Interest in using lipases as enzymatic catalyst for the production of alkyl fatty acid esters continuously grows. Some people work on the triglyceride by converting them to methyl esters, while some work on fatty acids. One of the main obstacles to the biocatalytic production of biodiesel is high cost of the enzyme; enzyme recy­cling might be the solution to this problem.

It was found that Pseudomonas fluorescens lipase immobilized on kaolinite lost one third of its activity when it is used for the second time, but no further decrease was observed in successive applications. The initial decrease in activity was put down to enzyme desorption from the solid support that was not observed after repeated (ten times) use [36 ] . Repeated batch reactions revealed that Rhizomucor miehei lipase had high stability, which retained about 70% of its initial conversion after 8 cycles (24 h each cycle). Meanwhile, under the same experimental conditions, Thermomyces lanuginosa retained only 35% of the initial conversion. This differ­ence was credited to factors such as inactivation of the biocatalyst in the oil phase, the type of carrier which is used for the immobilization, or enzyme sensitivity to long-term methanol exposure [79].

It is observed that various substances can slow down lipase activity (methanol, glycerol, phospholipids); however, a number of ways have been proposed to over­come these problems [71]. Commonly, biocatalytic process does not produce soaps or other by-products. If the reaction completes, only esters and glycerol are pro­duced. This makes purification steps much simpler and consequentially lowers plant costs. Once immobilized, lipases can be used many times or even in continuous processes. This resolves the major disadvantage related to high cost. In conclusion, if obtained by biocatalysis, biodiesel is an environmentally friendly fuel which will contribute to reducing negative impacts to the environment.

Modi et al. [56] used Propan-2-ol as an acyl acceptor for immobilized lipase- catalyzed preparation of biodiesel. The optimum conditions set for transesterification of crude Jatropha oil were 10% Novozym 435 (immobilized Candida antarctica lipase B) based on oil weight and alcohol to oil molar ratio of 4:1 at 50°C for 8 h. The maximum conversion reached using propan-2-ol was 92.8% from crude Jatropha oil. Reusability of the lipase was preserved over 12 repeated cycles with propan-2-ol as it reached to zero by the seventh cycle when methanol was utilized as an acyl acceptor, under standard reaction condition [55] .

Modi et al. [55, 57] explored ethyl acetate as an acyl acceptor for immobilized lipase-catalyzed preparation of biodiesel from the crude oil of J. curcas. The opti­mum reaction conditions set for interesterification of the oils with ethyl acetate were 10% of Novozym 435 (immobilized C. antarctica lipase B) which is based on oil weight, ethylacetate to oil molar ratio of 11:1, and the reaction period of 12 h at 50°C. Under the above optimum conditions, the maximum result of ethyl esters was 91.3% of purity. Reusability of the lipase over repeated cycles in interesterification and ethanolysis was also examined under standard reaction conditions. The relative activity of lipase could be well preserved over 12 repeated cycles with ethyl acetate as it reached to zero by the sixth cycle when ethanol was used as an acyl acceptor.

Shah and Gupta [73] conducted a process of optimization of monoethyl esters of the long chain fatty acids (biodiesel) by alcoholysis of Jatropha oil using lipase. The process includes (a) screening of various commercial lipase preparations, (b) pH tuning, (c) immobilization, (d) varying water content in the reaction media, (e) vary­ing amount of enzyme used, and (f) varying temperature of the reaction. The best yield 98% (w/w) was obtained by using Pseudomonas cepacia lipase which was immobilized on celite at 50°C in the presence of 4-5% (w/w) water in 8 h. The yield was not affected if analytical grade alcohol was replaced by commercial grade alco­hol. This biocatalyst could be applied four times without loss of any activity.

In order to lower the cost of biodiesel fuel production from Jatropha, Tamalampudi et al. [ 85] used lipase producing whole cells of Rhizopus oryzae which was immobilized onto biomass support particles. The activity of R. oryzae was compared to Novozym 435, the most effective lipase. Methanolysis of Jatropha oil progresses faster than other alcoholysis regardless of any lipase used. The maxi­mum methyl ester content in the reaction mixture reached 80 wt.% after 60 h using R. oryzae, but 76% after 90 h using Novozym 435. Both lipases could be used for repeated batches. They also exhibited more than 90% of their initial activities after 5 cycles. Whole-cell immobilized R. oryzae is a promising biocatalyst for produc­ing biodiesel from oil [85] .

Figure 6 shows the comparison of environmental impact of biodiesel production in each step of life cycle. Biodiesel production from palm oil has bigger total environ­mental impact than that of Jatropha oil. Cultivation contributes to the highest envi­ronmental impacts compared with other stages in the life cycle impact.

Figure 5 shows the downstream processing steps required to produce biodiesel from microalgae, while Table 2 provides a list of different technologies currently avail­able for each step. After the microalgal culture is harvested from the bioreactor, it is concentrated in the dewatering step to yield a wet paste. The microalgal pellets then undergo a pre-treatment step for preparation towards lipid extraction where lipids are extracted and separated from the cellular materials. The extracted lipids are then purified in the fractionation step before they are converted to biodiesel in the methy — lation step.

Large scale cultivation of microalgae is generally performed in either a raceway pond or a photobioreactor [11]. Even though there are numerous systems for microalgal cultivation, they can generally be classified into outdoor or indoor systems. In an outdoor system, the microalgae are grown in the open environment with variable culturing conditions, such as temperature and light intensity, and can

potentially exhibit inconsistent growth rate and unpredictable biochemical evolu­tion. On the other hand, the microalgae grown in an indoor system are placed in a greenhouse-type structure where cultivation conditions and biochemical evolution of the microalgal biomass can be tightly controlled. Even though the indoor system provides a more sound protection against local species invasion, it is not preferred due to its high operating cost. In either system, the culture must be aerated with CO2 supply and replenished with growth medium consisting of essential elements such as nitrogen, phosphorous and iron [11].

Once the microalgal culture is harvested from its cultivation system, it exists as a very dilute aqueous suspension (between 0.1 and 1 wt.%). Since such large vol­umes of water are undesirable for downstream processing, the culture needs to be substantially concentrated [15, 45] . Different solid-liquid separation techniques, such as centrifugation, fi. tration, flocculation, are used to dewater the microalgal

culture up to a solid concentration of 10 wt.% (Table 2). A cost-viable and energy — efficient concentrating technology is a current research endeavour.

After dewatering, the semi-wet microalgal paste undergoes pre-treatment steps before lipid extraction. Even though this pre-treatment step can be omitted, it is generally undertaken since the alteration of biomass conditions occurring in this step could lead to enhanced efficiency of subsequent lipid extraction [32, 33]. The treatment can be performed in multiple steps or as a single process. As one alterna­tive, residual water which is known to prohibit effective lipid mass transfer during the extraction step is completely removed from the paste through simple drying. The desiccated cells can then be milled into powder before being extracted with eluting solvents. In a different scenario, the paste can be exposed to disruption methods which disintegrate cellular structures and force the release of intracellular lipids to the surrounding medium, thereby easing the lipid extraction process. Effects of cell pre-treatment on lipid extraction are further discussed in Sect. 5.3.

After pre-treatment, the microalgal biomass undergoes a lipid extraction step that results in the separation of lipids from the cellular matrix. The principles and

processes involved in lipid extraction are discussed more thoroughly in Sect. 5. The isolated lipids are fractionated in order to remove unwanted non-lipid contaminants (such as carbohydrates, proteins and chlorophylls) and unusable lipid fractions (such as polar lipids and non-saponifiable neutral lipids) from the acylglycerols. Even though experimental methods, such as liquid chromatography and urea crys­tallizations, are available to perform this step on a bench scale [39], none of them is yet economically feasible to be retrofitted to a commercial scale.

In the transmethylation (also known as transesterification) step, the acylglycerols in the lipid extract are converted to fatty acid methyl ester (FAME) or biodiesel. Although a laboratory method to transmethylate polar lipids to biodiesel exists using toxic boron trifluoride or BF3 [14, 61], the industrial applicability of this method has not been tested. The method described here employs an alkaline catalyst (either NaOH or KOH) specific to acylglycerols, and is used to industrially transm­ethylate plant and animal oils which consist mainly of TG. The lipids are reacted with methanol and the select alkaline catalyst in a specific ratio depending on the concentration of TG and the presence of free fatty acids. The final FAME conver­sion is a function of reaction temperature and duration. During the transmethyla­tion, the alkaline catalyst cleaves the ester bonds holding the fatty acids to the glycerol backbone (Fig. 6) [11], The liberated fatty acids are then reacted with methanol to form new molecules with lower viscosity (FAME). The by-products (glycerol, reformed catalyst and excess methanol) are separated from the desired FAME via gravity settling as well as repeated water washings [17, 19, 31].

In order for biodiesel from microalgae to be environmentally sustainable, the total CO2 emitted in the downstream processes to produce usable biodiesel needs to be lower than or at least equal to the total CO2 originally captured by the microal­gae. Therefore, processes selected in each step should aim at minimizing energy consumption.

Pyrolysis systems cause many changes to the initial feedstock that inevitability is reflected in the biochars structural and elemental composition. These intensive thermal conditions during pyrolysis cause decomposition of organic structures from the raw feedstock through dehydrogenation, demethylation, and finally decar­boxylation resulting in the release of a variety of organic compounds, including volatile C compounds, CH4, and CO [7]. By assessing the elemental composition of the raw feedstock and the biochar, a determination of these released volatile com­pounds containing C, H, and O will result in major shifts in their atomic O/C and H/C ratios (Fig. 7).

The Van Krevelen diagram is a convenient way to show that the raw feedstocks are rich in H and O, and as the pyrolysis temperature increases, loss of volatile elements cause biochars to have decreasing O/C and H/C atomic ratios (Fig. 7). Consequently, manufacturers can quickly assess the degree of biochar production by examining for changes in the elemental concentrations of C, H, O, and N, and their associated ratios. For example, low H/C and O/C ratios indicate that the biochar is higher in aromatic structures [7, 48] . Biochars with O/C and H/C ratios in the

0. 3-1.2 range indicate that it contains lignin and polysaccharide-like compounds [48]. Krull et al. [56] has listed atomic ratios, including the OC contents, in biochars processed from several feedstocks and pyrolysis temperatures.

Computation of a biochars atomic ratio requires that a sample be digested result­ing in its loss for future experiments. Alternative, nondestructive methods for

biochar characterization are available, such as solid-state 13C nuclear magnetic reso­nance (NMR, [56]), and Fourier transformed infrared spectroscopy (FT-IR; [80, 84]). If 13C NMR spectroscopy is used, each sample analyses may take several hours to 1 day to acquire the spectral pattern. As presented below, 13C NMR spectroscopy is a more practical tool for examining progressive structural changes in biochars with increasing pyrolysis temperatures. Research has shown that plant-based feedstocks pyrolyzed between 350 and 400°C, cellulose and hemicellulose degradation occurs

[7] . In the mid-range temperature of 400-500°C, additional structural modifications can occur through condensation of aromatic molecules in the basal sheets followed by loss of functional groups as a result of decarboxylation and demethylation reac­tions. At the higher pyrolysis temperature regime (500-700°C), biochars will be dominated by aromatic-C groups, with minor contributions of carbonyl-C, 0-alky l-C, and alkyl-C moieties [56, 74]. The dominance of C in aromatic groups in high tem­perature pyrolyzed biochar is evident when plotting the 1 3C distribution in each biochars aliphatic, aromatic, and carbonyl region of the NMR spectral patterns (Fig. 8). Biochars pyrolyzed from switchgrass and peanut hull feedstocks at 500°C had the highest aromatic-C character (82%) among the 11 biochars evaluated. Lower temperature pyrolyzed biochars (250-350°C) have more C as aliphatic struc­tures because their polysaccharide-like compounds have not been lost to thermal degradation [5] .

As shown in Fig. 9 (top), the 13C NMR spectra of cotton gin trash biochar (500°C) was dominated by a peak at 128 ppm due to resonance of aromatic C structures,

while minor spectral peaks were recorded in the aliphatic-C (0-50 ppm), polysac — charide-C (60-110 ppm) and carboxylic-C (194 ppm) region. Integrating the area of the spectral region revealed that the cotton gin trash biochar contained 65% aromatic — C with only 12% occurring as polysaccharides. Most of the polysaccharide-like compounds in the cotton gin trash biochar were lost during pyrolysis at the higher temperature regime (500°C).

Biochar (Fig. 9, bottom), which has been produced from hardwoods using a fast pyrolysis system, had minor peaks at 56 and 75 ppm, respectively, which is indica­tive of methoxy and C-O groups in polysaccharides. Similar to gin trash biochar, the hardwood biochar was dominated by an aromatic-C peak (126 ppm) which accounted for 52% of the C distribution. A minor amount (20%) of the total C structures occurred in polysaccharide-like compounds.

Fourier transformed infrared spectroscopy can determine the presence of types of organic compounds in biochars [80, 84]. It is a robust system and uses the mid-infrared spectrum (4,000-500 cm-1) to examine for sorption peaks that are diagnostic of rotational and vibrational movements of molecular structures and bonds within those structures [101]. On the one hand, there are issues with FT-IR analyses includ­ing broad peaks due to sorbed moisture [101] and sorption overlap that complicates ascribing the organic compound responsible for the sorption peaks [78] . On the other hand, very little sample is needed (few mg), it is nondestructive, and the results are more rapidly obtained when compared to 13C NMR spectroscopy.

Fig. 10 FT-IR spectra of (bottom) hardwood biochar (fast pyrolysis) and (top) cotton gin trash biochar (500°C)

These properties make FT-IR an acceptable analytical tool for examination of biochar properties during manufacturer and for biochar mineralization studies. For example, FT-IR spectroscopy has been employed to determine structural and func­tional group changes during biochar mineralization in soils [23, 24, 75]. The FT-IR spectral analysis of biochar pyrolyzed from cotton gin trash (Fig. 10, top) and hard­wood biochar (bottom) show broad peaks between 3,500 and 2,000 cm-1, but also a few sharp peaks between 1,600 and 1,620, and 1,170-975 cm-1. Surface hydroxyls and or sorbed water and C-H stretching are responsible for the broad beak between

3,500 and 2,000 cm-1. Peaks at 1,620 and 1,600 cm-1 are ascribed to aromatic C=C and H-bonded C=O and peaks at 1,170, 1,070 and 975 cm-1 are indicative of C-O stretching of polysaccharides and OH deformation of COOH groups [ 101]. The aromatic peak in the FT-IR spectra of the hardwood biochar is more distinct than in the gin trash, which is consistent with 13C NMR results.

Lignin is the amorphous material that surrounds cellulose fibers and cements them together. It is a complex, heterogeneous polymer formed by the polymerization of three phenyl propane monomers, i. e., guaiacyl (4-hydroxy-3-methoxyphenyl), syringyl (3,5-dimethoxy-4-hydroxyphenyl), andp-hydroxyphenyl units. Lignin is the most complicated, least understood, and most thermally stable component of biomass.

Primary pyrolysis of lignin begins with thermal softening at around 200°C, while most lignin pyrolysis occurs at higher temperatures, higher than the fast decomposi­tion of cellulose. Fast pyrolysis of lignin will obtain higher char yield and lower liquid yield than holocellulose, and the liquid product can be classified into three groups, the large molecular oligomers (known as pyrolytic lignins), the monomeric phenolic compounds, as well as light compounds (such as methanol, HAA, acetic acid). The pyrolytic lignins are formed in much higher yield than the other two classes of products, usually account for 13.5-27.7 wt% of crude bio-oils on a water-free basis [37, 38, 65]. Several studies have been conducted to analyze them by various wet chemical and spectroscopic methods as well as pyrolysis-gas chromatography/mass spectrometry [12,13,38,83,84], and to find out that their average molecular weight is between 650 and 1,300 g/mol, and they are typically characterized by biphenyl, phenyl coumaran, diphenyl ethers, stilbene, and resinol structures.

Hydrothermal degradation of cellulose is a heterogeneous and pseudo-first-order reaction for which detailed chemistry and mechanism have been proposed earlier [9,80,81,107]. Cellulosereactioninhydrothermalandcatalystfreemediummainly proceeds via hydrolysis of glycosidic linkages. The long chain of cellulose starts breaking down in such condition to smaller molecular weight water soluble compounds (oligomers) and further to glucose (monomer). Glucose is water soluble and undergoes rapid degradation in hydrothermal medium at elevated temperature. The key products from the glucose decomposition are shown in Fig. 9. These prod­ucts are formed mainly via dehydration, isomerization, reverse aldol condensation, and fragmentation reactions.

Hydrolysis of cellulose in supercritical water is very sensitive to residence time. A high residence time enhances the formation of organic acids such as acetic acid, formic acid, and lactic acid. Formation of acids makes the reaction medium more acidic, which is conducive for further degradation of hydrolysis products. Indeed after hydrolysis of cellulose in water at 320°C and 25 MPa for 9.9 s, more than half of the cellulose was converted to organic acids [100] . The solid cellulose-like

residues have been inevitably observed because of the rapid change in the polarity of water in going from reaction condition to room temperature. These residues have been reported to have a lower viscosity-average degree of polymerization (DPv) with no significant change in crystallinity as compared to untreated cellulose [63].

The objective of this approach is to convert feedstock into FT liquid as much as possible. The heat and particularly electricity generations are of secondary impor­tance. This approach requires a large production capacity. Since FT process has a high fixed cost, the economy scale is important to produce FT liquid competitively with other biofuels such as bioethanol, biodiesel, as well as FT liquids derived from fossil fuels [46]. This means gasifiers of the size at least greater than 1,000 MWth are required. Fundamental assumptions and other elements of this approach are:

1. FT off-gas is recycled to the biomass gasifier to achieve maximum syngas con­version. Electricity is produced as a “by-product” from the relatively small recy­cle bleed stream.

2. The yield of syngas H2 + CO in the FT feed gas must be as high as possible.

3. Since significant energy remains in CH4, C2H4, BTX, and tar, the process requires tar cracker and shift reactor or a reformer to convert all hydrocarbons into CO and H2. The increase in feed H2 + CO and increase in their yields result in much higher production of FT liquids and wax and higher overall efficiency of 63.5%.

4. Both gasifier and tar cracker must be oxygen blown to reduce dilution of recycle stream by N2 and other inert materials.

5. Unlike in the front-end approach, the part of gas conditioning system, CO2 removal step is essential.

The performances of Zn, K, and Cu-promoted Fe catalysts were studied under CO2-free and CO2-containing syngas, respectively. The CO2 selectivity based on converted CO is given in Fig. 7. The catalysts can be divided into two categories.

For catalysts Z2K2C2/Fe and Z4K2C4/Fe, the CO2 added into reactants does not influence the CO2 selectivity, i. e., CO2 is inert to react on these two catalysts. But the CO2 selectivity decreases with the addition of CO2 to reactants on catalysts Z2K1C2/Fe, Z4K1C4/Fe, and Z6K2C6/Fe (Not shown) which have lower K content or higher Zn/K ratio in them. The characteristic of CO2 selectivity among these cata­lysts has a corresponding reflection in their CO2-TPD. The catalyst showing CO des­orption peak around 930 K (Fig. 6) is sensitive to the composition of reactants and added CO2 brings forth decreased CO2 selectivity during FT synthesis reaction.

It has been found that CO2 is inert at low temperature [34, 37]. The result of Z2K2C2/Fe and Z4K2C4/Fe is consistent with this conclusion, but the result of Z6K2C6/Fe, Z4K1C4/Fe, and Z2K1C2/Fe suggests that higher Zn/K ratio or lower K content leads to the decrease of CO2 selectivity. There are two possibilities for this phenomenon. The added CO2 may inhibit CO2 formation from WGS reaction or be converted into hydrocarbons itself. Both of the routes can result in decreased apparent CO2 selectivity based on converted CO. Correspondingly, more CO is converted into hydrocarbons and the hydrocarbon selectivity can be increased. Although it cannot be discerned which route is in effect here, the results from Z6K2C6/Fe, Z4K1C4/Fe, and Z2K1C2/Fe show the possibility to use CO2 contained in syngas for hydrocarbon synthesis at low temperature.

Production of butanol in a fed-batch reactor is another system where hydrolysis, fermentation, and recovery can be combined (SSFR). Using such a system, butanol was produced from wheat straw [58]. The reactor was loaded with 86 g/L pretreated wheat straw, enzymes, and the butanol producing culture. The system was operated for 533 h at a pH 5.0 and temperature 35°C. During the initial period of 120 h, wheat straw was hydrolyzed completely by the added hydrolytic enzymes. In order to ascertain that the culture was not deficient in sugar, a mixture containing glucose, xylose, arabinose, galactose, and mannose was fed to the reactor to mimic their proportion in wheat straw. In a 1 L reaction mixture a total (including sugar present in WS) of 430 g sugar was used thus producing 192 g total ABE with a yield of 0.44 (Table 4). In this reactor, a productivity of 0.36 g/L h was achieved which is 20% higher than achieved (0.30 g/L h) in a control reactor. One of the major problems associated with this fermentation was that the culture had difficulty utilizing xylose in the later part of fermentation. In this system, gas stripping was used to agitate the treated biomass and recover ABE.

Increased cellulose accessibility and hydrolysis has been achieved by chemical removal of hemicelluloses [135]. Therefore, hemicelluloses can be viewed as con­tributors to the recalcitrance of lignocellulosic material to degradative processes. Hemicelluloses are crosslinked to lignin by three types of covalent linkages. The first type involves p-coumaric or ferulic acid, ether linked to lignin, and ester linked to hemicellulose sugars [43]. The second type relies on spontaneous ether linkage formation between OH-groups of saccharides and lignin alcohols [129]. The third type involves ester links between uronic acid residues of glucuronoxylans and hydroxyl groups of lignin alcohols [130]. Such crosslinks hinder cell wall degrada­tion by solvents, enzymes and microbes and reduce plant digestibility. Hence, improved cell wall solubility and degradation can theoretically develop through cleavage of the links between lignin and hemicelluloses.

The first type of crosslinking is abundant among the hemicelluloses of grasses and related monocot cell walls and form lignin-carbohydrate complexes. The matrix properties of these complexes are further influenced by the character and number of hemicellulose side chains and by the nature of the crosslinking agents, such as arabi — nose or ferulic acid [126]. To date, only ferulic acid esterase (FAE) has been geneti­cally introduced into plants cell walls in an attempt to enhance the accessibility of hydrolyzing enzymes to hemicellulose fibers by removing ferulic acid side chains and crosslinking bonds [12—14,133]. Expression of a fungal FAE gene in transgenic ryegrass and tall fescue rendered the cell walls more accessible to endoxylanases with higher levels of fermentable sugars released upon cell wall hydrolysis [12-14].

Glucuronoxylans are the primary hemicellulosic component of hardwoods and account for ~20% of the woody cell wall [4]. Approximately 60-70% of the xylose residues in hardwood xylan are acetylated at the C2 and/or C3 positions I 73].

O-acetylation of xyloglucan has often been considered an additional barrier to enzy­matic hydrolysis of polysaccharides and decreased polysaccharide solubility [91]. Thus, the reduction in the degree of acetylation of cell wall polysaccharides may allow for enhanced rates of saccharification. The deacetylation of GX in vitro increases its water solubility [51]. In addition, acetylxylan esterases (AXEs) liber­ates acetic acid from partially acetylated 4-O-methyl glucuronoxylan [79] and may increase hemicellulose solubility and increase xylan degradation rates.