Since distillation is a highly energy-consuming process, several processes have been developed for purification of ethanol from fer­mentation broth: for example, solvent extraction, CO2 extraction, vapor recompression systems, and low-temperature blending with gasoline [9]. However, these processes are not established in the industrial pro­duction of ethanol.

Crop description. Virola surinamensis and V sebifera (see Fig. 4.25)— commonly known as ucuhuba, ucuiba, ucuba, muscadier porte-suif, and yayamadou—belong to the family Myristicaceae and grow in tropical swampy forests. Major producing countries are Brazil, Costa Rica, Ecuador, French Guiana, and Guyana. A typical tree is of medium height and can produce 60-90 L of oil each year. The seeds contain 65-76% oil. The yellow-brown aromatic oils from both varieties are very similar. Other related species, such as V otoba, which grows in Colombia and Peru, yield a fat similar to ucuuba, which is known as otoba butter or American nutmeg butter. Major fatty acids present in the oil are lauric

acid (15-17.6%), myristic acid (72.9-73.3%), palmitic acid (4.4-5%), and oleic acid (5.1-6.3%) [77, 87].

Main uses. This fat has been used traditionally in candle manufacture. The fat and pulverized kernels find use in traditional medicines. The tree has been proposed as a potential source of isopropyl myristate, which is used in cosmetic manufacture [186]. However, no references related to its use as a raw material to produce biodiesel have been found to date.

Corrosion of the engine parts has been one of the main reasons for not using alcohols as fuels. The problem of corrosion is severe during start­ing and idling; but once the engine starts and gets heated, corrosion does not take place. Severe corrosion is noticed with Zn, Pb, Cu, Mg, and Al. This problem has been solved by using a methanol-resistant filter before the carburetor. Corrosion by methanol has been prevented by using the corrosion inhibitor LZ541 manufactured by M/S Lubrizol India. Being solvent, it swells or softens many parts of plastic or rubber commonly used for gaskets or floats in the carburetor. This is solved by using elas­tomers instead of rubber or plastic. American Motors’ Gremlin model of 1970 has been used continuously for 9 years using pure methanol with­out facing any difficulty of corrosion. Two 1972 Plymouth Valiants have been used for 7 years: one using pure methanol and the other using a methanol blend without any difficulty. None of these vehicles has had a failure of engine components or fuel system components.

Prospects of ethanol and biodiesel as substitutes for conventional fuels will not be discussed here; these two aspects are presented in sufficient detail in Chaps. 3, 4, 5, and 6. One of the promising approaches for future fuel is, perhaps, hydrogen and methane, both of which could be obtained from living, particularly microbial resources.

Photosynthesis is the main route through which oxidized carbon is reduced and again oxidized back to carbon dioxide for the generation of energy. Based on this principle, we can utilize a few steps from this life chain. This topic could be called biophotolysis—alternatively, photobiolysis.

In the system, direct electron transport from water to hydrogen has not been demonstrated as a technically feasible reaction. For this, con­tinued research is required to elucidate the basic nature of FeS (PEA, ferredoxin, and hydrogenase). This may lead ultimately to the practical feasibility of production of hydrogen (ideally 20 ^L/h). Section 1.16 dis­cusses hydrogen in detail. One inherent problem is the stability of the hydrogenase system because of its sensitivity to molecular oxygen pro­duced during photosynthesis.

However, one may design a two-step or two-compartment system. Reduced Co II is the oxygen-stable electron carrier between photo­synthesis and hydrogenase. A higher ratio of reduced Co II or Co II helps the evolution of hydrogen, in spite of the unfavorable redox potential of the coenzyme. Only Co II (reduced) can be pumped or transported from one stage (compartment) to the other. Photosynthesis and hydrogenase systems have to be encapsulated or immobilized sep­arately in order to retain their respective activity; the two stages or compartments may be connected through fiber filters. An example could be to use appropriate algae to produce reduced organic com­pounds which can be pumped into bath of photosynthetic bacteria of hydrogen fermentation.

One partial modification will be to collect oxygen during the day and hydrogen at night, at the expense of accumulated reduced coenzymes, made operative by anaerobically adapted microalgae or nonheterocystous nitrogen — fixing blue-green algae. For product separation, the enzyme technology or immobilization is inapplicable for biophotolysis. However, there are potential practical applications of immobilized hydrogenase in biochemical hydrogen—oxygen fuel cells. If such enzymes can be immobi­lized on an electrode surface, an inexpensive fuel cell might be developed, which would increase the energy recoverable for hydrogen to save fuels.

Awareness of the limitations due to efficiency, engineering, and the economy of these principles will save disappointment and encourage con­tinued research. Geographical location and frequency of weather change limits the insolation. The best photosynthetic efficiency is only 6% of the total incident solar radiation, i. e., 5 kg/(m2 • yr) of H2 by biophotolysis. Half of this could be a very satisfactory achievement.

Chemical hydrolysis involves exposure of lignocellulosic materials to a chemical for a period of time, at a specific temperature, and results in sugar monomers from cellulose and hemicellulose polymers. Acids are predominantly applied in chemical hydrolyses. Sulfuric acid is the most investigated acid, although other acids such as hydrochloric acid (HCl) have also been used. Acid hydrolyses can be divided into two groups: concentrated-acid hydrolysis and dilute-acid hydrolysis [18].

Concentrated-acid hydrolysis. Hydrolysis of lignocellulose by concen­trated sulfuric or hydrochloric acids is a relatively old process. Concentrated-acid processes are generally reported to give higher sugar and ethanol yield, compared to dilute-acid processes. Furthermore, they do not need a very high pressure and temperature. Although this is a successful method for cellulose hydrolysis, concentrated acids are toxic, corrosive, and hazardous, and these acids require reactors that are highly resistant to corrosion. High investment and maintenance costs have greatly reduced the commercial potential for this process. In addi­tion, the concentrated acid must be recovered after hydrolysis to make the process economically feasible. Furthermore, the environmental impact strongly limits the application of hydrochloric acid [12, 15].

Dilute-acid hydrolysis. Dilute-sulfuric acid hydrolysis is a favorable method for either the pretreatment before enzymatic hydrolysis or the conversion of lignocellulose to sugars. This pretreatment method gives high reaction rates and significantly improves enzymatic hydrolysis.

Depending on the substrate used and the conditions applied, up to 95% of the hemicellulosic sugars can be recovered by dilute-acid hydrolysis from the lignocellulosic feedstock [2, 13]. Of all dilute-acid processes, the processes using sulfuric acid have been the most extensively studied. Sulfuric acid is typically used in 0.5-1.0% concentration. However, the time and temperature of the process can be varied. It is common to use one of the following conditions in dilute-acid hydrolysis:

■ Mild conditions, i. e., low pressure and long retention time

■ Severe conditions, i. e., high pressure and short retention time

In dilute-acid hydrolysis, the hemicellulose fraction is depolymerized at temperatures lower than the cellulose fraction. If higher temperature or longer retention times are applied, the monosaccharides formed will be fur­ther hydrolyzed to other compounds. It is therefore suggested that the hydrolysis process be carried out in at least two stages. The first stage is carried out at relatively milder conditions during which the hemicellulose fraction is hydrolyzed, and a second stage can be carried out by enzymatic hydrolysis or dilute-acid hydrolysis, at higher temperatures, during which the cellulose is hydrolyzed [13]. These first and second stages are some­times called “pretreatment” and “hydrolysis,” respectively.

Hydrolyzates of first-stage dilute-acid hydrolysis usually consist of hemicellulosic carbohydrates. The dominant sugar in the first-stage hydrolyzate of hardwoods (such as alder, aspen, and birch) and most agri­cultural residues such as straw is xylose, whereas first-stage hydrolyzates of softwoods (e. g., pine and spruce) predominantly contain mannose. However, the dominant sugar in the second-stage hydrolyzate of all lig­nocellulosic materials, either by enzymatic or dilute-acid hydrolysis, is glu­cose, which originates from cellulose.

Detoxification of acid hydrolyzates. In addition to sugars, several by-products are formed or released in the acid hydrolysis process. The most impor­tant by-products are carboxylic acids, furans, and phenolic compounds (see Fig. 3.6).

‘ Mannan—► Mannose —► HMF ► Acids

Xylan—- ► Xylose— ►Furfural—► Acids

I Glucan — ► Glucose— ► HMF — ► Acids

——————— ► Phenolic Compounds

Acetyl groups————————— ► Acetic acid

Figure 3.6 Formation of inhibitory compounds from ligno­cellulosic materials during acid hydrolysis.

Acetic acid, formic acid, and levulinic acid are the most common car­boxylic acids found in hydrolyzates. Acetic acid is mainly formed from acetylated sugars in the hemicellulose, which are cleaved off already at mild hydrolysis conditions. Since the acid is not further hydrolyzed, for­mation of acetic acid is dependent on the temperature and pressure of dilute-acid hydrolysis, until the acetyl groups are fully hydrolyzed. Therefore, the acetic acid yield in the hydrolysis does not significantly depend on the severity of the hydrolysis process [13, 19].

Furfural and HMF are the only furans usually found in hydrolyzates in significant amounts. They are hydrolysis products of pentoses and hexoses, respectively [13]. Formation of these by-products is affected by the type and size of lignocellulose, as well as hydrolysis variables such as acid type and concentration, pressure and temperature, and the retention time.

A large number of phenolic compounds have been found in hydrolyzates. However, reported concentrations are normally a few milligrams per liter. This could be due to the low water solubility of many of the phenolic com­pounds, or a limited degradation of lignin during the hydrolysis process. Vanillin, syringaldehyde, hydroxybenzaldehyde, phenol, vanillic acid, and 4-hydroxybenzoic acid are among the phenolic compounds found in dilute- acid hydrolyzates [18].

Biological (e. g., using enzymes peroxidase and laccase), physical (e. g., evaporation of volatile fraction and extraction of nonvolatile fraction by diethyl ether), and chemical (e. g., alkali treatment) methods have been employed for detoxification of lignocellulosic hydrolyzates [20, 21].

Detoxification of lignocellulosic hydrolyzates by overliming is a common method used to improve fermentability [22-25]. In this method, Ca(OH)2 is added to hydrolyzates to increase the pH (up to 9-12) and keep this condition for a period of time (from 15 min up to several days), followed by decreasing the pH to 5 or 5.5. Recently, it has been found that time, pH, and temperature of overliming are the effective param­eters in detoxification [26]. However, the drawback of this treatment is that part of the sugar is also degraded during the overliming process. Therefore, it is necessary to optimize the process to achieve a fer­mentable hydrolyzate without any loss of the sugar [21, 26].

Crop description. J. curcas—commonly known as pourghere, ratanjyot, Barbados nut, physic nut, parvaranda, taua taua, tartago, saboo dam, jarak butte, or awla—belongs to the family Euphorbiaceae and grows in hot, dry, tropical climates (see Fig. 4.6). It originated from South America and is now found worldwide in tropical countries. It grows wild especially in West Africa, and is grown commercially in the Cape Verde Islands and Malagasy Republic. The tree reaches a height of 8 m and is a tough, drought-resistant plant that bears oil-rich seeds prolifically under optimum growing conditions [75]. The seeds contain about 55% oil [76]. The oil contains a toxic substance, curcasin, which has a strong purging effect. Major fatty acid composition consists of myristic acid (0-0.5%), palmitic acid (12-17%), stearic acid (5-6%), oleic acid (37-63%), and linoleic acid (19-40%) [77].

Main uses. It has been cultivated as a drought-resistant plant in mar­ginal areas to prevent soil erosion. The oil has been commercially used

Figure 4.6 Jatropha curcas. (Photo courtesy of Piet Van Wyk and EcoPort [www. ecoport. org].)

for lighting purposes, as lacquer, in soap manufacture, and as a textile lubricant. It is also used for medicinal purposes for its strong purging effect. The leaves are used in the treatment of malaria. Products useful as plasticizer, hide softeners, and hydraulic fluid have been obtained after halogenation [75]. The wood is used for fuel. The cake, after oil extraction, cannot be used for animal feed due to its toxicity, but is a good organic fertilizer. The wood is very flexible and is used for basket making. A water extract of the whole plant has molluscicide effects against various types of snail, as well as insecticidal properties [77].

Recently, there has been considerable interest in the use of the oil in small diesel engines. This oil has great potential for biodiesel production [78-80]. Foidl et al. transesterified J. curcas oil, using a solution of KOH (0.53 mol) in methanol (10.34 mol) and stirring at 30oC for 30 min [81]. The ester fuel has high quality and meets the existing standards for vegetable-oil-derived fuels. Some researchers have proposed the use of immobilized enzymes such as Chromobacterium viscosum, Candida rugosa, and Porcine pancreas as a catalyst [82, 83]. Modi et al. have proposed the use of propan-2-ol as an acyl acceptor for immobilized Candida antarctica lipase B. Best results have been obtained by means of 10% Novozym-435 based on oil weight, with a alcohol-oil molar ratio of 4:1 at 50oC for 8 h [84]. Zhu et al. have proposed the use of a heterogeneous solid superbase cat­alyst (catalyst dosage of 1.5%) and calcium oxide, at 70oC for 2.5 h, with a methanol-oil molar ratio of 9:1 to produce biodiesel [85]. The lubrication properties of this biodiesel have also been taken into consideration [51].

Different techniques adopted for converting vegetable oils to biodiesel are (a) degumming of vegetable oils, (b) transesterification by acid or alkali, and (c) enzymatic transesterification.

5.2.1 Degumming of vegetable oils

Degumming is an economical chemical process involving acid treatment to improve the viscosity and cetane number up to a certain limit so that the blends of nonedible oils with diesel can be used satisfactorily in a diesel engine. It is a very simple process by which the gum of the veg­etable oil is removed to decrease the viscosity of oil by using an appro­priate acid that can be optimized for reduction in viscosity. The quantity of acid and the duration of the process are very important to obtain optimum results. Compared to transesterification, the process of degum — ming is simple, very easy, and less costly, and the reduction in viscosity of vegetable oil is very small.

Nag et al. [25] degummed karanja, putranjiva, and jatropha oils by phosphoric acid treatment. Before degumming the oils, the fuel properties of three oils have been measured and compared with diesel (Table 6.1). Acid concentrations of 1%, 2%, 3%, 4%, and 5% were used at 40°C with vigorous stirring. The stirring was continued for 10 min after adding the acid. After stirring, the mixtures were held for 1 week to complete the reactions and to settle the gum materials. Then the mixtures were filtered through a packed bed filled with charred sawdust. Viscosities of the fil­trate were then measured.

Performance and emission measurement. After studying the properties of the jatropha, karanja, and putranjiva oils, they were degummed. In this context, the Ricardo variable-compression engine (Ricardo & Co. Engineers Ltd., England, single cylinder, 3-in bore, 35/8 in stroke) was run with 10%, 20%, 30%, and 40% blends of degummed karanja, jatropha, and putranjiva oils with diesel at different loads (0-2.7 kW) and different timings (45°, 40°, 35°, and 30° bTDC [before top dead center]). To meas­ure emissions, an automotive exhaust monitor (model PEA205) and smoke meter (model OMS103, Indus Scientific Pvt. Ltd., India) were used.

Degumming by acid treatment lowers the viscosity. Viscosities of karanja, jatropha, and putranjiva oils degummed at 40°C and at various acid concentrations are shown in Fig. 6.1. Karanja oil with 4% acid treatment had the lowest viscosity, whereas jatropha and putranjiva oils both had the lowest viscosities with 1% acid treatment.

Effect of timing. By observing the performance data at various timings (45°, 40°, and 35° bTDC) in Fig. 6.2, it was concluded that at 45° bTDC timing, the nonedible karanja, jatropha, and putranjiva oils gave the highest yields, whereas at 40° bTDC timing, diesel gave the highest yield. That may have been due to the different ignition temperatures of the nonedible oils from diesel.

40
30
25
0 1 2 3 4 5
Acid concentration (%)
Figure 6.1 Viscosity versus acid concentration of jatropha, karanja, and putranjiva oils at 40°C.
Diesel Jatropha Karanja Putranjiva Oils Figure 6.2 Brake thermal efficiency at various timings of diesel and 20% vegetable oil blends at 1-kW brake power, 1200 rpm, and 20 compression ratio.

Performance of various blends. Performances of blends of degummed vegetable oil with diesel are shown in Figs. 6.3 and 6.4. The 20% blends of jatropha, karanja, and putranjiva oils with diesel gave quite satis­factory performance related to BSFC and brake thermal efficiency (^bt). Beyond the 20% blends, the cetane numbers and viscosities of the blends were not so effective.

Comparison of the performance of blends. As per Figs. 6.5 and 6.6, engine performance using jatropha and karanja oils was better than diesel but the use of putranjiva oil gave reverse results at all loads, although the results were more or less the same. Degummed karanja oil blends gave better performance, but at high loads, the performance of jatropha oil blends was better in comparison to the performance of karanja oil blends. The performance data showed that all three vegetable oils could be used as alternative fuels for diesel engines.

Effect of loads on emissions of vegetable oil blends and comparison. As per

Figs. 6.7 and 6.8, it is interesting to note that for the karanja, jatropha, and putranjiva oils, in every case, smoke and particulates decreased, which was very favorable in terms of their environmental impact on human beings. The rate of increase in smoke and particulate generation with the load of jatropha oil, in comparison to karanja and putranjiva

Diesel

IWWl 10% blend 20% blend 30% blend

Figure 6.6 Brake thermal efficiency versus brake power of diesel, 20% karanja oil, 20% jatropha oil, and 20% putranjiva oil blends at 1200 rpm, 45° bTDC, and 20 compression ratio.

oils, was very low. It is very interesting to observe that although the par­ticulates and smoke for all the oils decreased, jatropha oil blends gave the highest reduction.

In Figs. 6.9 and 6.10, the CO, CO2, NOx, and HC (hydrocarbon) emis­sions for the three nonedible oils were less in comparison to diesel at high loads. However, at low loads, emissions from the nonedible oils are almost parallel to diesel. Because of the higher ignition temperature of nonedible oils than diesel, the better combustion of these oils gave less exhaust emissions.

Thus, degumming is an economic chemical process for a 20% blend of karanja, jatropha, and putranjiva oils with diesel to have very satisfactory results. The degumming method, therefore, offers a potential low-cost method with simple technology for producing an alternative fuel for CI engines. Out of the three nonedible oils, jatropha oil was the most prom­ising to yield good performance and emissions at high loads in all respects. Comparing CO, CO2, NOx, HC, smoke, and particulate emis­sions from using the three nonedible oils, jatropha oil was very encour­aging (see Fig. 6.11). Considering the above-mentioned points, it can be concluded that the diesel engine can be run very satisfactorily using a 20% blend of vegetable oil with diesel at 45° bTDC, 1200 rpm, and 20 compression ratios. Any diesel engine can be operated with a 20% blend

of degummed vegetable oils as a prime mover for agriculture purposes without any modification of the engine.

The chemical nature of conversion products depends both on the structure or type of the zeolite used and the reaction temperatures, because restructuring occurs at the inner surface, which acts as a reaction vessel at the molecular scale. Specific reactions depend on the diameters of pores, the resident time of molecules within the pores or channels and voids of the microporous zeolite, and the temperature. The penetration of lipids into a zeolite is depicted in Fig. 8.5. The scheme is based on [22].

O

II C H

‘ C—— C17H35

HC O C C17H35

O

H2C ^-C^

O C17H35

O

Diffusion

Figure 8.5 Scheme of restructuring triglycerides with shape — selective H-ZSM-5 to aromatic hydrocarbons.

To demonstrate this influence of catalysts and reaction temperature on yields and products, Table 8.4 considers a shape-selective zeolite type H-ZSM-5, commercially available as Pentasil, PZ-2/50H, and Y — zeolite (DAY-Wessalith). The physical characteristics of oils formed from the conversion of animal fat (rendering plant) are depicted [56]. Yields are between 30% and 70%, depending on the type of zeolite and tem­perature. Net calorific values are in the range of 40 MJ/kg compared to

35 MJ/kg of animal fat. All reaction products show relatively low vis­cosity and densities.

Products at T = 400°C. Again, the chemical nature of products formed from animal fat was analyzed by spectroscopic methods (see Fig. 8.6). The IR spectrum reveals the hydrocarbon nature of products. The strong C-H stretching vibrations (frequencies) at 2900 cm-1 is characteristic for alkanes. Functional groups are widely missing. The comparison to diesel from a commercial gas filling station (imprinted spectrum) shows a sim­ilar pattern [37].

Proton resonance spectroscopy depicts the chemical environment of pro­tons in the product formed from the conversion of animal fat. Figure 8.7 shows the dominance of aliphatic protons at chemical shifts of 0.9-2.25 ppm. Aromatic protons absorb at 6.5-8 ppm. The inspection of the ratio of the integral of absorptions reveals 5% aromatics for catalysis at T = 450oC. This is also reflected in the 13C-NMR spectrogram (see Fig. 8.8). However, with increasing temperature in the catalytic bed, the content aromatic alkylbenzenes increase.

Using 13C-NMR spectroscopy in-depth mode (see Fig. 8.9), negative signals at 30-20 ppm are characteristic for CH2-groups. The intensity indicates the presence of long-chain hydrocarbons. Peaks between 140 and 120 ppm denote carbon atoms of aromatic systems. The low inten­sity reflects the low content. Obviously, catalytic cracking over a Y — zeolite widely preserves hydrocarbon moiety in vegetable oil.

Figure 8.6 IR spectrum of hydrocarbons derived from animal fat at 4000C (Y-zeolite catalyst, DAY-Wessalith).

9	4 ppm	1

These spectroscopic findings are confirmed by gas chromatogra­phy (GC) [56]. Pyrolyzates (see Fig. 8.10) and commercial diesel (see Fig. 8.11) have a similar GC pattern. However, crude conversion products contain more volatile hydrocarbons.

GC separation on an OV101 capillary [column: 20 m X 0.3 mm, split 1:25; temperature program: 25°C (2 min), 4°C/min to 320°C] reveals double peaks in more detail (see Fig. 8.12). The first peak is for the alkene with a double bond of a given C number. The second peak is for the alkane having the same C number.

cfi
О >

You may use these hydrocarbons as a base for biofuels. However, there are markets for certain fractions of this hydrocarbon mixture. For example, the C-12 to C-18 fraction is a raw material widely used for bulk com­modities. As mineral oil prices increase, it is becoming more financially viable to produce chemical feedstock for commodities and specialities from wastes. Wastes are an energy and carbon source of the future.

Products at T = 550°C. For a given H-ZSM-5-zeolite, the nature of con­version products of lipids (animal fat) shifts to more aromatic com­pounds as the temperature increases. This is demonstrated by different NMR findings [56] for animal fat as a substrate at a reaction temperature of T = 550°C (see Figs. 8.13 through 8.15). Especially, DEPT-135 13C-NMR pattern of oil from catalytic conversion of animal fat at 550°C shows the dominance of aromatic protons and a very low amount of CH2 groups. Chromatographic separation revealed alkylbenzenes (especially 1,3,5-trimethybenzene) as main products [38].

Heating oil and a conversion product from animal fat have been used in a commercial burner (Buderus, Germany). Both oils resulted in emis­sions within legal limits (see Table 8.5).

A straightforward approach to apply vegetable oil in the most-talked — about biomass-to-liquid-fuel scheme is to use it as a co-substrate in mineral oil refineries. Advantages are low investments for peripheral facilities such as loading and storage and use of an existing infrastruc­ture for distribution and marketing. The processing of rapeseed oil as a feed component in a hydrocracker was described in 1990 [39]. The results are summarized in Table 8.6.

It is worth mentioning that rapeseed oil is converted in the hydrotreat­ment step to paraffins. The oxygen content of the vegetable oil causes an increased consumption of hydrogen to form water. Changes in quality

occur in the middle distillate. A lower density and a higher cetane number are a quality-enhancing advantage. A drawback is the susceptibility to freezing point of the fuel. This kind of cold flow behavior would make its use in winter impossible unless special additives are supplemented [40].

Many or most organic cellulosic matter, after proper mechanical treat­ment (homogenizing), can be put to microbial conversion for (a) bio — methanation and/or (b) hydrogen production.

1. Biomethanation can utilize human or animal excreta as well as mixed green/organic wastes. This part has been discussed earlier in Secs. 1.12 and 1.13.

2. Hydrogen production is discussed hereafter.

Biohydrogen. Major routes are

1. Enzymatic (partly microbial) through microbial routes

2. Klebsiella and Clostridium groups of microbes

3. Different cyanobacteria (blue-green algae)

4. Various photosynthetic bacteria

5. Many aerobes, i. e., bacilli and alkaligenes

6. Facultative groups, i. e., enterobacters, and coli forms

7. Various anaerobes, i. e., rumens, methanogenic, methylotropes, and clostridia

Enzymatic. Glucose dehydrogenase oxidizes glucose into gluconic acid and NADPH, which helps the reduction of H+ by hydrogenase. Glucose dehydrogenase and hydrogenase are purified from Thermoplasma aci- dophylium and Pyrococcus furiosus (optimal growth at 59°C and 100°C, respectively) (Woodward).

Based on metabolic patterns, the microbial systems may be of four types:

1. Photosynthetic microbes evolving H2 mediated through NADPH (Nicotine Adenine Dinucleotide Phosphate [Coenzyme II-reduced]) by photoenergy.

2. Cytochrome systems operating in facultative anaerobes that convert mainly formates to H2.

3. Cytochrome containing strict anaerobe, Desulfovibrio desulfuricans.

4. Clostridia, micrococci, methanobacteria, and others, without cytochrome, anaerobic heterotrophs.

Klebsiella oxytocae. ATCC (American Type Culture Collection) 13182 can convert formates to H2 (100%), but only 2 moles of H2 for each mole of glucose (5%). C. butyricum can convert glycerol to 1,3-propanediol, butyric acid, 2,3-butanediol, formic acid, and CO2 and H2. Klebsiella pneumoniae can convert glycerol into 1,3-propanediol, acetic acid, formic acid, and CO2 and H2. The presence of acetate enhances the production of butyrates and H2, and less propanediol.

Before discussing cyanobacteria and photosynthetic bacteria, we should review the basic reactions involved in photosynthesis, i. e., steps in so-called photophosporylation:

H2O + NADP+ + PO4 + ADP—^ O2 + NADPH + H+ + ATP

CO2 + NADPH + H+ + ATP———— ► (HCOH)n + NADP+ + ADP + PO4

+hv

Aerobic: 6CO2 + 6H2O ———- * C6H12O6 + 6O2

+hv

Anaerobic: Isopropanol or H2S + CO2————- ► Acetone or S +

(CH2O)n H2O

Cyanobacteria. Popularly known as blue-green algae, and justifiably so (they consume CO2 and evolve O2), they are bacteria (absence of nuclei, mitochondria, chloroplasts, etc.) as well as algae.

Cyanobacteria are oxygenic photoautotrophs, possessing photo I and II systems. Cyanobacteria have been well studied, and the details of their physiology and biochemistry are available in reviews and books. They are held by many scientists as potential sources of chemicals, bio­chemicals, food, feed, and fuel. Most of them are molecular nitrogen fixers and possess a nitrogenase system for H2 production. They are found to be symbiotic to cycads, lichens, and so forth. Some are hetero­cystous, lacking photolysis of water, and produce H2 through the nitro — genase step (when N2 is low). The nonheterocystous species produce H2 at higher efficiency at low N2 and O2 concentrations. Some of the species favor anoxic and dark conditions, but with the presence of organic sub­strates. They may even use sulfides as a source of electrons under an anaerobic environment. They are highly adaptable to a changing envi­ronment and are widely found in salty or sweet water, deserts, hot springs (up to 75°C), as well as Antarctica. Some heterocystous Anabaena exhibit H2 production in an atmosphere of argon and absence of molecular nitrogen. This was the clue to the knowledge that the enzyme nitrogenase, the main biocatalyst for molecular nitrogen fixa­tion, is present in cyanobacteria and is the key route of H2 production:

N2 + 8H+ + 8e~ + 12ATP ^ 2NH3 + H2 + 12ADP + 12Pi

A “reversible hydrogenase” (in photolysis of water, 2H2O ^ 2H2 + O2), is present in both heterocyst and vegetative cells and produces H2 at a lower rate than a nitrogenase. An “uptake hydrogenase” also operates (minor) connected to cytochrome chain, providing both H+ and elec­trons. H2 evolution is common, but the photolytic O2 is inhibitory to nitrogenases, which is protected by other biochemical and structural alternatives existing in heterocysts.

Large amounts of ATP, which is required for the reaction are gener­ated in the event of photosynthesis and respiration. The electron (reduc — tant) supply in the nitrogenase equation comes from metabolites, i. e., amino acids, mainly from carbohydrates (maltose, glucose, fructose, other pentoses, tetroses, etc.), produced and stored in the vegetative cells through photo I and II systems.

Nitrogenase Co II, i. e., NADPH (gained through the pentose phosphate route) happens to be an electron donor through NADP oxidoreductase/ ferredoxin or flovodoxin. Other electron-supplying batteries are also envisaged.

1. Through uptake hydrogenase-ferredoxin (photoactivated)

2. Through pyruvate-ferredoxin oxidoreductase

3. Reduced ferredoxin from isocitrate dehydrogenase

4. NADH generated in the glycolytic route

Under anaerobic or low aerobic conditions, nitrogenase activity may exist in vegetative cells, but H2 generation is of poor order.

Photosynthetic bacteria. Hydrogen production is guided by the surplus of ATP and reductant organic metabolites (carbon sources from the

Krebs cycle) and reduced nitrogen sources (glutamate/aspartate). Interactions of hydrogenase and nitrogenase may be complementary or competitive in different species or mutants. Nitrogenase (Mo, Ni, or Fe) also with mixed isozymes are reported. Some mutants liberate H2 more efficiently, utilizing DL-malate, D-malate, and L-lactate. Photoautotrophic growth is found to be less efficient in producing H2 than photoheterotrophic growth with limited nitrogen in nutrients. Normally, in photosynthetic bacteria, hydrogenase utilizes the hydrogen as a reductant for CO2 fix­ation and also for fixing molecular nitrogen. Nitrogenase reduces molec­ular nitrogen, along with the production of molecular hydrogen at the expense of almost six stoichiometric equivalents of ATP. This means that concurrent nitrogenase activity during photosynthesis competitively con­sumes the ATP that is produced and lowers the CO2-fixing efficiency.

Rhodospirillum and Rhodopseudomonas grow aerobically in the dark. But Rhodospirillum rubrum growing on glutamate (a nitrogen source) exhibit good hydrogen release during photosynthesis. Quantitative pro­duction of hydrogen has also been observed, growing on acetate, succi­nate, fumarate, and malate, by photosynthesis, initially in the presence of limited ammonium salts.

In Rhodopsuedomonas acidophilla, hydrogenase and nitrogenase are genetically linked. Several species of Rhodospirillaceae can perform nonnitrogenase-mediated hydrogen production in the absence of light, using glucose and organic acids including formates. Different strains of Rhodopseudomonas gelatinus and Rhodobacter sphaerolides exhibit highly efficient production of hydrogen [90 pL/(h • mg) cell] grown in a glutamate—malate medium.

In some cultures of Rhodopseudomonas capsulata, R. rubrum, and Rhodomicrobium vannielli, replacement of glutamate by N2 gas improved productivity of H2 (760 mL/d, 10 days) decreasing a little on aging. The model of a nozzle loop bioreactor, with immobilized R. rubrum KS—301 in calcium alginate, initial glucose concentration of 5.4 g/L, 70 h at 30°C, showed production of hydrogen 91 mL/h (dilu­tion rate of 0.4 mL/h). Improvement was suggested by using an agar gel for immobilization.

Aerobes.

1. Bacillus licheniformis isolated from cattle dung showed production of H2 in mixed culture media. Immobilized on brick dust, the aerobe maintained H2 production for about 2 months in a continuous system, with an average bioconversion ratio of 1.5 mole of H2 per mol of glucose.

2. Alcaligenes eutrophus, when grown on gluconates or fructose anaer­obically, produces H2. Hydrogenase directly reduces the coenzyme using hydrogen, and the excess hydrogen is spilled out. Higher con­centration of formate reduced hydrogen production.

HCOOH ^ CO2 + H2

Facultative anaerobes.

1. Enterobacter: Enterobacter aerogenes, as an example, can use varied and mixed nutrients, i. e., glucose, fructose, galactose, mannose, pep­tones, and salts (pH 4.0, 40°C); and may show activity for about a month in a continuous culture; evolution of hydrogen was about 120 mL/h/L of medium; 0.8 mol/mol of glucose. Accumulation of acetic, lactic, or succinic acids is likely to cause antimetabolic suppression in older cultures.

2. Escherichia coli: Anaerobically, it can use formate to produce CO2 and H2. Carbohydrates as nutrient sources usually end up with mixed products, i. e., ethanol, acetate, hydrogen, formate, carbon dioxide and succinate.

Various anaerobes.

1. Ruminococcus albus mostly converts cellulose to CO2, H2, HCOOH, C2H5OH, CH3, and COOH. Pyruvatelyase may be functional in the production of H2 (237 mol/mol of glucose). Further details are not available.

2. P. furiosus (thermophilic archeon) possesses nickel-containing hydro — genase and produces hydrogen using carbohydrate and peptone, at 100°C. The metabolic system seems to be uncommon to those of non — thermophiles.

3. Methanobacterium (Methanotrix) soehngenii (methanogens) can grow on acetate and salts media, but can split formate into hydrogen and carbon dioxide. M. barkeri, in the presence of bromoethane sulphonate, has suppressed methane production; instead, hydrogen, carbon dioxide, carbon monoxide, and water were produced.

4. Methylomonas albus BG8 and Methylosinus trichosporium OB3b (methylotrophs) used various substrates, i. e., methane, methanol, formaldehyde, formate, pyruvate, and so forth. But formate was found to be most useful for production of hydrogen under anaerobic conditions.

5. C. butyricum, C. welchii, C. pasturianum, C. beljerinscki, and so forth are very efficient in utilizing different carbohydrate sources and even effluents to produce hydrogen (see Fig. 1.14). Immobilization of these cells has also been successful

Removal of or reducing concentrations of either CO2 or H2 or the com­bination of both is likely to favor a forward reaction, i. e., to improve pro­duction of H2. Attempts to remove CO2 by collecting the evolved gases through 25% (w) NaOH solution, using E. aerogenes (E 82005), showed better production of H2, which improved further by enriching nitroge­nous nutrients in the culture media—from 0.52 moles of H2 per mole of glucose, increased to 1.58 moles [9].

Similar attempts are made using E. cloacae and reducing the partial pressure of H2 during the production of gases, by reducing the operat­ing pressure of the reactor and simultaneous removal of CO2 [through 30% (w/v) KOH], maintaining an anoxic condition by flushing Ar at the onset [10]; by reducing the operating pressure to 0.5 atm, the molar ratio of H2 yield per mole of substrate doubled (1.9-3.9). Other technical and economic benefits were also cited. There are other similar claims of improved biohydrogen production [11], using altered nutrients (20 g of glu­cose, 5 g of yeast extract, and 5 g/L of tryptone) and different mutants of E. aerogenes HU-101. HU-101 and mutants A1, HZ3, and AAY, respec­tively yielded 52.5, 78, 80, and 101.5 mmol of hydrogen per liter of media.

In batch processes, all nutrients required for fermentation are present in the medium prior to cultivation. Batch technology had been preferred in the past due to the ease of operation, low cost of controlling and moni­toring system, low requirements for complete sterilization, use of unskilled labor, low risk of financial loss, and easy management of feed­stocks. However, overall productivity of the process is very low, because of long turnaround times and an initial lag phase [9].

In order to improve traditional batch processes, cell recycling and application of several fermentors have been used. Reuse of produced cells can increase productivity of the process. Application of several fermentors operated at staggered intervals can provide a continuous feed to the distillation system. One of the successful batch methods applied for industrial production of ethanol is Melle-Boinot fermen­tation. This process achieves a reduced fermentation time and increased yield by recycling yeast and applying several fermentors operated at staggered intervals. In this method, yeast cells from pre­vious fermentation are separated from the media by centrifugation and up to 80% are recycled [9, 68]. Instead of centrifugation, the cells can be filtered, followed by the separation of yeast from the filter aid using hydrocyclones and then recycled [69].

In well-detoxified or completely noninhibiting acid hydrolyzates of lignocellulosic materials, exponential growth will be obtained after inoc­ulation of the bioreactor. If the hydrolyzate is slightly inhibiting, there will be a relatively long lag phase during which part of the inhibitors are converted. However, if the hydrolyzate is severely inhibiting, no conversion of the inhibitors will occur, and neither cell growth nor fer­mentation will occur. A slightly inhibiting hydrolyzate can thus be detoxified during batch fermentation. However, very high concentration of the inhibitors will cause complete inactivation of the metabolism [18].

Several strategies may be considered for fermentation of hydrolyzate to improve the in situ detoxification in batch fermentation and obtain higher yield and productivity of ethanol. Having high initial cell den­sity, increasing the tolerance of microorganisms against the inhibitors by either adaptation of cells to the medium or genetic modification of the microorganism, and choosing optimal reactor conditions to minimize the effects of inhibitors are among these strategies.

Volumetric ethanol productivity is low in lignocellulosic hydrolyzates when low cell-mass inocula are used due to poor cell growth. Usually, high cell concentration, e. g., 10 g/L dry cells, have been used in order to find a high yield and productivity of ethanol in different studies. In addition, a high initial cell density helps the process for in situ detox­ification by the microorganisms, and therefore, the demand for a detox­ification unit decreases. In situ detoxification of the inhibitors may even lead to increased ethanol yield and productivity, due to uncoupling by the presence of weak acids, or due to decreased glycerol production in the presence of furfural [21]. Adaptation of the cells to hydrolyzate or genetic modification of the microorganism can significantly improve the yield and productivity of ethanol. Optimization of reactor condi­tions can be used to minimize the effects of inhibitors. Among the dif­ferent parameters, cell growth is found to be strongly dependent on pH [18, 21].