Cell Wall Engineering by Heterologous Expression of CellWall-Degrading Enzymes for Better Conversionof Lignocellulosic Biomass into Biofuels

Halbay Turumtay1,2

# Springer Science+Business Media New York 2015

Abstract Huge energy demand with increasing population isaddressing renewable and sustainable energy sources. A solu-tion to energy demand problem is to replace our current fossilfuel-based economy with alternative strategies that do notemit carbon dioxide. Plant biomass is one of the best candi-dates for this issue. Plants use solar power to convert carbondioxide and water into sugars, which can be used in fermen-tation reactions to produce both energy and materials.However, the desired sugars are trapped in the highly recalci-trant cell wall as building blocks of cellulose chains.Moreover, the complexity of the plant cell wall structure hin-ders the hydrolysis of cellulose into fermentable sugar mono-mers. Although pretreatments are used to change the physicaland chemical properties of the lignocellulosic biomass andimprove hydrolysis rates, these pretreatments often use harshand polluting chemicals and severely increase the cost of bio-fuel production. The goal of the review is to summarize recentresearches, which describe generating plants with a modifiedcell wall and improve hydrolysis of cellulose without applyingany or less pretreatment methods. Since pretreatment of lig-nocellulosic biomass is the most cost effective step in biofuelproduction, generating autodigestible plants could reduce theproduction cost of biofuels and bio-based biomaterials. One ofthe strategies to improve biomass conversion efficiency is themodification of the cell wall by heterologous expression ofcell wall-modifying proteins. These cell wall-modifying

proteins could alter the cell wall structure and reduce cell wallrecalcitrance. The use of such transgenic technologies wouldconsume less energy and chemicals when cellulose is moreaccessible for enzymatic hydrolysis.

Keywords Lignocellulosic biomass . Cell wall engineering .

Fermentable sugars . Biorefinery

Introduction

The availability of energy has been key in the development ofour modern way of life, and the importance of energy willonly increase in the years to come. World energy consump-tion is expected to grow by 56 % between 2010 and 2040from 524 quadrillion British thermal units (Btu) to 820 qua-drillion Btu. This huge increase is a direct consequence of theexponential growth of new industrialized countries such asChina, Malaysia, and India [1]. Most of our current energy isderived from fossil sources that are limited and nonrenew-able. These limited resources are also needed for chemicals/material production such as polymers and pharmaceuticals.Each year, 330 million tons petrochemical-based products areintroduced in our daily life [2]. In addition, fossil fuel com-bustion produces greenhouses gases such as CO2 and meth-ane, which are released into the atmosphere, causing globalwarming and climate changes.

This, together with the unequal geographical distribution offossil resources and the political instability in regions withimportant fossil energy reserves, has prompted the focus onalternative renewable and sustainable resources of energy(e.g., wave and wind power, solar, geothermal, and hydroelec-tric energy). Whereas these new technologies can supply en-ergy, they are inadequate in the replacement of fossil resourcesfor products and materials.

* Halbay Turumtayhalbay.turumtay@gmail.com

1 Department of Molecular Biotechnology, Ghent University, Coupurelinks 653, 9000 Ghent, Belgium

2 Present address: Department of Energy System Engineering,Karadeniz Technical University, 61830 Trabzon, Turkey

Bioenerg. Res.DOI 10.1007/s12155-015-9624-z

Plant biomass has the tremendous advantage that it canserve as a source for both products and energy. Production ismeasured in biomass per hectare (dmt/ha) and combined withthe high heating value (maximum amount of energy potential-ly derived from a given biomass source) of the biomass; theenergy yield of the cultivated crop can be calculated. Table 1shows the range of energy yields for a number of types ofbiomass [3].

Plants use photosynthesis to capture CO2 from the atmo-sphere and store the carbon into their biomass. These sugarscan subsequently serve as a source to feed fermentation reac-tions to produce a plethora of products and materials amongwhich is bioethanol. Because the combustion of biofuelsreturns as much CO2 into the atmosphere as was taken upby the plant to produce the biomass, the concept is consideredto be carbon neutral [4]. Biodiesel from grapeseed oil or palmoil and bioethanol produced from sucrose (sugar beet, sugar-cane, etc.) or starch (potato, corn, cereal, etc.) are called first-generation biofuels. Selection of the most suitable bioenergycrops largely depends on the geographical location of thefield, and each area has its own specific feedstock that couldbe used for biofuel production. For instance, the production ofbioethanol is derived from sugarcane in Brazil whereas in theUnited States (US), corn is being used [1]. Although first-generation biofuel sources have the advantage to be easilyconverted toward ethanol and biodiesel, they are in directcompetition with food and feed production [5]. In addition,sugar and starch crops require the use of fertilizers and pesti-cides, and therefore, the production of bioethanol from cornprovides only a limited reduction in green house gas (GHG)emission [6, 7]. Furthermore, first-generation biofuels are ac-celerating deforestation and compete for scarce water re-sources in some regions. These concerns make the use offirst-generation biofuels insufficient to address the world’shuge demand for sustainable energy.

In contrast to first-generation biofuels, second-generationbiofuels are produced from cheap and abundant nonfood plantmaterials [8]. This largely refers to the lignocellulosic plantcell wall containing different types of polysaccharides such ascellulose, hemicellulose, and pectins. Preferentially, thesesecond-generation bioenergy crops should produce substantialamounts of biomass over a short period of time, with no or lessrequirements for fertilizers or pesticides. Hence, their

production is considered cheap. Moreover, as second-generation bioenergy crops can be grown on marginal lands,their growth is not in direct competition with food production.Finally, the intelligent use of second-generation biofuels canbe truly carbon neutral or even carbon negative as it should bepossible to capture more carbon dioxide from the atmospherethan is released upon combustion and sequester the residualcarbon of the biomass in the soil [9]. One of the best crops forproduction of second-generation bioenergy in Europe and inthe USA is poplar (Populus spp.). Poplar properties which arementioned above as an ideal bioenergy crop plus does notrequire annual planting, and new branches will easily sproutfrom the main stem after harvesting make them an excellentcandidate in the bioenergy field [10].

In addition, poplar is more desirable for biofuels thanmanyother woody crops because of its fast growth and the produc-tion of a significant amount of biomass with high fermentablesugar content in a short period of time [11]. However, the useof lignocellulose biomass is not cost-effective at the moment,and this is mainly due to the complexity and recalcitrance ofthe plant cell wall, which limits the release of glucose fromcellulose fibers [12]. To overcome this limitation, the plantcell wall is being studied extensively, and the ultimate goalis to modify the plant cell wall to reduce the recalcitrance andthe downstream processing costs.

Although, second-generation biofuels are still in full devel-opment and still not economically viable, the usage of third-and fourth-generation biofuels are being explored. Oil extract-ed from cultured algae is considered the biofuel of the thirdgeneration, and it is expected that algae will have the potentialto produce more energy per acre than conventional crops [13].

Fourth-generation biofuels are still in the conceptual phaseand are based on the idea to use the plants’ photosyntheticmechanisms to produce hydrogen and electricity [14].

The Plant Cell Wall

The plant cell wall consists of a complex but dynamic networkof polysaccharides and other polymers around the plasmamembrane of plant cells (Fig. 1). This characteristic structureplays a key role in plant growth and development. In addition,it supports the cell as well as the plant and provides protectionagainst pathogens.

Despite the large diversity in cell wall morphology andcomposition, plant cell walls are generally classified as beingprimary or secondary. Primary cell walls are formed byexpanding cells and are mainly composed of polysaccharides(approximately 25% cellulose, 25% hemicellulose, and 35%pectin) and some proteins. Adjacent cells are joined togetherby a thin layer called the middle lamella, which is rich inpectin and wall proteins [15]. Unlike primary cell walls, sec-ondary cell walls are formed after cell growth ceased.

Table 1 Energy yieldfrom selected biomass,adapted from McKendry[3]

Biomass Energy yield (GJa/hab)

Wheat 123

Poplar 173–259

Switchgrass 139

Miscanthus 222–555

aGJ represents the unit gigajouleb ha represents the unit hectare

Bioenerg. Res.

Secondary cell walls differ both in structure and compositionfrom the primary wall. The multilayered and thick secondarywall is deposited against the primary cell wall and providesstrength to the cell in order to support the plant. It consistsof approximately 50 % cellulose, 25 % hemicelluloses, and20 % lignin [16].

Cellulose

Cellulose is known to be the most abundant organic com-pound on earth. It is a polymer of β-1,4-coupled glucose mol-ecules that are non-covalently linked by hydrogen bonds intomicrofibrils. The polymerization of the glucose monomers isperformed by plasma membrane-embedded protein com-plexes each composed of six cellulose synthase subunits.Each subunit transfers glucose residues from a sugar nucleo-tide donor (uridine diphosphate D-glucose) to the growingglucose chain [17]. The formedmicrofibrils can be longer than1 μm [18, 19] with a width that varies from 5 to 30 nm de-pending on the plant species [20, 21]. This variation indicatesthat the cellulose fibrils of different species may differ in thenumber of chains. Current models for cellulose microfiberorganization suggest that it has a substructure consisting ofhighly crystalline domains where glucose molecules arehighly ordered and bound to each other. These crystallinedomains are linked together by less-organized glucosechains, considered an amorphous region [15]. Its insolubilityin water, chemical stability, and relative resistance to hydro-lysis make it an excellent compound to support the cell wall.The cellulose microfibrils form spatially oriented overlap-ping layers around the cell and prevent it from burstingunder the osmotic pressure.

Hemicelluloses

Hemicelluloses are polysaccharides that form a strong butflexible network with cellulose; hence, they are consideredto be important for the load-bearing glycan network. In con-trast to cellulose, they are mostly composed of different hex-oses (D-glucose, D-galactose, and D-mannose) or pentoses (D-xylose and L-arabinose). Hemicelluloses have β-1, 4 linkedbackbones with an equatorial configuration. They are

synthesized by glycosyltransferases (GTs) in the Golgi andsubsequently transported in vesicles toward the plasma mem-brane. Upon fusion with the plasma membrane, these vesicleswill release their content in the apoplast where the polysac-charides can be incorporated in the cell wall [22]. The majorhemicelluloses in grasses are glucuronoarabinoxylan andarabinoxylan. In dicots, the major hemicellulose isxyloglucan, although in some tissues and in the secondary cellwall, the primary hemicellulose is xylan with a varying sidechain composition [23].

Pectin

The cellulose-hemicellulose network is embedded in a matrixcomposed of pectins, which are considered the most complexclass of cell wall polysaccharides. Similar to hemicelluloses,pectin is synthesized in the Golgi apparatus, delivered to thecell membrane via small vesicles, and released into the apoplastonce the Golgi vesicles fuse with the plasma membrane.Synthesis of pectin polysaccharides is estimated to involve atleast 67 different enzyme activities, including acetyltransfer-ases, methyltransferases, and glycosyltransferases [24, 25].Pectin is mainly present in the middle lamella and the primarycell wall and makes only a minor portion of the secondary cellwall. Non-commelinoid monocots and dicots contain up to35 % pectin in their primary walls and approximately 5 % insecondary walls whereas commelinoid monocot plants areknown to be relatively poor in pectins [26]. The primary func-tion of the pectin is to regulate intercellular adhesion, which isimportant for different physiological processes such as growth,development, defense response, seed hydration, leaf abscission,and fruit development [27]. Despite its low abundance in thesecondary cell wall, recent findings suggest a prominent role forpectin in the secondary cell wall [28]. Typical for pectins is thebackbone enriched in galacturonic acid (GalA). Depending onthe backbone composition and site chain decoration, severaldomains can be distinguished that are covalently linked to builda pectin network. The homopolymer of (1→4)-α-linked D-galacturonic acid is named homogalacturonan (HG) and makesthe smooth region of the pectin. Seventy to eighty percent of thegalacturonic acid residues are methyl esterified at the C-6 car-boxyl group, and the rest might be O-acetylated mostly at C-3

Fig. 1 Schematic representation of the molecular structure of the plant cell wall. Cellulose is organized into microfibrils while hemicelluloses andpectins crosslink cellulose microfibrils. Lignin covers cellulose microfibrils and fills the space between the polysaccharide matrix

Bioenerg. Res.

but sometimes at C-2. In order to form a stable gel with otherpectic molecules, the unmethylated C-6 site of HG residues isnegatively charged and might interact with Ca+2. This HG-calcium complex is referred to as the Begg box^ model. Thegalacturonic acid can also be substituted at its C-3 with a D-xylose resulting in a xylogalacturonan (XGA) structure [29].The second pectin domain is rhamnogalacturonan I(RG I) characterized by a backbone consisting of alternatinggalacturonic acid and rhamnose [(1→2)-α-L-rhamnose-α-(1→4)-α-D-galacturonic acid]. This backbone can be O-acetylated on C-2 and/or C-3 and arabinose and galactoseoligomer site chains can anchor to the RGI via rhamnosesugars [29, 30].

Pe rhaps the mos t complex pec t in domain i srhamnogalacturonan II (RG II). The RG II domain has at leasteight 1,4-linked α-D-GalpA residue backbones with four dif-ferent oligosaccharide side chains (named A to D) containingup to 13 different sugars linked by at least 22 possible glyco-sidic linkages [31]. RG II molecules are forming dimers via aboron diester in a way similar to the calcium-mediated eggbox structures [32]. This function of boron makes it an impor-tant microelement for plants [33].

Lignin

Lignin is crucial for the structural integrity of the secondarycell wall. The rigid and recalcitrant polymer supports theplant and provides protection against microorganisms [34].In addition, its hydrophobicity contributes to the imperme-ability of the cell wall, making transport of water and nutri-ents through the vascular system possible over longer dis-tances. Most likely, the emergence of lignin was crucial inthe development of land plants. In contrast to previouslydescribed polysaccharides, it is a complex aromatic polymercomposed of hydroxycinnamyl alcohol monomers that differin their degree of methoxylation, namely p-coumaryl,coniferyl, and synapyl alcohol.

When incorporated into the lignin polymer, the alcoholswill give rise to p-hydroxyphenyl (H) guaiacyl (G) andsyringyl (S) units, respectively. Incorporation of the alcoholmonomers into the lignin occurs through radical reactions,which results in a highly complex polymer, and it is mainlythis level of complexity that gives lignin its recalcitrant char-acter. Most of these units are linked via ether bonds (in so-called β–O–4-structures) and carbon-carbon bonds [in resinol(β–β) and phenylcoumaran (β–5) structures] [35]. These car-bon linkages make the polymer an energy-rich carbon sinkthat has higher energy content than cellulose or hemicellu-loses. For instance, burning 1 g of lignin produces 30 % morethan the energy of cellulosic carbohydrates [36, 37]. The com-position and amount of lignin varies with the location in thecell wall, the developmental phase of the cell, the cell type, thetissue, the species, and the environmental circumstances [34,

38]. The lignin of angiosperms is mainly composed of G- andS-units with traces of H-units, while the gymnosperm lignincontains mostly G-units with traces of H-units. Grasses incor-porate G- and S-units in equal amounts as well as H-units intheir lignin [34, 39]

Production Process of Second-generation Biofuels

Production of biofuels from lignocellulosic biomass isachieved in three steps. (1) A pretreatment step is used toopen the cell wall. (2) Saccharification releases free sugars(preferentially glucose) from the polysaccharides via hydro-lysis. (3) The final step—fermentation—converts the re-leased sugars to ethanol [40].

Biomass Pretreatment

To open up the cell wall structure and lower its recalcitrance,biomass pretreatment is applied. Biomass pretreatment can bedone in various ways such as chemical pretreatment (acid,alkali, ionic liquid, ozonolysis), physical pretreatment (micro-wave, grinding, and milling), and physicochemical pretreat-ment (liquid hot water, steam explosion, wet oxidation, CO2

explosion) [41]. Normally, a combination of these processes isused to increase the efficiency [42]. One example of suchcombination is the steam explosion treatment where the bio-mass is mixed with dilute sulfuric acid, heated with steam, andquenched by rapid depressurization. During pretreatment,mainly the matrix polysaccharides and, to a lesser extent, lig-nin are solubilized. Besides the positive effects of the pretreat-ment on the cell wall structure, some inhibitory compoundscan be formed during this process. These compounds can havenegative effects on microbial activity in downstream fermen-tation processes [43, 44]. It is therefore common that theresulting biomass is washed extensively to remove these in-hibitors and to neutralize the pH.

Enzymatic Saccharification

Enzymatic saccharification is the preferred strategy to degradethe polysaccharide polymers of lignocellulosic biomass intofermentable sugars. Different necrotrophic or biotrophic mi-croorganisms secrete cellulose-degrading enzymes into theplant tissue (i.e., endoglucanases (EC.3.2.1.4), exoglucanasesor cellobiohydrolases (EC.3.2.1.91), and β-glucosidases(EC.3.2.1.21). The cost of heterologous-produced enzymeshas long been approximately 30 % of the total productioncosts for bioethanol (US$ 0.38/gal ethanol) [45].

In order to reduce this cost, extensive research was per-formed to optimize the production process of enzymes andto optimize enzymatic cocktails used for saccharification.However, the real cost contribution of enzymes to the produc-tion of lignocellulosic ethanol varies significantly, for

Bioenerg. Res.

instance, $0.12/gal [46], $0.10/gal [47], $0.30/gal [48], $0.32/gal [49], and $0.35/gal (Klein-Marcuschamer et al. [45]), and$0.40/gal [50]. Similar to the pretreatment process, there is noideal enzyme mixture that works on all kinds of biomass.Instead, dedicated cocktails are being developed to treat aspecific type of biomass under optimal conditions [51].Different approaches have been applied to improve the prop-erties of individual enzymes as well as mixtures to improvebiomass degradation. For instance, random mutagenesis hasfamously been applied to Trichoderma reseei to create strainRUT-C30 that secretes up to 20 g/L cellulase protein [52].Thanks to this progress in the field, extremely efficient en-zyme cocktails were obtained to degrade biomass. As a result,not the enzyme but rather the pretreatment cost is determiningthe costs for biofuel production these days [53, 54].

During the saccharification process, potential inhibitors forthe next biofuel conversion step (fermentation)—such as ace-tate—can accumulate in the mixture [55]. These are mainlyderived from deacetylation of pentose sugars of xylan andpectins; therefore, partial reduction of cell wall acetylationby modulating pectin acetylesterase/ acetyltransferase activi-ties might increase the conversion efficiency of lignocellulosicbiomass to biofuels [28].

Fermentation

Through fermentation, the sugars released after saccharifi-cation are converted to various products such as ethanol,antibiotics, food additives, pharmaceuticals, and surfac-tants. The microorganism most commonly used in fermen-tation is the yeast Saccharomyces cerevisiae. This yeast isable to grow both on simple sugars, such as glucose aswell as on the disaccharide sucrose. Furthermore, easymanipulation of S. cerevisiae in industrial fermentationprocesses makes it a perfect candidate microorganism forbioethanol production [56].

S. cerevisiae has high resistance to ethanol, consumes sig-nificant amounts of substrate in adverse conditions, and showshigh resistance to inhibitors present in the medium [57].Although pentoses are as good candidates for biofuel produc-tion as hexoses, fermentation of pentoses is more difficult thanapplying the same procedure to hexoses. Recently, Demekeet al. [58] circumvented this problem by creating a transgenicstrain of industrial yeast. Using an evolutionary engineeringstrategy, the researchers were able to improve the yeasts abil-ity to convert D-xylose to ethanol.

Plant Cell Wall Modification Strategies

There is a tight and complex interaction between the differentpolymers of the plant cell wall. In a typical secondary cellwall, lignins and hemicelluloses cover the cellulose fibersforming an important factor for the reduction in

saccharification efficiency. In order to maximize the produc-tion of cellulosic biofuels while minimizing costs associatedwith energy and chemical inputs, different approaches havebeen developed to reduce cell wall recalcitrance (e.g., modifi-cation of hemicellulose and monolignol biosynthesis path-ways, producing water soluble polymers, modification of cel-lulose crystallinity, and overexpressing genes coding for cellwall-modifying proteins [28, 59–62]. Using biotechnologicalapproaches to alter cell wall recalcitrance and increase biofuelyields from lignocellulosic biomass by expressing cell walldegrading enzymes derived from different organisms (plantparasitic nematodes, fungi, bacteria, insects, or molluscs)(Fig. 2) is an attractive topic for biofuel researchers.Heterologous expression of different genes coding for cellwall-degrading enzymes effect on plant phenotype has beenlisten in Table 2. Since the substrates for the enzymes accu-mulate in the apoplastic region, the enzymes had to be targetedtoward the extracellular space or in different plant cell com-partments. Therefore, different signal peptides have been usedfor compartmentalization of enzymes at different parts of theplant cell (Table 2.). As mentioned before, plant cell wallpentoses like xylose and arabinose could also be fermentedinto biofuels. However, to efficiently metabolize pentosesugars at an industrial scale, particularly D-xylose, whichmakes up to 35 % of total sugars in xylan-rich lignocellulosicbiomass is the field of intense research [58].

Cellulose Modification

In general, cellulose consists of crystalline and amorphousdomains in varying proportions depending on the source.The physical characteristics of cellulose as well as their chem-ical behavior and reactivity are strongly influenced by thearrangement of these domains. Complete depolymerizationof cellulose needs synergistic activity of different types ofenzymes. These enzymes include endoglucanases that cleaveinternal bonds of amorphous cellulose and create new chainends, cellobiohydrolases, or exoglucanases that cleave cello-biose from the ends of the cellulose polymer, and finally, β-glucosidases or cellobiose hydrogenases hydrolyzing cellobi-ose into monosaccharides [111].

In addition, lytic polysaccharide mono-oxygenases(LPMOs), e.g., AA9 endoglucanases will cleave crystallinecellulose through reductive elimination. The electrons usedfor this reaction can be delivered by the cellobiose dehydro-genase [112].

Plants are excellent sources for large-scale/low-cost pro-duction of high-value enzymes. The cost for enzymes pro-duced from bioengineered plants has been reported to be 5-to 80-folds lower than those from other sources [113]. Sincecellulases are relatively costly enzymes, different cellulaseshave been successfully expressed in plants (Table 1). Majordrawbacks of heterologous expression of cellulases are

Bioenerg. Res.

dramatic yield penalty and detrimental effect on plant devel-opment. To avoid these negative effects on plant growth anddevelopment, cellulases have been targeted to different cellu-lar compartments [63, 67–78]. For instance, Mahadevan et al.[63] successfully generated tobacco plants expressingendoglucanases (CEL5) originated from Thermotogamaritima with an Arabidopsis rubisco activase signal peptidefor chloroplast targeting. These transgenic plants increaseautohydrolytic efficiency with 33 %. Endocellulase E1(CEL5A) from Acidothermus cellulolyticus was expressed inboth Nicotiana tabacum and Zea mayswith secretion targetedto the cell wall. Tobacco andmaize plants were phenotypicallynormal compared to wild-type (WT) plants. After pretreat-ment, these transformed plants were clearly more digestiblethanWT requiring less severe pretreatments to achieve a com-parable conversion level [114]. Another strategy to overcomedeleterious effects is the use of thermostable enzymes, whichare not active during plant growth and might be activatedduring post-harvesting treatments [115]. For example, theendoglucanase (E1) gene from the thermophilic bacteriumAcidothermus cellulolyticus was overexpressed in rice.Hydrolysis of transgenic rice straw yielded 43 % more reduc-ing sugars than wild-type rice straw [64]. These studies indi-cate that the heterologous expression of cellulases in the plantmight reduce the requirement of cellulases for saccharificationand increase cell wall amenability to biomass conversion.

In order to improve the value of bioenergy crops, one of thechallenging approaches is increasing cellulose synthesis.Though, the molecular mechanisms of cellulose synthesisand deposition are not completely identified [116]. In additionto transgenic approaches, modification of plant cell wall poly-mers might also be drastically altered by specific mutations.For instance, mutations in genes involved in cellulose

synthesis like KORRIGAN (KOR) and CELLULOSE SYNTHASE A (CesA) have a lower cellulose crystallinity and in-creased conversion rates to fermentable sugars with only alittle decrease in growth rate [117]. On the other hand, hybridpoplars expressing Arabidopsis KOR reduced cellulose crys-tallinity without deleterious effect on plant growth [65].

Hemicellulose Modification

In plants, up to 35 % of the secondary cell wall consists ofhemicelluloses (mainly xylan), and it has been shown thatchemical removal of hemicelluloses could increase celluloseaccessibility and hydrolysis. Also, higher esterification of xy-lans with acetate reduces enzymatic hydrolysis efficiency. Theimportance of hemicellulose for the bio-based economy is alsoreflected on the global market of industrial enzymes, wherexylanases comprise approximately 25–28 % of total enzymesales [118]. Hence, plants with reduced xylan content could bevaluable for biofuel production. Studies on Arabidopsis re-vealed several GTs involved in xylan biosynthesis. The irreg-ular xylem 7 (irx7), irx8, and irx9 T-DNA insertion mutantsare deficient in glucoronoxylan biosynthesis and show a col-lapsed xylem phenotype. These mutants exhibit a severedwarf phenotype, which is not desirable for biomass produc-tion [119]. One strategy to overcome this problem isreintroducing xylan biosynthesis into these irx-mutants usingvessel specific promoters (e.g., VND6 or VND7). In theselines, the morphological phenotype is rescued, but hemicellu-lose and lignin content remains low. As a result, the plantsshow a 42 % increase in saccharification yield after hot waterpretreatment [61].

The complete hydrolysis of xylan requires the combinedactions of various enzymes such as endoxylanase (EC

Fig. 2 Biotechnological approaches for the modification of plant cellwalls by expressing cell wall-degrading enzymes derived from differentorganisms (plant parasitic nematodes, fungi, bacteria, insects, or mol-luscs) to increase bioethanol yields from lignocellulosic biomass. a Be-fore genetic manipulation, complex cell wall polymers are tightly con-nected with each other. b Gene coding for cell wall-degrading enzymes

were cloned, and suitable constructs are made with a signal peptide forsecretion and c introduced into the plant under control of a constitutivepromoter and depending on the substrate type, the secreted enzyme altersthe plant cell wall structure (lignin in brown, hemicellulose in red, pectinin green, cellulose in blue) and e boosts bioethanol yields

Bioenerg. Res.

Table 2 Literature overview of the effects of heterologous expression of gene coding for cell wall-degrading enzymes on the phenotypes and cell wall composition of the plant with the main focus onArabidopsis, poplar, tobacco, maize, rice, and potato plants

Polymer Gene Source ofenzymes

Organism Localization Phenotype Reference

Cellulose Endoglucanase (CEL5) B Tobacco Chloroplast Increased autohydrolytic efficiency Mahadevan et al. [63]

Cellulose Endoglucanase (CEL5A) B B Tobacco and maize Apoplast Improved cell wall digestibility Mahadevan et al. [63]

Cellulose Endoglucanase (E1) B Rice Apoplast 43 % more yielded sugar Chou et al. [64]

Cellulose KORRIGAN P Poplar Reduced cellulose crystallinity Maloney and Mansfield [65]Cellulose Glucanase B Tobacco Cytosol Increased digestibility of cell wall fibers Kawazu et al. [66]

Cellulose Glucanase B Tobacco and alfalfa Cytosol Transgenic plants exhibited normal growth Ziegelhoffer et al. [67]

Cellulose Glucanase B Tobacco Chloroplast Transgenic plants exhibited normal growth Dai et al. [68]

Cellulose Glucanase B Potato ER, vacuole, apoplast,chloroplast

Transgenic plants exhibited normal growth Dai et al. [69]

Cellulose Glucanase B Arabidopsis Apoplast Transgenic plants exhibited normal growth Ziegler et al. [70]

Cellulose Glucanase B Tobacco Cytosol, apoplast, chloroplast Transgenic plants exhibited normal growth Ziegelhoffer et al. [67]

Cellulose Glucanase B Tobacco Chloroplast Transgenic plants exhibited normal growth Jin et al. [71]

Cellulose Glucanase B Barley Cytoplasm Transgenic plants exhibited normal growth Xue et al. [72]

Cellulose Glucanase B Tobacco ER, vacuole, apoplast,chloroplast

Transgenic plants exhibited normal growth Dai et al. [73]

Cellulose Glucanase B Maize Apoplast Transgenic plants exhibited normal growth Biswas et al. [74]

Cellulose Glucanase B Duckweed Cytosol Transgenic plants exhibited normal growth Sun et al. [75]

Cellulose Glucanase B Rice Apoplast Increased conversion of plant biomass into glucose Oraby et al. [76]

Cellulose Glucanase B Maize Apoplast Increased autohydrolytic efficiency Ransom et al. [77]

Cellulose Glucanase B Tobacco Chloroplast Transgenic plants exhibited normal growth Yu et al. [78]

Hemicellulose Xylanase B Brassica napus Oil bodies Liu et al. [79]

Hemicellulose Xylanase B Barley Cytoplasm Patel et al. 80

Hemicellulose Xylanase A B Rice Cytosol No phenotypic effect was noted Kimura et al. [81]

Hemicellulose Xylanase III B Arabidopsis Cytosol, chloroplast, peroxisome No phenotypic effect was noted Hyunjong et al. [82]

Hemicellulose Feruloyl esterase B Arabidopsis Apoplast Increased biomass degradability Pogorelko et al. [83]

Hemicellulose B-xylosidase/α-arabinosidase B Arabidopsis Apoplast Increased biomass degradability Pogorelko et al. [83]

Hemicellulose Xylanase (xyn2) B Arabidopsis Golgi, apoplast, and vacuole Apoplast localized lines reduced xylose andincreased arabinose and ferulate content

Buanafina et al. [84]

Hemicellulose Carbohydrate esterase (CE15) F Arabidopsis Apoplast Transgenic lines reduced glucose and xylose content Tsai et al. [85]

Hemicellulose Xylanase B B Arabidopsis Apoplast Decreased in molecular weight of xylans Borkhardt et al. [86]

Hemicellulose Thermophilic xylanase B Tobacco Apoplast ND Chatterjee et al. [87]

Hemicellulose Thermostable xylanase B Tobacco Apoplast No growth penalty in transgenic lines Herpes et al. [88]

Pectin Polygalacturanase (PG) F Tobacco, Arabidopsis, and wheat Apoplast Increased saccharification yield Lionetti et al. [89]

Pectin Pectin methyl esterase inhibitors (PMEI) P Tobacco, Arabidopsis, and wheat Apoplast Increased saccharification yield Lionetti et al. [89]

Pectin Pectin acetyl esterase (PtPAE1) P Tobacco Apoplast Lower digestibility of pectin Gou et al. [90]

Pectin Pectin acetyl esterase (PAE) B Potato tubers Apoplast Cell wall of transgenic tubers were mechanically stronger Orfila et al. [91]

Pectin Pectate lyase (PtxtPL1) B Aspen Apoplast Increased solubility of pectin, xylan, and otherhemicelluloses

Biswal et al. [92]

Pectin EndoPG P Tobacco Apoplast Transgenic tobacco did not result in pectin degradationin vivo

Osteryoung et al. [93]

Bioenerg.R

es.

Table 2 (continued)

Polymer Gene Source ofenzymes

Organism Localization Phenotype Reference

Pectin EndoPG P Apple Apoplast Reduced cell wall rigidity and cell adhesion Atkinson et al. [94]

Pectin Rhamnogalacturonanase Flax ND Reduction in RGI content. No visual phenotypic changes Musialak et al. [95]

Pectin Galactanase Arabidopsis Apoplast Initiation of bolting was delayed Obre et al. [96]

Pectin Polygalacturanase Y Arabidopsis Apoplast Early stage development conspicuous malformation. Nophenotype differences at the later developmental stage

Prieto-Alcedo et al. [97]

Pectin Polygalacturanase II F Arabidopsis, tobacco Apoplast Transgenic Arabidopsis and tobacco showed yieldpenalty

Capodicasa et al. [98]

Pectin Endo-1.4-β-galactosidase F Potato Apoplast Increased solubility of pectins. Reduction in galactosylcontent by 30 %

Oxenboll Sorensen et al. [99]

Pectin Arabinanase (RGI) F Potato Apoplast Arabinose content was decreased by approximately 70 % Skjøt et al. [100]

Pectin endoRGL F Potato Apoplast Large reduction in galactosyl and arabinosyl residues intransgenic tubers

Oomen et al. [101]

Pectin endoPG F Tobacco Apoplast 30 % decrease in the GalUA content of the cell walls Boudart et al. [102]

Pectin PME F BY2 Apoplast Significant rise of PME and the level of methanol in thetransformant increased by 28.7 %

Hasunuma et al. [103]

Pectin PME F Tobacco Apoplast Alteration in cell wall metabolism and a dwarftrangenic line

Hasunuma et al. [104]

Pectin F Potato Apoplast Transgenic plants have stiff stems and thickened, curled,and red brownish leaves. Do not produce side shootor flowers

Borkhard et al. [86]

Pectin PGI and RGA F Flax Apoplast Significant reduction of pectin content Musialak et al. [95]

Pectin PGs F Arabidopsis, tobacco Apoplast Increase plant resistance to the pathogens and reduceauxin sensitivity

Ferrari et al. [105]

Lignin Lig-D B Arabidopsis Apoplast Lignin composition altered Tsuji et al. [106]

Lignin QsuB B Arabidopsis, tobacco Apoplast Lignin content reduced and biomass saccharificationefficiency improved

Eudes et al. [107]

CBM CBM B Tobacco Apoplast Reduced stem elongation, prolonged juvenility, delayedflowering

Obembe et al. [108]

CBM CBM B Tobacco Apoplast No observable changes were detected Obembe et al. [108]

CBM CBM N Arabidopsis Apoplast Increased root length, enhanced susceptibility toH. schachtii

Hewezi et al. [109]

CBM CBM B Potato Apoplast Fast elongation of the main stem, high and big leaf areasof transgenic lines

Safra-Dassa et al. [110]

Source of enzymes: B Bacteria, F Fungi, N Nematode, P Plant, Y Yeast

ND not determined

Bioenerg.R

es.

3.2.1.8), β-d-xylosidase (EC 3.2.1.37), α-glucuronidase (EC3.2.1.139), acetyl xylan esterase (EC 3.2.1.72), ferulic/coumaric acid es terase (EC 3.2 .1 .73) , and α - l -arabinofuranosidase (EC 3.2.1.55) [120, 121]. Large numbersof xylanases have been found in a variety of bacteria andfungi, some of which have been cloned, characterized[121–123], and expressed in plants [86, 87].

The reduction of xylan content in plants to increase cellwall degradability by heterologous expression of xylanaseswas shown by several research groups [84, 86]. For instance,Buanafina et al. [84] studied the production of Trichodermareesei xylanase (xyn2) targeted toward different cell compart-ments such as the Golgi, apoplast, and vacuole.

The lines with an apoplast localized xylanase showed anincreased ferulate and arabinose content with a decreasedamount of xylose. However, these plants had a 30% reductionin enzymatic saccharification efficiency.

An alternative approach to avoid detrimental effects causedby highly active xylanases is the use of thermophilic endo-xylanases. In Arabidopsis, this strategy resulted in plants withnormal growth characteristics, indicating that the enzyme isnot active during plant development [86]. Cell extractionfrom dry bioengineered stems showed a high enzymaticactivity during incubation at 85 °C for 24 h, indicatingthat the enzyme is not degraded during stem senescence.Engineered plants showed a decrease in molecular weightof xylans compared to WT lines; however, sugar releaseof these plants has not been reported [86]. Similarly,Chatterjee et al. [87] successfully expressed a thermophilicxylanase in tobacco without any deleterious effect. In thisstudy, the enzyme activity and transcript analysis havebeen identified, but there was no information regardingthe cell wall composition.

Besides the use of xylanases, other hemicellulosedegrading enzymes have been expressed in different plants.One of these papers describes the expression of feruloyl ester-ase and β-xylosidase/α-arabinosidase derived fromAspergillus nidulans in Arabidopsis thaliana. These transgen-ic plants did not show any visible phenotype; yet, they haveincreased biomass degradability [83]. Tsai et al. [85] demon-strated that in planta expression of carbohydrate esterase(CE15) from the white-rot basidiomycete Phanerochaetecarnosa resulted in a leaf-yellowing phenotype and reducedxylose and glucose content. This study revealed that transgen-ic expression of a fungal glucuronoyl esterase in plants couldalter the extent of intermolecular cross-linking within plantcell walls [85]. Heterologous expression of the glucuronoylesterase in bioenergy crops in the context of biofuels has beenstudied by Latha Gandla et al. [124]. Expressing fungalglucuronoyl esterase in Populus increased lignin content andreduced cellulose content and saccharification yield.However, the enzymatic hydrolysis following the acid pre-treatment, which efficiently removed the inhibitory effects of

lignin, resulted in increased cellulose conversion by about12 % in the transgenic lines [124].

Furthermore, several hemicellulose-degrading enzymeshave been expressed in different plants. Unfortunately, inmost of these studies, the effect on saccharification yieldand cell wall composition was not investigated. Most likely,these strategies will have a positive effect on the saccharifi-cation. This can be deduced from the observation that theaddition of α-xylosidase to commercial cellulase prepara-tions, which lack α-xylosidase activity, improved ferment-able sugar yield [122]. In addition, Alvira et al. [123] em-phasized that application of endoxylanase (XlnC) fromAspergillus nidulans and α-Larabinofuranosidase fromAspergillus niger increased enzymatic hydrolysis for ethanolproduction on wheat straw by 10 %.

Pectin Modification

Despite the low abundance of pectin in the secondary cellwalls that comprise the majority of lignocellulosic biomass,recent studies suggest that pectin affects secondary cell wallformation [28]. Pectins are secreted into the cell wall in ahighly methyl-esterified form and are subsequently de-esterified in the apoplast by pectin methylesterase (PME).As de-methylesterified homogalacuronic acid interacts withcalcium to form the egg box structure, methyl-esterificationinfluences the number of the calcium bridges between pectinmolecules and hence determines to some extent cell wall ri-gidity [125, 126]. Expression of PME in poplar was shown tostimulate the length of wood fibrils in transgenic lines [127].In addition, expression of fungal polygalacturanase (PG) orpectin methylesterase inhibitor (PMEI) in Arabidopsis, tobac-co and wheat increased saccharification yield [89]. Moreover,mutant Arabidopsis plants affected in gene-encoding proteinsinvolved in methylesterification (pectin methylesterase 3;pme3) and homogalacturonan biosynthesis (quasimodo2-1;qua2-1) revealed that reduction in methylesterification ofHGA level improves saccharification yield [128]. Also, theacetylation of pectin affects cell wall recalcitrance with highlyacetylated pectin being less susceptible to degradation andhence increasing biomass recalcitrance [55]. Expressing pop-lar pectin acetyl esterase (PtPAE1) in tobacco led to lowerdigestibility of pectin [90]. Furthermore, side chains of RGIsuch as galactan, arabinan, and arabinogalactan are supposedto modulate cell wall porosity by filling wall gaps [96, 129].Therefore, engineering of RGI side chains might result intomore porous cell wall structures characterized by a reducedrecalcitrance [130].

Lignin Modification

Lignin is one of the major limiting factors reducing sacchari-fication efficiency [131]. Modification of lignin pathway is

Bioenerg. Res.

one of the well-studied subject to reduce cell wall recalci-trance, and an increasing number of examples show that ligninmodification can improve the downstream processing of plantbiomass [60, 131–134]. For example, the Arabidopsis cse(caffeoyl shikimate esterase) mutant deposits less lignin thanwild-type plants, and the remaining lignin is enriched in p-hydroxyphenyl units. This lignin modification enhances theconversion of cellulose to glucose in cse mutants up to four-fold as compared to that in the wild type without pretreatment[135]. Since lignin is essential to support the plant and trans-port water and nutrients from the roots toward the leaves, largereductions in lignin content result in undesired phenotypessuch as dwarfism [136]. Nevertheless, rather than reducinglignin content, changing the structure of the lignin polymermight be another elegant way in order to diminish cell wallrecalcitrance [137, 138]. Lignin enriched in S-units is moreeasily removed and more accessible to hydrolytic enzymes,and Arabidopsis plants with high S-lignin content releasedmore glucose after pretreatment compared to G-ligninenriched plants [139]. Alternatively, there is a novel and prom-ising strategy that involve expression of specific proteins toalter lignin building blocks composition andmake lignin poly-mer more prone to degradation [107, 140, 141]. For instance,expression of bacterial 3-dehydroshikimate dehydratase incombination with specific promoters in Arabidopsis reducedlignin content and improved saccharification efficiency [107].

Expressing Carbohydrate-Binding Modules

Carbohydrate-binding proteins (e.g., carbohydrate-bindingmodules (CBM), expansins, and swolenins) are non-catalyticproteins, which might interfere with the tightly connectedpolysaccharide network and affect cell growth and expansion.By loosening cell wall structures, they might increase the cel-lulose accessibility [142, 143]. The heterologous expressionof a CBM derived from Clostridium cellulovorans targeted tothe cell wall in transgenic poplar, tobacco, and potato resultedin plants with accelerated plant growth [144–146].

A similar increased growth rate was observed whenexpansins were expressed in Arabidopsis, poplar, or rice[108, 147, 148]. In contrast, the heterologous expression ofCBMderived from Piromyces inNicotiana bentamina reducedstem elongation and delayed flower development withoutchanges in cellulose content [149]. Unfortunately, these excel-lent studies do not include any data concerning of the effect ofexpressing non-catalytic proteins on saccharification yield.

Production of Water-Soluble Polymers Within the CellWall

The classical approach to improved feedstock is enhancingcellulose accessibility by reducing the concentration of a spe-cific cell wall polymer (either polysaccharide or lignin). These

modifications most often result in reduced plant fitness sincethey alter the mechanical properties of the cell wall. In order toovercome this negative effect, a new strategy has been devel-oped (called ‘Trojan Horse’) whereby soluble polysaccharidesare introduced into the plant cell wall. These polymers behavenormally during plant growth and development but displayenhanced solubility during downstream processing. The ideais to create soluble Bpockets,^ enabling channel formation andpermitting rapid solvent and enzyme penetration during thepretreatment resulting in plants with higher saccharificationyields [150]. Alginate, carrageenan, acetan, hyaluronan, chi-tosan, and levan polymers derived from algae cell wall orbacterial exopolysaccharides could be used as a source ofsoluble polysaccharides. Although this approach is still in itsinfancy, some of the polymers were successfully expressed inplants already. For example, levansucrase (a water-solublefructan polymer synthesized from sucrose) of Erwiniaamylovora was produced in Eucalyptus resulting in highlyporous cell wall structures compared to their wild-type coun-terparts by using elongation-specific and secondary cell walldevelopment promoters since the use of constitutive pro-moters appeared to be lethal and no transformants were ob-tained [151]. Although this could be due to the modified cellwall, according to the author, this is a direct result of theextensive use of sucrose to produce water-soluble fructanpolymer making sucrose limiting for cellulose synthesis.Other promising results were obtained by the production ofresilin in the apoplast of tobacco. The accumulation of thiselastomeric rubber-like protein of insects in the cell wall im-proved sugar release from plant material [152]. Surprisingly,expression of NodC-expressing Arabidopsis plants accumu-lating N-acetylglucosamine (GlcNAc) oligomers showed in-creased cell wall recalcitrance and reduced saccharificationyield [128]. Reducing cell wall digestibility could be due tothe GlcNAc oligomers competing with non-covalent interac-tions in the apoplast.

Concluding Remarks

Since fossil fuels are limited and their combustion has a sig-nificant effect on global climate change, the introduction ofrenewable and sustainable energy sources in our economy is amust. Lignocellulosic biomass is an interesting source for bothenergy and products; however, the challenge lies in the effi-cient extraction of fermentable sugars from the highly recalci-trant plant cell wall. Lignocellulosic biomass saccharificationrequires pretreatment to diminish substrate recalcitrance andhigh enzyme loadings, which would otherwise make the costof ethanol production from lignocellulose more expensivethan fossil-based fuels. An ideal Arabidopsis cell wall com-position for biofuel research has been investigated by VanAcker et al. [131], which contains less lignin and more

Bioenerg. Res.

hemicelluloses that comprise less xylose, more galactose,arabinose units, and these cell wall properties. Genetic manip-ulation of plants to alter their cell walls is one of the promisingapproaches to reduce cell wall recalcitrance and improve bio-fuel production potential. However, these genetic modifica-tions (including reducing lignin contents and producing gly-coside hydrolases in plant cells) create major concerns forplant growth and development resulting in plants sufferingfrom severe growth defects (i.e., dwarfing, collapsed xylem,and low mechanical strength). For in planta expression ofglycoside hydrolases, tissue-specific gene expression ortargeting the enzyme to specific subcellular compartments isattractive choices to reduce undesired activities in the plant orincrease enzyme production. Alternatively, thermophilic en-zymes, which are not active under normal plant growth con-ditions, are successfully used to avoid growth defects. Thus,heterologous expression of cell glycoside hydrolases in plantsis a potential way to make lignocellulosic biomass suitable forthe biorefineries and to reduce the cost of biomass processing.The process of developing an ideal plant, in order to furtherreduce cell wall complexity and get more yields, different cellwall polymers should be targeted in one plant.

Acknowledgements The author would like to thank Bartel Vanholmeand Godelieve Gheysen for help in formatting the manuscript and AliAkyüz for formatting drawing. HT is indebted to the Scientific and Tech-nological Research Council of Turkey (TUBITAK) for a partial predoc-toral fellowship.

References

1. IEA (2013) World energy outlook 2013. IEA Publications. www.worldenergyoutlook.org

2. De Jong E, Langeveld H, Van Ree R. (2009) IEA bioenergy task42 on biorefinery

3. McKendry P (2002) Energy production from biomass (part 1):overview of biomass. Bioresour Technol 83:37–46

4. Gray KA, Zhao L, Emptage M (2006) Bioethanol. Curr OpinChem Biol 10(2):141–6

5. Young AL (2009) Finding the balance between food and biofuels.Environ Sci Pollut Res Int 16(2):117–9

6. Hill J, Nelson E, Tilman D, Polasky S, Tiffany D (2006)Environmental, economic, and energetic costs and benefits of bio-diesel and ethanol biofuels. Proc Natl Acad Sci U S A 103(30):11206–10

7. Ford C, Runge BS (2007) Biofuel: corn isn’t the king of thisgrowing domain. Nature 450:478

8. Gomez LD, Steele-King CG, McQueen-Mason SJ (2008)Sustainable liquid biofuels from biomass: the writings on thewalls. New Phytol 178(3):473–85

9. TilmanD,Hill J, LehmanC (2006) Carbon-negative biofuels fromlow-input high-diversity grassland biomass. Science 314(5805):1598–600

10. Davis J (2008) Genetic improvement of poplar (Populus spp.) as abioenergy crop. In: Vermerris W (ed) Genetic improvement ofbioenergy crops. Springer, New York, pp 397–419

11. Gordon JC (2001) Poplars: trees of the people, trees of the future.For Chron 77(2):217–9

12. Wingren A, Galbe M, Zacchi G (2003) Techno-economic evalua-tion of producing ethanol from softwood: comparison of SSF andSHF and identification of bottlenecks. Biotechnol Prog 19(4):1109–17

13. Chisti Y (2007) Biodiesel from microalgae. Biotechnol Adv25(3):294–306

14. Antizar-Ladislao B, Turrion-Gomez JL (2008) Second-generationbiofuels and local bioenergy systems. Biofuels Bioprod Bioref2(5):455–69

15. Cosgrove DJ (1997) Assembly and enlargement of the primarycell wall in plants. Annu Rev Cell Dev Biol 13:171–201

16. Cosgrove DJ (2005) Growth of the plant cell wall. Nat Rev MolCell Biol 6(11):850

17. Peng L, Kawagoe Y, Hogan P, Delmer D (2002) Sitosterol-beta-glucoside as primer for cellulose synthesis in plants. Science295(5552):147–50

18. Brett CT (2000) Cellulose microfibrils in plants: biosynthe-sis, deposition, and integration into the cell wall. Int RevCytol 199:161

19. Brown DM, Zeef LA, Ellis J, Goodacre R, Turner SR (2005)Identification of novel genes in Arabidopsis involved in secondarycell wall formation using expression profiling and reverse genet-ics. Plant Cell 17(8):2281–95

20. Ha MA, Apperley DC, Evans BW, Huxham IM, Jardine WG,Viëtor RJ et al (1998) Fine structure in cellulose microfibrils:NMR evidence from onion and quince. Plant J 16:183–90

21. Herth W (1983) Arrays of plasma-membrane rosettes involved incellulose microfibril formation of spirogyra. Planta 159(4):347–56

22. Scheller HV, Ulvskov P (2010) Hemicelluloses. Annu Rev PlantBiol 61:263–89

23. Ebringerová A, Hromádková Z, Heinze T (2005) Hemicellulose186:1–67

24. Mohnen D (2008) Pectin structure and biosynthesis. Curr OpinPlant Biol 11:266–77

25. Harholt J, Suttangkakul A, Vibe SH (2010) Biosynthesis of pectin.Plant Physiol 153(2):384–95

26. Wiethölter N, Graeßner B, Mierau M, Mort AJ, MoerschbacherBM (2003) Differences in the methyl ester distribution ofhomogalacturonans from near-isogenic wheat lines resistant andsusceptible to the wheat stem rust fungus. Mol Plant MicrobeInteract 16(10):945–54

27. Caffall KH, Mohnen D (2009) The structure, function, and bio-synthesis of plant cell wall pectic polysaccharides. Carbohydr Res344(14):1879–900

28. Xiao C, Anderson CT (2013) Roles of pectin in biomass yield andprocessing for biofuels. Front Plant Sci 4:67

29. O’Neill M, Albersheim P, Darvill A (1990) The pecticpolysaccharides of primary cell walls. In: CarbohydratesPMD, Harborne JB (eds) Methods in plant biochemistry.Academic, London, pp 415–41

30. Albersheim P, Darvill A, O’Neill M, Schols H, Voragen A (1996)An hypothesis; the same six polysaccharides are components ofthe primary cell walls of all higher plants. In: Visser J, VoragenAGJ (eds) Pectins and pectinases. Elsevier Science BV,Amsterdam, pp 47–55

31. O’Neill MA, Ishii T, Albersheim P, Darvill AG (2004)Rhamnogalacturonan II: structure and function of a boratecross-linked cell wall pectic polysaccharide. Annu Rev PlantBiol 55:109–39

32. Ishii T, Matsunaga T (1996) Isolation and characterization of aboron-rhamnogalacturonan-II complex from cell walls of sugarbeet pulp. Carbohydr Res 284(1):1–9

33. Shorrocks VM (1997) The occurrence and correction of borondeficiency. Plant Soil 193(1–2):121–48

34. BoerjanW, Ralph J, BaucherM (2003) Lignin biosynthesis. AnnuRev Plant Biol 54:519–46

Bioenerg. Res.

35. Vanholme R, Demedts B, Morreel K, Ralph J, Boerjan W (2010)Lignin biosynthesis and structure. Plant Physiol 153(3):895–905

36. Shafizadeh F, Chin P (1977) Thermal degradation of wood. In ISGoldstein (ed), Wood technology: chemical aspects Americanchemical society symposium series, Washington DC, 57–81

37. White RH (1987) Effect of lignin content and extractives on thehigher heating value of wood. Wood Fiber Sci 19:446–52

38. Ralph J, Lundquist K, Brunow G, Lu F, Kim H, Schatz P et al(2004) Lignins: natural polymers from oxidative coupling of 4-hydroxyphenyl-propanoids. Phytochem Rev 3:29–60

39. Vogel J (2008) Unique aspects of the grass cell wall. Curr OpinPlant Biol 11(3):301–7

40. Jordan DB, Bowman MJ, Braker JD, Dien BS, Hector RE, LeeCC et al (2012) Plant cell walls to ethanol. Biochem J 442:241–52

41. Mood SH, Golfeshan AH, Tabatabaei M, Abbasalizadeh S,Ardjmand M (2013) Comparison of different ionic liquids pre-treatment for barley straw enzymatic saccharification. Biotech3(5):399–406

42. Zhang D, VanFossen AL, Pagano RM, Johnson JS, Parker MH,Pan S et al (2011) Consolidated pretreatment and hydrolysis ofplant biomass expressing cell wall degrading enzymes. BioEnergyRes 4(4):276–86

43. Klinke HB, Thomsen AB, Ahring BK (2004) Inhibition ofethanol-producing yeast and bacteria by degradation products pro-duced during pre-treatment of biomass. Appl MicrobiolBiotechnol 66:10–26

44. Palmqvist E, Hahn-Hagerdal B (2000) Fermentation of lignocel-lulosic hydrolysates. II: inhibitors and mechanisms of inhibition.Bioresour Technol 74:25–33

45. Klein-Marcuschamer D, Oleskowicz-Popiel P, SimmonsBA, Blanch HW (2012) The challenge of enyzme cost inthe production of lignocellulosic biomass. BiotechnolBioeng 109(4):1083–7

46. Merino ST, Cherry J (2007) Progress and challenges in enzymedevelopment for biomass utilization. Biofuels 108:95–120

47. Aden A, Foust T (2009) Technoeconomic analysis of the dilutesulfuric acid and enzymatic hydrolysis process for the conversionof corn stover to ethanol. Cellulose 16:535–45

48. Lynd LR, LaserMS, Bransby D, Dale BE, Davison B, Hamilton Ret al (2008) How biotech can transform biofuels. Nat Biotechnol26:169–72

49. Dutta A, Dowe N, Ibsen KN, Schell DJ, Aden A (2010) An eco-nomic comparison of different fermentation configurations to con-vert corn stover to ethanol using Z. mobilis and Saccharomyces.Biotechnol Prog 26:64–72

50. Kazi FK, Fortman JA, Anex RP, Hsu DD, Aden A, Dutta A et al(2010) Techno-economic comparison of process technologies forbiochemical ethanol production from corn stover. Fuel 89:20–8

51. Levine SE, Fox JM, Clark DS, Blanch HW (2011) A mechanisticmodel for rational design of optimal cellulase mixtures.Biotechnol Bioeng 108:2561–70

52. Peterson R, Nevalainen H (2012) Trichoderma reesei RUT-C30—thirty years of strain improvement. Microbiology 158(Pt 1):58–68

53. Alvira P, Tomas-Pejo E, Ballesteros M, Negro MJ (2010)Pretreatment technologies for an efficient bioethanol productionprocess based on enzymatic hydrolysis: a review. BioresourTechnol 101(13):4851–61

54. Haghighi Mood S, Hossein Golfeshan A, Tabatabaei M, SalehiJouzani G, Najafi GH, Gholami M et al (2013) Lignocellulosicbiomass to bioethanol, a comprehensive review with a focus onpretreatment. Renew Sust Energ Rev 27:77–93

55. Gille S, Pauly M (2012) O-acetylation of plant cell wall polysac-charides. Front Plant Sci 3:12

56. Lin Y, Tanaka S (2006) Ethanol fermentation from biomass re-sources: current state and prospects. Appl Microbiol Biotechnol69(6):627–42

57. Hector R, Qureshi N, Hughes S, Cotta M (2008) Expression of aheterologous xylose transporter in a Saccharomyces cerevisiaestrain engineered to utilize xylose improves aerobic xylose con-sumption. Appl Microbiol Biotechnol 80(4):675–84

58. Demeke MM, Dietz H, Li Y, Foulquié-Moreno MR, Mutturi S,Deprez S et al (2013) Development of a D-xylose fermenting andinhibitor tolerant industrial Saccharomyces cerevisiae strain withhigh performance in lignocellulose hydrolysates using metabolicand evolutionary engineering. Biotechnol Biofuels 6:89

59. Abramson M, Shoseyov O, Shani Z (2010) Plant cell wall recon-struction toward improved lignocellulosic production and process-ability. Plant Sci 178(2):61–72

60. Eudes A, Liang Y, Mitra P, Loqué D (2014) Lignin bioengineer-ing. Curr Opin Biotechnol 26:189–98

61. Petersen PD, Lau J, Ebert B, Yang F, Verhertbruggen Y, Kim JSet al (2012) Engineering of plants with improved properties asbiofuels feedstocks by vessel-specific complementation of xylanbiosynthesis mutants. Biotechnol Biofuels 5:84

62. Garvey M, Klose H, Fischer R, Lambertz C, Commandeur U(2013) Cellulases for biomass degradation: comparing recombi-nant cellulase expression platforms. Trends Biotechnol 31(10):581–93

63. Mahadevan SA, Wi SG, Kim YO, Lee KH, Bae HJ (2011) Inplanta differential targeting analysis of Thermotoga maritimaCel5A and CBM6-engineered Cel5A for autohydrolysis.Transgenic Res 20:877–86

64. Chou HL, Dai Z, Hsieh CW, KuMS (2011) High level expressionof Acidothermus cellulolyticus beta-1, 4-endoglucanase in trans-genic rice enhances the hydrolysis of its straw by cultured cowgastric fluid. Biotechnol Biofuels 4:58

65. Maloney VJ, Mansfield SD (2010) Characterization and variedexpression of a membrane-bound endo-beta-1,4-glucanase in hy-brid poplar. Plant Biotechnol J 8:294–307

66. Kawazu T, Sun JL, Shibata M, Kimura T, Sakka K, Ohmiya K(1999) Expression of a Bacterial Endoglucanase Gene in TobaccoIncreases Digestibility of Its Cell Wall Fibers. J Biosci Bioeng88(4):421–425

67. Ziegelhoffer T, Raasch JA, Austin-Phillips S (2001) Dramatic ef-fects of truncation and sub-cellular targeting on the accumulationof recombinant microbial cellulase in tobacco. Mol Breed 8(2):147–58

68. Dai Z, Hooker BS, Anderson DB, Thomas SR (2000) Expressionof Acidothermus cellulolyticus endoglucanase E1 in transgenictobacco: biochemical characteristics and physiological effects.Transgenic Res 9(1):43–54

69. Dai Y, Hooker BS, Anderson DB, Thomas SR (2000)Improved plant-based production of E1 endoglucanase usingpotato: expression optimization and tissue targeting. MolBreed 6:277–85

70. Ziegler MT, Thomas SR, Danna KJ (2000) Accumulation of athermostable endo-1, 4-β-D-glucanase in the apoplast ofArabidopsis thaliana leaves. Mol Breed 6(1):37–46

71. Jin R, Richter S, Zhong R, Lamppa GK (2003) Expression andimport of an active cellulase from a thermophilic bacterium intothe chloroplast both in vitro and in vivo. Plant Mol Biol 51:493–507

72. Xue GP, Patel M, Johnson JS, Smyth DJ, Vickers CE (2003)Selectable marker-free transgenic barley producing a high levelof cellulase (1,4-β-glucanase) in developing grains. Plant CellRep 21:1088–94

73. Dai Z, Hooker BS, Quesenberry RD, Thomas SR (2005)Optimization of Acidothermus cellulolyticus endoglucanase(E1) production in transgenic tobacco plants by transcriptional,post-transcription and post-translational modification.Transgenic Res 14:627–43

Bioenerg. Res.

74. Biswas GCG, Ransom C, Sticklen M (2006) Expression of bio-logically active acidothermus cellulolyticus endoglucanase intransgenic maize plants. Plant Sci 171(5):617–23

75. Sun Y, Cheng JJ, Himmel ME, Skory CD, Adney WS, ThomasSR et al (2007) Expression and characterization of Acidothermuscellulolyticus E1 endoglucanase in transgenic duckweed Lemnaminor 8627. Bioresour Technol 98:2866–2872

76. Oraby H, Venkatesh B, Dale B, Ahmad R, Ransom C, Oehmke Jet al (2007) Enhanced conversion of plant biomass into glucoseusing transgenic rice-produced endoglucanase for cellulosic etha-nol. Transgenic Res 16(6):739–49

77. Ransom C, Balan V, Biswas G, Dale B, Crockett E, Sticklen M(2007) Heterologous Acidothermus cellulolyticus 1, 4-β-endoglucanase E1 produced within the corn biomass convertscorn stover into glucose. Appl Biochem Biotechnol 137(140):207–19

78. Yu LX, Gray BN, Rutzke CJ,Walker LP,Wilson DB, HansonMR(2007) Expression of thermostable microbial cellulases in thechloroplasts of nicotine-free tobacco. J Biotechnol 131(3):362–9

79. Liu JH, Selinger LB, Cheng KJ, Beauchemin KA, Moloney MM(1997) Plant seed oil-bodies as an immobilization matrix for arecombinant xylanase from the rumen fungus Neocallimastixpatriciarum. Mol Breed 3(6):463–470

80. Patel M, Johnson JS, Brettell RIS, Jacobsen J, Xue GP (2000)Transgenic barley expressing a fungal xylanase gene in the endo-sperm of the developing grains. Mol Breed 6(1):113–124

81. Kimura T, Mizutani T, Tanaka T, Koyama T, Sakka K, Ohmiya K(2003) Molecular breeding of transgenic rice expressing axylanase domain of the xynA gene from Clostridiumthermocellum. Appl Microbiol Biotechnol 62(4):374–379

82. Hyunjong B, Lee DS, Hwang I (2006) Dual targeting of xylanaseto chloroplasts and peroxisomes as a means to increase proteinaccumulation in plant cells. J Exp Bot 57(1):161–169

83. Pogorelko G, Fursova O, Lin M, Pyle E, Jass J, ZabotinaOA (2011) Post-synthetic modification of plant cell wallsby expression of microbial hydrolases in the apoplast. PlantMol Biol 77(4–5):433–45

84. BuanafinaMM, Langdon T, Dalton S,Morris P (2012) Expressionof a Trichoderma reesei beta-1,4 endo-xylanase in tall fescue mod-ifies cell wall structure and digestibility and elicits pathogen de-fence responses. Planta 236(6):1757–74

85. Tsai AY, Canam T, Gorzsas A, Mellerowicz EJ, Campbell MM,Master ER (2012) Constitutive expression of a fungal glucuronoylesterase in Arabidopsis reveals altered cell wall composition andstructure. Plant Biotechnol J 10(9):1077–87

86. Borkhardt B, Harholt J, Ulvskov P, Ahring BK, Jorgensen B,Brinch-Pedersen H (2010) Autohydrolysis of plant xylans byapoplastic expression of thermophilic bacterial endo-xylanases.Plant Biotechnol J 8(3):363–74

87. Chatterjee A, Das NC, Raha S, Babbit R, Huang Q, Zaitlin D et al(2010) Production of xylanase in transgenic tobacco for industrialuse in bioenergy and biofuel applications. In Vitro Cellular andDevelopmental Biology Plant 46(2):198–209

88. Herbers K,Wilke I, Sonnewald U (1995) A thermostable xylanasefrom Closteridium thermocellum expressed at high levels in theapoplast of transgenic tobacco has no detrimental effects and iseasily purified. Biotechnology 13:63–66

89. Lionetti V, Francocci F, Ferrari S, Volpi C, Bellincampi D, GallettiR et al (2010) Engineering the cell wall by reducing de-methyl-esterified homogalacturonan improves saccharification of planttissues for bioconversion. Proc Natl Acad Sci U S A 107(2):616–21

90. Gou JY, Miller LM, Hou G, Yu XH, Chen XY, Liu CJ (2012)Acetylesterase-mediated deacetylation of pectin impairs cell elon-gation, pollen germination, and plant reproduction. Plant Cell24(1):50–65

91. Orfila C, Dal Degan F, Jorgensen B, Scheller HV, Ray PM,Ulvskov P (2012) Expression of mung bean pectin acetyl esterasein potato tubers: effect on acetylation of cell wall polymers andtuber mechanical properties. Planta 236:185–196

92. Biswal AK, Soeno K, Gandla ML, Immerzeel P, Pattathil S,Lucenius J et al (2014) Aspen pectate lyase PtxtPL1-27 mobilizesmatrix polysaccharides fromwoody tissues and improves sacchar-ification yield. Biotechnol Biofuels 7:11

93. Osteryoung KW, Toenjes K, Hall B, Winkler V, Bennett AB(1990) Analysis of Tomato Polygalacturonase Expression inTransgenic Tobacco. Plant Cell 2:1239–1248

94. Atkinson RG, Schroder R, Hallett IC, Cohen D, MacRae EA(2002) Overexpression of polygalacturonase in transgenic appletrees leads to a range of novel phenotypes involving changes incell adhesion. Plant Physiol 129(1):122–133

95. Musialak M, Wróbel-Kwiatkowska M, Kulma A, Starzycka E,Szopa J (2008) Improving retting of fibre through genetic modi-fication of flax to express pectinases. Transgenic Res 17:133–147

96. Obro J, Borkhardt B, Harholt J, Skjot M,Willats WGT, Ulvskov P(2009) Simultaneous in vivo truncation of pectic side chains.Transgenic Res 18:961–9

97. Prieto-Alcedo M, Veiga-Crespo P, Poza M, Coronado C, Zarra I,Villa TG (2011) Expression of a yeast polygalacturonase gene inArabidopsis thaliana. Biol Plant 55(2):349–352

98. Capodicasa C, Vairo D, Zabotina O, McCartney L, Caprari C,Mattei B et al (2004) Targeted modification of homogalacturonanby transgenic expression of a fungal polygalacturonase alters plantgrowth. Plant Physiol 135(3):1294–1304

99. Oxenboll Sorensen S, Pauly M, Bush M, Skjot M, McCann MC,Borkhardt B et al (2000) Pectin engineering: modification of po-tato pectin by in vivo expression of an endo-1,4-beta-D-galactanase. Proc Natl Acad Sci U S A 97(13):7639–7644

100. Skjøt M, Kauppinen S, Kofod LV, Fuglsang C, Pauly M, DalbøgeH et al (2001) Functional cloning of an endoarabinanase fromAspergillus aculeatus and its heterologous expression in A. ororyzae and tobacco. Mol Gen Genomics 265(5):913–921

101. Oomen RJ, Doeswijk-Voragen CH, Bush MS, Vincken JP,Borkhardt B, van den Broek LAM et al (2002) In muro fragmen-tation of the rhamnogalacturonan I backbone in potato (Solanumtuberosum L.) results in a reduction and altered location of thegalactan and arabinan side-chains and abnormal periderm devel-opment. Plant J 30:403–413

102. Boudart G, Charpentier M, Lafitte C, Martinez Y, Jauneau A,Gaulin E et al (2003) Elic i tor act ivi ty of a fungalendopolygalacturonase in tobacco requires a functional catalyticsite and cell wall localization. Plant Physiol 131:93–101

103. Hasunuma T, Fukusaki E, Kobayashi A (2003) Methanol produc-tion is enhanced by expression of an Aspergillus niger pectinmethylesterase in tobacco cells. J Biotechnol 106:45–52

104. Hasunuma T, Fukusaki E, Kobayashi A (2004) Expression offungal pectin methylesterase in transgenic tobacco leads to alter-ation in cell wall metabolism and a dwarf phenotype. J Biotechnol111(3):241–251

105. Ferrari S, Galletti R, Pontiggia D, Manfredini C, Lionetti V,Bellincampi D et al (2008) Transgenic expressionof a fungalendo-polygalacturonase increases plant resistance to pathogensand reduces auxin sensitivity. Plant Physiol 146(2):669–681

106. Tsuji Y, Vanholme R, Tobimatsu Y, Ishikawa Y, Foster CE,Kamimura N et al (2015) Introduction of chemically labile sub-structures into Arabidopsis lignin through the use of LigD, theCα-dehydrogenase from Sphingobium sp. strain SYK-6. PlantBiotechnol J. doi:10.1111/pbi.12316

107. Eudes A, Sathitsuksanoh N, Baidoo EEK, George A, Liang Y,Yang F et a l (2015) Express ion of a bac te r i a l 3 -dehydroshikimate dehydratase reduces lignin content and

Bioenerg. Res.

improves biomass saccharification efficiency. Plant Biotechnol J.doi:10.1111/pbi.12310

108. Obembe OO, Jacobsen E, Timmers J, Gilbert H, Blake AW, KnoxJP et al (2007) Promiscuous, non-catalytic, tandem carbohydrate-binding modules modulate the cell-wall structure and develop-ment of transgenic tobacco (Nicotiana tabacum) plants. J PlantRes 120(5):605–17

109. Hewezi T, Howe P, Maier TR, Hussey RS, Mitchum MG, DavisEL et al (2008) Cellulose binding protein from the parasitic nem-atode Heterodera schachtii interacts with Arabidopsis pectinmethylesterase: cooperative cell wall modification during parasit-ism. Plant Cell 20(11):3080–3093

110. Safra-Dassa L, Shani Z, Danin A, Roiz L, Shoseyov O, Wolf S(2006) Growth modulation of transgenic potato plants by heterol-ogous expression of bacterial carbohydrate-binding module. MolBreed 17(4):355–364

111. Horn SJ, Vaaje-Kolstad G, Westereng B, Eijsink VG (2012)Novel enzymes for the degradation of cellulose. BiotechnolBiofuels 5:45

112. Fushinobu S (2014) Metalloproteins: A new face for biomassbreakdown. Nat Chem Biol 10:88–99

113. Menkhaus TJ, Bai Y, Zhang Z, Nikolov L, Glatz CE (2004)Considerations for the recovery of recombinant proteins fromplants. Biotechnol Prog 20(4):1001–14

114. Brunecky R, SeligMJ, Vinzant TB, HimmelME, Lee D, BlaylockMJ et al (2011) In planta expression of A. Cellulolyticus Cel5Aendocellulase reduces cell wall recalcitrance in tobacco and maize.Biotechnol Biofuels 4(1):1

115. Klose H, Röder J, Girfoglio M, Fischer R, Commandeur U (2012)Hyperthermophilic endoglucanase for in planta lignocelluloseconversion. Biotechnol Biofuels 5:63

116. Guerriero G, Fugelstad J, Bulone V (2010) What do we reallyknow about cellulose biosynthesis in higher plants? J IntegrPlant Biol 52(2):161–75

117. Harris D, Stork J, DeBolt S (2009) Genetic modification incellulose-synthase reduces crystallinity and improves bio-chemical conversion to fermentable sugar. Glob ChangBiol Bioenergy 1:51–61

118. Shrinivas D, Savitha G, Raviranjan K, Naik GR (2010) A highlythermostable alkaline cellulase- free xylanase fromthermoalkalophilic bacillus sp. JB 99 suitable for paper and pulpindustry: purification and characterization. Appl BiochemBiotechnol 162:2049–57

119. Wu AM, Hornblad E, Voxeur A, Gerber L, Rihouey C, Lerouge Pet al (2010) Analysis of the Arabidopsis IRX9/IRX9-L andIRX14/IRX14-L pairs of glycosyltransferase genes reveals criticalcontributions to biosynthesis of the hemicellulose glucuronoxylan.Plant Physiol 153:542–54

120. Beg QK, Kapoor M, Mahajan L, Hoondal GS (2001) Microbialxylanases and their industrial applications: a review. ApplMicrobiol Biotechnol 56:326–38

121. Subramaniyan S, Prema P (2002) Biotechnology of microbialxylanases: enzymology, molecular biology, and application. CritRev Biotechnol 22:33–64

122. Jabbour D, Borrusch MS, Banerjee G, Walton JD (2013)Enhancement of fermentable sugar yields by alpha-xylosidasesupplementation of commercial cellulases. Biotechnol Biofuels6(1):58

123. Alvira P, Negro MJ, Ballesteros M (2011) Effect of endoxylanaseand a-L-arabinofuranosidase supplementation on the enzymatichydrolysis of steam exploded wheat straw. Bioresour Technol102:4552–8

124. Latha Gandla M, Derba-Maceluch M, Liu X, Gerber L, MasterER, Mellerowicz EJ et al (2014) Expression of a fungalglucuronoyl esterase in populus: effects on wood properties andsaccharification efficiency. Phytochemistry 112:210–20

125. Brummel DA, Harpster MH (2001) Cell wall metabolism in fruitsoftening and quality and its manipulation in transgenic plants.Plant Mol Biol 47:311–40

126. Willats WG, Orfila C, Limberg G, Buchholt HC, van Alebeek GJ,Voragen AG et al (2001) Modulation of the degree and pattern ofmethyl-esterification of pectic homogalacturonan in plant cellwalls. Implications for pectin methyl esterase action, matrix prop-erties, and cell adhesion. J Biol Chem 276(22):19404–13

127. Pelloux J, Rusterucci C, Mellerowicz EJ (2007) New insights intopectin methylesterase structure and function. Trends Plant Sci12(6):267–77

128. Francocci F, Bastianelli E, Lionetti V, Ferrari S, De Lorenzo G,Bellincampi D et al (2013) Analysis of pectin mutants and naturalaccessions of Arabidopsis highlights the impact of de-methyl-esterified homogalacturonan on tissue saccharification.Biotechnol Biofuels 6:163

129. Vincken JP (2003) If homogalacturonan were a side chain ofrhamnogalacturonan I. Implications for cell wall architecture.Plant Physiol 132(4):1781–9

130. Ulskov P,WiumH, Bruce D, Jorgensen B, Qvist KB, SkjøtM et al(2005) Biophysical consequences of remodeling the neutral sidechains of rhamnogalacturonan I in tubers of transgenic potatoes.Planta 230:609–20

131. Van Acker R, Vanholme R, Storme V, Mortimer JC, Dupree P,Boerjan W (2013) Lignin biosynthesis perturbations affect sec-ondary cell wall composition and saccharification yield inArabidopsis thaliana. Biotechnol Biofuels 6:46

132. Vanholme R, Morreel K, Ralph J, Boerjan W (2008) Lignin engi-neering. Curr Opin Plant Biol 11(3):278–85

133. Vanholme R, Morreel K, Darrah C, Oyarce P, Grabber J, Ralph Jet al (2012) Metabolic engineering of novel lignin in biomasscrops. New Phytol 196:978–1000

134. Poovaiah CR, Nageswara-Rao M, Soneji JR, Baxter HL, StewartCN (2014) Altered lignin biosynthesis using biotechnology toimprove lignocellulosic biofuel feedstocks. Plant Biotechnol J12(9):1163–73

135. Vanholme R, Cesarino I, Rataj K, Xiao Y, Sundin L, Goeminne Get al (2013) Caffeoyl shikimate esterase (CSE) is an enzyme in thelignin biosynthetic pathway. Science 341:1103–1106

136. Chen F, Dixon RA (2007) Lignin modification improves ferment-able sugar yields for biofuel production. Nat Biotechnol 25(7):759–61

137. Grabber JH (2005) How do lignin composition, structure, andcross-linking affect degradability? A review of cell wall modelstudies. Crop Sci 45(3):820

138. Ciesielski PN, Resch MG, Hewetson B, Killgore JP, Curtin A,Anderson N et al (2014) Engineering plant cell walls: tuning ligninmonomer composition for deconstructable biofuel feedstocks orresilient biomaterials. Green Chem 16(5):2627–35

139. Li X, Ximenes E, Kim Y, Slininger M, Meilan R, Ladisch M et al(2010) Lignin monomer composition affects arabidopsis cell-walldegradability after liquid hot water pretreatment. BiotechnolBiofuels 3:27

140. Wilkerson CG, Mansfield SD, Lu F, Withers S, Park J-Y,Karlen SD et al (2014) Monolignol ferulate transferase in-troduces chemically labile linkages into the lignin back-bone. Science 344(6179):90–3

141. Cosgrove DJ (2000) Loosening of plant cell walls by expansins.Nature 407(6802):321–6

142. Shoseyov O, Shani Z, Levy I (2006) Carbohydrate binding mod-ules: biochemical properties and novel applications. MicrobiolMol Biol Rev MMBR 70(2):283–95

143. Levy I, Shoseyov O (2002) Cellulose-binding domains: biotech-nological applications. Biotechnol Adv 20:191–213

144. Shani Z, Shpigel E, Roiz L, Goren R, Vinocur B, Tzfira Tet al (1999) Cellulose-binding domain increases cellulose

Bioenerg. Res.

synthase activity in acetobacter xylinum and biomass oftransgenic plants. In: Altman A, Ziv M, Izhar S (eds) Plantbiotechnology and in vitro biology in the 21st century.Kluwer Academic Publishers, Dordrecht, The Netherlands,pp 213–8

145. Shoseyov O, Levy I, Shani Z, Mansfield SD (2003)Modulation ofwood fibers and paper by cellulose-binding domains. In S DMansfield and J N Saddler (eds), Application of enzymes to lig-nocellulosics ACS symposium series 855 American ChemicalSociety, Washington DC, 116–31

146. Cho HT, Cosgrove DJ (2000) Altered expression of expansinmodulates leaf growth and pedicel abscission in Arabidopsisthaliana. Proc Natl Acad Sci U S A 97:9783–8

147. Choi D, Lee Y, Cho HT, Kende H (2003) Regulation of expansingene expression affects growth and development in transgenic riceplants. Plant Cell 15:1386–98

148. Madoka Gray M, Blomquist K, McQueen-Mason SJ, Teeri TT,Sundberg B, Mellerowicz EJ (2008) Ectopic expression of a

wood-abundant expansin PttEXPA1 promotes cell expansionin primary and secondary tissues in aspen. Plant Biotechnol J6(1):62–72

149. Shoseyov O, Shani Z, Abramson M, Barimboim N, Dekel M,Lapidot S (2008) Transgenic plants containing soluble cell wallpolysaccharides. WO 2008120194 A2

150. Preis I, Lapidot S, Abramson M, Shoseyov (2013) Modificationsof cell wall properties by production of recombinant resilin com-posites in transgenic plants. XIII Cell Wall Meeting, Nantes,France

151. Barimbiom-Moshe N (2008) Cloning and characterization oflevansucrase from Erwina amylovora in Eucalyptus, Ph.D. thesis,Hebrew University of Jerusalem

152. Vanholme B, Vanholme R, Turumtay H, Goeminne G, CesarinoI, Goubet F et al (2014) Accumulation of N-acetylglucosamineoligomers in the plant cell wall affects plant architecture in adose-dependent and conditional manner. Plant Physiol 165(1):290–308

Bioenerg. Res.