Purple non-sulfur (PNS) photosynthetic bacteria are a physiological group of different kinds of gram negative aquatic bacteria. They are considered to be as old as the first photototrophic organisms on our planet. The common characteristics of this group are their ability to perform an anaerobic type of photosynthesis without the production of oxygen. Depending on the degree of anaerobiosis, and avail­ability of carbon and light source they can grow as photoheterotrophs, photoau — totrophos or chemoheterotrophs. Because they do not use hydrogen sulfide as electron donor while growing photoautotrophically they are called purple non­sulfur bacteria. While growing facultative anaerobically they give purple to deep red pigments. Under photoheterotrophic conditions (light, anaerobiosis, organic electron donor) they can produce hydrogen. The mostly studied species are the members of Rhodobacter, Rhodopseudomonas and Rhodospirillum [93]. The enzyme systems, the carbon flow (specially TCA cycle) and photosynthetic membrane apparatus make up the overall production of hydrogen by intercon­necting the exchange of electrons, protons and ATP [94]. Because of the reasons like; high substrate/product conversion yields, lack of oxygen evolving activity, a wide range of light can be used and different kinds of organic substrates can be used; photofermentative hydrogen production has been an interest of many researchers [93]. Because of not producing hydrogen during photosynthesis in anaerobic conditions both nitrogenase and hydrogenase in bacterial membrane are active. Hydrogen production is mainly associated with nitrogenase action (Fig. 10.3).

These nitrogen-fixing bacteria can utilize the enzyme nitrogenase to catalyze the reduction of molecular nitrogen (N2) to ammonia (NH3) while producing hydrogen. Mo-nitrogenase is the most common and the most efficient nitrogenase for converting N2 to NH3 (Eqs. 10.1-10.3). It is also found in all nitrogen-fixing bacteria and is thus the most studied [95]. Three different kinds of nitrogenase and the reactions are:

Mo — nitrogenase: N2 + 8H+ + 8e~ + 16ATP! 2NH3 + H2 + 16ADP + 16Pi

(10.10)

V — nitrogenase: N2 + 12H+ + 12e“ + 24ATP! 2NH3 + 3H2 + 24ADP + 24Pi

(10.11)

Fe — nitrogenase: N2 + 24H+ + 24e~ + 48ATP! 2NH3 + 9H2 + 48ADP + 48Pi

(10.12)

In the absence of molecular nitrogen this enzyme catalyzes the hydrogen production with the reaction below (Eq. 3.4) [94]:

2H+ + 2e~ + 4ATP! H2 + 4ADP + 4Pi (10.13)

For efficient operation of nitrogenase large amounts of ATP and reducing power are needed. Oxygen is a potent inhibitor of nitrogenase that can destroy the enzyme irreversibly. Ammonium is a second important inhibitor because it can repress the synthesis of nitrogenase and can inhibit nitrogenase activity. The inhibition is reversible because nitrogenase can recover activity after consuming or removing ammonium [96].

Hydrogenase enzyme is a common feature of photosynthetic bacteria and it can be responsible for both hydrogen production and consumption. Because hydrogen production is by nitrogenase hydrogen producing activity by hydrogenase can be ignored. Studies have shown that hydrogen producing activity of hydrogenase is less than hydrogen consuming activity [97, 98]. Hydrogenase is generally accepted as a metabolic antagonist of nitrogenase. Since it is very critical to eliminate the hydrogen uptake property of hydrogenase studies on mutations on organisms to eliminate hydrogenase synthesis have been reported. Hup-mutants have been found to have more hydrogen production capacity [99-101]. Another way is to make inhibitions chemically, using carbon monoxide or oxygen, limiting the amount of nickel since hydrogenases are nickel enzymes [102] and finally the presence of ethylendiamintetraacetic acid (EDTA) is known to inhibit hydrogenase activity [101, 102]. For reducing 2H+ to H2 nitrogenase re-oxidizes electron carriers and also another reductive process can compete with hydrogen production. A good and common example is the formation of the carbon storage polymer poly — b-hydroxybutyrate (PHB) from acetate [103].

9nCH3COOH ! 4(CH3CH2CHOO~)n+2nCO2 + 6nH2O (10.14)

In 1937 Salvo Chemical Corporation started the industrial production of vanillin from lignin oxidation using spent liquors from pulp and paper industry by the Howard’s [96] patented technology [91, 97]. In Canada, Howard Smith Chemicals, Ltd. also began the industrial conversion of concentrated spent sulfite liquor to vanillin with technology based on the process developed by Hibbert and Tomlinson [98]. In the same country, one other industrial unit started up at about 1945 by Ontario Paper Co., Ltd. that, besides vanillin, recovered also fermentable sugars from liquor [91]. In the beginning of 1950s, Monsanto Chemical Company substituted the process of synthetic vanillin from eugenol by the lignosulfonates oxidation process using fermented spent sulfite liquor [97].

Until 1980s, the major vanillin supply market share was provided by the oxidation of lignin from the sulfite pulping [99]. After that, industrial units of lignin-derived vanillin faced constrains that forced them to close [91]. Addition­ally, since the 1980s, changes introduced in the pulp and paper industry processes led to a decrease of lignin availability: at that time, kraft process emerges as the competing pulping process that includes the recovery boiler for burning the spent liquor allowing the recovery of pulping chemicals and producing energy. Since then, guaiacol-based vanillin has gained relevance [97]. Nowadays the synthesis of vanillin from petrochemical guaiacol accounts for 85% of the world supply, with the remaining 15% being produced from lignin [99].

The high market prices of vanilla beans and their limited supply and the increasing concern for alternatives to the non-renewable raw materials, have encouraged the research for alternative pathways for natural flavor production. A detailed review on biotechnological routes of vanillin production using different substrates and biosynthesis methods can be found elsewhere [100, 101]. Com­mercially, vanillin obtained by fermentation from ferulic acid is considered the only profitable biocatalytic route [101, 102].

As future prospect, the concern for alternatives to the non-renewable raw materials and the emerging lignocellulosic-based biorefineries could lead to a promising future for lignin-derived vanillin; the condition is the profitability of the production process using as raw material spent liquor lignins or lignins from new biomass conversion technologies. Due to constant efforts of several R&D centers around world, many challenges for the profitable large scale application of lignin as source of aromatics, and vanillin in particular, are being progressively overcome.

Some microbes float freely in the solution around the minerals and some attach themselves to the mineral particles forming a biofilm. The microbes, whether they are freely floating or whether they are in the biofilm, continuously devour their food sources—iron (chemically represented as Fe2+) and sulfur. The product of the microbial conversion of iron is ‘‘ferric iron’’, chemically represented as ‘‘Fe3+’’. Ferric iron is a powerful oxidizing agent, corroding metal sulfide minerals (for example, pyrite, arsenopyrite, chalcocite, and sphalerite) and degrading them into a dissolved metal, such as copper, zinc, and more iron—the latter is the food source for the microbes.

The reaction of the biological oxidation involved in leaching of a mineral sulfide is

MS + 2O2 ! MSO4 where, M is a bivalent metal.

There are two major mechanisms involved in microbial metal solubilization of sulfide minerals. One is a direct mechanism that involves physical contact of the organism with the insoluble sulfide.

Microorganisms oxidize the metal sulfides obtaining electrons directly from the reduced minerals. Another, indirect mechanism, involves the ferric-ferrous cycle. The oxidation of reduced metals is mediated by the ferric (III) ion and this is formed by microbial oxidation of ferrous (II) ion present in the minerals. Ferric (III) ion acts as an oxidant and oxidizes metal sulfides and is reduced to ferrous (II) ion that, in turn, can be microbially oxidized. Both direct and indirect mechanisms of bacterial leaching are shown schematically in Fig. 14.9.

11.3.3.1 Cellulose/Pulp

The obtained cellulose has a high amount of cellulose, high proportion of para-crystalline and amorphous cellulose and lower DP as compared to the native material. Lower DP of the obtained material can improve the enzymatic hydrolysis due to the two factors [52]: (1) increasing the amount of the reducing ends of cellulose chain; and (2) making cellulose more amenable to enzymes. Cellulose with short chains allows it to be attacked by enzymes more easily because they form weaker networks rather than strong hydrogen bonding [19, 53]. This suggests that the cellulose-rich residue is amenable to the enzymatic deconstruction for the production of ethanol. Generally, high conversion of cellulose to glucose can be achieved up to 90-100% for most softwood and hardwood after ethanol fractio­nations [20, 45]. For instance, the ethanol/water treated B. davidii has been subjected to enzymatic hydrolysis applying cellulose (Cellucast 1.5 l) and b-glucosidase (Novozm 188). The data indicates that high conversion of cellulose to glucose up to 98% was achieved under the optimal conditions [39].

Most studies show that lignin removal enhances the enzymatic deconstruction of cellulose, since lignin inhabits cellulose activity [54-57]. While reducing the lignin content from 30 to 19%, the enzymatic hydrolysis was enhanced hugely, whereas further reduction in the lignin content to 9%, only negligible increase of enzymatic hydrolysis was observed [39]. An exception was the treatment of B. davidii, decreasing the lignin content of the sample did not increase the enzymatic hydrolysis. It seems that other factors influenced the enzymatic hydrolysis in addition to lignin content [39].

It is generally believed that amorphous cellulose is more easily attacked than crystalline cellulose [55]. A comparative study conducted by Jeoh et al. [58] showed that amorphous cellulose exhibited high enzymatic hydrolysis as compared to crystalline cellulose, due to formation of more extensive bonding between the reducing ending groups of amorphous cellulose and cellulase (Trichoderma reesei). With respect to enzymatic hydrolysis of the ethanol pretreated B. davidii, it was considered that a low CrI value of 0.55 was already efficient for hydrolysis, and further reduction of the value did not afford additional benefits for hydrolysis [39].

Compared to the conventional chemical pulping process, the obtained pulp from ethanol fractionation has a higher yield, easier bleachability and comparable pulp properties.

Poplar was subjected to ethanol pulping to optimize the process by varying the ethanol concentration, pulping time, pulping temperature and usage of catalyst (H2SO4). Even using 0.02% acid catalyst, the obtained pulp yield and viscosity were lower than the acceptable level; therefore, acid catalyst should not be added. This was due to the serious degradation of carbohydrates in an acid medium. Under optimal conditions, i. e., cooking at 180°C for 90 min with 50% ethanol, pulp was obtained with yield around 45%, viscosity 892 ml/g and kappa number 67 [59].

A significant feature of the ethanol/water produced pulp is that the pulp is easy to be bleached even with rather high kappa number. It has been reported that ethanol aspen pulp of a kappa number 30 was bleached to 81-86% ISO brightness applying a chlorine bleaching sequence (CEH) and to an higher brightness of 90% ISO via a chlorine dioxide bleaching (DED) sequence, whereas ethanol birch pulp of a kappa number 40 was bleached to 83% ISO brightness with a CEH bleaching sequence [60]. With respect to the ability of delignification in oxygen delignifi — cation process, hardwood ethanol pulp showed more extensive delignification extent than the corresponding kraft pulp. A delignification up to 75% was achieved without a significant reduction of pulp viscosity, and pulp was bleached to a brightness level greater than 92% ISO after either an ECF sequence or a totally chlorine free (TCF) sequence [61]. On the contrary, delignification extent of kraft pulp in oxygen delignification stage is below 50% to avoid the extensive degra­dation of carbohydrates.

It has been reported that ethanol pulp was also suitable for alkaline extraction and alkaline oxygen delignification [62]. Reduction of residual lignin prior to bleaching by alkaline extraction can reduce the amount of bleaching chemicals thus reducing the environmental impact of the bleaching process. After the wheat straw ethanol pulp with kappa number around 60 was extracted with 1% NaOH aqueous solution for 1 h, a large proportion of lignin was removed from the fiber [62].

However, an increase of alkaline concentration resulted in an increase of the lignin concentration on the fiber surface due to the enhanced adsorption of the dissolved lignin back on the fiber surface, similar to the phenomenon observed in alkaline extraction of kraft pulp.

Sugar bagasse pulps produced from ethanol/water organosolv process were used to produce carboxymethyl cellulose [63]. In this process, the acid-catalyzed ethanol pulp (prepared with 0.02 mol/l sulfuric acid at 160°C for 1 h) was bleached with sodium chlorite, and then was used to prepare carboxymethyl cel­lulose (CMC). The CMC yield was 35% (based on the pulp) with substitution up to 0.70 groups CH2COONa per unit of glucose residue.

Surface modification of cellulose fractionated from ethanol/water was conducted by heterogeneous esterification with octadecanoyl and dodecanoyl chloride [64]. After esterification, the modified cellulose showed strong reduction in the values of the polar parameter cpp, i. e., 4.4 and 1.8 for dodecanoyl and octadecanoyl cellulose, as compared to a high value of 35.7 for the original ethanol extracted cellulose. Since the esterified cellulose had a good fiber/matrix interfacial compatibility and low moisture uptake, it was a potential feedstock as reinforcing elements used in composite materials.

Sisal (Agave sisalana) ethanol pulp prepared from cordage residues was used as reinforcement to cement-based composites [65], and the prepared pulp/cement composites could further combine with polypropylene (PP) fibers. Compared to that added by kraft pulp, the composites with the addition of ethanol pulp showed lower modulus of rupture (MOR), limit of proportionality (LOP) and toughness. However, the performance of ethanol pulp reinforced composites was improved through a further optimization of pulping process. After 100 aging cycles (without fast carbonation), the ethanol pulp composites showed lower water absorption and apparent voids volume than that combined with PP.

After the extensive research work on lignin oxidation in batch mode, the contin­uous process is the natural next step in view of an eventual industrial application. An experimental pilot set up was built for lignin oxidation in continuous operating mode at the LSRE [119, 120]: a schematic diagram of the operational pilot installation is shown in Fig. 12.11. The bubble column reactor has a capacity of 8 l and is possible to fill it with modules of structured packing to improve the overall mass transfer performance of the system. This setup allows the operation with different flow conditions, for example in semi-batch mode (closed to gas or liquid), and is designed to work in very strong alkaline media (pH of 14), temperatures up to 170°C and pressures up to 15 bar. More details on the reactor set up can be found in the literature [119, 120, 122].

In a typical run, the alkaline solution of kraft lignin (60 g/l in NaOH 2 M) is fed to the reactor at 1-2.5 l/h, the temperature is selected (e. g. 403 K) and N2 is used to pressurize the system. The oxidation is initiated when the operating tempera­ture, pressure, and flow rates stabilized. Two different reactor configurations were tested: structured packed bubble column reactor (SPBCR) and bubble column reactor (with no internals, BCR).

The steady state condition was attained at approximately 6 h of operation for both types of reactor configuration. BCR was tested for O2 flow rates (QO2) of 1 L and 2 LNTP/min, reaching to the vanillin yields of 0.56 and 0.67 g/l, respectively. The higher yield of vanillin for higher QO2 is probably due to the higher pO2 which improved the oxygen solubility in the liquid phase. However, the continuous oper­ation led to about 25-30% of the maximum of vanillin concentration produced in the

Feed Deposit

Fig. 12.11 Layout of the experimental set-up of continuous lignin oxidation with O2 in alkaline medium. The design and construction were performed at LSRE within the PhD work of Daniel Araujo (advisor Professor Alirio Rodrigues) [119] (image was a courtesy of Dr. Daniel Araujo, FEUP, Portugal) batch process. In the SPBCR the hydrodynamics environment (dispersion coeffi­cient, phase hold up or even heat transfer coefficients) is quite different from the simple bubble column reactor. The oxidations performed at QO2 = 2,000 mlNTP/ min and feed rates of 2.0 l/h and 1.0 l/h lead, at steady state, to the final yields in vanillin of 0.73 and 0.89 g/l, respectively. The improvement on the vanillin content in the SPBCR configuration is due to an increase in the mass transfer of oxygen. The mass transfer coefficient was 35% higher for SPBCR than for BCR [119]. However, in both cases, the insufficient mass transfer of oxygen from the gas phase to the liquid phase was stated as the main reason for the low conversion.

To improve the performance of the continuous reactor and reach the production levels obtained in batch mode, the influence of some operating conditions was studied using a model developed by Araujo [119]. In this approach, pure oxygen in the gas feed was considered, decreasing simultaneously the residence time to avoid vanillin oxidation. From the SPBCR it was possible a vanillin concentration in the exit stream of 1.8 g/l, which is 85% of the maximum levels of vanillin concen­tration obtained in the batch reactor. Besides the low rates of oxygen transfer, the amount of vanillin produced is probably also limited by the type of lignin used.

Dr. Chinnappan Baskar is an Associate Professor of Chemistry and Academic In-charge, THDC Institute of Hydropower Engineering and Technology, Uttarakhand Technical University, Dehradun, India. He has received his M. Sc. Chemistry from the Indian Institute of Technology Madras and PhD in Organic and Materials Chemistry from the Department of Chemistry, National University of Singapore (NUS), Singapore under the direction of Prof. Suresh Valiyaveettil. He has joined the faculty in the Department of Chemistry, Lovely Professional University (LPU) as a Reader then promoted to Head of the Department (2006­2009). He moved to the Department of Environmental Engineering and Biotechnology, Myongji University, South Korea in September 2009 as a Brain Korean 21 (BK21) Research Professor and co-researcher in Energy and Environmental Fusion Technology Center, Myongji University. He has worked as Director (Academic Affairs), Dev Bhoomi Group of Institutions, Dehradun, Uttarakhand. Dr. Baskar research interests include synthetic organic chemistry, conducting polymers, green chemistry, production of biofuels and fine chemicals from biomass, ionic liquids, and membrane science separation. He has published several research papers in reputed international journals and conference proceedings. He was invited to attend and deliver lectures/seminars in international and national conferences & workshops. He serves on the Editorial Advisory Board member and referee for many international chemistry, materials science, biotechnology and energy journals.

Dr. Shikha Baskar obtained her PhD in Organic Chemistry from the Department of Biochemistry and Chemistry, Punjab Agricultural University, Ludhiana, Punjab, India under the guidance of Prof. Ranjit S. Dhillon and received postdoctoral training at Myongji University. She has joined as Sr. Lecturer/Assistant Professor and Head of Laboratory in the Department of Chemistry, Lovely Professional University, Phagwara, Punjab. She is currently Visiting Faculty of Chemistry at THDC Institute of Hydropower Engineering and Technology, Tehri, Uttarakhand. Her current research interests are in the areas of synthetic organic chemistry, green chemistry, ionic liquids, and production of biofuels. She has authored few peer-reviewed journal articles and attended many national and international conferences and workshop.

Dr. Ranjit S. Dhillon is a retired Professor of Organic Chemistry at the Punjab Agricultural University (PAU), Ludhiana, Punjab, India. He received his PhD from PAU under the direction of Prof. P. S. Kalsi. He spent three years as a Post­doctoral Fellow with Nobel Laureate Prof. Akira Suzuki at Hokkaido University, Sapporo, Japan. Dr. Dhillon has supervised 12 PhD students and 20 M. Sc. students in the areas of chemoselective green methodologies, natural products and their bioactive studies, and synthesis of eco-friendly Agrochemicals. His research work mainly ‘‘versatile boranes and borohydrides’’ carried out at PAU was cited by Nobel Laureate Late Prof. Herbert C. Brown in his research articles and many other eminent scientists. Professor Dhillon has published over 60 peer-reviewed papers and one book chapter. He is the author of Hydroboration and Organic Synthesis (Springer-Verlag Heidelberg, 2007).

Tugba Keskin and Patrick C. Hallenbeck

10.1 Introduction

The sustainability of economic growth and the ecology of the environment are under threat by rising petroleum prices and global warming. Much research is going into finding alternative reliable and effective energy sources. Among the energy sources under development, hydrogen is recognized as the most promising alternative to fossil fuels, and it is assumed that it will play a major role in the future energy supply because it is recyclable, energy efficient and clean energy carrier [1]. Hydrogen can be produced from fossil fuels by steam reforming or thermal cracking of natural gas, partial oxidation of hydrocarbons, coal gasifica­tion or pyrolysis. A second option is to produce hydrogen from water by elec­trolysis, photolysis, thermochemical processes or thermolysis. A third option is to produce hydrogen by biological processes such as biophotolysis of water by algae and cyanobacteria, photofermentation of organic substrates, or dark fermentation of organic substrates [2]. Among the various options for hydrogen production, biological production would appear to be the most efficient due to its low energy demand compared with physical and thermochemical processes, and due to its ability to use organic wastes. When biological hydrogen production is coupled with the treatment of organic wastes two main problems, the reduction of pollution from the uncontrolled degradation of waste, and the production of a clean fuel, can be solved [3].

T. Keskin • P. C. Hallenbeck (H)

Departement de Microbiologie et Immunologie, Universite de Montreal, CP 6128 Succursale Centre-ville, Montreal, QC H3C 3J7, Canada e-mail: patrick. hallenbeck@umontreal. ca

T. Keskin

Environmental Biotechnology and Bioenergy Laboratory, Bioengineering Department, Ege University, 35100 Bornova, Izmir, Turkey

C. Baskar et al. (eds.), Biomass Conversion,

DOI: 10.1007/978-3-642-28418-2_10, © Springer-Verlag Berlin Heidelberg 2012

Dark fermentation processes are well-known biohydrogen production methods. Acidogenic bacteria like Enterobacter, Bacillus and Clostridium are the main groups of hydrogen producing bacteria which convert organic substrates (e. g. glucose and sucrose) into soluble metabolites, i. e. volatile fatty acids (VFAs) and alcohols, as well as hydrogen. On the other hand dark fermentation by mixed anaerobic consortia is assumed to be more economical since this process does not incur sterilization costs, and hydrogen can be produced continuously since there is no direct light requirement. Proper pretreatment method allows the hydrogen producing species to dominate in the mixed culture, leading to utilization of carbohydrates with the formation of hydrogen and VFAs, as well as biomass, growth which lowers the experimental yields below theoretical values [4].

Photofermentation is hydrogen production from organic acids in the presence of photoheterotrophic bacteria under illumination with visible light. Purple non-sulfur bacteria are the main hydrogen producing organisms capable of photofermenta­tion. The bottleneck of photofermentation systems is the source of organic acids used as substrate. Pure organic acids are too expensive for the practical sustainable production of hydrogen. Organic wastes can be inexpensive, but they are not pure and can have a variable composition, affecting systems operation. Organic wastes like starch and cellulose cannot be metabolized by photofermentative bacteria. Using dark fermentation as a first step to convert waste and complex products into organic acids, and then using the produced organic acids for photofermentation can increase the maximum yield of hydrogen production from 4 to 12 mol of H2/mol glucose theoretically [5]. Dark fermentation is an incomplete oxidation of substrate, therefore converting the remaining organic acids by photofermentation can improve overall hydrogen yields. For an economically viable process, it is very important to combine dark fermentation with photofermentation to obtain high yields of hydrogen production.

Organosolv fractionation is considered to be an environmentally friendly process to afford substantially cellulose, hemicelluloses/degraded sugars and lignin for further process that is specific to each component. After organosolv fractionation, the recalcitrance of lignocellulosic material is destroyed to some extent regarding cellulose crystallinity, degree of polymerization, lignin structure, lignin removal, hemicelluloses solubilization, etc. The obtained cellulosic residue is an enzyme hydrolyzable substance for the production of biofuels. In addition, it can also be converted into pulp for the production of paper, silk and other modified products through further process. The efficient degradation and dissolution of lignin in organic solvents allow the highly selective delignification of lignocellulosic material without the addition of large amounts of inorganic catalyst. Due to the mild conditions in the extraction process, the lignin dissolved in the liquor is easy to be recovered without complicated purification schemes. The obtained sulfur — free organosolv lignin is an ideal renewable and alternative feedstock for a variety of petrochemical-based chemicals and materials, which have great potential markets as well as high value applications. The dissolved carbohydrates, furfural and HMF, can also be served as feedstocks for some chemicals and polymers.

Acknowledgements The authors wish to express their gratitude for the financial support from the State Forestry Administration (200804015/2010-0400706), the National Natural Science Foundation of China (30930073 and 31070526), Major State Basic Research Projects of China (973-2010CB732204), Ministry of Education (111), and Hei Long Jiang Province for Distin­guished Young Scholars (JC200907).

The original birch and spruce wood samples, as well as the biochar samples were analyzed by FTIR spectroscopy (Figs. 13.2, 13.3, respectively). Infrared band assignments for wood and biochar samples are given in Table 13.1 [30-32]. The spruce and birch biochar FTIR spectra were similar to those reported by Sharma et al. [32] of lignin-derived biochar, suggesting that the biochar was lignin derived.

The band at * 1,390 cm-1 was attributable to H-bonded hydroxyl groups (O-H stretch) in both wood and biochar samples. The OH peak intensity decreased after SCM treatment of both birch and spruce. The aliphatic bands (2,850-2,950 cm-1) increased after SCM treatment due to dehydration reactions. The C=O ester band at 1,730-1,735 cm-1 was eliminated after SCM treatment as a result of hemi — cellulose deacetylation. The presence of a new C=O signal at *1,698 cm-1 suggests the presence of a conjugated ketone (Hibbert’s ketones) resulting from lignin demethylation of aromatic methoxy groups or b-O-4 cleavage after SCM treatment [33]. The new peak in the biochar samples at *1,640 cm-1 was assigned to a C=C bond. The increase in peak intensities of bands at 1,600 and 1,510 cm-1 (aromatic skeletal vibrations) suggests that cleavage of the aliphatic side chains and condensation reaction occurred in lignin during biochar formation [31, 34]. Other bands in the biochar spectra correspond to a lone aryl C-H wag (*860 cm-1). The band at 1,373 cm-1 is probably due to both OH in-plane bending and CH bending [32].

To assess the level of lignin cross-linking during biochar formation lignin CIs were determined. For spruce and birch wood the CI was 0.511 and 0.433, respectively. The lower CI value for birch wood is due to the less condensed structure of hardwood lignin relative to the spruce softwood lignin [31]. The CI values for the SCM biochar from spruce and birch were respectively, 0.681 and 0.666 suggesting lignin cross-linking had occurred to a significant extent. For

comparison softwood kraft lignin (Indulin AT, Mead Westvaco) has a CI of 0.624 which is known to be highly condensed [29, 35].

PNS bacteria can use a wide variety of substrates as carbon and nitrogen sources. Different kinds of strains can have different pathways for using substrates to produce hydrogen. Lactate, acetate, butyrate, propionate and succinate are the mostly known simple organic molecules that serve as suitable electron donors.

Glucose can be used as substrate for photofermentative hydrogen production. Photofermentation by Rhodobacter sphaeroides; Rhodobacter capsulatus from glucose achieved rates of 250 and 88 ml/g/h hydrogen, respectively [104, 105]. Using sucrose as the sole carbon source for Rhodobacter capsulatus resulted in 60 ml/g/h hydrogen yield [105]. Lactate and malate are the most studied carbon sources for photofermentative hydrogen production [106]. These two substrates are known as the best carbon sources for PNS bacteria for hydrogen production. By using Rhodobacter sphaeroides as bacterium and lactate as carbon source hydrogen rates varying between 16.7 and 240 ml/l/h have been reported in dif­ferent batch reactor studies [107-110]. Rhodobacter capsulatus can metabolize lactate to hydrogen at rates of 105 and 130 ml/g/h [106, 111]. Rhodopseudomonas palustris and Rhodobacter rubrum can metabolize the lactate with hydrogen production rates of 82.7 and 20 ml/l/h, respectively [112, 113]. Malate is another important carbon source for PNS bacteria. Studies have shown that up to 58 ml/l/h hydrogen rates can be achieved by using different PNS bacteria [107, 112]. Since PNS bacteria can use volatile fatty acids produced in dark fermentation, acetate, butyrate or mixtures of different organic acids were used as carbon sources in different studies. Hydrogen rates changing between 1.6 and 26 ml/l/h were obtained with mainly Rhodobacter capsulatus and Rhodobacter palustris [99, 114].

Non-sulfur photo-heterotrophic bacteria are known to be very effective bio­catalysts for hydrogen production from food, alcohol distilling or sugar industry wastewaters. Environmental benefits could be gained by using wastewaters as substrates. While using the wastewater as a substrate for photofermentative hydrogen production it is necessary to choose a proper pre-treatment that will not damage the main substrate component by sterilization.

Dairy wastewaters can be a good candidate for biohydrogen production with a yield of 3.6 l H2/l-reactor [115]. Since brewery wastewaters contain useful com­pounds like amino acids, proteins, organic acids, sugar as well as vitamins they can be assumed as useful substrates for hydrogen production. By using Rhodobacter sphaeroides 2.24 l H2/l medium hydrogen rate was reported. The best hydrogen production is achieved by R. sphaeroides RV with a yield of 1.23 mol H2/mol glucose from ground wheat [116]. From crude glycerol by Rhodopseudomonas palustris 6 mol of H2/mol glycerol can be produced [117]. In addition, photofer­mentation of glycerol should be used as substrate in a continuous process [117]. Olive mill effluent (OME) with a high content of sugar, volatile fatty acids, polyalcohols and fats and an advantage of low nitrogen content can give a rate of 0.009 l H2/l/h by photofermentation with Rhodobacter sphaeroides [118]. It can be easily seen that PNS bacteria are capable of efficient conversion of organic acids to H2. Using the industrial wastewater with proper modifications of the system can be an ecologically viable solution. Most PNS bacteria cannot metabolize sugar totally but they can metabolize organic acid easily which can be a good solution for dark fermentation end products.