Fungal Pretreatment of Lignocellulosic Biomass

Biotechnology Advances 30 (2012) 1447–1457

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Research review paper

Fungal pretreatment of lignocellulosic biomass

Caixia Wan, Yebo Li ⁎Department of Food, Agricultural, and Biological Engineering, The Ohio State University/Ohio Agricultural Research and Development Center, 1680 Madison Ave, Wooster,OH 44691-4096, USA

⁎ Corresponding author. Tel.: + 1 330 263 3855; fax:E-mail address: [email protected] (Y. Li).

0734-9750/$ – see front matter © 2012 Elsevier Inc. Alldoi:10.1016/j.biotechadv.2012.03.003

a b s t r a c t
a r t i c l e i n f o
Available online 10 March 2012

Keywords:FungalPretreatmentWhite rot fungiBiodelignificationBiomassBiofuel

Pretreatment is a crucial step in the conversion of lignocellulosic biomass to fermentable sugars and biofuels. Com-pared to thermal/chemical pretreatment, fungal pretreatment reduces the recalcitranceof lignocellulosic biomass bylignin-degrading microorganisms and thus potentially provides an environmentally-friendly and energy-efficientpretreatment technology for biofuel production. This paper provides an overview of the current state of fungal pre-treatment by white rot fungi for biofuel production. The specific topics discussed are: 1) enzymes involved in bio-degradation during the fungal pretreatment; 2) operating parameters governing performance of the fungalpretreatment; 3) the effect of fungal pretreatment on enzymatic hydrolysis and ethanol production; 4) efforts forimproving enzymatic hydrolysis and ethanol production through combinations of fungal pretreatment and physi-cal/chemical pretreatment; 5) the treatment of lignocellulosic biomass with lignin-degrading enzymes isolatedfrom fungal pretreatment, with a comparison to fungal pretreatment; 6) modeling, reactor design, and scale-up ofsolid state fungal pretreatment; and 7) the limitations and future perspective of this technology.

© 2012 Elsevier Inc. All rights reserved.

Contents

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1448Lignin-degrading microorganisms and degrading enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1448

Lignin-degrading microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1448Degrading enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1449

Ligninolytic enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1449Hydrolytic enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1449

Gene expression of ligninolytic enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1450Solid state fungal pretreatment process . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1450

Inoculum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1450Moisture content . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1451Particle size . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1451Supplements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1451Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1451Aeration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1451Decontamination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1451Time . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1452

Enzymatic hydrolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1452Non-selective lignin degrading fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1452Selective lignin degrading fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1452

Combination of fungal pretreatment and physical/chemical pretreatments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1453Enzymatic treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1454Modeling and scale-up . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1454Limitations and potentials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1455

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1448 C. Wan, Y. Li / Biotechnology Advances 30 (2012) 1447–1457

Conclusions and future perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1455Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1455References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1455

Introduction

In order to reduce dependence on fossil fuels and thus alleviateassociated economic and environmental concerns, biofuels derivedfrom renewable and domestic sources have received extensive interestfor displacement of fossil transportation fuels in many countries.Lignocellulosic biomass, mostly from agricultural and forestry sources,is rich in carbohydrates (55–75% dry basis) and widely available, thusproviding attractive feedstocks for ethanol production. To maximallyutilize carbohydrates in the biomass, a pretreatment process is neededto overcome the biomass recalcitrance and subsequently improve itsaccessibility to hydrolytic enzymes (Mosier et al., 2005). Thermal/chemical pretreatment methods have been regarded as the currentleading pretreatment technologies; however, they usually need expen-sive corrosion resistant reactors, processing large volumes of the wastestream, extensive washing of treated solids, and detoxification of com-pounds inhibitory to ethanol-fermenting microorganisms. Thus, pre-treatment still remains one of the most costly steps in cellulosicethanol production and is a significant barrier to its commercialization(Mosier et al., 2005).

From both economic and environmental perspectives, fungal pre-treatment with lignin-degrading microorganisms, preliminary whiterot fungi, has received renewed interest as an alternative to ther-mal/chemical pretreatment for cellulosic ethanol production. The feasi-bility of fungal pretreatment for improving enzymatic digestibility ofvarious biomass feedstocks, such as corn stover (Keller et al., 2003; Xuet al., 2010), wheat straw (Dias et al., 2010), rice straw (Bak et al.,2009), cotton stalks (Shi et al., 2009), and woody biomass (Yu et al.,2009a), has been reported. Moreover, most researchers are interestedin solid state fungal pretreatment as limitations of liquid state cultiva-tion are apparent, mainly due to a considerably low substrate loading(b5%). The advantages of this technology over thermo-chemical pre-treatments include simple techniques, low energy requirements, no or

Table 1The effect of fungal pretreatment on enzymatic hydrolysis and ethanol production.

Fungus Substrate Su

Phaerochaete chrysosporium Cotton stalk RePhaerochaete chrysosporium Corn stover NPhaerochaete chrysosporium Rice straw 50

50Phaerochaete chrysosporium Beechwood 9.Phaerochaete chrysosporium Corn fiber RePleurotus ostreatus Rice straw 33Pleurotus ostreatus Rice hull 38Pleurotus ostreatus, Pycnoporus cinnabarinus 115 Wheat straw 27Euc-1 Wheat straw 22Cyathus stercoeus Corn stover 36Irpex lactues Corn stover 66Ceriporiopsis subvermispora Corn stover 56Pheblia tremellosus Aspenwood 11Polyporus giganteus Aspenwood 55Stereum hirsutum Japanese red pine 13Echinodontium taxodii 2538 Chinese willow, China-fir 5–Echinodontium taxodii 2538, Coriolus versicolor Bamboo culm 37

a % of theoretical yield of glucan in the original material, unless stated otherwise.b Results were obtained from submerged fungal pretreatment.c % of dry mass of the treated material.d % of dry mass of the original material.e % of theoretical yield of holocellulose in the original material.f % of theoretical yield of holocellulose in the treated material.

reduced output of waste streams, reduced downstream processingcosts, and no or reduced inhibitors to ethanol fermentation (Keller etal., 2003; Nigamand Pandey, 2009). Despite the advantages, substantialholocellulose (cellulose and hemicellulose) loss and long pretreatmenttime are the major issues associated with fungal pretreatment. Toensure a cellulose-rich but highly delignified biomass for biofuel pro-duction, white rot fungi, highly selective in lignin degradation, are pre-ferred for fungal pretreatment. In addition, cultivation parameters alsoaffect the pretreatment performance. Scale-up related issues likedecontamination and reactor design also need to be addressed for in-dustrial applications.

This review provides an overview of recent studies on solid statefungal pretreatment with white rot fungi for biofuels production, fo-cusing on critical pretreatment parameters affecting the effectivenessof fungal pretreatment, enzymes involved in the degradation of bio-mass feedstocks, and sugar and ethanol yields resulting from fungalpretreatment. An overall comparison of the effectiveness of fungaland enzymatic pretreatments, synergistic effects of combined fungaland physical/chemical pretreatment, and feasibility and limitationsof scale-up of fungal pretreatment for biofuels production are alsodiscussed.

Lignin-degrading microorganisms and degrading enzymes

Lignin-degrading microorganisms

Microorganisms, including white-, brown-, and soft-rot fungi, andsome ruminant bacteria, are capable of degrading lignocellulosic bio-mass. Among them, white rot fungi are most effective for delignifica-tion due to their unique ligninolytic systems (Eriksson et al., 1990).Traditionally, white rot fungi have been employed for biopulping, for-age upgrading, and bioremediation of soil and wastewater by oxidizinglignin and awide range of lignin analogous compounds (Sanchez, 2009;

gar/ethanol yielda References

duced glucose yield Shi et al. (2009)o improvement on glucose yield Keller et al. (2003)% glucose yield b

% ethanol yield bBak et al. (2009)

5% total sugar yield c Sawada et al. (1995)duced sugar yield or no significant improvement Shrestha et al. (2008)% glucose yield Taniguchi et al. (2005).9% glucose yield Yu et al. (2009b)–28% glucose yield d Hatakka (1983).5% total sugar yield c Dias et al. (2010)% glucose yield Keller et al. (2003).4% total sugar yield e Xu et al. (2010)–66% glucose yield and 57.8% ethanol yield Wan and Li (2010b).6% glucose yieldc Mes-Hartree et al. (1987).2% glucose yield Kirk and Moore (1972).7% glucose yieldc Lee et al. (2007)35% glucose yield Yu et al. (2009a)% total sugar yield f Zhang et al. (2007a, 2007b)

1449C. Wan, Y. Li / Biotechnology Advances 30 (2012) 1447–1457

Wesenberg et al., 2003;Winquist et al., 2008). As listed in Table 1, sev-eral white rot fungi, such as Phanerochaete chrysosporium, Pleurotusostreatus, Coriolus versicolor, Cyathus stercoreus, and Ceriporiopsis sub-vermispora, have been studied for pretreatment of a wide range ofbiomass feedstocks. Although delignification is of significance to re-duce the biomass recalcitrance, white rot fungi with a high selectiv-ity for lignin degradation over cellulose loss, as aforementioned anddiscussed in the later sections, aremost important to fungal pretreat-ment for biofuel production.

Analysis of white rot-degraded wood indicates that lignin degra-dation by these microorganisms is highly oxidative and may involvechemical oxidants such as singlet oxygen and hydroxyl radicals(Eriksson et al., 1990). The following reactions are generally involved:(i) oxidative, (ii) demethylation (or demethoxylation), (iii) side-chain oxidation, and (iv) propyl side-chain cleavage. In contrast towhite rot fungi, brown rot fungi are able to circumvent the ligninbarrier, remove the hemicellulose and cellulose with only minormodifications to the lignin (Eriksson et al., 1990). Consequently, ligninremains a major component of the degraded plant cell wall. Theremaining lignin is demethylated on arylmethoxy groups and containsa greater number of ring hydroxyl groups.

Degrading enzymes

Ligninolytic enzymesIt is well known that lignin peroxidase (LiP), manganese peroxidase

(MnP), and laccase are the three major oxidative enzymes secreted bywhite rot fungi (Eriksson et al., 1990). They are responsible for oxida-tion of lignin and a wide range of lignin analogous compounds(Winquist et al., 2008). However, not all of these enzymes are detectedfrom fungal cultures. For example, P. chrysoporium produced LiP andMnP, but no laccase (Ruttimann-Johnson et al., 1993). C. subvermisporaonly producedMnP and laccasewhereas LiPwas not detected, althougha lip-like genewas revealed in this fungus by a southern-blot hybridiza-tion technique using the same probe coding as lip (Rajakumar et al.,1996). Lignin peroxidase (also “ligninase”, EC 1.11.1.14) is a hemepro-tein, involving in the oxidative cleavage of non-phenolic aromaticlignin moieties and similar compounds. Manganese peroxidase (EC1.11.1.13) is an enzyme secreted to aid lignin degradation, catalyzingthe chemical reaction that oxidizes numerous phenolic compounds,especially syringyl (3, 5-dimethoxy-4-hydroxyphenyl) and vinylside-chain substituted substrates in the presence of Mn2+. Laccases(EC 1.10.3.2) are copper-containing oxidase enzymes that act onphenols and similar molecules, performing one-electron oxidations.Ligninolytic enzymes may be non-specific on different lignin sub-strates and have similar degradation reactions. For example, bothLiP and MnP were found to degrade non-phenolic lignin by one-electron oxidation of the aromatic ring (Kirk et al., 1986; Srebotniket al., 1997). Versatile peroxidase (VP) isolated from Pleurotus andBjerkandera species is regarded as the third peroxidase, a LiP-MnPhybrid as it showsMn2+ independent activity but is capable of degrad-ing phenolic and non-phenolic compounds (Martinez et al., 1996;Mester and Field, 1998). However, due to the complicated and hetero-geneous nature of lignocellulosic biomass, the role of ligninolytic en-zymes on the early stage decay of biomass feedstocks still remainsunclear. A study conducted by Guerra et al. (2002) showed no evidencefor a correlation between oxidative enzymes and lignin degradation byC. subvermispora. Moreover, the mineralization of lignin occurred afterthe lignin modification, which was indicative of a series of reactions in-volved in lignin depolymerization.

Studies with immunocytochemical techniques revealed that lignin-degrading enzymes did not diffuse into sound and unaltered cellwalls due to their large molecular weight (Blanchette et al., 1997;Flournoy et al., 1993; Srebotnik et al., 1988). However, the oxidativeenzymes could be active at the surface of the cell wall and induce theformation of low molecular mass agents such as radicals and oxalic

acids (Enoki et al., 1999; Kapich et al., 1999). These lowmolecular com-pounds are diffusible and could initiate wood decay and facilitate thepenetration of lignin-degrading enzymes (Galkin et al., 1998). For in-stance, C. subvermispora produces a variety of low molecular com-pounds including glyoxylic acids (Urzua et al., 1998), oxalic acid(Galkin et al., 1998; Urzua et al., 1998), and several unsaturated fattyacids (Enoki et al., 1999; Gutierrez et al., 2002) in liquid or solid stateculture. Radical agents (e.g., peroxyl and acyl radicals), generatedfrom MnP-dependent peroxidation of white rot fungi, have been sug-gested as playing an important role in non-phenolic lignin oxidation(Kapich et al., 1999; Watanabe et al., 2000).

Hydrolytic enzymesHydrolytic enzymes play important roles by providing easily

digestible carbon sources to fungal growth and metabolism. However,non-selectivewhite rot fungi, due to their high cellulolytic and hemicel-lulolytic activity, cause substantial cellulose loss. In contrast, selectivewhite rot fungi mainly secrete hemicellulolytic enzymes and utilizehemicellulose-derived sugars as the main carbon sources for self-growth and metabolism. Cellulase-deficient mutants developed fromfast-growing but non-selective white rot fungi, such as P. chrysosporium,were also evaluated for lignin degradation. Although some P. chrysospor-ium mutants can act effectively in lignin and lignin model compounds(Eriksson et al., 1983), no degradation on lignocellulosic biomass bythemutantswas ever reported relative towild species (Akin et al., 1993).

Due to the low permeability of sound wood cells, it is difficult forsuch large molecule hydrolytic enzymes to penetrate. Similar to ligni-nolytic enzymes, low molecular mass agents have also been sug-gested as a mechanism to induce cellulose depolymerization. TheFenton system (Fe2+ and H2O2), known to depolymerize cellulose,can be generated by the action of some enzymes such as cellobiohy-dranase and MnP in white rot fungal cultures (Dumonceaux et al.,2001; Henriksson et al., 2000; Xu et al., 2009). In contrast to cellulaseand peroxidase, hemicellulases, such as xylanase, did not act in thesynergism of low molecular compounds (Ferraz et al., 2003). There-fore, diffusion of xylanase into the plant cell wall is suggested to be fa-cilitated by lignin degradation which increases cell wall permeability(Machuca and Ferraz, 2001; Vicentim and Ferraz, 2007).

Due to the lack of a complete cellulolytic enzyme complex,C. subvermispora has become well known as a selective white rotfungus that preserves most cellulose during fungal decay (Ferraz et al.,2003). In the study of Ferraz et al. (2003), no significant glucoseloss was detected from C. subvermispora culture on the wood speciesEucalyptus grandis until 60 d. Maximum glucan loss of 7.3% wasobserved after 90 d. Low cellulase activity revealed by filter paperactivity explained the low glucan loss caused by this fungus. Anotherwood species, Pinus taeda, cultured with this fungus was observedwithonly 2% glucan loss after 90 d (Guerra et al., 2003). During 42-d fungalpretreatment of corn stover by this fungus, there were no detectablefilter paper and carboxymethylcellulose activities and the celluloseloss was less than 5% (Wan and Li, 2010a). However, cellulose degrada-tion up to 26%was observed in Pinus radiate during 200-d pretreatmentby C. subvermispora, probably due to slowly induced cellulolytic activity(Ferraz et al., 2001).

Unlike C. subvermispora, P. ostreatus is a widely studied white rotfungus that is able to produce a hydrolytic enzyme complex in differentlignocellulosic biomass or under different fermentation strategies(Elisashvili et al., 2008; Sánchez, 2009). Several studies have focusedon hydrolytic enzyme production by this fungus and detected multiplecellulolytic enzymes, including cellulases, endoglucanase, and cellobio-hydrolase, from its culture on agricultural residues (Elisashvili et al.,2008; Garzillo et al., 1994; Marnyye et al., 2002; Okamoto et al.,2002). Similar results were also reported for other Pleutorus strains(Elisashvili et al., 2008).With high hydrolytic and ligninolytic activities,P. ostreatus has been widely used for upgrading crop residues, such aswheat straw, for animal feed (Cohen et al., 2002a). However, as


discussed in the enzymatic hydrolysis section, based on the effective-ness of fungal pretreatments, P. ostreatus is regarded as a moderately se-lective lignin degrader, due to its substantial consumption of cellulose inlignocellulosic substrates, especially with prolonged pretreatment time.

Gene expression of ligninolytic enzymes

Numerous studies have demonstrated that expression of white rotfungi genes encoding ligninolytic enzymes is differentially regulatedat the transcriptional level by the culture conditions as reviewed inthe solid state fungal pretreatment process section. Expression ofLiP isoenzyme genes of P. chrysosporium is strongly influenced by ni-trogen or carbon limitation. Holzbaur and Tien (1988) reported thatlipA transcripts of P. chrysosporium were relatively abundant innitrogen-starved media while lipD transcripts were dominant incarbon-starved cultures. Both lipC and lipJ gene transcripts were dra-matically upregulated under nitrogen-deficient conditions (Stewartet al., 1992). In contrast, lipE transcript levels were not affected by ei-ther carbon or nitrogen deficiency in the culture media (Reiser et al.,1993). Culture substrates such as soil and wood also largely affecttranscript patterns and levels. For example, lipF transcripts wereabsent in soil culture but were abundant in aspen culture (Bogan etal., 1996; Janse et al., 1998). lipD and lipE transcripts showed almostthe opposite expression, being highly expressed in soil but not inaspen wood. Mn2+ also had an indirect effect on LiP gene expressionof white rot fungi Bjerkandera sp. Strain BOS 55 and P.chrysosporiumBKM-F-1767 by its regulation of veratryl alcohol levels in culturemedia (Mester et al., 1995). Relative to LiP gene expression, morecomplex regulation by culture conditions, such as nutrient limitation,Mn2+, H2O2 concentration, and other physiological factors (e.g. tem-perature, agitation, moisture), has been observed with MnP gene ex-pression (Janse et al., 1998).mnp1 andmnp2 of P. chrysosporiumwerestrongly dependent on Mn2+ and also differentially regulated by cul-ture agitation while mnp3 had no response to Mn2+ (Gettemy et al.,1998). Similarly, mnp2 isolated from P. ostreatus grown on saw dustappeared not to be regulated by Mn2+ (Giardina et al., 2000). Mn2+

amendment also affected MnP and VP expression of P. ostreatus. Unlikemultiple lip genes, all three mnp genes were detected in both woodand soil cultures (Janse et al., 1998). Laccase encoding genes were alsostudied for constitutive expression and inducible expression. Mansuret al. (1998) studied expression patterns of the laccase gene familyfrom Basidomycete I-62 (CECT 20197) and reported that lcc1 and lcc2were induced by veratryl alcohol at different stages of growth whilelcc3 was non-induced by veratryl alcohol. RT-PCR demonstratedthat lcc transcription in Trametes versicolor was activated by copper,nitrogen and certain aromatic compounds tested (i.e. c2,5-xylidineand 1-hydroxybenzotriazole) (Collins and Dobson, 1997).

Several regulatory elements in promoter regions of genes encod-ing ligninolytic enzymes have been postulated to be responsiblefor transcriptional activation (Collins and Dobson, 1997; Have andTeunissen, 2001). Putative metal response elements (MREs) havebeen identified as target sequence for transcription factors respond-ing to heavy metals (e.g. Mn2+, Cu2+, Ag 2+). High dependence ofexpression of certain mnp genes on Mn2+ reveals MREs regulationat the transcription level (Cohen et al., 2001, 2002b). Activation pro-tein of cup1 expression (ACE) elements provide specific binding sitesfor the ACE1-like transcription factor which activates transcriptionof target genes especially in response to copper (Manubens et al.,2007). As P. chrysosporium does not produce laccase, ACE1 identifiedin this fungus activated expression of mco1 gene encoding a differenttype of multicopper oxidase (Canessa et al., 2008). Alvarez et al.(2009) first isolated and characterized an ACE1-like copper-fist tran-scription factor fromgenes encoding laccase andMnP in basidiomycetes.Receptors or binding proteins of putative xenobiotic responsive ele-ments (XREs) are associated with transcription activation of genes inresponse to nonpolar carbon compounds (Collins and Dobson, 1997).

Putative XREs have been identified in the promoter of lip genes andlcc genes in a number of white rot fungi (e.g., T. versicolor, P. ostreatus),which explains inducible expression of these genes by aromatic com-pounds. Activator protein-2-binding sequence identified in upstreamregulatory region of lip and mnp genes of white rot fungi has beenproposed to be involved in differential regulation of these gene familiesbynitrogen (Dhawale, 1993). cAMP response elements (CRE) could alsoplay a role in inducing gene expression during carbon or nitrogen limi-tation (Have and Teunissen, 2001). Other regulatory elements identi-fied, such as cis-regulatory elements, are proposed to respond tophysiological factors, in this case to heat shock. More research isongoing to provide insight into the mechanism involved in differentialregulation of gene expression at transcription levels.

Most studies reported that transcription levels are collinear to en-zyme activity profiles in the culture media (Cohen et al., 2001; Collinsand Dobson, 1997). However, it should be noted that some genesencoding ligninolytic enzymes have not yet been cloned or needsome posttranslational modification to be active (Morgenstern etal., 2010). Unidentified factors hindering free secretion of expressedgenes to the surrounding media could also cause the discrepancy be-tween enzymatic activity and transcription levels. Not readily de-monstrable enzyme activities have also been observed for white rotfungi with putative ligninolytic genes. For example, Phanerochaetesordida and C. subvermispora were detected with lip-like genes butnot Lip activities (Rajakumar et al., 1996). It is also widely quotedthat P. chrysosporium lacks laccase even with putative lcc genes.However, Srinivasan et al. (1995) first reported that a low level lac-case was produced by P. chrysosporium BKM-F1767 in a defined cul-ture medium containing cellulose (10 g/l) and either 2.4 or 24 mMammonium tartrate.

Heterologous expression of active lignin and Mn peroxidases wasstudied with isolation of cDNA encoding many peroxidases isoen-zymes. Baculovirus systems have been successfully used for expres-sion of MnP isozymes H4 (Pease et al., 1991), LiP isozymes H2(Johnson et al., 1992), and LiP H8 of P. chrysosporium H2 (Johnsonet al., 1992). lcc1 of T. versicolor was also expressed by Pichia pastoris(Jönsson et al., 1997). Although properties and activity of these heter-ologously expressed genes are similar to that of native enzymes,yields obtained are mostly too low to be feasible for large scale pro-duction or convenient for biochemical characterization. Fermenta-tion strategies have been reported to markedly enhance expressionlevels (Hong et al., 2002).

Solid state fungal pretreatment process

Inoculum

Inoculum for solid state fungal pretreatment can be prepared by dif-ferent methods, e.g., mycelium grown in liquid or agar medium, spawngrown in cereal grains, or fungal-precolonized substrate (Reid, 1989b).Phanerochaete chrysosporium yields spores which enable convenientinoculum preparation and mixing with the substrate. In contrast,most white rot basidiomycetes do not produce spores. Instead, theprecolonized lignocellulosic biomass is generally used for inocula-tion. Similar to liquid fermentation, fermented materials in solidstate reactors can also serve as inoculum and the fresh substrate canbe fed to partially replace fermented materials. A minimum level ofinoculum is generally required for effective colonization and subse-quent delignification (Reid, 1989a). However, a further increase in inoc-ulum level may only have a marginal effect on the fungal colonizationrate and subsequent growth. Akhtar et al. (1998) tested different levelsof inoculum using precolonized wood chips by P. chrysosporium for thedecay of aspen wood as alternatives to mechanical pulping. The resultsshowed that a 2–5% inoculation level gave good performance and ener-gy savings while increasing the inoculation level to 20% did not corre-spondingly increase energy savings for mechanical pulping.


Moisture content

Initial moisture content of the substrate is important to the fungalestablishment and growth and affects secondary metabolism infungal pretreatment (Reid, 1989b). Previous studies suggested thatinitial moisture ranging from 70 to 80% was the optimal level for lig-nin degradation and ligninolytic activities of most white rot fungi.Shi et al. (2008) observed that after 14-d cultivation of cotton stalksby P. chrysosporium, lignin degradation of 27.6% was obtained at 75%moisture content in the substrate, which was approximately 7%higher than that at 65% moisture content but was not significantlydifferent from that at 80% moisture content. Asgher et al. (2006) in-vestigated the solid state cultivation of P. chrysosporium on corn cobswith a moisture content ranging from 40% to 90%. The highest ligni-nase activity was obtained at 70% moisture content. During fungalpretreatment of corn stover with C. subvermispora, no fungal growthand degradation occurred at 45% moisture content while lignin deg-radation reached 19.48%, 29.54%, and 31.33% when the moisturecontents increased to 60%, 75%, and 85%, respectively (Wan andLi, 2010b). In general, high moisture content is favorable forformation of fungal mycelia but not necessarily for increased deligni-fication (Zadrazil and Brunnert, 1981). Another side effect ofhigh moisture content is a reduced solid loading for fungal pretreat-ment. On the other hand, too low a moisture content could hamperfungal delignification without providing sufficient water to fungalgrowth.

Particle size

Particle size of the substrate is also a major factor affecting theperformance of solid state fungal pretreatment. Large particle sizecan hamper the penetration of fungi into cellulosic biomass and alsoprevent the diffusion of air, water, and metabolite intermediatesinto the particles. However, the reduced particle size with a decreasedsize of interparticle channel may adversely affect interparticle gas cir-culation (Reid, 1989a; Zadrazil and Puniya, 1995), thus not necessar-ily giving an enhanced delignification rate. Reid (1989a) found thatduring the cultivation of Phlebia tremellosa with active aeration, 40-mesh aspen wood had similar lignin degradation and subsequent en-zymatic digestibility to 10-mesh aspen, while the least effectivenesswas observed with aspen chips. Membrillo et al. (2008) also reportedthe highest level of lignocellulolytic enzyme production on sugar canebagasse at 2.9 mm particle size among the three particle sizes tested(0.92, 1.68, and 2.9 mm). Sarikaya and Ladisch (1999) reported that afine particle size of less than 0.42 mm caused less lignin degradationcompared to 0.42–0.82 mm during 32-d treatment of rapeseed with P.ostreatus. Substantially lower lignin degradation (12.83–24.90%) wasobtained with 15 mm corn stover compared to that with particle sizesof 5 and 10 mm (19.48–31.59%) while there was no significant differ-ence between the lignin degradation of 5 and 10 mm corn stover(Wan and Li, 2010b).

Supplements

Supplementation of inducers, such as Mn2+, H2O2, and aromaticcompounds, can potentially stimulate secretion of ligninolytic en-zymes and lignin degradation on biomass feedstock. Shrestha et al.(2008) tested effects of Mn2+, H2O2, and veratryl alcohol on lignindegradation of corn fiber by P. chrysosporium and found Mn2+ addi-tion resulted in the highest lignin degradation. Some studies, howev-er, showed the addition of Mn2+ did not improve lignin degradationon cotton stalk by P. chrysosporium (Shi et al., 2008). The addition ofnutrients generally increased formation of the fungal biomass andalso facilitated fungal colonization in the deeper areas of feedstocks(Messner et al., 1998). As a low cost carbon/nitrogen source, cornsteep liquor has been used for biopulping to increase fungal biomass

production and reduce energy consumption of mechanical pulping(Messner et al., 1998). For most white rot fungi, the depolymerizationof lignin is induced under nitrogen starvation (Akhtar et al., 1998;Ruttimann-Johnson et al., 1993). In other words, addition of supple-mental nitrogen may inhibit lignin degradation while stimulatinggrowth of white rot fungi and consumption of carbohydrates (Reid,1985).

Temperature

In general, white rot ascodiomycetes grow well around 39 °Cwhile white rot basidiomycetes can grow between 15 and 35 °C andtheir high delignification rate is generally obtained within an optimaltemperature range between 25 and 30 °C (Reid, 1985; Reid, 1989b).Wan and Li (2010b) tested a wide range of temperature (4–37 °C)for its effect on pretreatment of corn stover with C. subvermispora in250-ml flasks and the results showed that 28 °C was the optimal tem-perature for improving enzymatic digestibility of corn stover. The me-tabolism of white rot fungi generates heat and causes temperaturegradients in solid state cultivation. The accumulated heat can kill orinhibit the fungal growth and metabolism. Therefore, in the scale-up of solid state cultivation, heat dissipation is one of the key factorsto be taken into account in bioreactor design.

Aeration

Aeration is known to markedly affect the performance of solidstate fermentation. Since lignin degradation is an oxidative process,oxygen availability is important for ligninolytic enzyme activity ofwhite rot fungi. In flask reactors (b500 ml), passive air diffusionthrough cotton plugs was as sufficient as forced air circulation for thedelignification process (Reid, 1989a). However, for reactors containingpacked feedstock, active aeration was necessary to provide uniformair diffusion throughout the substrate. Messner et al. (1998) tested airaeration (0.001, 0.022, 0.1 vv−1 min−1) for P. chrysosporium treatmentof aspen chips for biopulping and concluded that the flow rate of0.022 vv−1 min−1 was enough to achieve good fungal performance.Hatakka (1983) reported that flushing with oxygen 3 times per weekshortened pretreatment time by approximately 1 week during pre-treatment of wheat straw by the fungi Pycnoporus cinnabarinus 115and Phanerochaete sorodida 37. As reviewed by Reid (1989a), high oxy-gen could increase the delignification rate but it did not increasedelignification selectivity. Thus, aeration needs to be controlled toensure the performance of fungal pretreatment.

Decontamination

Decontamination of feedstocks (e.g., gas, steam, chemicals) caneffectively kill or inhibit indigenous microorganisms in the feedstockand is generally required prior to fungal pretreatment, especiallywith white rot basidiomycetes. P. chrysosporium was found to beefficiently competitive against fungal and bacterial infection whileC. subvermispora was more vulnerable to contaminates (SrebotnikandMessner, 1994). Akin et al. (1995) tested the influence of contamina-tion on activity of two white rot fungi (C. subvermispora andC. steroreus). The results showed that abundant bacteria and unknownfungi were prevalent on both control and contaminated Bermudagrass stems. However, contamination did not affect either fungal per-formance or the resulting digestibility of pretreated materials. It indi-cated that activities of white rot fungi were not suppressed by fungaland bacterial infection. However, in scaled up applications, decon-tamination poses one of the major costs for fungal pretreatment. Inthe study of Akhtar et al. (1998), it was found that complete steriliza-tion was not necessary and short atmospheric steaming (~15 s) wassufficient to allow the establishment of white rot fungi and to outcom-pete indigenous fungi and bacteria. Instead of decontamination by


atmospheric steam or autoclave, inexpensive chemicals such as sodiumbiosulfite, sodium meta-bisulfite, and sodium hydrosulfite were alsoreported to be effective at concentrations of 600–1200 ppm for decon-tamination of wood chips for their pretreatment by the biopulpingfungi such as P. chrysosporium and C. subvermispora (Akhtar et al., 1998).

Time

Long pretreatment time, due to low delignification rates, is oneof the major barriers to large scale application of fungal pretreat-ment. Generally, several weeks to months are needed to obtain a highdegree of lignin degradation. P. chrysosporium is a fast-growing whiterot fungus, only requiring a fewdays toweeks for vigorous degradation;however, it degrades lignin and holocellulose unselectively, which isundesirable for biofuel production (Keller et al., 2003; Sawada et al.,1995; Shi et al., 2009). In contrast, for P. ostreatus, one of the widelystudied white rot fungi, a few weeks only resulted in 10.3% lignindegradation in beechwood (Itoh et al., 2003). After 72 d of cultiva-tion with P. ostreatus, 32% of the carbohydrates in rice straw wereconverted to soluble sugars (Taniguchi et al., 2005). Locci et al.(2008) observed that substantial degradation in wheat bran oc-curred during 62 d of fungal treatment by P. ostreatus. For pretreat-ment time as long as 90 d, 51% lignin was degraded from wheatstraw by this fungus (Lindenfelser et al., 1979). For the feedstocks re-sistant to fungal attack, such as softwood and bamboo, even longerpretreatment time is needed. Zhang et al. (2007a, 2007b) reportedthat 60–120 d was required for pretreatment of bamboo with Echino-dontium taxodii and C. versicolor to obtain significant improvement onsugar yields.

Enzymatic hydrolysis

Non-selective lignin degrading fungi

As summarized in Table 1, cellulose digestibility of different biomassfeedstocks resulting from fungal pretreatment has been reported in thepast. P. chrysosporium, a well-known non-selective lignin-degradingfungus, had little or no effect on improvement on enzymatic hydrolysis.The fungus itself consumes a large amount of readily accessible carbo-hydrates due to the simultaneous degradation of cellulose, hemicellu-lose, and lignin. The remaining cellulose might be less digestible andthus resistant to the subsequent hydrolysis. Moreover, the reduced cel-lulose content after fungal pretreatment also contributes to the reducedglucose yield. The longer time the biomass feedstock is pretreated byP. chrysosporium, the lower the expected saccharification yield com-pared to that from the non-treated feedstock. This expectation wasobserved during a 100-d pretreatment of aspen wood by this fungus,where the maximum saccharification yield was reached on day 28and thereafter the saccharification yield decreased (Sawada et al.,1995). The lignin degradation was increased from 42% to 50% as pre-treatment time increased from 28 to 100 d while the correspondingholocellulose loss was increased from 17% to 50%. Although the degra-dation selectivity was high on day 28, the fungal pretreatment wasnot sufficient to increase the saccharification yield of aspen wood.Pretreatment time beyond 2 weeks also resulted in a reduced sacchari-fication yield of corn fiber compared to that of the control (Shresthaet al., 2008). Keller et al. (2003) reported that pretreatment of cornstover with P. chrysosporium for 29 d did not significantly increase sac-charification yield compared to the control, probably due to too long apretreatment time. Even for a 14-d fungal pretreatment of cotton stalkby P. chrysosporium, a reduced saccharification yield was obtainedwith both submerged and solid state fungal-treated cotton stalks (Shiet al., 2009). The cellulose loss from both methods was as high as40.49%, while lignin degradation was in the range of 19.38–35.53%.Hot water washing, for removal of fungal biomass/protein and ligninderivatives, improved the saccharification yield of solid state treated

stalks but not the submerged-treated stalks. Nevertheless, the sacchar-ification yield was not significantly different from that of non-treated.

There was an exception for P.chrysosporiumwith respect to the de-gree to which saccharification yield was improved. Bak et al. (2009)serially optimized the culture media for submerged pretreatment ofrice straw with P. chrysosporium in order to achieve high ligninolyticenzymes. It was found that glucose yield was improved by at least 2times during a 30-d pretreatment, reaching a maximum glucoseyield of 64.9% (equivalent to 50% of glucose yield of the raw feed-stock) when cellulase and β-glucosidase (Novozyme 188) loadingswere 60 FPU and 30 CBU per gram glucan, respectively. Similar im-provement was also reported for ethanol yield from their study. How-ever, taking into consideration both considerable cellulose loss andlow solid loading of submerged cultivation, such non-selective fungusmay not be effective for fungal pretreatment to improve the sacchar-ification yield of biomass feedstocks.

Selective lignin degrading fungi

Selective lignin-degrading white rot fungi degrade larger amountsof lignin relative to cellulose. However, fungal selectivity variesamong species and with pretreatment time. In addition, fungal per-formance on degradation and the resulting digestibility varies withdifferent feedstocks. For agricultural residue, Keller et al. (2003)reported that a saccharification yield of 36% was obtained from cornstover pretreated with C. stercoreus for 29 d when the enzymatichydrolysis was conducted at 60 FPU/g glucan for 136 h, which wasabout 4 times that of the untreated. In contrast, the saccharificationyield of 66.4% was obtained with 25-d Irpex lacteus CD2 treated cornstover at a cellulase loading of 20 FPU/g solid for enzymatic hydroly-sis (Xu et al., 2010). In their study, the saccharification yield offungal-treated corn stover decreased slightly during 25–60 d of pre-treatment and then decreased dramatically due to holocellulose loss.It was found that holocellulose degradation, especially hemicellulosedegradation, was dominant during the early stage (0–5 d) while nolignin degradation was observed during this period. Thereafter,active lignin degradation was observed on days 5 to 10 with a ratehigher than that of holocellulose degradation. In contrast, glucoseyield as high as 67% was obtained from corn stover pretreated withC. subvermispora as a result of 35.81% lignin degradation when cellu-lase loading for enzymatic hydrolysis was 10 FPU/g solid (Wan andLi, 2010b). These studies indicated that delignification of corn stoverwas fungus-specific. P. ostreatus was reported to be more effectivewith straw materials than other fungi (Taniguchi et al., 2005). Wheatstraw pretreated with this fungus gave 27–33% cellulose digestibilityduring 72-h enzymatic hydrolysis with cellulase loading of about10 FPU/g solid (Hatakka, 1983; Taniguchi et al., 2005). It should benoted that this fungus was less selective in lignin and cellulose deg-radation when pretreatment time was extended to several weeks.Thus, the cellulose digestibility tended to level off during the laterstage of cultivation (Taniguchi et al., 2005; Yu et al., 2009b). Fungalpretreatment also greatly improved the digestibility of hardwoodssuch as aspen and birch but with a longer pretreatment time. A desir-able polysaccharide digestibility of aspenwood (50–55%) was obtainedafter pretreatment with Polyporus giganteus, Polyporus berkeleyi, orPolyporus resinosus for 63–99 d while the digestibility of birch woodwas not substantially improved with these fungi (Kirk and Moore,1972). For Chinese willow after pretreatment with E. taxodii 2538 for120 d, about 37% of the polysaccharides were converted when enzy-matic hydrolysis was conducted at a cellulase loading of 20 FPU/gsolid. Similar saccharification yieldwas also observedwith bamboo res-idues under the same pretreatment and enzymatic hydrolysis condi-tions (Zhang et al., 2007b). Compared to other types of feedstocks,softwood generally required a longer pretreatment but its resultingsaccharification rate appeared to be unattractive. Yu et al. (2009a)reported that the saccharification yield of China fir was 17% after


pretreatment with E. taxodii for 120 d. Lee et al. (2007) also reporteda 21% sugar yield from Japanese red pine pretreated with Stereumhirsutum for 8 weeks, of which 14% was glucose. As a result, it is sug-gested that thermo-chemical pretreatment be used rather than fun-gal pretreatment for softwood.

Combination of fungal pretreatment andphysical/chemical pretreatments

Fungal pretreatment of wood with white rote fungi has beenwidely applied in the pulp industry. This so-called biopulping processcan potentially overcome the problems associated with mechanicaland chemical pulping methods (e.g., intensive energy input for me-chanical pulping, high chemical loadings for chemical pulping). Theadvantages include significant energy savings in mechanical pulping,low severity for subsequent chemical pulping, improved paperstrength properties, and reduced toxicity of pulping waste (Akhtaret al., 1998; Kang et al., 2003). Messner et al. (1998) observed that arelatively short incubation time substantially favored the chemicalpulping process due to appreciable pre-modification of lignin. Similarto benefits of biopulping, fungal pretreatment combined with mildmechanical or physical/chemical pretreatment has been of interestto improve the digestibility of lignocellulosic biomass.

Fungal pretreatment followed by physical/chemical pretreatmentis summarized in Table 2. Similar to biopulping, fungal pretreatmentimproved the performance of the subsequent non-fungal pretreat-ment. Yu et al. (2010) reported that the 15-d fungal pretreatment ofcorn stalks with I. lacteus modified the lignin structure and signifi-cantly facilitated lignin degradation and xylan removal during mildalkaline pretreatment (1.5% NaOH, 30–75 °C for 15–120 min). Thesynergic effect largely depended on the severity of alkaline pretreat-ment. The less the severity of the alkaline pretreatment, the morethe cellulose digestibility was improved by fungal pretreatment. Inother words, the fungal pretreatment reduced the required severityof alkaline pretreatment. For example, at the enzyme loading of30 FPU/g solid, an 80% glucose yield was obtained with corn stalkpretreated by alkaline solution at 60 °C for 120 min, while, with

Table 2Combined white rot fungal pretreatment and physical/chemical pretreatment.

Fungal pretreatment Physical/chemical pretreatment Substrate E

Prior a Alkaline (NaOH) Corn stalk Fr

Prior Diluted acid Water hyacinth Cb

Prior Ethanolysis Beech wood chips Cie

Prior Ethanolysis Japanese cedar wood Ct

Prior Steam explosion Beech wood meal CPost b Alkaline (NaOH) Wheat straw S

oPost H2O2 Rice hull H

ys

Post Acid (Sulfuric acid) Beechwood and pine sawdust Ap

Post Alkaline (Ammonia) Beechwood and pine sawdust Ap

Post Ultrasound Beechwood and pine sawdust UPost Ultrasound Rice hull U

ih

Post Hot water extraction wheat straw Hr

Post Liquid hot water pretreatment Soybean straw La

a Prior indicates that fungal pretreatment is the first step of combined pretreatment.b Post indicates that fungal pretreatment is the subsequent step of combined pretreatme

fungal pretreatment for 15 d, the similar yield was obtained by lesssevere alkaline pretreatment conditions (60 °C for 30 min or 30 °C for120 min). Similar results were also reported by Ma et al. (2010), whoobserved that acid pretreatment (0.25% H2SO4, 28–100 °C for 15–60 min) following 15-d fungal pretreatment improved enzymatichydrolysis and ethanol yields of water hyacinth by 1–2 times over acidpretreatment alone. Ethanol yields of woody biomass such as Japanesecedar and beech wood resulting from combined C. subvermispora pre-treatment and ethanolysis were about 1–2 fold higher than that with-out fungal pretreatment (Baba et al., 2011; Itoh et al., 2003). Fungalpretreatment of beech wood meal with P. chrysosporium and thensteam explosion also increased saccharification yield compared to asingle pretreatment (Sawada et al., 1995). However, the effectivenessof combined pretreatment depended on the operational conditions ofsteam explosion. A severe steam explosion at a higher pretreatmenttemperature and time could cause condensation of Klason lignin withcarbohydrate oligomers and methanol soluble lignin (Sawada et al.,1995). As a result, these newly formed compounds can lower the sus-ceptibility of the co-treated substrate to hydrolytic enzymes. Therefore,regardless of chemical or physical pretreatment following fungal pre-treatment, a moderate subsequent pretreatment method is suggestedto achieve a more synergic effect exerted by fungal pretreatment.

Fungal pretreatment as the second step of combined pretreatment isalso summarized in Table 2. As one physical pretreatment method, ul-trasonic pretreatment has been applied to degrade α-O-4 and β-O-4linkages in lignin (Seino et al., 2001), oxidize hydroxyl groups by radi-cals and H2O2 formed during ultrasonic cavitation (Tan et al., 1985),and increase fiber wall porosity (Laine and Goring, 1977). The syner-gistic effect of ultrasonic pretreatment on the subsequent fungal pre-treatment was evident. As reported in the study of Kadimaliev et al.(2003), prior ultrasonic pretreatment (22 kHz, 10 min) improved fun-gal delignification of beech and pine sawdust with Panus (Lentinus)tigrinus. Yu et al. (2008) combined more powerful ultrasonic pretreat-ment (250 w, 40 kHz, 30 min) with fungal pretreatment and foundthat during 18-d fungal pretreatment with P. ostreatus, lignin degrada-tion on ultrasonic-modified rice hull was much higher than that ofraw hull. The substrate resulting from combined pretreatment had

ffectiveness References

ungal pretreatment improved delignification and xylanemoval during mild alkaline pretreatment

Yu et al. (2010)

ombined pretreatment increased sugar and ethanol yieldsy 1–2 folds over single acid pretreatment

Ma et al. (2010)

ombined pretreatment saved 15% electricity andncreased the ethanol yield by up to 1.6 times overthanolysis alone.

Itoh et al. (2003)

ombined pretreatment increased the sugar yield by 7imes over ethanolysis alone

Baba et al. (2011)

ombined pretreatment improved overall sugar yield Sawada et al. (1995)trong alkaline pretreatment masked the synergistic effectf fungal pretreatment on the combined process

Hatakka (1983)

2O2 enhanced fungal delignification, resulting in a sugarield that is comparable to that obtained from a long-termole fungal pretreatment.

Yu et al. (2009b)

cid pretreatment reduced lignin degradation on fungalretreatment

Kadimaliev et al. (2003)

lkaline pretreatment reduced lignin degradation on fungalretreatment

Kadimaliev et al. (2003)

ltrasound accelerated fungal lignin degradation Kadimaliev et al. (2003)ltrasound accelerated fungal delignification while slightlyncreased cellulose and hemicellulose loss, resulting in aigher sugar yield compared to the fungal pretreatment alone.

Yu et al. (2009b)

ot water extraction improved delignification and theesulting sugar yield

Wan and Li (2011)

iquid hot water pretreatment improved delignificationnd the resulting sugar yield

Wan and Li (2011)

nt.


a 20% higher glucose yield than the sole fungal-pretreated material.Moreover, this glucose yield was comparable to that obtained with42-d fungal-pretreated, which indicated that pre-modification of thefeedstock by ultrasound led to a shortened fungal pretreatment time.In the same study, H2O2 pretreatment (2% w/v, 48 h) performed betteron the following fungal delignification than ultrasound pretreatment.Hydrothermal treatment improved the fungal degradation of wheatstraw and soybean straw that appeared to be strongly resistant toC. subvermispora degradation (Wan and Li, 2011). The hot water extrac-tion (85 °C for 10 min at atmospheric pressure) facilitated fungal pre-treatment of wheat straw by removing water soluble extractives butnot of soybean straw. A pressurized hot water treatment (170 °C for3 min at 110 psi), altering rather than degrading the cell wall structure,finally facilitated the fungal degradation of soybean straw, and the glu-cose yield of the combined liquid hot water and fungal pretreatmentreached about 65% when the cellulase loading for enzymatic hydrolysiswas 10 FPU/g solid (Wan and Li, 2011) .

In contrast to the effectiveness of mild chemical pretreatment,severe prior chemical pretreatment could mask the effect of fungalpretreatment and result in no or less improvement on lignin degrada-tion and saccharification yield. Ammonia (5%, 165 °C for 10 min) andsulfuric acid (2.5%, 165 °C for 10 min) before fungal pretreatmentresulted in negative impacts on lignin degradation of both beechand pine sawdust (Kadimaliev et al., 2003). This problem was pre-sumed to be due to partial degradation of polysaccharides caused bychemical pretreatment, which provided more easily accessible com-pounds for the fungal growth; thus, more carbohydrates were con-sumed during fungal pretreatment while fungal lignin degradationactivity was probably also depressed. Similarly, NaOH pretreatment(0.4 g NaOH/g straw, 115 °C for 10 min) prior to fungal pretreatmentdid not improve sugar yields of wheat straw (Hatakka, 1983) due todegradation caused by severe alkaline pretreatment. Therefore, itcan be concluded that the severity of physical or chemical pretreat-ment strongly affects the performance of the combined pretreatmentand a mild prior pretreatment resulting in sufficient modification onbiomass could favor fungal pretreatment.

Enzymatic treatment

As mentioned in the above section, ligninolytic enzymes, mainlylignin peroxidase (LiP), manganese peroxidase (MnP), and laccase,are responsible for delignification by white rot fungi. However, theslow fungal growth (several weeks to months) and holocelluloseloss are the major problems related to fungal pretreatment. The useof ligninolytic enzymes for direct treatment of lignocellulose, whichonly takes several hours to days (Ramos et al., 2004), has the poten-tial to overcome the problems of fungal pretreatment.

Ligninolytic enzymes were shown to cleave lignin model com-pounds oxidatively in vitro (Tuor et al., 1992). The oxidation by per-oxidase is highly dependent on oxygen as the final electron acceptor(Ibarra et al., 2006) and H2O2 is usually used as a source of O2

(Khazaal et al., 1993). For oxidation of lignin by laccase, chemicalmediators, such as 1-hydroxybenzotrizole (HBT) and N-hydroxy-N-phenylacetamide (NHA), are extensively used for radical formationduring enzyme oxidation (Palonen and Viikari, 2004). These radicalscan react with aromatic acid compounds of lignin. Therefore, the lig-ninolytic enzyme-based treatment can be considered as an enzyme-catalyzed oxidative treatment (Ibarra et al., 2006). Temperature, pH,and enzyme concentration are important factors for enzyme treat-ment. Ramos et al. (2004) tested the effect of treatment of defibratedsugarcane bagasse with crude enzyme extract from P. chrysosporiumon the production of mechanical pulps. It was found that 36-hourenzymatic treatment with the addition of H2O2 resulted in higherpulp yield than the 2-week fungal pretreatment. The treatment withenzymatic extract isolated from white rot fungi also improved invitro digestibility of wheat straw as reported by Rodrigues et al.

(2008). However, this effect seemed strongly dependent on synergis-tic effects between extracted enzymes. For enzymatic treatment of di-luted acid-pretreated wheat straw, laccase had a synergistic effect oncellulase for enhancing glucose yield but was less effective thanxylanse and feruloyl esterase (Tabka et al., 2006). A maximal increaseof 21% in the saccharification rate of steam-treated softwood wasobtained from sequential combination of laccase-mediator treatmentand commercial cellulase hydrolysis (Palonen and Viikari, 2004).

White rot fungi may serve as a good producer of extracellularenzymes including oxidative enzymes and polysaccharide-degradingenzymes. The isolated enzyme complex has the potential to replace fun-gal pretreatment to avoid long pretreatment and the possible concom-itant degradation of lignin and holocellulose. However, enzymatictreatment is immature and the efficiency of oxidative enzymes largelyrelies on chemical mediators. On the other hand, the fungal-pretreated substrate containing the enzyme complex could be directlyused for the subsequent digestion and fermentation if it acts on the syn-ergism between commercial enzymes.

Modeling and scale-up

Solid state fungal pretreatment involves the degradation of thesubstrate taking place in absence (or near absence) of free water,due to release of extracellular enzymes or cell bound enzymes tothe external environment. Both interparticle and intraparticle masstransfer occurs in solid state fungal pretreatment. The intraparticlemass transfer refers to the transfer of nutrients and enzymes withinthe substrate solids. However, in aerobic digestion, the transfer ofoxygen to the growing microorganisms is the major interparticlemass transfer. Modeling of microscale phenomena, such as themicroorganism growth behavior, mass diffusion, and particle sizereductions, can describe howmicroscale processes influence growthkinetics of microorganisms. The growth kinetics models, such as lin-ear, exponential, logistic, and Monod equation, have been used todescribe microorganism growth. The logistic model, which is mostcommonly used, assumes the growth rate (μ) is independent of sub-strate concentration (Saucedo-Castaneda et al., 1990). Some studieshave proposed to express the growth rate as a function of temperature,which varies during growth (Fanaei and Vaziri, 2009; Saucedo-Castaneda et al., 1990). Due to the difficulty in separating the fila-mentous fungal biomass from the substrate, various indirect measure-ments, such as drymass loss, oxygen consumption/CO2 production, andcell specific compounds, have been proposed as fungal biomass indica-tors (Mitchell et al., 2006). Cell specific compounds such as glutamate orergosterol content are often used for indirect measurements of fungalbiomass (Joergensen, 2000; Mitchell et al., 2006).

Due to lack of free water and low conductivity of solid particles,heat generation related to metabolic activities of the microorganismgrowth causes temperature gradients in solid state fungal pretreat-ment; thus, modeling of temperature gradient and optimizing heatremoval are the major considerations in the design of solid state bio-reactors. Various bioreactors have been developed for solid state fer-mentation, including tray reactors, packed-bed, rotating drums, andstirred bioreactors (Mitchell et al., 2006). Although mixing can im-prove heat dissipation, it is only useful for solid state fungal pretreat-ment in which the fungus does not bind the solid particles together.For fungal fermentation with the interparticle hyphal bridges acrossthe substrate, mixing is deleterious to bioreactor performance be-cause of disruption of the hyphae between particles and the shearforce resulting from mixing (Fanaei and Vaziri, 2009; Mitchell et al.,2000). Instead, static operation is preferred for binding the substratebed with fungi that does not tolerate mixing. Two types of reactors,tray and packed-bed bioreactors, are commonly used for this operation.The tray reactors are simple but suitable for low volume production dueto a limited loading capacity. In packed-bed reactors, modelingis mostly focused on temperature gradients and heat transfer while


neglecting oxygen transfer because heat transfer has a greaterimpact on microorganism growth than oxygen transfer (Mitchellet al., 2000). Heat transfer with a water jacket is impractical for alarge bioreactor as it requires a large diameter water jacket. Instead,convective heat transfer, such as forced humid air, is more practicalto remove the heat and also prevents the substrate bed from dryingout. The traditional packed-bed bioreactor has the problem of axialtemperature gradients. It causes significant water evaporation andhigh temperature at the outlet of the bed even if saturated air is sup-plied. In contrast, the Zymotis packed-bed bioreactor has internalheat transfer plates, which has advantages over traditional packed-bed bioreactors, including a decrease in axial temperature gradientsand evaporation rates (Mitchell et al., 2006). However, the substrateloading and unloading is difficult in the Zymotis bioreactor due to thepresence of the heat transfer plates. Also, the water condensation onthe exposed surface limits oxygen on the top of the bed.

Scott et al. (1998) tested both the tubular reactor and chip pilesfor the scale-up of fungal pretreatment of wood chips for biopulps.The tubular reactors such as PVC tubes and silos were used aspacked-bed reactors. The larger scale-up trial of up to 40 t was donein an outdoor wood chip pile; however, the heat gradient was stilla problem. Compared to the tubular reactors, the chip piles aremore difficult to control due to 2- to 3-dimensional airflow. In addi-tion, the surface of the chip piles is more exposed to undesirable fun-gal species, and air flow and weather changes. Less controlled chippiles could also lead to reduction and variation of pulp quality dueto more degradation of cellulose which was caused by undesirablefungi and bacteria.

Limitations and potentials

Generally speaking, fungal pretreatment with selective lignin-degrading white rot fungi has been shown to have a significant im-pact on degradation and digestibility improvement for a wide rangeof feedstocks. Lignin in biomass is one of the major barriers to enzy-matic hydrolysis, and lignin removal can increase pore size in the sub-strate and provide more accessible surface area to cellulase(Taniguchi et al., 2005; Yu et al., 2009a). Therefore, lignin degradationis the key indicator to the performance of fungal pretreatment, andgood correlations between extent of lignin and polysaccharide digest-ibility were reported in many studies (Sawada et al., 1995; Taniguchiet al., 2005; Zhang et al., 2007b). However, fungal degradation mayoccur slowly and thus long pretreatment time is required to achievea relatively high lignin removal and cellulose saccharification. Con-cerns have been raised about increased costs and contaminationdue to very long pretreatment time. Combined fungal pretreatmentwith other pretreatment methods has been shown to synergisticallyimprove enzymatic digestibility and thus has the potential to shortenthe fungal pretreatment time.

Fungal pretreatment as an application of on-farm wet storage is apromising direction that addresses both the long pretreatment timeand biomass storage concerns. Currently, wet storage (>45%moisture)is receiving interest for on-farm storage of biomass, and has advan-tages over dry storage including low risk of fire, reduced weather re-lated delay during harvest, reduced harvesting costs, and improvedfeedstock uniformity (Atchison and Hettenhaus, 2003; Digmanet al., 2007; Shinners et al., 2007). Chen et al. (2007) studied ensilingagricultural residues (barley straw, triticale straw, wheat straw, cot-ton straw, and triticale hay) for bioethanol production. The ensilagewas conducted in sealed jars at room temperatures for 96 d. It wasfound that holocellulose loss ranged from 1.31% to 9.93% as a resultof ensiling while no lignin degradation was observed. About 5–10%increase in sugar yield over untreated feedstocks was observedwith ensiled feedstocks. In contrast, enzymatic digestibility resultingfrom fungal pretreatment was much higher as a result of substantiallignin degradation (Table 1). For example, the overall glucose yield

of corn stover after 35-d fungal pretreatment reached 67%, whichwas more than 45% higher than the untreated (Wan and Li, 2010b).The glucose yield obtained from optimized enzymatic hydrolysis iscomparable to that from thermo-chemical pretreatment (Wan andLi, 2010a).

Conclusions and future perspectives

Compared to current leading thermal or chemical pretreatment pro-cesses, fungal pretreatment with white rot fungi is an environmental-friendly and energy-efficient process.White rot fungi with a high selec-tivity of lignin degradation over cellulose loss are important for fungalpretreatment. Moisture and particle size of the feedstock, aeration,and pretreatment time are critical for fungal growth and metabolismto achieve good performance. Complete decontamination may not benecessary since white rot fungi can survive in contamination andactively act on degradation. Fungal pretreatment prior to mild physicaland chemical pretreatment has shown synergism on the improvementof cellulose digestibilitywith advantages similar to that of the biopulpingprocess.

Long pretreatment time is a major and common barrier for the ap-plication of fungal pretreatment. Using fungal treatment concurrentlywith on-farm wet storage is a promising option to solve the long pre-treatment time issue. Another option is to apply fungal pretreatmentprior to physical or thermo-chemical pretreatment. As short-term fun-gal pretreatment can modify the cell walls before evident degradationtakes place, the required pretreatment severity of thermo-chemicalpretreatment can be substantially reduced.

Acknowledgments

This work was supported by funding from North Central Sun GrantProgram (No. GRT00013735), USDA NIFA 1890 Capacity BuildingProgram, and Ohio Agricultural Research and Development CenterSeeds Program. The authors wish to thank Mrs. Mary Wicks andDr. Jian Shi in the Department of Food, Agricultural and BiologicalEngineering of the Ohio State University for reading through themanuscript and providing useful suggestions.

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Fungal Pretreatment of Lignocellulosic Biomass

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