and *C. carboxidivorans* [55, 112]. A patent documenting a new ethanologenic species, "*C. coskatii*" was also recently filed by Coskata [238]. Backed by a conditional US\$250 million loan guarantee from the US Department of Agriculture (USDA), Coskata has announced that it is planning to build a commercial plant with the capacity to produce 55 million gallon fuel grade ethanol per annum in Alabama [234, 239]. While the initial strategy saw biomass as feedstock, the company recently announced its first commer‐ cial plant will be switched to 100% natural gas as feedstock [240]. A planned IPO with the aim to tap into private investors to finance the plant was put on hold [241]. In 2012, Coskata and INEOS Bio were involved in a trade secret dispute which culminated in a settlement that see INEOS Bio receiving US\$2.5 million cash payment, shares and right to receive 2.5% of future ethanol royalties from Coskata [242].

LanzaTech is a NZ/US based company that has developed a gas fermentation technology to utilize industrial off-gases from steel making and other sources, as well as syngas pro‐ duced from biomass as feedstocks. The company has reported the development of a pro‐ prietary Clostridial biocatalyst that is able to convert the CO-rich waste gas with minimal gas conditioning into bioethanol and the platform chemical 2,3-butanediol. The use of in‐ dustrial off-gases as feedstock not only helps to reduce the carbon footprint of the steelmaking operations but also allows the production of valuable commodities without the costs associated with feedstock gasification. The company has estimated that up to 30 bil‐ lion gallon of bioethanol per year can be produced from the CO-rich off gases produced through steel manufacturers globally [243]. Founded in 2005, LanzaTech has successfully demonstrated bioethanol production at a pilot plant at BlueScope Steel in Glenbrook, NZ, since 2008 and the company has recently started operating its 100,000 gallon bioethanol per year demonstration facility in Shanghai, China, using waste gas collected from an ad‐ jacent steel mill plant owned by its partner Baosteel Group [243, 244]. LanzaTech is plan‐ ning to build a commercial facility with the capacity to produce 50 million gallon of bioethanol per annum in China by 2013 [243]. The recent acquisition of a biorefinery fa‐ cility developed by the US-based gasification technology company Range Fuels in Geor‐ gia, and a milestone signing of its first commercial customer, Concord Enviro Systems (India), highlighted LanzaTech's intention to utilize MSW and lignocellulosic waste as feedstocks for biofuel and chemical production [243, 244].

**Author details**

**References**

2009.

January 2012.

2004;86(5):587-94.

Fung Min Liew, Michael Köpke and Séan Dennis Simpson

[1] Vasudevan PT, Gagnon MD, Briggs MS. Environmentally sustainable biofuels – The case for biodiesel, biobutanol and cellulosic ethanol. In: Singh OV, Harvey SP, edi‐ tors. Sustainable Biotechnology: Sources of Renewable Energy: Springer Science

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[3] European\_Union. Directive 2009/28/EC of the European Parliament and of the Coun‐ cil of 23 April 2009 on the promotion of the use of energy from renewable sources and amending and subsequently repealing Directives 2001/77/EC and 2003/30/EC.

[5] Demirbas A. Political, economic and environmental impacts of biofuels: A review. Applied Energy. 2009 Nov;86:S108-S17. PubMed PMID: WOS:000271170300013.

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LanzaTech NZ Ltd., Parnell, Auckland, New Zealand

+Business Media B.V.; 2010. p. 43-62.

#### **14. Conclusion**

One of the fundamental factors that govern the environmental and economical sustaina‐ bility of biofuel production is feedstock. Through gasification, a spectrum of renewable non-food feedstock such as agricultural wastes, dedicated energy crops, forest residues, and MSW can be converted into syngas. This article presents a detailed examination of gas fermentation technology in capturing the carbon and energy from syngas and pro‐ duce biofuels and chemicals. In comparison to indirect fermentation of lignocellulose via enzymatic hydrolysis, and thermo-chemical FTP, gas fermentation offers several advan‐ tages such as good product yield and selectivity, operation in ambient conditions, high tolerance to gas impurities, and elimination of expensive pre-treatment steps and costly enzymes. Furthermore, some industries such as steel mill, natural gas steam reforming, oil refining and chemical production generate large volumes of CO-rich off-gas. Gas fer‐ mentation can access these existing feedstocks and generate valuable products from these while reducing carbon emissions. Pivotal to gas fermentation is acetogens such as *C. ljungdahlii*, *C. carboxidivorans*, "*C. ragsdalei*" and *C. autoethanogenum,* which are able to me‐ tabolize CO, and CO2/H2 into a range of products such as ethanol, butanol, isopropanol, acetone, 2,3-butanediol, acetic acid and butyric acid. Sustained effort in studying the physiology and biochemistry using advanced molecular techniques such as genomics, transcriptomics, proteomics, metabolomics and systems biology are essential to further the understanding of these microbes. Furthermore, recent advances in Clostridial genetic tools offer endless opportunities to engineer strains that have improved product yield, sub‐ strate utilization, no strain degeneration, and synthesis of new products.

The main challenges associated with commercialization of gas fermentation have been identified as gas-to-liquid mass transfer limitation, product yield, substrate utilization effi‐ ciency, low biomass density and product recovery. Further development of bioreactor is necessary to improve the availability of gas substrates and maintain high cell density for higher productivity. Improvement in integrated product recovery technology is also es‐ sential to lower the costs of product recovery and alleviate product inhibition. Gas fer‐ mentation appears to be mature enough for commercialization since several companies have already demonstrated their technologies at pilot scale and are moving towards com‐ mercialization in the near future.

#### **Author details**

Fung Min Liew, Michael Köpke and Séan Dennis Simpson

LanzaTech NZ Ltd., Parnell, Auckland, New Zealand

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leases.


**Chapter 6**

**The Promising Fuel-Biobutanol**

Hongjuan Liu, Genyu Wang and Jianan Zhang

In recent years, two problems roused peoples' concern. One is energy crisis caused by the depleting of petroleum fuel. The other is environmental issues such as greenhouse effect, global warming, etc. Therefore, renewable sources utilization technology and bioenergy pro‐ duction technology developed fast for solving such two problems. Bioethanol as one of the biofuel has been applied in automobiles with gasoline in different blending proportions (Zhou and Thomson, 2009; Yan and Lin, 2009). Biobutanol is one of the new types of biofuel. It continuously attracted the attention of researchers and industrialists because of its several

Butanol is a four carbon straight chained alcohol, colorless and flammable. Butanol can be mixed with ethanol, ether and other organic solvent. Butanol can be used as a solvent, in cosmetics, hydraulic fluids, detergent formulations, drugs, antibiotics, hormones and vita‐ mins, as a chemical intermediate in the production of butyl acrylate and methacrylate, and additionally as an extract agent in the manufacture of pharmaceuticals. Butanol has a 4-car‐ bon structure and the carbon atoms can form either a straight-chain or a branched structure, resulting in different properties. There exist different isomers, based on the location of the– OH and carbon chain structure. The different structures, properties and main applications

Although the properties of butanol isomers are different in octane number, boiling point, viscosity, etc., the main applications are similar in some aspects, such as being used as sol‐ vents, industrial cleaners, or gasoline additives. All these butanol isomers can be produced from fossil fuels by different methods, only n-butanol, a straight-chain molecule structure

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

© 2013 Liu 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.

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

Additional information is available at the end of the chapter

http://dx.doi.org/10. 5772/52535

**1. Introduction**

distinct advantages.

**1.1. Property of butanol**

are shown as Table 1.

can be produced from biomass.

### **Chapter 6**

### **The Promising Fuel-Biobutanol**

Hongjuan Liu, Genyu Wang and Jianan Zhang

Additional information is available at the end of the chapter

http://dx.doi.org/10. 5772/52535

#### **1. Introduction**

In recent years, two problems roused peoples' concern. One is energy crisis caused by the depleting of petroleum fuel. The other is environmental issues such as greenhouse effect, global warming, etc. Therefore, renewable sources utilization technology and bioenergy pro‐ duction technology developed fast for solving such two problems. Bioethanol as one of the biofuel has been applied in automobiles with gasoline in different blending proportions (Zhou and Thomson, 2009; Yan and Lin, 2009). Biobutanol is one of the new types of biofuel. It continuously attracted the attention of researchers and industrialists because of its several distinct advantages.

#### **1.1. Property of butanol**

Butanol is a four carbon straight chained alcohol, colorless and flammable. Butanol can be mixed with ethanol, ether and other organic solvent. Butanol can be used as a solvent, in cosmetics, hydraulic fluids, detergent formulations, drugs, antibiotics, hormones and vita‐ mins, as a chemical intermediate in the production of butyl acrylate and methacrylate, and additionally as an extract agent in the manufacture of pharmaceuticals. Butanol has a 4-car‐ bon structure and the carbon atoms can form either a straight-chain or a branched structure, resulting in different properties. There exist different isomers, based on the location of the– OH and carbon chain structure. The different structures, properties and main applications are shown as Table 1.

Although the properties of butanol isomers are different in octane number, boiling point, viscosity, etc., the main applications are similar in some aspects, such as being used as sol‐ vents, industrial cleaners, or gasoline additives. All these butanol isomers can be produced from fossil fuels by different methods, only n-butanol, a straight-chain molecule structure can be produced from biomass.

© 2013 Liu 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 Liu 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.


Butanol appeared the good properties compared with it's homologues such as 2-butanol, iso-butanol and tert-butanol and other fuels such as Gasoline and ethanol. Actually, when ethanol is mixed with gasoline (less than 10%), there exists some disadvantages. Firstly, the heating value of ethanol is one sixth of gasoline. The fuel consumption will increase 5% if the engine is not retrofitted. Secondly, acetic acid will be produced during the burning proc‐ ess of ethanol, which is corrosive to the materials of vehicle. The preservative must be added when the ethanol proportion upper than 15%. Thirdly, ethanol is hydroscopic and the liquid phase separation may be occurring with high water proportion. Furthermore, ethanol as fuel cannot be preserved easily and it is more difficult in the process of allocation, storage, transi‐

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 177

Compared with ethanol, butanol overcomes above disadvantages and it shows potential ad‐ vantages. For example, Butanol has higher energy content and higher burning efficiency, which can be used for longer distance. The air to fuel ratio and the energy content of butanol are closer to gasoline. So, butanol can be easily mixed with gasoline in any proportion. Buta‐ nol is less volatile and explosive, has higher flash point, and lower vapor pressure, which makes it safer to handle and can be shipped through existing fuel pipelines. In addition, Bu‐ tanol can be used directly or blended with gasoline or diesel without any vehicle retrofit

Actually, the first-time synthesis of biobutanol at laboratory level was reported by Pasteur in 1861 (Durre, 1998) and the industrial synthesis of biobutanol was started during 1912– 1914 by fermentation (Jones and Woods, 1986). However, before 2005, butanol was mainly used as solvent and precursor of other chemicals due to the product inhibition and low bu‐ tanol productivity. To bring awareness to butanol's potential as a renewable fuel, David Ra‐ mey drove his family car from Ohio to California on 100% butanol (http:// www.consumerenergyreport.com /2011/02/09/reintroducing-butanol/). And then, two giant companies DuPont and BP have declared to finance development of a modernize produc‐ tion plant supported by research and development. (http://biomassmagazine.com/articles/ 2994 /eu-approves-bp-dupont-biobutanol-venture) The economy of biobutanol production also was revaluated. The research of a continuous fermentation pilot plant operating in Aus‐ tria in the 1990s introduced new technologies and proved economic feasibility with agricul‐

Butanol can be obtained using chemical technologies, such as Oxo-synthesis and aldol con‐ densation. It is also possible to produce butanol in the process of fermentation by bacteria and butanol as one of the products called biobutanol. The most popular bacteria species used for fermentation is *Clostridium acetobutylicum*. Because the main products of this proc‐ ess containing acetone, butanol and ethanol, the fermentation is called ABE fermentation

tion than that of gasoline.

(Durre, 2007; Pfromm et al., 2010).

tural waste potatoes. (Nimcevic and Gapes, 2000).

**2. Production methods of butanol**

(Qureshi and Maddox, 1995).

**Table 1.** Structures, properties and main applications of n-butanol, 2-Butanol, iso-Butanol and tert-Butanol

#### **1.2. Advantages of butanol as fuel**

Except the use of solvent, chemical intermediate and extract agent, butanol also can be used as fuel, which attracted people's attention in recent years. Because of the good properties of high heat value, high viscosity, low volatility, high hydrophobicity, less corrosive, butanol has the potential to be a good fuel in the future. The properities of butanol and other fuels or homologues are compared as Table 2. (Freeman et al., 1988; Dean, 1992)


**Table 2.** Properities of butanol and other fuels

Butanol appeared the good properties compared with it's homologues such as 2-butanol, iso-butanol and tert-butanol and other fuels such as Gasoline and ethanol. Actually, when ethanol is mixed with gasoline (less than 10%), there exists some disadvantages. Firstly, the heating value of ethanol is one sixth of gasoline. The fuel consumption will increase 5% if the engine is not retrofitted. Secondly, acetic acid will be produced during the burning proc‐ ess of ethanol, which is corrosive to the materials of vehicle. The preservative must be added when the ethanol proportion upper than 15%. Thirdly, ethanol is hydroscopic and the liquid phase separation may be occurring with high water proportion. Furthermore, ethanol as fuel cannot be preserved easily and it is more difficult in the process of allocation, storage, transi‐ tion than that of gasoline.

Compared with ethanol, butanol overcomes above disadvantages and it shows potential ad‐ vantages. For example, Butanol has higher energy content and higher burning efficiency, which can be used for longer distance. The air to fuel ratio and the energy content of butanol are closer to gasoline. So, butanol can be easily mixed with gasoline in any proportion. Buta‐ nol is less volatile and explosive, has higher flash point, and lower vapor pressure, which makes it safer to handle and can be shipped through existing fuel pipelines. In addition, Bu‐ tanol can be used directly or blended with gasoline or diesel without any vehicle retrofit (Durre, 2007; Pfromm et al., 2010).

Actually, the first-time synthesis of biobutanol at laboratory level was reported by Pasteur in 1861 (Durre, 1998) and the industrial synthesis of biobutanol was started during 1912– 1914 by fermentation (Jones and Woods, 1986). However, before 2005, butanol was mainly used as solvent and precursor of other chemicals due to the product inhibition and low bu‐ tanol productivity. To bring awareness to butanol's potential as a renewable fuel, David Ra‐ mey drove his family car from Ohio to California on 100% butanol (http:// www.consumerenergyreport.com /2011/02/09/reintroducing-butanol/). And then, two giant companies DuPont and BP have declared to finance development of a modernize produc‐ tion plant supported by research and development. (http://biomassmagazine.com/articles/ 2994 /eu-approves-bp-dupont-biobutanol-venture) The economy of biobutanol production also was revaluated. The research of a continuous fermentation pilot plant operating in Aus‐ tria in the 1990s introduced new technologies and proved economic feasibility with agricul‐ tural waste potatoes. (Nimcevic and Gapes, 2000).

#### **2. Production methods of butanol**

Butanol can be obtained using chemical technologies, such as Oxo-synthesis and aldol con‐ densation. It is also possible to produce butanol in the process of fermentation by bacteria and butanol as one of the products called biobutanol. The most popular bacteria species used for fermentation is *Clostridium acetobutylicum*. Because the main products of this proc‐ ess containing acetone, butanol and ethanol, the fermentation is called ABE fermentation (Qureshi and Maddox, 1995).

#### **2.1. Chemical process**

Butanol can be produced by chemical synthesis. One process is Oxo-synthesis, which in‐ volves the reaction of propylene with carbon monoxide and hydrogen in the presence of co‐ balt or rhodium as the catalyst. The mixture of n-butyraldehyde and isobutyraldehyde are obtained and then the mixture can be hydrogenated to the corresponding n-butanol and iso‐ butyl alcohols (Park, 1996).The reactions are as following:

$$\text{CH}\_3\text{CH-CH}\_2\text{+CO} + \text{H}\_2 \rightarrow \text{CH}\_3\text{CH}\_2\text{CH}\_2\text{CHO} \star \text{(CH}\_3\text{)}\_2\text{CHCHO} \tag{1}$$

$$\begin{array}{l}\text{CH}\_3\text{CH}\_2\text{CH}\_2\text{CHO} + \text{H}\_2 \rightarrow \text{CH}\_3\text{CH}\_2\text{CH}\_2\text{CH}\_2\text{OH} \text{ (a)}\\\text{B}\begin{pmatrix}\text{CH}\_3\end{pmatrix}\_2\text{CHCHO} \text{+H}\_2 \rightarrow \begin{pmatrix}\text{CH}\_3\end{pmatrix}\_2\text{CHCH}\_2\text{OH}\end{array}\tag{2}$$

straw, corn core, switch grass, etc. Furthermore, biological process has high product se‐ lectivity, high security, less by-products. Furthermore, the fermentation condition of buta‐ nol production is milder than that of chemical ways and the products are easier to separate. The process of biobutanol production with Lignocellulosic feedstocks is as fol‐

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 179

For the first step, biomass containing lignocellulosics should be pretreated before they were used as the substrate for the fermentation, except for a few high cellulase activity strains (Ezeji and Blaschek, 2008). The pretreatment methods are different according to the different types of biomass used. There often use dilute sulfuric acid pretreatment, alkaline peroxide pretreatment, steam explosion pretreatment, hydrothermal pretreatment, organic acid pre‐ treatment etc. Some inhibitors such as acetic acid, furfural, 5- hydroxymethyl furfural, phe‐ nols etc. that need to be further detoxified. The ordinary detoxification methods are using activated charcoal (Wang et al., 2011), overliming (Sun and Liu, 2012; Park et al., 2010), elec‐ trodialysis (Qureshi et al., 2008c), membrane extraction (Grzenia et al., 2012) to remove the inhibitors. This step is determined by different feed stock and different pretreatment meth‐ ods. After the fermentation, the desired product is recovered and purified in the down‐ stream process. Biological ways has been set up for many years while it was inhibited for industrial application for economic reasons. So, as an alternative fuel, biomass feedstock for biobutanol production must be widely available at low cost (Kent, 2009). Therefore, by using agricultural wastes for butanol production such as straw, leaves, grass, spoiled grain and fruits etc are much more profitable from an economic point of view. Recently, other sources

lowing (Fig. 1):

**Figure 1.** Butanol production process from lignocellulosic feedstocks

When using cobalt as the catalyst, the reaction processes at 10∼20MPa and 130∼160°°C, the products ratio of n-butyraldehyde and isobutyraldehyde is 3. Rhodium as the catalyst used in industry from 1976 and the reaction processes at 0.7-3MPa and 80-120°°C.The products ratio of n-butyraldehyde and isobutyraldehyde can reach 8-16. Hydrogenaration processes by using the catalyst of nickel or copper in gaseous phase or nickel in liquid phase. Some byproducts can be transferred into butanol at high temperature and high pressure that will en‐ hance the product purity.

Another route is aldol condensation, which involves the reaction of condensation and dehy‐ dration from two molecules of acetic aldehyde. And then, the product crotonaldehyde was transformed into n-butanol by hydrogenation at 180°°C and 0.2MPa. The reaction is as fol‐ lowing: CH3CH=CHCHO+2H2─→CH3CH2CH2CH2OH

Comparing the two processes, Oxo-synthesis route has the advantages of materials easily obtained, comparable moderate reaction conditions, enhanced ratio of n-butanol to isobutyl alcohol. So, Oxo-synthesis process is the main industrial route for n-butanol production. There are also some other fossil oil derived raw materials such as ethylene, propylene and triethylaluminium or carbon monoxide and hydrogen are used in butanol production (Zver‐ lov, et al., 2006).

#### **2.2. Biological process**

Except the chemical ways, butanol can also be obtained from biological ways with the re‐ newable resources by the microorganism through fermentation. The *Clostridia* genus is very common for butanol synthesis under anaerobic conditions, and the fermentation products are often the mixture of butanol, acetone and ethanol. A few kinds of *Clostridium* can utilize cellulose and hemicellulose with the ability of cellulolytic activities (Mitchell et al., 1997; Be‐ rezina et al. 2009).

Compared with the chemical ways for butanol production, biological ways has the dis‐ tinct advantages. For example, it can utilize the renewable resources such as wheat straw, corn core, switch grass, etc. Furthermore, biological process has high product se‐ lectivity, high security, less by-products. Furthermore, the fermentation condition of buta‐ nol production is milder than that of chemical ways and the products are easier to separate. The process of biobutanol production with Lignocellulosic feedstocks is as fol‐ lowing (Fig. 1):

**Figure 1.** Butanol production process from lignocellulosic feedstocks

For the first step, biomass containing lignocellulosics should be pretreated before they were used as the substrate for the fermentation, except for a few high cellulase activity strains (Ezeji and Blaschek, 2008). The pretreatment methods are different according to the different types of biomass used. There often use dilute sulfuric acid pretreatment, alkaline peroxide pretreatment, steam explosion pretreatment, hydrothermal pretreatment, organic acid pre‐ treatment etc. Some inhibitors such as acetic acid, furfural, 5- hydroxymethyl furfural, phe‐ nols etc. that need to be further detoxified. The ordinary detoxification methods are using activated charcoal (Wang et al., 2011), overliming (Sun and Liu, 2012; Park et al., 2010), elec‐ trodialysis (Qureshi et al., 2008c), membrane extraction (Grzenia et al., 2012) to remove the inhibitors. This step is determined by different feed stock and different pretreatment meth‐ ods. After the fermentation, the desired product is recovered and purified in the down‐ stream process. Biological ways has been set up for many years while it was inhibited for industrial application for economic reasons. So, as an alternative fuel, biomass feedstock for biobutanol production must be widely available at low cost (Kent, 2009). Therefore, by using agricultural wastes for butanol production such as straw, leaves, grass, spoiled grain and fruits etc are much more profitable from an economic point of view. Recently, other sources such as algae culture (Potts et al., 2012; Ellis et al., 2012) also is studied as one substrate for butanol production.

#### **3. Biobutanol production by fermentation**

#### **3.1. Microbes**

*Clostridium* is a group of obligate, Gram positive, endospore-forming anaerobes. There are lots of strains used for ABE fermentation in different culture collections, such as ATCC (American Type Culture Collection), DSM (German Collection of Microorganisms, or Deut‐ sche Sammlung Von Mikroorganismen), NCIMB (National Collections of Industrial & Ma‐ rine Bactria Ltd), and NRRL (Midwest Area National Center for Agriculture Utilization Research, US Department of Agriculture). The different strains share similar phonotype such as main metabolic pathway and end products. Molecular biology technology offers ef‐ ficient method for classification. The butanol-producing clostridium can be assigned to four groups according to their genetic background, named *C. acetobutylicum, C. beijerinckii, C. sac‐ charoperbutyl acetonicum*, and *C. saccharobutylicum*, respectively. *C. acetobutylicum* is phyloge‐ netically distinct from the other three groups.

**Figure 2.** Wood-Ljungdahl pathway in Carboxdivorans Strain P7T. (Bruant et al. 2010, http://creativecommons. org/ licenses/by/3. 0/)Wood-Ljungdahl pathway key enzymes and protein identified in C. carboxidivorans strain P7T. 1, for‐ mate dehydrogenase; 2, formate-tetrahydrofolate ligase; 3 and 4, bifunctionalmethenyl-tetrahydrofolatecyclohydro‐ lase/methylene-tetrahydrofolate dehydrogenase (NADP+); 5, 5, 10-methylene-tetrahydrofolate reductase; 6, 5 methyl-tetrahydrofolate:- corrinoid iron-sulfur protein methyltransferase; 7, carbon monoxide dehydrogenase; 8, acetyl-CoA synthase; CFeSP, corrinoid iron-sulfur protein; CODH, additional carbon monoxide dehydrogenase com‐ plex. Reactions from the western branch are indicated in blue, those from the eastern branch are indicated in red. The

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 181

The ABE producing strains can hydrolyze starch to glucose or other hexose by amylases. Glucose was firstly converted to pyruvate through the Embden-Meyerhoff pathway (EMP, or glycolysis). Pyruvate was then cleaved to acetyl-CoA by pyruvate ferredoxin oxidoreduc‐ tase. Acetyl-CoA is the common precursor of all the fermentation intermediate and end products. The enzyme activity and the coding genes have been widely assayed and descri‐

The ABE fermentation process can be divided into two successive and distinct phase as acidogenesis phase and solvetogenesis phase. The acidogenesis phase is accompanied with cell exponential growth and pH drop, accumulation of acetate and butyrate. Solventogenesis phase begins with endospore forming and the cells entering stationary state. The products of acidogenesis phase include acetate and butyrate. Acetate forms from Acetyl-CoA, which is catalyzed by two enzymes, phosphotransacetylase (PTA, or phosphate acetyltransferase, endoced by *pta* gene) and acetate kinase (AK, encoded by *ak* gene). The butyrate synthesis is a little complicated with more steps. At first, two molecular of acetyl-CoA is catalyzed by thiolase (thl, or acetyl-CoA acetyltransferase, encoded by *thl* gene) and transforms into one molecular C4 unit acetoacetyl-CoA, which is another important node and precursor of buty‐

bed in butanol-producing strains (Dürre et al., 1995; Gheshlaghi et al., 2009).

corresponding genes in strain P7T genome are indicated below the enzyme.

**3.2. Metabolic pathway**

The common substrate for the solvent production by these strains is soluble starch. The orig‐ inal starch-fermenting strains belong to *C. acetobutylicum*. A recently isolated butanol-pro‐ ducing strain *C. saccharobutylicum* showed high hemicellulotic activity (Berezina et al., 2009). All of the four group strains can ferment glucose-containing medium to produce solvent. In 4% glucose TYA medium, *C. beijerinckii* gave the lowest solvent yield (28%), while the sol‐ vent yield was upper than 30% compared to the other three groups (Shaheen et al., 2000). In standard supplement maize medium (SMM), *C. acetobutylicum* is the best strain for maize fermentation, and the total solvent concentration can reach 19g/L. The solvent yield was 16, 14, and 11 for that of *C. beijerinckii, C. saccharoperbutyl acetonicum*, and *C. saccharobutylicum* respectively. However, *C. acetobutylicum* can't ferment molasses well and it produces bright yellow riboflavin in milk, which is different from other groups and easy identified. The best molasses-fermenting strains belong to *C. saccharobutylicum* and *C. beijerinckii* (Shaheen et al., 2000). *C. saccharoperbutyl acetonicum* can utilize sugar, molasses and maize. Comparing to *C. acetobutylicum*, *C. beijerinckii* was more tolerant to acetic acid and formic acid (Cho et al., 2012), which suggests the advantage when using lignocellulosic hydrolysate treated with acetic and formic acid as substrate.

There are also some *C. beijerinckii* strains produce isopropanol instead of acetone (George et al., 1983). Some microorganisms can produce biobutanol from carbon monoxide (CO) and molecular hydrogen (H2), including acetogens, *Butyribacterium methylotrophicum*, *C. autoetha‐ nogenum*, *C. ljungdahlii* and *C. carboxidiworans*. The *C. carboxidivorans strain P7(T)* genome possessed a complete Wood-Ljungdahl pathway gene cluster which is responsible for CO, hydrogen fixation and conversion to acetyl-CoA(Fig.2) (Bruant et al., 2010).

**Figure 2.** Wood-Ljungdahl pathway in Carboxdivorans Strain P7T. (Bruant et al. 2010, http://creativecommons. org/ licenses/by/3. 0/)Wood-Ljungdahl pathway key enzymes and protein identified in C. carboxidivorans strain P7T. 1, for‐ mate dehydrogenase; 2, formate-tetrahydrofolate ligase; 3 and 4, bifunctionalmethenyl-tetrahydrofolatecyclohydro‐ lase/methylene-tetrahydrofolate dehydrogenase (NADP+); 5, 5, 10-methylene-tetrahydrofolate reductase; 6, 5 methyl-tetrahydrofolate:- corrinoid iron-sulfur protein methyltransferase; 7, carbon monoxide dehydrogenase; 8, acetyl-CoA synthase; CFeSP, corrinoid iron-sulfur protein; CODH, additional carbon monoxide dehydrogenase com‐ plex. Reactions from the western branch are indicated in blue, those from the eastern branch are indicated in red. The corresponding genes in strain P7T genome are indicated below the enzyme.

#### **3.2. Metabolic pathway**

The ABE producing strains can hydrolyze starch to glucose or other hexose by amylases. Glucose was firstly converted to pyruvate through the Embden-Meyerhoff pathway (EMP, or glycolysis). Pyruvate was then cleaved to acetyl-CoA by pyruvate ferredoxin oxidoreduc‐ tase. Acetyl-CoA is the common precursor of all the fermentation intermediate and end products. The enzyme activity and the coding genes have been widely assayed and descri‐ bed in butanol-producing strains (Dürre et al., 1995; Gheshlaghi et al., 2009).

The ABE fermentation process can be divided into two successive and distinct phase as acidogenesis phase and solvetogenesis phase. The acidogenesis phase is accompanied with cell exponential growth and pH drop, accumulation of acetate and butyrate. Solventogenesis phase begins with endospore forming and the cells entering stationary state. The products of acidogenesis phase include acetate and butyrate. Acetate forms from Acetyl-CoA, which is catalyzed by two enzymes, phosphotransacetylase (PTA, or phosphate acetyltransferase, endoced by *pta* gene) and acetate kinase (AK, encoded by *ak* gene). The butyrate synthesis is a little complicated with more steps. At first, two molecular of acetyl-CoA is catalyzed by thiolase (thl, or acetyl-CoA acetyltransferase, encoded by *thl* gene) and transforms into one molecular C4 unit acetoacetyl-CoA, which is another important node and precursor of buty‐ rate, acetone, and butanol synthesis. The acetoacetyl-CoA is subjected to three enzymes in turn and another C4 unit butyryl-CoA is the intermediate product. The three enzymes are hydroxybutyryl-CoA dehydrogenase (encoded by *hbd* gene) (Youngleson et al., 1995), croto‐ nase (CRT, or hydroxybutyryl-CoA dehydrolase, encoded by *crt* gene), and butyryl-CoA de‐ hydrogenase (BCD, encoded by *bcd* gene). Accordingly, three encoded genes coexist in the BCS operon with additional two genes coding for the α and β subunit of electron transfer protein (Bennett and Rudolph, 1995). Butyryl-CoA was then catalyzed by phosphotransbu‐ tylase (PTB, or phosphate butyltransferase, encoded by *ptb* gene) and butyrate kinase (BK, encoded by *bk* gene) to form butyrate during acidogenesis phase.

As the organic acid accumulation, pH drop to the lowest point during the fermentation. This leads to the switch of acidogenesis phase to solventogenesis phase. Acetate and butyrate are reassimilated and participate in the solvent formation. Under the catalyzing of CoA transfer‐ ase (CoAT, two unit encoded by *ctfα* and *ctfβ*), acetate and butyrate was transformed into acetyl-CoA and butyryl-CoA respectively again. The alcohols formation share the same key enzymes, NAD(P)H dependent aldehyde/alcohol dehydrogenases (encoded by *adh*1 and *adh*2 gene) (Chen, 1995). In addition, Butanol owns its unique butanol dehydrogenase (en‐ coded by *bdh* gene) (Welch et al., 1989). The formation of acetone from acetoacetyl-CoA is a two-step reaction. Acetoacetyl-CoA is catalyzed to acetoacetate by CoA transferase. Acetone is produced after a molecular CO2 released from acetoacetate by decarboxylase (AADC, en‐ coded by *aadc* gene) (Janati-Idrissi et al., 1988; Cary et al., 1993). Both acid reassimilation and acetone formation utilize CoA transferase, however, the butyrate uptake was not concomi‐ tant with the production of acetone (Desai et al., 1999). The metabolic pathway accompanied by electron transfer and reduction force forming. The main ABE fermentation pathway was illustrated in Fig.3.

Solventogenic genes *aad*, *ctfA*, *ctfB* and *adc* constitute the *sol* operon (Durre et al., 1995). In some conditions, butanol producing strains lose the ability to produce solvents after repeat‐ ed subculturing, called as degenerated (DGN) strain. In *C. acetobutylicum* ATCC 824, the plasmid pSOL1 carrying the *sol* operon was found missing during degenerating process (Cornillot et al., 1997). For *C. saccharoperbutyl acetonicum* strain N1-4, the *sol* genes main‐ tained in degenerated DGN3-4 strain, while the *sol* operon was hardly induced during sol‐ ventogenesis. Extract from the culture supernatants of wild-type N1-4 is enough to induce the transcription of the *sol* operon in DGN3-4 (Kosaka et al., 2007). It suggested that the de‐ generation maybe caused by the incompetence of the induction mechanism of the *sol* oper‐ on. The transcription of *sol* operon may be under the control of the quorum-sensing mechanism in *C. saccharoperbutyl acetonicum*.

**Figure 3.** Metabolic pathway of Acetone-butanol-ethanol fermentation. EMP: Embden-Meyerhoff pathway (glycoly‐ sis); AK, acetate kinase; PTA, phosphotransacetylase; CoAT, CoA transferase; AADC, acetoacetate decarboxylase; THL, thiolase; BK, butyrate kinase; PTB, phosphotransbutylase; HBD, hydroxybutyryl-CoA dehydrogenase; CRO, crotonase; BCD, butyry-CoA dehydrogenase; AAD, aldylde/ alcohol dehydrogenase; BdhA, butyryl-CoA dehydrogenase A; BdhB,

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 183

The increasing genetic knowledge provides feasible technique for the strain modification. Many efforts have been made to construct the strain with high butanol tolerance, superior butanol yield, productivity and less byproduct. The process can be classified into pathway-

Except butanol, acetone and ethanol are main products in ABE fermentation. The byprod‐ uct, especially acetone is low valuable and undesirable. Blocking the expression key enzyme gene for acetone is thought perfect to decrease the split flux and enhance butanol yield. However, the results were not ideal as expected. Knocking out the C. acetobutylicum EA 2018 *adc* gene, the acetone is still produced in low level (Jiang et al., 2009). In *C. beijerinckii* 8052, the strain with *adc* gene disruption produced similar acetone with the original wild type strain (Han et al., 2011). To block acetate and acetone pathway by knocking out gene

butyryl-CoA dehydrogenase B.

**3.3. Metabolic engineering**

based construction and regulation-based construction.

Though the metabolic pathway is clear, the underlying regulation mechanism is poorly un‐ derstood, such as the phase switch of fermentation, the relationship between solventogene‐ sis and sporulation. Answering these questions is critical to improve the efficiency of butanol producing fundamentally. Proteomics and transcriptomics can provide more un‐ known details, which will be helpful for solving these problems (Sivagnanam et al., 2011; Sivagnanam et al., 2012).

**Figure 3.** Metabolic pathway of Acetone-butanol-ethanol fermentation. EMP: Embden-Meyerhoff pathway (glycoly‐ sis); AK, acetate kinase; PTA, phosphotransacetylase; CoAT, CoA transferase; AADC, acetoacetate decarboxylase; THL, thiolase; BK, butyrate kinase; PTB, phosphotransbutylase; HBD, hydroxybutyryl-CoA dehydrogenase; CRO, crotonase; BCD, butyry-CoA dehydrogenase; AAD, aldylde/ alcohol dehydrogenase; BdhA, butyryl-CoA dehydrogenase A; BdhB, butyryl-CoA dehydrogenase B.

#### **3.3. Metabolic engineering**

The increasing genetic knowledge provides feasible technique for the strain modification. Many efforts have been made to construct the strain with high butanol tolerance, superior butanol yield, productivity and less byproduct. The process can be classified into pathwaybased construction and regulation-based construction.

Except butanol, acetone and ethanol are main products in ABE fermentation. The byprod‐ uct, especially acetone is low valuable and undesirable. Blocking the expression key enzyme gene for acetone is thought perfect to decrease the split flux and enhance butanol yield. However, the results were not ideal as expected. Knocking out the C. acetobutylicum EA 2018 *adc* gene, the acetone is still produced in low level (Jiang et al., 2009). In *C. beijerinckii* 8052, the strain with *adc* gene disruption produced similar acetone with the original wild type strain (Han et al., 2011). To block acetate and acetone pathway by knocking out gene *adc* and *ctfA* reduced solvent production (Lehmann et al., 2012). These results demonstrated that the butanol metabolic mechanism is more complicated than expected.

There also some strategies aim at the upstream regulation. Global transcription machinery engineering (gTME) is thought to be a promising method to improve the butanol-producing performance (Alper et al., 2006; Papoutsakis, 2008). By regulating the transcription factor, the gTME strategy is thought to be able to change the metabolic strength and direction. gTME has been shown an efficient solution to improve substrate utilization, product toler‐ ance, and production in yeast (Alper et al. 2006) and *E. coli* (Chen et al., 2011). In butanolproducing *Clostridium*, the metabolic pathway have been described clearly, however, the mechanism of metabolism regulation is still not fully understood. This situation keeps the gTME strategy away from butanol-producing strains. Much effort should be devoted on the proteomics and transcriptomics etc. that will increase more details behind the appearance of ABE fermentation. A true gTME strategy will bring fresh and effective innovation to the bu‐

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 185

The concept of metabolic engineering is to develop strains as "cell factory" which is efficient for desired products production from renewable sources (Na et al., 2010). Some microbes at‐ tracted interests because they are more tolerant to butanol than *Clostridium*, although these bacteria haven't natural solvent-producing ability. Some kinds of Lactic acid bacteria can grow in 3-4% butanol (Liu et al., 2012) after long term adaption, that makes them promising host for butanol producing. The synthetic biology strategy has been implemented by con‐ structing the whole butanol-producing pathway in *Escherichia coli*, *Bacillus subitilis*, *Saccharo‐ myces cerevisiae* and *Pseudomonas putida* (Shen and Liao, 2008; Nielsen et al., 2009). This

ABE fermentation can be conducted as batch, fed-batch, and continuous under anaerobic conditions. Batch fermentation is the simplest mode. The substrate is typical 40-80g/L and the efficiency decreased as substrate concentration upper than 80g/L (Shaheen et al, 2000). With optimized physiological and nutritional parameters, 20g/L n-butanol was obtained by *C. beijerinckii* ATCC 10132 in 72h (Isar and Rangaswamy, 2012). Fed-batch fermentation was adopted to avoid substrate inhibition. However, because of product inhibition, the substrate feeding seems ineffective. The solvent must be removed from the broth to decrease the product toxicity. The solvent can be removed by several ways such as liquid-liquid extrac‐ tion, perstraction, gas-stripping, and pervaporation etc. (Qureshi and Maddox, 1995; Qure‐ shi and Blaschek, 2001b). The whole systemic technique of high productivity was constructed by continuous feeding combined with product removal (Qureshi et al., 1992), such as using membrane reactor (Qureshi et al., 1999a). With these techniques, the fermenta‐ tion can be continuing for a long time and resulting in higher productivity. To improve the utilization efficiency of cells, the immobilization system is used (Huang et al., 2004; Qureshi et al., 2000; Lienhardt et al., 2002). Comparing with the free cell system, the immobilization system is easier to separate cells from product, can reach high cell concentration and pro‐

Co-culture is another important way for butanol fermentation (Abd-Alla and El-Enany, 2012). *C. beijerinckii* NCIMB 8052 was entangled with ATCC 824 and thought as *C. acetobuty‐*

strategy deserves further attempts in spite of the poor final butanol concentration.

ductivity, and can decrease nutrient depletion and product inhibition.

tanol fermentation.

**3.4. Fermentation application**

Acetate and butyrate are produced during acidogenesis, and then they are transformed into acetyl-CoA and butyl-CoA to participate the solvent formation during solventogenesis phase. It seems an ineffective loop. In fact, the "inefficiency" loop is necessary for acid accu‐ mulation and switching to solventogenesis, at the same time, energy and reduction force were reserved. Disruption of acetate and butyrate pathway didn't enhance butanol produc‐ tion. Knocking out acetate biosynthetic pathway gene by Clos Tron had no significant influ‐ ence on the metabolite distribution (Lehmann et al., 2012). Disruption of *ptb* gene blocked the butyrate synthesis and led to acetate and lactate accumulation. Some mutant strain with‐ out *bk* gene even can't survival (Sillers et al., 2008). It indicated that the pathways seeming useless were necessary for butanol synthesis. What's more, it is not possible to improve per‐ formance by decrease acid formation.

The genes participate in butanol synthesis including of *thl*, BCS operon, and *add*, *bdh*. Over‐ expression these genes are thought useful to increase the butanol yield. Overexpression of *aad* gene alone could enhance butanol production (Nair and Papoutsakis, 1994; Tummala et al., 2003). Transformed strain M5 (*sol* operon deficient because of lose of plasmid pSOL) with a plasmid carrying *aad* gene restored butanol-producing capability (Nair and Papoutsa‐ kis, 1994). Overexpression of *aad* gene and down-regulated *ctf* gene increased the butanol and ethanol production. To boost the butyryl-CoA pool, the strain with both *thl* and *aad* overexpression was constructed. However, butyrate and acetone concentration were in‐ creased, not butanol. The *thl* overexpression with *ctf* knock down didn't change the product significantly (Sillers et al., 2009). So, the metabolic is more complicated than it seems. Theo‐ retical analyses also suggested alteration single solvent-associated gene is not sufficient to increase butanol yield (Haus et al., 2011).

Low butanol tolerance of the strains is another problem of butanol production. Although butanol synthesis is spontaneous in clostridium, the wild type strains can't endure high bu‐ tanol concentration upper than 2%. Butanol stress influence gene expression of amino acid, nucleotide, glycerolipid biosynthesis and the cytoplasmic membrane composition (Janssen et al., 2012). Cells have heat shock response system will protect it from heat or other stress (Bahl, Müller et al. 1995). Overexpression of grosESL improved the strain tolerance and bu‐ tanol titer (Tomas et al., 2003).

The utilization of xylose and other carbon sources was inhibited by glucose is a phenomen‐ on called as Carbon catabolite repression (CCR). CCR limited the efficiency of butanol fer‐ mentation with lignocellulosic material as substrate. The utilization rate of pentose was improved efficiently by knocking out pleiotropic regulator gene *ccpA*, *glcG* (responsibility for phosphoenoopyruvate-dependent phophotransferase system, PTS) and overexpressing the genes of xylose utilization (Ren et al., 2010; Xiao et al., 2012). By heterogonous expres‐ sion transaldolase gene talA in ATCC 824, the xylose utilization was improved significantly (Gu et al., 2009). Knocking out xylose repressor gene *XylR* also increased the fermentation efficiency (Xiao et al., 2012).

There also some strategies aim at the upstream regulation. Global transcription machinery engineering (gTME) is thought to be a promising method to improve the butanol-producing performance (Alper et al., 2006; Papoutsakis, 2008). By regulating the transcription factor, the gTME strategy is thought to be able to change the metabolic strength and direction. gTME has been shown an efficient solution to improve substrate utilization, product toler‐ ance, and production in yeast (Alper et al. 2006) and *E. coli* (Chen et al., 2011). In butanolproducing *Clostridium*, the metabolic pathway have been described clearly, however, the mechanism of metabolism regulation is still not fully understood. This situation keeps the gTME strategy away from butanol-producing strains. Much effort should be devoted on the proteomics and transcriptomics etc. that will increase more details behind the appearance of ABE fermentation. A true gTME strategy will bring fresh and effective innovation to the bu‐ tanol fermentation.

The concept of metabolic engineering is to develop strains as "cell factory" which is efficient for desired products production from renewable sources (Na et al., 2010). Some microbes at‐ tracted interests because they are more tolerant to butanol than *Clostridium*, although these bacteria haven't natural solvent-producing ability. Some kinds of Lactic acid bacteria can grow in 3-4% butanol (Liu et al., 2012) after long term adaption, that makes them promising host for butanol producing. The synthetic biology strategy has been implemented by con‐ structing the whole butanol-producing pathway in *Escherichia coli*, *Bacillus subitilis*, *Saccharo‐ myces cerevisiae* and *Pseudomonas putida* (Shen and Liao, 2008; Nielsen et al., 2009). This strategy deserves further attempts in spite of the poor final butanol concentration.

#### **3.4. Fermentation application**

ABE fermentation can be conducted as batch, fed-batch, and continuous under anaerobic conditions. Batch fermentation is the simplest mode. The substrate is typical 40-80g/L and the efficiency decreased as substrate concentration upper than 80g/L (Shaheen et al, 2000). With optimized physiological and nutritional parameters, 20g/L n-butanol was obtained by *C. beijerinckii* ATCC 10132 in 72h (Isar and Rangaswamy, 2012). Fed-batch fermentation was adopted to avoid substrate inhibition. However, because of product inhibition, the substrate feeding seems ineffective. The solvent must be removed from the broth to decrease the product toxicity. The solvent can be removed by several ways such as liquid-liquid extrac‐ tion, perstraction, gas-stripping, and pervaporation etc. (Qureshi and Maddox, 1995; Qure‐ shi and Blaschek, 2001b). The whole systemic technique of high productivity was constructed by continuous feeding combined with product removal (Qureshi et al., 1992), such as using membrane reactor (Qureshi et al., 1999a). With these techniques, the fermenta‐ tion can be continuing for a long time and resulting in higher productivity. To improve the utilization efficiency of cells, the immobilization system is used (Huang et al., 2004; Qureshi et al., 2000; Lienhardt et al., 2002). Comparing with the free cell system, the immobilization system is easier to separate cells from product, can reach high cell concentration and pro‐ ductivity, and can decrease nutrient depletion and product inhibition.

Co-culture is another important way for butanol fermentation (Abd-Alla and El-Enany, 2012). *C. beijerinckii* NCIMB 8052 was entangled with ATCC 824 and thought as *C. acetobuty‐* *licum* before the 16S rDNA based method was exploited (Johnson and Chen, 1995). These data implied that they could be cocultured before isolation. A microflora of four strain iso‐ lated from hydrogen-forming sludge of sewage performed a little high solvent yield (Cheng et al., 2012). Different strains possess various advantages, either with larger carbon sub‐ strate, higher butanol yield, or with high substrate and product tolerance. The co-culture should possess potential benefits and be harnessed fully after all the details are disclosed for each individual strain.

mentation broth, it captures the solvents. The solvents then condensed in the condenser and are collected in a receiver. Ezeji applied gas stripping on the fed-batch fermentation, 500 g glu‐ cose was consumed and 233 g/l solvent was produced with the productivity of 1.16 g/(Lh) and the yield of 0.47 g/g.When combined with continuous fermentation with gas stripping, 460g/l solvent was obtained with 1163g glucose consuming (Ezeji et al., 2004a; Ezeji et al., 2004b).

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 187

Pervaporation is a membrane-based process that allows selective removal of volatile com‐ pounds from fermentation broth. The membrane is placed in contact with the fermentation broth and the volatile liquids or solvents diffuse through the membrane as a vapor which is recovered by condensation. A vacuum applied to the side of permeate. Polydimethylsilox‐ ane membranes and silicon rubber sheets are generally used for the pervaporation process. Selection of a suitable polymer forming the active part of the membrane is a key factor in this case. In the batch fermentation, Evans and Wang increased the solvent concentration and productivity from 24.2g/l and 0.34g/(lh) to 32.8g/l and 0.5g/(lh) with pervaporation (Evans and Wang, 1988). Groot et al. applied pervaporation on the fed-batch fermentation and the solvent productivity and concentration reached 0.98g/lh and 165.1g/l (Groot et al., 1984). The Reverse osmosis is another recovery technique that based on membranes. Before the reverse osmosis is carried out, the suspended vegetative organisms must be removed us‐ ing the hollow-fiber ultra-filter. After the pretreatment, reverse osmosis starts to dewater the fermentation liquor by rejecting solvents but allowing water to pass through the membrane.

Liquid–liquid extraction can be used to remove solvents from the fermentation broth. In this process, the water-insoluble organic extractant is mixed with the fermentation broth. Buta‐ nol is more soluble in the organic (extractant) phase than in the aqueous (fermentation broth) phase. So, butanol can be selectively concentrated in the organic phase. As the extrac‐ tant and fermentation broth are immiscible, the extractant can easily be separated from the fermentation broth after butanol extraction. (Qureshi and Blaschek, 1999a). However, there still some problems with liquid–liquid extraction such as toxicity of extractant, extraction solvent losing, the formation of an emulsion, etc. Oleyl alcohol as a good extractant with rel‐ atively low-toxic has been used widely by the researchers (Karcher et al., 2005; Ezeji, 2006).

The butanol extraction process using conventional solvents may be useful, but the solvents used are often volatile, toxic and dangerous. In recent years, a growing interest in ionic liq‐ uids(IL) which also can be used in butanol recovery. Ionic liquids are organic salts present in the liquid state at room conditions, have very low vapor pressure and low solubility in water. Hence, Ionic liquids is valuable solvent in the extraction process from aqueous solutions (Fa‐ deev and Meagher, 2001; Garcia-Chavez et al., 2012). Ionic liquids as the non-volatile, environ‐ ment friendly solvents have been used in various chemical processes. With the development of the technology, ionic liquids extraction would be more promising for butanol recovery.

**4.4. Butanol recovery by pervaporation**

**4.5. Liquid–liquid extraction**

**4.6. Application of ionic liquids**

And then, the products are concentrated (Zheng et al., 2009).

#### **4. Separation of butanol product**

Because butanol has a higher boiling point than water, therefore, distillation is not suitable for butanol recovery. Other processes such as adsorption, pervaporation, membrane pertrac‐ tion, reverse osmosis and gas stripping have been developed to improve recovery perform‐ ance and reduce costs (Oudshoorn et al., 2009; Ezeji et al., 2004b).

#### **4.1. Adsorption process**

Adsorption is the technology operating easily for the butanol separation. Butanol can be ad‐ sorpted by the adsorbents in the fermenter and then the butanol was obtained by desorp‐ tion. A variety of materials can be used as adsorbents for butanol recovery and silicalite is the common one used (Qureshi et al., 2005b; Ezeji et al., 2007). Silicalite is a form of silica with a zeolite-like structure and hydrophobic properties, it can selectively adsorb small or‐ ganic molecule like C1–C5 alcohols from dilute aqueous solutions (Zheng et al., 2009). How‐ ever, adsorption separation process is not suitable on an industrial or semi-technical scale because the capacity of adsorbent is very low.

#### **4.2. Butanol recovery by membrane reactor**

Immobilization of microorganisms in the membrane or using membrane reactors is another option of butanol removal. The productivity can be enhanced obviously by this way. Huang et al. reported the continuous ABE fermentation by immobilized *C. acetobutylicum* cells with the fibrous as carrier and a productivity of 4.6 g/L/h was obtained (Huang et al., 2004). Qure‐ shi et al. studied the butanol fermentation by immobilized *C. beijerinckii* cells with different carriers such as clay brick, the reactor productivity was enhanced to 15.8 g/(lh) (Qureshi and Blaschek, 2005a). Although the butanol productivity increased by using immobilized cell fermentation, leakage of cells from the matrices is a frequent problem for the industrial ap‐ plication. There still some other problems such as poor mechanical strength and increase mass transfer resistance etc.

#### **4.3. Butanol recovery by gas stripping**

Gas stripping seems to be a promising technique that can be applied to butanol recovery com‐ bined with ABE fermentation. When the gas (ordinary N2 or CO2 ) are bubbled through the fer‐ mentation broth, it captures the solvents. The solvents then condensed in the condenser and are collected in a receiver. Ezeji applied gas stripping on the fed-batch fermentation, 500 g glu‐ cose was consumed and 233 g/l solvent was produced with the productivity of 1.16 g/(Lh) and the yield of 0.47 g/g.When combined with continuous fermentation with gas stripping, 460g/l solvent was obtained with 1163g glucose consuming (Ezeji et al., 2004a; Ezeji et al., 2004b).

#### **4.4. Butanol recovery by pervaporation**

Pervaporation is a membrane-based process that allows selective removal of volatile com‐ pounds from fermentation broth. The membrane is placed in contact with the fermentation broth and the volatile liquids or solvents diffuse through the membrane as a vapor which is recovered by condensation. A vacuum applied to the side of permeate. Polydimethylsilox‐ ane membranes and silicon rubber sheets are generally used for the pervaporation process. Selection of a suitable polymer forming the active part of the membrane is a key factor in this case. In the batch fermentation, Evans and Wang increased the solvent concentration and productivity from 24.2g/l and 0.34g/(lh) to 32.8g/l and 0.5g/(lh) with pervaporation (Evans and Wang, 1988). Groot et al. applied pervaporation on the fed-batch fermentation and the solvent productivity and concentration reached 0.98g/lh and 165.1g/l (Groot et al., 1984). The Reverse osmosis is another recovery technique that based on membranes. Before the reverse osmosis is carried out, the suspended vegetative organisms must be removed us‐ ing the hollow-fiber ultra-filter. After the pretreatment, reverse osmosis starts to dewater the fermentation liquor by rejecting solvents but allowing water to pass through the membrane. And then, the products are concentrated (Zheng et al., 2009).

#### **4.5. Liquid–liquid extraction**

Liquid–liquid extraction can be used to remove solvents from the fermentation broth. In this process, the water-insoluble organic extractant is mixed with the fermentation broth. Buta‐ nol is more soluble in the organic (extractant) phase than in the aqueous (fermentation broth) phase. So, butanol can be selectively concentrated in the organic phase. As the extrac‐ tant and fermentation broth are immiscible, the extractant can easily be separated from the fermentation broth after butanol extraction. (Qureshi and Blaschek, 1999a). However, there still some problems with liquid–liquid extraction such as toxicity of extractant, extraction solvent losing, the formation of an emulsion, etc. Oleyl alcohol as a good extractant with rel‐ atively low-toxic has been used widely by the researchers (Karcher et al., 2005; Ezeji, 2006).

#### **4.6. Application of ionic liquids**

The butanol extraction process using conventional solvents may be useful, but the solvents used are often volatile, toxic and dangerous. In recent years, a growing interest in ionic liq‐ uids(IL) which also can be used in butanol recovery. Ionic liquids are organic salts present in the liquid state at room conditions, have very low vapor pressure and low solubility in water. Hence, Ionic liquids is valuable solvent in the extraction process from aqueous solutions (Fa‐ deev and Meagher, 2001; Garcia-Chavez et al., 2012). Ionic liquids as the non-volatile, environ‐ ment friendly solvents have been used in various chemical processes. With the development of the technology, ionic liquids extraction would be more promising for butanol recovery.

ideal for environment problem solving.

#### **5. Biobutanol production from renewable resources** organic salts present in the liquid state at room conditions, have very low vapor pressure and low solubility in water. Hence, Ionic liquids is valuable solvent in the extraction process fromaqueous solutions (FadeevandMeagher, 2001; Garcia-Chavez et al., 2012).

Biobutanol is no doubt a superior candidate renewable energy facing the exhausted fossilenergy. The clostridium can incorporate simple and complex soluble sugar, such as corn, molasses, cassava, and sugar beet. The ABE fermentation is also a solution to deal with agri‐ culture residue, spoilage material, and domestic organic waste (Table 3). Additionally, using renewable resources is also ideal for environment problem solving. development of the technology, ionic liquids extraction would be more promising for butanol recovery. **5. Biobutanol production from renewable resources**  Biobutanol is no doubt a superior candidate renewable energy facing the exhausted fossil-energy. The clostridium can incorporate simple and complex soluble sugar, such as corn, molasses, cassava, and sugar beet. The ABE fermentation is also a solution to deal with agriculture residue,spoilage material, and domestic organic waste (Table 3). Additionally, using renewable resources is also

The butanol extraction process using conventional solvents may be useful, but the solventsused are often volatile, toxic and dangerous.In recent years, a growing interest in ionic liquids(IL) which also can be used in butanolrecovery.Ionic liquids are

Ionic liquids as the non-volatile, environment friendly solvents have been used in various chemical processes. With the

reached 0.73-1.07 US\$/kg when grass-rooted plant was used as substrate (Qureshi and Bla‐ schek, 2001a). A promising solution is co-culture of butanol-producing and cellulolytic strains. However, many obstacles must be cleared before the system is constructed. It's diffi‐ cult for different strains to play a role in turn in the substrate medium. Firstly, strain with high hydrolysis activity must be obtained. Secondly, the procedure must also be optimized.

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 189

Some strains can use CO2, H2, and CO as substrate (Tracy et al., 2012). The celluloses sub‐ strate can be transformed into CO (2) and H2 firstly. The simple substrates then are used by *C. carboxidivorans* to produce butanol. The more simple and feasible process is still need to

Due to the excessive exploitation, the fossil fuels are facing scarce and they cannot be generated. On the other hand, most of the carbon emissions result from fossil fuel combustion. Reducing the use of fossil fuels will considerably reduce the amount of carbon dioxide and other pollutants produced. Renewable energy has the potential to provide energy services with low emissions of both air pollutants and greenhouse gases. Currently, renewable energy sources supply over 14% of the total world energy demand. Biofuels as the important renewable energy are generally considered as sustainability, reduction of greenhouse gas emissions, regional development, so‐ cial structure and agriculture, and security of supply (Reijnders, 2006). Biodiesel and bioethanol are presently produced as a fuel on an industrial scale, including ETBE partially made with bioe‐

Biobutanol also has a promising future for the excellent fuel properties. It has been demon‐ strated that n-butanol can be used either 100% in unmodified 4-cycle ignition engines or blended up with diesel to at least 30% in a diesel compression engine or blended up with kerosene to 20% in a jet turbine engine in 2006 (Schwarz et al., 2006). The production of bio‐ butanol from lignocellulosic biomass is promising and has been paid attention by many companies. Dupont and BP announced a partnership to develop the next generation of bio‐ fuels, with biobutanol as first product (Cascone, 2007). In 2011, Cobalt Technologies Compa‐ ny and American Process Inc. (API) have been partnering to build an industrial-scale cellulosic biorefinery to produce biobutanol. Additionally, the companies agreed to jointly market a GreenPower+ biobutanol solution to biomass power facilities and other customers worldwide. The facility is expected to start ethanol production in early 2012 and switch to biobutanol in mid-2012. The annual production of biobutanol is estimated to 470, 000 gal‐ lons. (http://www.greencarcongress.com/2011/04/cobalt-20110419.html, http://www.renewa‐ bleenergyfocususa.com/view/17558/cobalt-and-api-cooperate-on-biobutanol/) Gevo, Inc. signed a Joint Development Agreement with Beta Renewables, a joint venture between Chemtex and TPG, to develop an integrated process for the production of bio-based isobuta‐ nol from cellulosic, non-food biomass, such as switch grass, miscanthus, agriculture resi‐ dues and other biomass will be readily available. (http://www.greencarcongress.com/ biobutanol/). Syntec company also is currently developing catalysts to produce bio-butanol

**6. The promising application and prospect of biobutanol**

thanol, these fuels make up most of the biofuel market (Antoni et al., 2007).

be further explored for different substrates.


**Table 3.** Butanol production with different raw materials

Table 3. Butanol production with different raw materials

need to be further explored for different substrates.

cellulose materials (Kumar et al.,2012). For the cellulose-based substrate, the crystal structure of cellulose is hard to use for normal ABE fermentation clostridium. The pretreatment of cellulose is costly, complex, and often leads to new environment problems. For example, using corn as substrate, the cost is 0.44-0.55 US\$/kg butanol by the hyper-butanol producing strain *C. beijerinckii*BA101 (Qureshi and Blaschek, 2000) by continuous fermentation combined with butanol separation. The cost reached 0.73-1.07 US\$/kg when grass-rooted plant was used as substrate (Qureshi and Blaschek, 2001a). A promising solution is co-culture of butanolproducing and cellulolytic strains. However, many obstacles must be cleared before the system is constructed. It's difficult for different strains to play a role in turn in the substrate medium. Firstly, strain with high hydrolysis activity must be obtained. Secondly, the procedure must also be optimized. Some strains can use CO2, H2, and CO as substrate (Tracy et al., 2012). The celluloses substrate can be transformed into CO (2) and Food-based substrate arouses many problems. The cost of butanol from glucose was four fold higher than that from sugarcane and cellulose materials (Kumar et al., 2012). For the cellulose-based substrate, the crystal structure of cellulose is hard to use for normal ABE fer‐ mentation clostridium. The pretreatment of cellulose is costly, complex, and often leads to new environment problems. For example, using corn as substrate, the cost is 0.44-0.55 US \$/kg butanol by the hyper-butanol producing strain *C. beijerinckii* BA101 (Qureshi and Bla‐ schek, 2000) by continuous fermentation combined with butanol separation. The cost

H2firstly. The simple substrates then are used by *C. carboxidivorans* to produce butanol. The more simple and feasible process is still

Food-based substrate arouses many problems. The cost of butanol from glucose was four fold higher than that from sugarcane and

reached 0.73-1.07 US\$/kg when grass-rooted plant was used as substrate (Qureshi and Bla‐ schek, 2001a). A promising solution is co-culture of butanol-producing and cellulolytic strains. However, many obstacles must be cleared before the system is constructed. It's diffi‐ cult for different strains to play a role in turn in the substrate medium. Firstly, strain with high hydrolysis activity must be obtained. Secondly, the procedure must also be optimized.

Some strains can use CO2, H2, and CO as substrate (Tracy et al., 2012). The celluloses sub‐ strate can be transformed into CO (2) and H2 firstly. The simple substrates then are used by *C. carboxidivorans* to produce butanol. The more simple and feasible process is still need to be further explored for different substrates.

#### **6. The promising application and prospect of biobutanol**

Due to the excessive exploitation, the fossil fuels are facing scarce and they cannot be generated. On the other hand, most of the carbon emissions result from fossil fuel combustion. Reducing the use of fossil fuels will considerably reduce the amount of carbon dioxide and other pollutants produced. Renewable energy has the potential to provide energy services with low emissions of both air pollutants and greenhouse gases. Currently, renewable energy sources supply over 14% of the total world energy demand. Biofuels as the important renewable energy are generally considered as sustainability, reduction of greenhouse gas emissions, regional development, so‐ cial structure and agriculture, and security of supply (Reijnders, 2006). Biodiesel and bioethanol are presently produced as a fuel on an industrial scale, including ETBE partially made with bioe‐ thanol, these fuels make up most of the biofuel market (Antoni et al., 2007).

Biobutanol also has a promising future for the excellent fuel properties. It has been demon‐ strated that n-butanol can be used either 100% in unmodified 4-cycle ignition engines or blended up with diesel to at least 30% in a diesel compression engine or blended up with kerosene to 20% in a jet turbine engine in 2006 (Schwarz et al., 2006). The production of bio‐ butanol from lignocellulosic biomass is promising and has been paid attention by many companies. Dupont and BP announced a partnership to develop the next generation of bio‐ fuels, with biobutanol as first product (Cascone, 2007). In 2011, Cobalt Technologies Compa‐ ny and American Process Inc. (API) have been partnering to build an industrial-scale cellulosic biorefinery to produce biobutanol. Additionally, the companies agreed to jointly market a GreenPower+ biobutanol solution to biomass power facilities and other customers worldwide. The facility is expected to start ethanol production in early 2012 and switch to biobutanol in mid-2012. The annual production of biobutanol is estimated to 470, 000 gal‐ lons. (http://www.greencarcongress.com/2011/04/cobalt-20110419.html, http://www.renewa‐ bleenergyfocususa.com/view/17558/cobalt-and-api-cooperate-on-biobutanol/) Gevo, Inc. signed a Joint Development Agreement with Beta Renewables, a joint venture between Chemtex and TPG, to develop an integrated process for the production of bio-based isobuta‐ nol from cellulosic, non-food biomass, such as switch grass, miscanthus, agriculture resi‐ dues and other biomass will be readily available. (http://www.greencarcongress.com/ biobutanol/). Syntec company also is currently developing catalysts to produce bio-butanol from a range of waste biomass, including Municiple Solid Waste, agricultural and forestry wastes. (http://www.syntecbiofuel.com/butanol.php). Utilization the waste materials im‐ prove the economy of butanol production that makes biobutanol great potential to be the next new type of biofuel in spite of the existing drawbacks.

[4] Badr HR, Toledo R, Hamdy MK. Continuous acetone ethanol butanol fermentation by immobilized cells of Clostridium acetobutylicum. Biomass Bioenergy. 2001,

The Promising Fuel-Biobutanol http://dx.doi.org/10. 5772/52535 191

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#### **7. Conclusions**

Biobutanol production has only recent years booming again after long time of silence. Quite a lot of progress has been made with the technology development of metabolic engineering in enhancing solvent production, increasing the solvent tolerance of bacteria, improving the selectivity for butanol. Fortunately, *Clostridia* have been tested being able to consume ligno‐ cellulosic biomass for ABE fermentation. The complex regulation mechanism of butanol synthesis is still need to be further study. For the strain improvement, for example, con‐ structing better butanol tolerance strains, more suitable hosts and genetic methods are re‐ quired to be set up. Furthermore, more efficient techniques for removing the inhibitors in the lignocellulosic hydrolysate need to be developed. In addition, from the economic point of view, the integrated system of hydrolysis, fermentation, and recovery process also are im‐ portant to be further developed to reduce the operation cost of butanol synthesis.

#### **Author details**

Hongjuan Liu\* , Genyu Wang and Jianan Zhang

\*Address all correspondence to: liuhongjuan@tsinghua.edu.cn; zhangja@tsinghua. edu. cn

Institute of Nuclear and New Energy Technology, Tsinghua University, Beijing, P. R., China

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**Chapter 7**

produces at 90% filling ratio 1053

**Biobutanol from Renewable Agricultural and**

**Alternative of Liquid Fuels**

Pradeep K. Sharma

**1. Introduction**

http://dx.doi.org/10.5772/52379

László Kótai, János Szépvölgyi, Mária Szilágyi, Li Zhibin, Chen Baiquan, Vinita Sharma and

Additional information is available at the end of the chapter

volume of wastewater. A bioreactor with a volume of 100 m3

**2. History of industrial biobutanol production**

results on methods of biobutanol production.

**Lignocellulose Resources and Its Perspectives as**

Biobutanol (n-C4H9OH, available as fermentation product of various carbohydrate derivatives obtained from different resources of agricultural production such as crops and wastes) is one of the most promising biofuels in the near future. It can be produced by the so-called ABE (acetone-butanol-ethanol) type anaerobic fermentation discovered by Pasteur [1, 2] and industrialized by Weizmann [3]. Main problems associated with industrial production of biobutanol include high energy demand for processing of dilute ferment liquors and high

kg of butanol, 526 kg of acetone and 175 kg of ethanol together with 2900 kg of carbon dioxide, 117 kg of hydrogen and 84150 kg of wastewater. Efforts to increase productivity and decrease production costs resulted in many new methods. This chapter summarizes some selected

During investigations aimed at discovering cheaper sources of acetone and butanol for chemical industry, Weizmann [3] isolated an organism which could ferment a fairly concen‐ trated corn mash with good yields of acetone and butanol. In 1915 the British Admirality took over the research and carried out large-scale tests in an improvised apparatus but without

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

© 2013 Kótai 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.

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