Lignin depolymerization in supercritical solvents next to water includes ethanol, methanol, CO2, and CO2/acetone/water. The supercritical properties of these fluids are displayed in Table 17.10.

The choice for using CO2 as solvent is obvious as CO2 is cheap, environmentally friendly and generally recog­nized as safe by the US Food and Drug Administration. scCO2 has other advantages because of its high diffusivity combined with its easily tunable solvent strength. To use CO2 under supercritical conditions, the temperature needed is low (>31 0C) and the pressure needed relatively low (>7.4 MPa) in comparison to other supercritical sol­vents (Table 17.10). Additionally, CO2 is a gas at room temperature and pressure, which leads to a solvent-free product after pressure expansion. A drawback of scCO2 is its low polarity, which is comparable to hexane, but this problem can be overcome by using cosolvents to change the polarity of the SCF (Herrero et al., 2010). Furthermore, SCF processing based on CO2 enables the easy recycling of CO2, which is advantageous for the development of a sustainable process. Research per­formed on supercritical processing of lignin to produce aromatic compounds has been summarized hereafter.

Depolymerization of lignin model compounds and organosolv lignin have been studied in supercritical al­cohols like methanol and ethanol in a temperature range of >239 0C and a pressure of >8.1 MPa. By using bases such as KOH and NaOH a high depolymerization con­version was obtained. The dominant depolymerization route is the solvolysis of ether linkages in the lignin structure while the carbon—carbon linkages are mostly stable (Miller et al., 1999; Minami et al., 2003).

Yuan et al. (2010) used BCD at mild temperatures (220—300 0C) of kraft lignin in water—ethanol into oligo­mers with a negligible char and gas production.

However, under the conditions applied lignin could not be completely degraded into monomers.

Oxidation of lignin and lignin model compounds with peroxide was studied under scCO2 conditions in the absence of alkali. The 5-5 biphenols were shown to be degraded and in this process mostly the formation of carboxylic acids from kraft lignin was observed (Argyropoulos et al., 2006).

Gosselink et al. (2012) found that hardwood and wheat straw organosolv lignins were depolymerized in supercritical carbon dioxide/acetone/water fluid at 300 0C and 100 bar into 10—12% monomeric aromatic compounds. Small amounts of formic acid were intro­duced as in situ hydrogen donor. Furthermore, lignin is converted into a phenolic oil consisting of both mono­meric and oligomeric aromatic compounds. Interest­ingly, maximum individual yields of 3.6% for syringol and 2.0% for syringic acid based on lignin were ob­tained. Depolymerized phenolic products and char were separated during this process by pressure expan­sion. As during this process competition occurs between lignin depolymerization and recondensation of frag­ments a substantial amount of char is formed.

High-value phytochemicals may be produced from cultured plant cells, which can be far more expensive and demand far more complex, highly specialized technologies (to promote and sustain the growth, prop­agation, vitality and productivity of the cells) in compar­ison with other methods. Such approaches are generally developed for phytochemicals of pharmaceutical uses, as exemplified by the production of paclitaxel or other taxoids from cultured Taxus plants cells. Anthocyanin production from Ajugareptans, Aralia, Euphorbia milli, Fragaria, Oxalis, Perilla, Vitis, grapes or carrot have also been explored (Chattopadhyay et al., 2008; Tripathi and Tripathi, 2003).

Production from Microbial Fermentation

Microbial fermentation may be a viable phytochemical-producing technology, alternative to those directly targeting plants. A fermentative process is independent of plant harvesting cycle, suits for pro­cess engineering and control, and could be more economical than plant cell culturing. Specifically selected wild-type or genetically engineered microbes are fed (in addition to N sources, growth-stimulators, and other medium ingredients) with inexpensive fermentable sugars such as isolated glucose, corn steep liquor, whey or other coproducts from various plants, crops or dairy processings (Gupta et al., 2011; Dufosse, 2006; Mapari et al., 2005; Adrio and Demain, 2003). After fermentation, various steps such as cell disruption and solvent extraction may be applied to obtain, enrich or purify phytochemical products.

Full or semicommercial processes for phytochemical production by microbial fermentation have been devel­oped, as exemplified by the production of riboflavin from fungi (Eremothecium ashbyii, Ashbya gossypi), yeast (Candida guilliermondii, Debaryomyces subglobosus), or bacteria (Clostridium acetobutylicum) (Chattopadhyay et al., 2008), as well as other vitamins (Shimizu, 2008). Carotenoids may also be produced by microbial fermen­tation, as exemplified by the production of b-carotene from B. trispora or Phycomyces blakesleeanus; lycopene from Fusarium sporotrichioides or bacterium Erwinia uredovora; zeaxanthin from a Flavobacterium sp.; astaxan — thin from Xanthophyllomyces dendrorhous, Rhodotorula glutinis, Rhodotorula gracilis, Rhodotorula rubra or Rhodotorula graminis; canthaxanthin from bacterium

Bradyrhizobium sp.; and isorenieratene from bacterium Brevibacterium aurartiacum, Streptomyces mediolani, or Mycobacterium aurum. Production of certain therapeutic phytochemicals in microbial fermentation has been reported as well (Demain and Adrio, 2008).

Production from Algae via Aquaculture

As known producers of many compounds identical or homologous to plant-derived phytochemicals of industrial interest, algae have been explored for phyto­chemical production. Currently, the majority of commercial b-carotene is produced from Dunaliella sal — ina and Dunaliella bardawil. Astaxanthin may be pro­duced from Haematococcus lacustris; canthaxanthin from H. lacustris, Coelastrella striolata or Chlorella zofin — giensis; and lutein from Muriellopsis sp., Scenedesmus almeriensis or Chlamydomonas zofingiensis (Skjanes et al., 2012; Chattopadhyay et al., 2008). Algae may also produce vitamins and bioactive or dietary amino acids, proteins (e. g. phycobiliproteins from Spirulina (Artho- spira) platensis), lipids or fatty acids, or phycocolloids (agar, carrageenan and alginate) (Brennan et al., 2012; Becker, 2004). Those algae may be grown and harvested either outdoor (aquaculture) or indoor inside factory tanks, as selected wild types or genetically engineered strains.

Butanol has many desirable properties as a fuel and thus is a suitable target for modification of

cyanobacteria. In fact, as a fuel it is superior to ethanol, being less corrosive and less volatile. Thus, it can easily be mixed with hydrocarbon-based fuels and used in the same infrastructure. A number of recent studies have shown that engineered cyanobacteria can in fact make surprisingly high levels of this compound, at rates that in fact surpass published rates for ethanol production by engineered cyanobacteria. Since cyanobacteria pro­duce this fuel directly through photosynthetically driven CO2 fixation, it is appropriate to compare the pro­ductivity per area of this process, as presently described, with that required for other biofuels, be it growing corn to produce the necessary sugars, or growing algae to produce biodiesel. Such a comparison shows that butanol production by cyanobacteria could be much bet­ter than making fuels from corn and very comparable to making biodiesel from microalgae (Sheehan, 2009).

Many cyanobacteria are capable of producing volatile compounds, including higher alcohols, but the natural production levels are miniscule (Hasegawa et al., 2012). In order to make fuel molecules at significant quantities, new pathways must be introduced as well as changes made to the native metabolic pathways. Studies on creating heterotrophic bacterial strains capable of producing butanol demonstrated that two possible routes were useful: the 2-ketoacid pathway, normally involved in amino acid biosynthesis, and the acetyl-CoA pathway, found in organisms such as Clos­tridium that naturally produce butanol during fermenta­tion (Figure 22.4).

The first successful attempt in this direction was to engineer S. elongates to produce isobutyraldehyde through the 2-ketoacid pathway (Atsumi et al., 2009). Isobutyraldehyde is a precursor for isobutanol and other chemicals of interest and has the advantage of being highly volatile, easing its recovery from the culture broth thus removing product inhibition. The strategy applied consisted of boosting carbon flux through the pathway from pyruvate to 2-ketoisovalerate by integra­tion into the genome of three foreign genes, alsS, ilvC and ilvD, catalyzing these steps, as well as kivd from Lactococcus lactis, the gene encoding the ketoacid decar­boxylase enzyme that converts 2-ketoisovalerate to iso — butyraldehyde. Overall carbon flux was then increased by integrating an additional copy of Rubisco (rbcLS) and the resulting strain produced 6230 mg isobutyralde — hyde per liter per hour, a production rate that is higher than any other fuel molecule made by cyanobac­teria to date. Additionally, it was demonstrated that isobutanol could be formed if a foreign alcohol dehydro­genase (YqhD from E. coli) was introduced, but titers were lower, presumably due to product inhibition.

However, the isomer that is made by the 2-ketoisovalerate pathway is isobutanol, a fuel additive, but not nearly as desirable in itself as a fuel as n-butanol, the product of the acetyl-CoA pathway or the 2-ketobutyrate pathway (Figure 22.4). Metabolic engineering was used to create an n-butanol-producing strain of S. elongatus by introducing the hbd, crt, and adhE2 genes from C. acetobutylicum, the ter gene from Treponema denticola, and the atoB gene (instead of thl) from E. coli (Lan and Liao, 2011). However, n-butanol was only produced by this strain under anaerobic con­ditions, either in the light when photosystem II was inhibited by DCMU, or in the dark, which gave the highest production, a meager 20.8 mg/L/h. It was sug­gested that anaerobic conditions were necessary since some of the enzymes introduced are oxygen sensitive, severely limiting its usefulness. On the other hand, metabolic fluxes are obviously different during dark metabolism than during photosynthesis and the differ­ence could be in the supply of a key metabolite. In line with this, in a more recent attempt to create an n-butanol-producing strain, flux through acetyl-CoA was increased by substituting an irreversible ATP hydrolysis step leading to the formation of acetoacetyl CoA (Lan and Liao, 2011). Other improvements con­sisted of substituting NADPH-requiring enzymes for NADH enzymes. With these changes, it was possible to demonstrate light-dependent n-butanol production, but at 62.5 mg/L/h this is well below (by a factor of 100) the initial promising results with butyraldehyde. This system would need very significant improvement before it could be considered for practical biofuel production.

A search made in the database Current Contents Con­nect® for key words "lignocellulosic biomass" and "pre­treatment" resulted in a total of 1217 articles published from 2000 to 2012. From the total, 91.2% corresponded to research papers and 8.8% to review papers (Figure 4.1). The number of published papers has increased exponentially from 2007 to 2012 as shown in Figure 4.1, indicating the relevance that the topic has gained in the recent years. Among the reviews docu­ments, 37.2% corresponded to reviews directly related to pretreatments, pointing out the importance of this step in the concept of biorefinery. The remaining docu­ments corresponded to reviews on the general topic of biofuels from lignocellulosic biomass.

For the research papers, the search was refined to select only papers that focus on the pretreatment pro­cesses obtaining a total of 692 papers and around 54% of these papers were published in 2012. The research papers related to pretreatment were classified into nine categories as shown in Figure 4.2. From this analysis we can conclude that alkaline, acid, thermal and IL pre­treatments are the most reported, the comparison and combination of them is also widely reported. However, this variety of studied pretreatments indicates that there is not a prevalent pretreatment suggesting that further investigation on the topic is required.

1 2

Rama Raju Baadhe, Ravichandra Potumarthi2’*, Vijai K. Gupta
^Department of Biotechnology, National Institute of Technology, Warangal, Andhra Pradesh, India,
^Department of Chemical Engineering, Monash University, Clayton, Victoria, Australia,
3Molecular Glycobiotechnology Group, Department of Biochemistry, School of Natural Sciences,
National University of Ireland Galway, Galway, Ireland
*Corresponding author email: ravichandra. potumarthi@monash. edu; pravichandra@gmail. com

OUTLINE

Introduction 119

Chemistry of Biodiesel 120

Transesterification 120

Disadvantages of Chemical Transesterification 120

Advantages of Using Lipases in

Biodiesel Production 121

Historical Background of Lipase 121

Lipase-Catalyzed Transesterification Done

in Two Approaches 121

Advantages of Immobilized Lipase 122

Technical Challenges 123

Feedstock 123

Vegetable Oils 123

INTRODUCTION

World’s commercial primary energy needs are mostly supplied through fossil fuels and accounts about 87% of total energy source (OPEC, 2011, 2012). Primary energy demand by 2035 increases to 54% and still fossil fuels con­tributes 82% of the global total by 2035 (OPEC, 2012). All fossil-fuel sources are finite and if the crude oil con­sumption continued at current usage rates, it will last
only for 54.2 years (British Petroleum Statistical Review, 2012). Projected demand for oil reaches 110 mb/day by 2035. Among the fossil fuel, diesel fuels have an essen­tial function in the industrial, transportation and agri­cultural sectors in developing countries. Gradual depletion of crude oil and emission of greenhouse gases in to the environment triggers the alarm for suitable alternative fuels for use in diesel engines (Ganesan et al., 2009). Biodiesel is one of the attractive and

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00008-5

alternative fuels along with bioethanol. Biodiesel or fatty acid methyl esters (FAMEs) are mono-alkyl esters of long-chain fatty acids, derived from transesterification of triglycerides (plant or animal or algal origin). It can be used directly in its pure form or as a blend with con­ventional diesel fuel in diesel engines (Ma and Hanna,

1999) . This fuel is biodegradable and nontoxic and has low emission profiles when compared to petroleum diesel (Krawczyk, 1996). But the cost of biodiesel, how­ever, is the main obstacle to commercialize the product. There are four primary ways of making biodiesel: direct use and blending (Ma and Hanna, 1999), micro emulsions (Schwab et al., 1987), thermal cracking (pyrol­ysis) (Sonntag, 1979) and transesterification. However, the first three have some limitations and drawbacks in case of physiochemical properties of biodiesel (Schwab et al., 1987). Transesterification is the well-known method and involves conversion of oils or fat to FAMEs or fatty acid ethyl esters in the presence of a catalyst such as acid, base or lipase (Bisen et al., 2010). The conventional method for producing biodiesel involves acid and base catalysts to form fatty acid alkyl esters. Processing expenses and environmental concerns associated with biodiesel production and difficulties connected with by-products recovery have led to the search for alterna­tive production methods and alternative sources (Bisen et al., 2010). Enzyme-mediated transesterification can be a moderate alternative to produce biodiesel in its pure form, which also makes its separation easy against the by-product (glycerol). But still due to the cost of enzyme, commercialization of biodiesel has not come to reality. Though there are many attempts made for bio­diesel production through enzyme-mediated method (Ranganathan et al., 2008; Sanchez and Vasudevan, 2006; Lai et al., 2005; Noureddini et al., 2005; De Oliveira et al., 2004; Xu et al., 2004; Sha et al., 2003; Belafi-Bako et al., 2002; Iso et al., 2001; Fukuda et al., 2001; Freedman et al., 1984), profitable commercial production was not achieved for industrial utilization. Recombinant DNA and protein engineering technologies improved the quantities and catalytic efficiency of lipase (Akoh et al.,

2007) . There are several technical challenges that need to be addressed to make biodiesel production profitable. Some of them associated with enzyme transesterification process. In this chapter, some of the technical challenges

О

II

r’co-ch2

о

II

R’CO-CH2 + 3 ROH

I Alcohol

R’CO-CH2

Triacylglycerol (vegetable oil)

3 CH2-O-CO-R I

2 H-C-O-CO-R’

I

1 CH2-O-CO-R»

1,2,3,-O-tri-acyl -glycerol R’, Rand R» are saturated or unsaturated chains

FIGURE 8.1 Chemical structure of triacylglycerol. Source: Bisen et al. (2010).

involved in the lipase-catalyzed biodiesel production were discussed.

CHEMISTRY OF BIODIESEL

Chemically biodiesel is defined as mono-alkyl (methyl or ethyl) esters of triacylglycerol. All vegetable oils, algal lipids and animal fats (triacylglycerol/triglyc — eride molecules) consist of a three-carbon chain forms the glycerol backbone, which consists of three long fatty acid chains (Figure 8.1). Amounts of each fatty acid pre­sent in molecules determine the properties of triacylgly — cerol (Knothe, 2001).

TRANSESTERIFICATION

In chemical transesterification process, fatty acid re­acts with any alcohol and forms mono-alkyl ester (bio­diesel) in the presence of a catalyst (acid, base and enzyme). General reaction scheme of biodiesel produc­tion is shown in Figure 8.2. The reaction has two inputs: triacylglycerol and the alcohol—commonly ethanol or methanol is used (Meher et al., 2004).

RENEWABLE ENERGY

Energy is an important currency for human society. The world population growth and rapid economic pro­gresses are expected to result in considerable increase in the demand for energy. In the reference scenario, the International Energy Agency has projected an increase in energy need by 55%, between 2005 and 2030, at an average annual rate of 1.8% (IEA, 2007). Driven by such increasing demand, and the dwindling fuel production, the cost of petroleum fuel has gone up sky high in recent times, which can jeopardize the eco­nomic progresses of a nation. Despite the fuel crisis, increasing concentrations of CO2 and other heat­trapping greenhouse gases (GHGs) in the atmosphere, primarily due to the combustion of fossil fuels, is clearly the prime reason for rapid warming of the
planet (Shay, 1993). The use of renewable energy is largely motivated from the standpoint of global energy crisis and environmental issues. Renewable energy is a form of energy that is produced from natural sources like sunlight, wind, hydropower, geothermal and biomass, which can be naturally replenished. Currently, renewable energy supplies only ~18% of the world’s energy consumption (Kumar et al., 2010). Most of these renewable energy sources (hydropower, wind, solar and geothermal) target the electricity market, while the majority of world energy consumption (about two-thirds) is derived from liquid fuels (Campbell, 2008; Hankamer et al., 2007). This has stimulated recent interest to explore alternative sources for petroleum — based fuels and much of the attention has been focused on biomass-derived liquid fuels or biofuels (Haag, 2007; Schneider, 2006).

Bioenergy Research: Advances and Applications http://dx. doi. org/10.1016/B978-0-444-59561-4.00011-5

Sauter (2012) circumvents this problem by spraying digestate on top of the scum layer using water canons. The patent of Rossow (2011) claims that the scum layer can be 5 m in thickness. It is to be expected that digested particles with a high lignin content will be washed down, sink to the bottom and can be collected there using a pump. The input material is loaded into the digester through an input shaft with its exit below the lower level of the scum layer in the digester. It will be acidified for a few days in the first step of the digestion process. The acids will be transformed into methane and carbon dioxide in the other parts of the digester. Schoenberg and Linke (2012) tested a 451 scum layer digester with whole plants (Silphium perfolia — tum) as substrate. A loading rate of 8kg/(m3d) was possible with a methane yield of 215 l/kg VS.

Solid Biomass Digester

VandeVivere et al. (2003) gave an overview of anaer­obic digesters for solid biomass. These "dry" (20—40% solids) systems are used for the digestion of kitchen and garden waste and the mechanically sorted fraction of municipal waste, but can be used for most types of solid farm and food processing by-products.

The Kompogas (VandeVivere et al., 2003) system consists of a cylindrical plug flow reactor in which the fermenting wastes are mixed and moved by pad­dles. Temperature is 55 °C and retention time 15 days. Capacity is limited to 10,000 tons/a due to the maximum dimensions of the axis with paddles. Capac­ity of these systems in Europe and Qatar is more than 1 million tons/a. Methane yields are 200—300 l/kg VS (VandeVivere et al., 2003).

There are some batch systems with leachate recircula­tion. Methane yield is up to 40% less due to channeling of the leachate in the substrate (Vandevivere). Mussoline et al., 2012 obtained 175 l/kg VS for rice straw and swine manure in a digestion period of 1 year. Tempera­tures ranged from 15 to 35 °C. The cell with 6500 m3 capacity was filled with cylindrical bales. Packing density was 100 kg/m3.

Storage of straw for 6 months is expensive (25 V/ton). In Western Europe storage cum digester tanks can be used. The idea is to fill these tanks in the period July— October with shredded straw and a fraction of old digestate together with macro — and micronutrients. Mussoline (2012) demonstrated this concept during one year with rice straw bales and swine manure. Daily power generation was directly correlated to the digester temperature.

German tanks with concrete tops are 120 V/m3 and with foil tops 70 V/m3. At a packing density of 10% this amounts to an investment of 700—1200 V/ton straw stored and digested.

Arabinoglucuronoxylans (AGXs) (arabino-4-O — metylglucuronoxylans) are the major components of nonwoody materials (e. g. agricultural crops) and a minor component of softwoods (5—10% of dry mass). They consist of a linear b-(1,4)-D-xylopyranose backbone containing 4-O-methyl-D-glucuronic acid and a-L-arabi — nofuranosyl linked by a-(1,2) and a-(1,3) glycosidic bonds (Table 17.2) (Girio et al., 2010). The xylopyranose backbone might be slightly acetylated (Peng). The typical ratio arabinose:glucuronic acid:xylose is 1:2:8. Conversely to hardwoods xylan, AGXs might be less acetylated, but may contain low amounts of galacturonic acid and rhamnose. The average DP of AGXs ranges be­tween 50 and 185 (26). In addition, because of their fur — anosidic structure, the arabinose side chains are easily hydrolyzed by acids (Peng et al., 2012).

Arabinogalactan

The heartwood of larches contains exceptionally large amounts of water-soluble arabinogalactan (AG), which is only a minor constituent in other softwood species (Peng et al., 2012). Its concentration and quality are not affected by seasonal variability. AGs are highly branched polysaccharides with molecular weights ranging from 10,000 to 120,000 Da. All larch AGs iso­lated from the Larix sp. are of the b-(3,6)-D-galactan type and consist of galactose and arabinose in a 6 to 1 ratio. Larch AG has a galactan backbone that features b-(1 / 3) linkages and galactose b-(1 / 6) and arabi­nose p-(1 / 6 and 1 / 3) side chains (Peng et al., 2012) (Table 17.2). The highly branched structure is responsible for the low viscosity and high solubility in water of this polysaccharide (Peng et al., 2012). It has the ability to bind fat, retain liquid, and dispersing prop­erties and AG also possesses a high biological activity. Larch AG is currently used in a variety of food, beverage, nutraceutical, and medicine applications (Peng et al., 2012).

Arabinoxylan

AXs are the main hemicelluloses of the grasses (Gra — mineae). AXs have been generally present in a variety of tissues of the main cereals: wheat, rye, barley, oat, rice, corn, and sorghum, as well as other plants (Peng et al., 2012). AXs are generally present in the starchy endo­sperm (flour) and outer layers (bran) of cereal grain. They are similar to hardwood xylan, but the amount of L-arabinose is higher. In AX, the linear b-(1 / 4)-d — Xylp backbone is substituted by a-L-Araf units in the positions 2-O and/or 3-O (Table 17.2). In addition, the AXs are also substituted by a-D-glucopyranosyl uronic unit or its 4-O-methyl derivative in the position 2-O, as can be found in wheat straw, bagasse and bamboo.

O-acetyl substituents may also occur (Peng et al., 2012). According to the amount of glucuronic acid and arabinose, the types of AXs are classified as AGX and glucuronoarabinoxylan (GAX), respectively (Ebringerova et al., 2005). AGXs are the dominant hemi — celluloses in the cell walls of grasses and cereals, such as sisal, corncobs and straw. Compared to AGXs, the GAXs have an AX backbone, which contains about 10 times fewer uronic acid side chains than arabinose, and also contains xylan that is double substituted by uronic acid and arabinose units. Ferulic acid and p-coumaric acid can occur esterified to the C-5 of arabinosyl units of GAXs (Peng et al., 2012). The physical and/or covalent interaction with other cell wall constituents restricts xylan extractability (Girio et al., 2010).

According to the Global Industry Classification Standard (GICS®) developed by Morgan Stanley Capital International and Standard & Poor’s, the global indus­tries can be divided into 10 sectors: energy, materials, in­dustrials, consumer discretionary, consumer staples, health care, financials, information technology, telecom­munication services, and utilities. After a review of each of the 10 GICS® industrial sectors, it should be noted that lignin finds applications in almost every sector of the industry with the exception of financials, telecom­munication services, and information technology. The latter illustrates the versatility of lignin chemistry and its potential. Overall, lignin applications can be divided into traditional and emerging. The traditional applica­tions of lignosulfonates and sulfonated kraft lignins are primarily lower value such as dust control (ca. 11% market), concrete admixtures (ca. 50% market), and oil well drilling muds (ca. 4% market) while most of the emerging applications target higher value applications, in which the chemical versatility of lignin could be fully leveraged, or very large volume applications such as the production of BTX and other petrochemicals.

Hydrogen photoproduction in green algae is catalyzed by [Fe—Fe]-hydrogenases. Earlier reports have suggested the existence of [Ni—Fe]-hydrogenase in S. obliquus (Zinn et al., 1994), but presently S. obliquus is considered as having only the [Fe—Fe]-hydrogenase (Wunschiers et al., 2001; Florin et al., 2001). Some green algal species do not show hydrogenase activity at all (Brand et al., 1989; Boichenko and Hoffmann, 1994). Currently, the presence of genes encoding [Fe—Fe]-hydrogenases has been proved in the following species: Chlamydomonas reinhard — tii (Happe and Kaminski, 2002; Forestier et al., 2003), Chlorellafusca (Winkler et al., 2002), Chlamydomonas nocti — gama (Skjanes et al., 2010), Volvox carteri (Prochnik et al., 2010), Tetraselmis subcordiformis (Yan et al., 2011) and Chlorella variabilis (Meuser et al., 2011). Algal [Fe—Fe]- hydrogenases in vivo interact with Fd, a terminal acceptor of photosynthetic electron transport chain (Chang et al.,

2007) . In contrast to [Ni—Fe]-hydrogenase enzymes, [Fe—Fe]-hydrogenases have significantly higher turnover rate (6000—9000/s) and usually catalyze H2 production instead of H2 uptake (Frey, 2002). However, the possible role of these enzymes in H2 uptake under high H2 partial

21. RECENT DEVELOPMENTS ON CYANOBACTERIA AND GREEN ALGAE FOR BIOHYDROGEN PHOTOPRODUCTION

pressure has also been suggested (Kosourov et al., 2012). Unfortunately, the [Fe—Fe]-hydrogenases are extremely sensitive to O2 that irreversibly inactivates purified en­zymes within seconds (Ghirardi et al., 1997).

The most studied green alga, C. reinhardtii has two monomeric [Fe—Fe]-hydrogenases: HydA1 and HydA2 with a molecular mass of around 48 kD (Happe and Kaminski, 2002; Forestier et al., 2003). Both proteins are nuclear encoded and contain putative transit se­quences that target them to the chloroplast. The hydAl gene shows 74% similarity to hydA2 and encodes protein that is 68% identical to HydA2. Two homologous hydrogenases are typically observed in almost all green algae showing hydrogenase activity (Winkler et al., 2004). Nevertheless, some species have three [Fe—Fe]- hydrogenase enzymes (Skjanes et al., 2010). The phy­siological basis for the presence of two and more hydrogenases in green algae has not been determined. In C. reinhardtii cells, HydA1 most probably participates in the light-dependent H2 production pathway (Happe and Naber, 1993). Examination of relative enzyme activ­ities by gene-silencing techniques indicate that HydA1 catalyzes the majority of the hydrogenase activity, but the role of HydA2 in algal H2 production has not been clearly resolved (Godman et al., 2010). Recently, Meuser and et al. (2012) using the single hydAl, hydA2 and double hydA1/hydA2 knockout mutants showed that HydA2 also participates in H2 photoproduction. However, according to the authors, its contribution in the light-dependent process does not exceed 25%. The next important step in this direction should be investiga­tion of the role of these enzymes in the H2 uptake, including the mechanisms of photoreduction (Kosourov et al., 2012).