Isosynthesis (for the production of branched hydrocarbons) is interesting because (a) under proper conditions it gives saturated branched-chain aliphatic hydrocarbons containing 4-8 carbon atoms, (b) compared to other types of synthesis, it is the only one that uses difficult reducible oxides as catalysts [24-27], and (c) it produces isob­utane for high octane gasoline. The oxide catalysts that are used for this synthesis are thorium oxide, aluminum oxide, tungstic oxide, uranium oxide, and zinc oxide. The thoria and alumina catalysts can be prepared by precipitation from nitrate solutions by adding a hot sodium carbonate solution. The other oxide catalysts can be pre­pared by addition of alkali to their respective nitrate solutions except in the case of tungsten which can be precipitated from Na2WO4 solution by nitric acid addition.

The most effective catalysts for isosynthesis are the tetravalent oxides, thoria, ceria, and zirconia. Alumina and the other oxides are not as effective in producing

isobutane. Other oxides such as chromia, lanthana, praseodymium oxide, magnesia, manganese oxide, titania, and berylia have been tested but they had lower activity. Of all the catalysts tested so far, thoria or promoted thoria has been the most promis­ing catalysts. Unlike catalysts of iron group, thoria is not poisoned by sulfur com­pounds. In addition, an activity decline due to carbon deposition can be easily overcome by simply passing air over the catalyst to generate CO2 at the synthesis temperature. Some typical results for one-component catalysts at various pressures are given in Table 5.

Thoria catalyzes the production of hydrocarbons with relatively high carbon number from synthesis gas. Furthermore, the so-called gasol hydrocarbons consisting of mainly C3-C4 also contain large amounts of isobutane. In contrast, the trivalent oxide alumina produced mainly methane and carbon with small amount of “gasol” and traces of isobutane. The most active divalent oxide, zinc oxide, produced no liquid hydrocarbons but mainly methane and alcohols. For thoria catalyst, the best temperature has been found to be between 375 and 475°C. Alcohol production dominated below 375°C and methane and LPG were the main products between pressure of 300 and 600 atm. Below 300 atm, gas conversion was small; above 600 atm, methane and dimethyl ether were obtained.

The effects of additions of various promoters on the performance of thoria cata­lysts have also been examined. Three promoters have been studied: (1) alkali to increase molecular weight of the product, (2) phosphoric acid to convert unsaturated hydrocarbons such as propane to я-butene to higher branched hydrocarbons particularly to dimers, and (3) zinc oxide and alumina to enhance the formation of alcohols and subsequently the dehydration of alcohols for the formation of branched hydrocarbons.

The alkali did not work well and the activity of thoria catalyst decreased with an increase in alkali content. The most effective promoter was alumina. For thoria — alumina catalyst, the literature data show [29] that the amount of isobutene increased with an addition of alumina; best results were obtained at 20% alumina on thorium oxide. Catalysts prepared by separate precipitation and mixing of the wet precipi­tates produced the highest i-C4 (isobutane) yields. While “gasol” increased with an increase in temperature, C5+ and alcohols (not shown in the table) decreased with an increase in temperature. i-C4 increased with an increase in temperature. In this study, thoria was precipitated from a nitrate solution with sodium carbonate while aluminum oxide was precipitated from a sodium aluminate solution using sulfuric acid. Finally, the best thoria-alumina-zinc oxide catalysts gave lower yields of isobutane and somewhat higher yield of C5+ hydrocarbons than thoria-alumina catalysts. Once again, just like for FT synthesis, most recent efforts on the catalyst development for isosynthesis are focused on improving product selectivity.

In Part 1.2, it is suggested that to use CO. — containing syngas directly for FT synthesis can solve the shortcoming of current FT technology. The studies of CO2 hydrogenation and CO. + CO hydrogenation on Fe catalysts are reviewed in the following.

In view of engineering, to use CO2-containing syngas is easy to control temperature of synthesis reactor because of the lower exothermicity of the overall reaction of CO2 as compared to CO [ 16] . Furthermore, a perceived disadvantage of using Fe-based catalysts for FT is that a large proportion of the CO in the syngas is con­verted to CO. rather than the desired hydrocarbons [21]. The addition of CO2 to syngas can inhibit CO2 formation from CO and may increase the ratio of CO to hydrocarbons rather than CO2.

In an interesting investigation, it was observed that some of the cellulosic hydrolyzate chemicals that are generated during the pretreatment or the neutralization process stimulate both cell growth and ABE production. These chemicals include sodium acetate, furfural, and HMF. In the presence of 8.9 g/L sodium acetate cell growth was slightly improved. In the control experiment 17.8 g/L ABE was produced while in the presence of 8.9 g/L acetate 20.3 g/L ABE was produced, showing an increase of 14%. An improvement in fermentation performance on supplementation of acetate has previously been documented [11,24]. Inclusion of furfural and HMF (0.3-2.0 g/L) in the fermentation medium improved both cell concentration and ABE production [18]. Itis suggested that acetate, furfural, and HMF are beneficial to this fermentation within a certain concentration range.

Meeting the growing demands for energy supply while simultaneously promoting sustainable development is one of the greatest challenges facing humanity today. Many strategies and technologies have been proposed to offer alternative energy sources, but no ultimate solution has been developed thus far.

Paradoxically, prior to the development of the almost total dependency on fossil fuels, conversion of biomass to energy provided a means of heating, illumination, and cooking for centuries. The plant is a cheap, highly efficient energy generator as it converts light energy to simple sugars through photosynthesis and CO2 fixation. Combustible and highly energetic polymers are then formed from these sugars, and generate a composite lignocellulosic secondary cell wall constituted of cellulose, hemicellulose, and lignin [102].

To date, most ethanol fuels have been generated from corn grain or sugarcane, often referred to as “first generation” biofuels. However, bioconversion of such feedstocks to biofuels exploits land sources necessary for food production, diverts essential crops from the human food chain and increases the cost of human and animal feed [36] . “Second generation” or biomass-derived biofuels are extracted from inedible plant tissues or from inedible plants and support sustainable develop­ment by processing the residual structural biomass after the food portions have been removed. Cellulosic ethanol can thus be formed from a variety of sources, including woody materials, agricultural residue and specially cultivated grasses. Ultimately, lignocellulosic-derived ethanol has the potential to provide for the vast majority of

global transportation fuel needs, while only slightly impacting food supplies, in a more cost-effective manner and with less net carbon dioxide emission than that from fossil fuel combustion [36, 108].

The plant cell wall is a composite material that provides structural integrity to plants as well as protection from pathogen invasion. The polyphenolic lignin portion, whose proportion and character can vary significantly among feedstock sources [30], sterically hinders access of enzymes and chemicals that degrade hemi — cellulose and cellulose. In addition to the degree of lignification, the cellulose crys­tallinity and its degree of polymerization [ 78] contribute to the recalcitrance of lignocellulosic feedstock to saccharifying chemicals or enzymes. These composite factors contribute to the result that decomposition of plant cell wall polysaccharides into fermentable sugars is more complicated than breaking down purified starch or sucrose to fermentable sugars. Conversion of lignocellulosic biomass to bioethanol requires cell wall breakdown in a pretreatment stage to degrade lignin and release polysaccharides before the conversion of such lignocellulosic biomass to bioetha­nol. The pretreatment requires toxic solvents and high energy inputs (e. g. high tem­peratures and pressure). These steps are then followed by saccharification of cellulose and hemicellulose to simple sugars via hydrolysis, and the free sugars are then fermented to ethanol. Thus, the leading technological barrier to effective employment of lignocellulosic biomass as a source of fermentable sugars lies in the economic breakdown of the polysaccharide wall [37].

Commercial realization of biofuel production requires an integrated solution combining appropriate biofuel crops, biomass modification techniques, and sophis­ticated process engineering. The application of recent advances in plant genetic engineering, carbohydrate chemistry, and increased knowledge of plant cell wall ultrastructure will together allow reengineering crop cell walls, which in turn facili­tates lowering of biomass saccharification and conversion costs, while significantly increasing biofuel production yields. This review discusses the most recent plant cell wall modification and plant engineering technologies designed to improve sugar yields upon enzymatic hydrolysis and to reduce energy demands and cost of the bioethanol production.

Over the years, a large number of assays have been developed that report on cellu — lase activity. Most assays are quantitative, although many of these do not evaluate activity in real-time, but require postprocessing of digests at fixed time points, limit­ing the extent to which these assays allow kinetic data to be interpreted using mech­anistic models (for example, models including Michaelis-Menten terms, processivity, prebinding, and diffusion). This type of treatment is further limited by the heteroge­neity of various cellulosic substrates, and to time-dependent enzyme inactivation that is sometimes observed (especially with insoluble substrates). Other qualitative assays have been developed to screen for activity in live cells. Such screens have the potential to allow large numbers of engineered cellulases to be assayed quickly, without having to purify each protein. In this section, we briefly touch on different assays for activity, highlighting strengths, weaknesses, and when relevant, relative sensitivities to exocellulolytic vs. endocellulolytic activity. For a comprehensive description of different cellulase assays, see a review by Zhang, Himmel, and Mielenz [81] .

2.3.1 An NADPH/NADH Conversion Mechanism

According to the photosynthetic butanol production pathway(s), to produce one molecule of butanol from 4CO2 and 5H2O is likely to require 14 ATP and 12 NADPH, both of which are generated by photosynthetic water splitting and photo­phosphorylation across the thylakoid membrane. In order for the 3-phosphoglycer — ate-branched butanol-production pathway (03-12 in Fig. 1) to operate, it is a preferred practice to use a butanol-production-pathway enzyme(s) that can use NADPH that is generated by the photo-driven electron transport process. Clostridium saccharoperbutylacetonicum butanol dehydrogenase (GenBank accession number: AB257439) and butyraldehyde dehydrogenase (GenBank: AY251646) are exam­ples of a butanol-production-pathway enzyme that is capable of accepting either NADP(H) or NAD(H). Such a butanol-production-pathway enzyme that can use both NADPH and NADH (i. e., NAD(P)H) can also be selected for use in this 3-phosphogly cerate branched and any of the other designer butanol-production pathway(s) (Fig. 1) as well. C. beijerinckii Butyryl-CoA dehydrogenase (GenBank: AF494018) and 3-hydroxybutyryl-CoA dehydrogenase (GenBank: AF494018) are examples of a butanol-production-pathway enzyme that can accept only NAD(H). When a butanol-production-pathway enzyme that can only use NADH is employed, it may require an NADPH/NADH conversion mechanism in order for this 3-phos- phoglycerate-branched butanol-production pathway to operate well. However, depending on the genetic backgrounds of a host organism, a conversion mechanism between NADPH and NADH may exist in the host so that NADPH and NADH may be interchangeably used in the organism. In addition, it is known that NADPH could be converted into NADH by a NADPH-phosphatase activity [21] and that NAD can be converted to NADP by a NAD kinase activity [22, 23]. Therefore, when enhanced NADPH/NADH conversion is desirable, the host may be genetically modified to enhance the NADPH phosphatase and NAD kinase activities. Thus, in one of the various embodiments, the photosynthetic butanol-producing designer plant, designer alga, or plant cell further contains additional designer transgenes (Fig. 2b) to induc- ibly express one or more enzymes to facilitate the NADPH/NADH interconversion, such as the NADPH phosphatase and NAD kinase (GenBank: XM_001609395, XM_001324239), in the stroma region of the algal chloroplast.

Another embodiment that can provide an NADPH/NADH conversion mecha­nism is by properly selecting an appropriate branching point at the Calvin cycle for a designer butanol-production pathway to branch from. To confer this NADPH/ NADH conversion mechanism by pathway design according to this embodiment, it is a preferred practice to branch a designer butanol-production pathway at or after the point of glyceraldehyde-3-phosphate of the Calvin cycle as shown in Fig. 1. In these pathway designs, the NADPH/NADH conversion is achieved essentially by a two-step mechanism: (1) use of the step with the Calvin cycle’s glyceraldehyde-3- phosphate dehydrogenase, which uses NADPH in reducing 1,3-diphosphoglycerate to glyceraldehyde-3-phosphate; and (2) use of the step with the designer pathway’s NAD+-dependent glyceraldehyde-3-phosphate dehydrogenase 01 , which produces NADH in oxidizing glyceraldehyde-3-phosphate to 1,3-diphosphoglycerate. The net result of the two steps described earlier is the conversion of NADPH to NADH, which can supply the needed reducing power in the form of NADH for the designer butanol-production pathway(s). For step (1), use of the Calvin cycle’s NADPH — dependent glyceraldehyde-3-phosphate dehydrogenase naturally in the host organ­ism is usually sufficient. Consequently, introduction of a designer NAD+-dependent glyceraldehyde-3-phosphate dehydrogenase 01 to work with the Calvin cycle’s NADPH-dependent glyceraldehyde-3-phosphate dehydrogenase may confer the function of an NADPH/NADH conversion mechanism, which is needed for the 3-phosphoglycerate-branched butanol-production pathway (03-12 in Fig. 1) to oper­ate well. For this reason, the designer NAD+ — dependent glyceraldehyde-3-phos — phate-dehydrogenase DNA construct (example 12, SEQ ID NO:12) is used also as an NADPH/NADH-conversion designer gene (Fig. 2b) to support the 3-phospho — glycerate-branched butanol-production pathway (03-12 in Fig. 1) in one of the vari­ous embodiments. This also explains why it is important to use a NAD+-dependent glyceraldehyde-3-phosphate dehydrogenase 01 to confer this two-step NADPH/ NADH conversion mechanism for the designer butanol-production path way (s). Therefore, in one of the various embodiments, it is also a preferred practice to use a NAD+-dependent glyceraldehyde-3-phosphate dehydrogenase, its isozymes, func­tional derivatives, analogs, designer modified enzymes, and/or combinations thereof in the designer butanol-production pathway(s) as illustrated in Fig. 1.

An environmentally kind process was developed for the production of biodiesel from Jatropha oil using a heterogeneous solid super base catalyst and calcium oxide. The results revealed that under the optimum conditions of catalyst calcina­tions, temperature of 900°C, reaction temperature of 70°C, reaction time of 2.5 h, catalyst dosage of 1.5%, and methanol/oil molar ratio of 9:1, the oil conversion was 93% [34]. Nazir [58] found that the yield of JCO FAME could reach up to 94.35% using the following reaction conditions: 79.33 min reaction time, 10.41:1 methanol:oil molar ratio, and 0.99% of CaO catalyst at reaction temperature 65°C.

Figure 7 summarises the different stages of production and downstream processing of biofuel-directed microalgae. Each stage itemises the applicable techniques based on current knowledge.

1.3 Harvesting Techniques

The harvesting of microalgae biomass is a critical step in the production of biofuels from microalgae, because it determines the final quality of extracts, and energy inputs, and therefore overall sustainability, and it also prevents fouling of the production process [106,174]. Harvesting probably accounts for the highest compo­nent of the total cost of microalgae biomass production [59, 173]. This high cost is due to the low cell densities (typically in the 0.3-0.5 g L-1 range) and the small size of the microalgal cells (typically <40 pm) [118], which results in low separation or dewatering efficiency and poor product quality under suboptimal growing conditions [49]. The selection of appropriate harvesting techniques is therefore critical

Fig. 7 Harvesting and processing pathways for microalgal-directed biofuels (Adapted from Schmid-Staiger [186])
to economic and environmentally sustainable harvesting [185]. For biofuel-directed production, harvesting process should not incur negative energy balance [187].

There is no universal harvesting method for microalgae, but typical processes used singly or in combinations include flocculation, flotation, filtration, sedimenta­tion, filtration and cell immobilisation (Fig. 7). Factors such as strain, cell density, culture condition, growth media and value of target products determine both the cost of and ease of harvesting [106]. For example, the cyanobacterium Spirulina’s long spiral shape (20-100 pm) naturally lends itself to cheaper and energy-efficient micro-screen harvesting method [14]. Overall, appropriate harvesting unit process should be able to: maximise the recovered biomass dry weight (varies with microal­gae species, biomass concentration and cell sizes), minimise the cost of operation and maintenance (including energy consumption), and yield quality extracts [49].

Flocculation: Flocculation process aggregates the microalgal cells to increase the overall cluster sizes [128], and is generally used as a preparatory step for other har­vesting methods [141]. It involves the addition of multivalent cations and cationic polymers to the culture medium to neutralise the negative charge that is carried on the surface of many microalgal cells [158] . Flocculation may also physically link one or more aggregates in a process referred to as bridging [141]. Since chemical flocculants are used, this process is unsuitable for food-grade products.

Flotation: Dissolved air flotation (DAF) process injects micro-air bubbles into the culture column, and as they rise to the surface, they trap microalgae cells [158,218]. Some microalgae production units such as the tubular PBRs promote flotation through high overflow rates which causes cells to move upwards [64,102]. Flotation has advantages in its moderate cost, low space requirement and rapid operation compared to other processes, but its technical viability has only been demonstrated in a few studies [33, 120] . The use of ozone produces more efficient solid-liquid separation compared to DAF [33]

Sedimentation: Gravity sedimentation is one of the most commonly used harvesting processes because of its capability to handle large volumes of culture, and suitabil­ity for low-value biomass [153] . However, it can only be applied to species with large cell size (>70 pm) like Spirulina [149]. Ultrasonic aggregation may be used as precursor to enhance sedimentation. The main advantages are that it is a non-fouling technique, therefore can be used for food-grade products, and it can also be operated continuously without inducing shear stress on the microalgal cells which causes cell destruction [21]. Centrifugal recovery (CR) sedimentation is only feasible for har­vesting of high-value metabolites and extended shelf-life concentrates for aquacul­ture [86]. The process is rapid but energy-intensive, which makes it unsuited to harvesting biofuel-directed algae biomass, but it may be more economic in large — scale production systems [24] .

Filtration: Conventional filtration processes with micro-strainers or rotating screen filters with backwash may be the most appropriate for harvesting of large cell algae (>70 pm) such as Spirulina [128], but they cannot be used for species ofbacterial dimensions like Scenedesmus, Nannochloropsis and Chlorella [140]. Conventional filtration is popular because of its mechanical simplicity, availability in large unit sizes, and filtration aids such as diatomaceous earth and cellulose can be used to improve efficiency [141]. Membrane filtration and ultra-filtration are possible alter­natives to conventional filtration for the recovery of smaller microalgae cells (<30 pm) [161], albeit a more expensive process because of the need for membrane replacement and pumping [128] . It is also a more suitable process for fragile cells like Dunaliella, because it pre-disposes low trans-membrane pressure and low cross­flow velocity conditions [19]. It can also be used to remove protozoans and viruses from used algal culture medium, while retaining nutrients for recirculation [234].

Cell immobilisation: This process prevents cells from moving independently of its neighbours to all parts of the aqueous phase of the system [203]. It has been presented as an option for harvesting microalgae biomass in wastewater treatment systems, but more research is needed on the effects of immobilisation on algal cell physiology and biochemistry [125]. Assessment for large-scale applications is still limited [106].

Thermal drying: Drying is sometimes required after the harvesting steps to prevent fouling of the final biomass product. The most common drying methods include: sun-drying, and low-pressure shelf-drying [163], spray-drying [58], drum-drying [163], fluidised bed-drying [111] and freeze-drying [143]. Sun-drying is the most popular method due to the low cost, but disadvantages include long drying times, the requirement for large surface areas, and high material loss [163]. Spray-drying and freeze-drying are expensive processes commonly used for the high-value products and oils [58, 143].

1.2 Biochar Production

Biochar is a byproduct of the biofuel industry [5, 58, 62] , It is produced by the pyrolysis of organic feedstocks at temperatures between 300 and 700°C in an oxygen — free or low oxygen noncombustible atmosphere. Different feedstocks are used to make biochars, including biomass energy crops, bioenergy residues, crop residues, manures, and kitchen wastes. During the pyrolytic process, these organic feedstocks thermally decompose, releasing volatile compounds, syngas and biochar. The vola­tile compounds can be recondensed and refined as bio-oil [11]. The biochar residual product has chemical and physical properties that depend on complex reactions during the pyrolysis process and are reported to vary with feedstock selection and pyrolysis conditions [38, 74] .

Biochars can be made using various thermochemical processes systems such as slow/fast pyrolysis, flash pyrolysis, and gasification [58, 94]. Slow and fast pyrolysis

technologies are featured in this review. A more detailed explanation of gasification technologies for syngas production is available [58].

In the slow/fast pyrolysis systems, the feedstock (depending on the delivery feed scheme) can remain in the pyrolysis reactor anywhere from a few seconds to 24 h [94]. Pyrolysis reaction times vary among manufacturers because of differences in reactor temperature ramp settings, choices of dehydration (100-150°C) and carbon­ization temperatures (300-700°C), and cooling time. Under these conditions, biochar yields can range from 51 to 72% on an oven-dry C basis and between 29 and 57% on an air-dry mass basis [74]. More biochar is recovered at lower pyrolysis temperatures (around 350°C) because less volatile material is driven off as bio-oil. If maximizing bio-oil production is the goal, the manufacturer can adjust the slow pyrolysis process to operate at a higher temperature range (500-700°C). While more bio-oil is recovered, biochar mass yields will decline because of dehydration of hydroxyl groups and thermal degradation of ligno-cellulose structures [4, 5] .

Biochars pyrolyzed at higher temperatures (500-700°C) tend to have greater ash contents, and hence, more alkaline pH values (Table 1). High temperature pyrolysis will concentrate the salts because of the loss of C-, O-, and H-containing com­pounds removed as volatiles [20, 43, 74]. Ash contents for several biochars pyro­lyzed at the higher (400-700°C) temperatures regime ranged from 5.62 to 52.9% while at the lower temperature (<350°C) biochar ash contents ranged from 2.4 to 35.9% (Table 1). Biochar pyrolyzed from poultry litter had the highest ash content because of excretion of unassimilated nutrients [92] and from chemical additives to the litter to reduce N volatilization [74]. The high ash content also contributed to the poultry litter biochar having a calcareous pH (Table 1).

The elemental composition in several biochars is heterogeneous (Tables 1 and 2) because of differences in nutrient uptake by the raw feedstock [21] and by chemicals added to manure feedstocks prior to pyrolysis [74]. If the ash contains elements like

N, P, and K, then it could serve as a low grade fertilizer with a corresponding low N-P-K ratio (Table 1). These ratios were calculated based on the total contents of elements in the biochar, and does not necessarily reflect their plant availability status. Poultry litter and peanut hulls have a modest N-P-K fertilizer ratio while biochar made from pine chip and pecan shells had the lowest ratio (Table 1). Results from Table 2 show that the biochars pyrolyzed from different feedstocks can contain size­able quantities of base cations such as Ca and Mg. While also being essential plant nutrients, the presence of Ca and Mg causes the biochar to act like a liming agent. As Novak et al. [73] reported pecan shell biochar had liming properties since it had an alkaline pH, and contained 3.6 and 0.7 g kg-1, respectively, of Ca and Mg. Another important property of these four plant-based biochars (Table 2) is the low concentra­tions of heavy metals (i. e., Cd, Cr, Ni, Pb, and V). If these biochars are used as a soil amendment, low metal concentrations should ease environmental concerns.

In fast pyrolysis, the feedstock is placed in a retort and subjected to a very short burst (1-2 s in duration) of heat (400-600°C) usually under pressure [94]. These conditions also maximize bio-oil production (75%); however, lower biochar mass yields are recovered (»12%, [94]). For comparative purposes, one biochar made from hardwood using the fast pyrolysis system was included in this review. Its ash content, pH value, and fertilizer ratio were fairly similar to characteristics of the low temperature (350°C) pecan shell biochar (Table 1).

There are considerable time advantages when using fast pyrolysis, including shorter residence, carbonization, and temperature squelching times. The choice of pyrolysis system (slow vs. fast) for biochar manufacturer will ultimately be decided by a balance between biochar, bio-oil, and syngas recovery [94] .

Atomic O/C

Fig. 7 Atomic ratio distribution shown in the Van Krevelen diagram for raw feedstocks and biochars pyrolyzed using two temperature ranges

Hemicelluloses are closely associated with cellulose in the cell wall as well as to lignin in the middle lamella. They are amorphous polysaccharides with building units belong either to hexoses (mainly D-glucose, D-mannose, and D-galactose) or to pentoses (mainly D-xylose and L-arabinose). The primary hemicellulose components are galactoglucomannans (glucomannans) and arabinoglucuronoxylan (xylan). Compared with cellulose, hemicelluloses have received less attention in their pyrolytic mechanism study.

Hemicelluloses are less thermally stable than cellulose, presumably due to the lack of crystallinity, and their pyrolysis is generally thought to be analogous to cel­lulose in the reaction mechanisms. Fast pyrolysis of glucomannans generates similar pyrolytic products as the cellulose, while the pyrolytic products from xylan differ more than those from cellulose [6]. Generally, xylan fast pyrolysis will obtain higher char yield than that from cellulose, and will not form a typical depolymerization product, like the LG from cellulose. A probable cause of this difference was explained by Ponder and Nichards [74]. During cellulose pyrolysis, the glucosyl cation from the scission of the common glucans can readily form a stable 1,6-anhydride with the free primary hydroxyl group at C-6, and finally yields the volatile LG. However, there is no feasible mechanism for intramolecular “stabilization” for the xylosyl cation

2CO + (Acetone)

Fig. 3 The major pyrolytic pathways during xylan fast pyrolysis (proposed by Shen et al. [88]). (a) The main chain of O-acetyl-4-methylglucurono-xylan, (b) O-acetylxylan and 4-O-methyl — glucuronic acid unit via anhydride formation, and hence, the xylosyl cation is more likely to enter the nonspecific dehydration pathways, resulting in the char formation rather volatiles.

In a recent study, Shen et al. [88] also proposed the detailed possible routes for the pyrolytic formation of several major products from main xylan chain, as well as the O-acetylxylan and 4-O-methylglucuronic acid unit, as shown in Fig. 3.