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Heterologous expression of Pycnoporus cinnabarinus cellobiose dehydrogenase in Pichia pastoris and involvement in saccharification processes

Background Cellobiose dehydrogenase (CDH) is an extracellular hemoflavoenzyme produced by lignocellulose-degrading fungi including Pycnoporus cinnabarinus. We investigated the cellulolytic system of P. cinnabarinus, focusing on the involvement of CDH in the deconstruction of lignocellulosic biomass. Results First, P. cinnabarinus growth conditions were optimized for CDH production. Following growth under cellulolytic conditions, the main components secreted were cellulases, xylanases and CDH. To investigate the contribution of P. cinnabarinus secretome in saccharification processes, the Trichoderma reesei enzymatic cocktail was supplemented with the P. cinnabarinus secretome. A significant enhancement of the degradation of wheat straw was observed with (i) the production of a large amount of gluconic acid, (ii) increased hemicellulose degradation, and (iii) increased overall degradation of the lignocellulosic material. P. cinnabarinus CDH was heterologously expressed in Pichia pastoris to obtain large amounts of pure enzyme. In a bioreactor, the recombinant CDH (rCDH) expression level reached 7800 U/L. rCDH exhibited values of biochemical parameters similar to those of the natural enzyme, and was able to bind cellulose despite the absence of a carbohydrate-binding module (CBM). Following supplementation of purified rCDH to T. reesei enzymatic cocktail, formation of gluconic acid and increased hemicellulose degradation were observed, thus confirming the previous results observed with P. cinnabarinus secretome. Conclusions We demonstrate that CDH offers an attractive tool for saccharification process enhancement due to gluconic acid production from raw lignocellulosic material.


Background
In natural environments, cellulolytic microorganisms secrete enzymes that function synergistically, in association with the microorganism or independently. Although it is not fully known how many enzymes are involved in cell wall deconstruction, three general categories of enzymes are considered necessary to hydrolyze native cell wall materials: cellulases, hemicellulases and accessory enzymes such as hemicellulose debranching enzymes, phenolic acid esterase, and possibly lignindegrading and modifying enzymes [1].
The main industrial source of cellulases and hemicellulases is the mesophilic soft-rot fungus T. reesei (teleomorph Hypocrea jecorina), valued for the high protein secretion capacity of its mutant strains obtained by random mutagenesis (producing up to 100 g of extracellular protein per liter of culture) [2,3].
Among fungal classes, basidiomycetes are known to be efficient degraders of cellulose, many species growing on dead wood or litter. The lignocellulolytic system of basidiomycetes has been studied intensively in the last decades. Genome sequencing and proteomic tools are often used, but the cellulolytic system is still not completely understood, especially the oxidative part of this system [4,5].
Although the role of CDHs is still unclear, it is established that CDHs are produced in cellulolytic conditions and are involved in cellulose and lignin degradation. CDHs have been shown to bind cellulose in different ways depending on species: a long aromatic-rich region for P. chrysosporium [31] or a cellulose-binding domain for ascomycetes and soft-rot fungi, similar to that observed for cellulases [32]. Their involvement in many reactions has been demonstrated, e.g. reduction of quinones [33,34], inhibition of phenol radical repolymerization [35,20], production of hydrogen peroxide [36,37] and one of the most often cited reactions, the production of hydroxyl radicals by a Fenton-type reaction, which may participate in the degradation of cellulose, lignin and xylan [38]. CDHs are known to enhance the action of cellulases on crystalline cellulose [39,40] and also to degrade wood components, but their role in complex lignocellulosic substrate degradation has never been investigated.
Here we examined the cellulolytic system of P. cinnabarinus and the involvement of CDH therein. Given its relevance to saccharification processes, we heterologously expressed the P. cinnabarinus CDH in Pichia pastoris. The recombinant enzyme was thoroughly characterized and assessed for its ability to degrade natural substrate as a supplement to commercial Trichoderma reesei cocktail.

Results
Production and characterization of P. cinnabarinus ss3 secretome in cellulolytic conditions CDH is produced by P. cinnabarinus when cellulose is added to the culture medium. The best production (355 U/L) appeared after 10 days of cultivation when cellulose was used as sole carbon source. To understand the role of CDH when secreted in cellulolytic conditions, we characterized the P. cinnabarinus secretome after 11 days of growth.
Main enzymatic activities present in P. cinnabarinus secretome in cellulolytic conditions were measured by assay on a range of substrates (Table 1). No significant laccase or peroxidase activities were detected under our experimental conditions. However, P. cinnabarinus secretome contained enzymes able to hydrolyze a broad range of polysaccharides. Significant levels of activities towards pNP-glucose, CMC and pNP-cellobiose were detected, corresponding to β-glucosidase (0.35 U/mg), endoglucanase (0.55 U/mg) and cellobiohydrolase (0.32 Results are expressed in U/mg of total proteins. a no activity detected U/mg). A variety of hemicellulases were also identified in P. cinnabarinus secretome. The two main endo-glycosidase activities were present corresponding to endomannanase and endo-xylanase with about 2 U/mg. Hemicellulase exoglycosidase enzymes were detected to a lesser extent: 0.85 U/mg of α-galactosidase and 0.01 U/mg of β-xylosidase were measured. Zymogram assays were performed on the culture extract of P. cinnabarinus to give insight into the number of isoforms present for the main enzymatic activities previously measured.
SDS-PAGE of P. cinnabarinus ( Figure 1, lane 2) grown in cellulolytic conditions presented two main differences when compared with reference culture supernatant grown in non-cellulolytic conditions: (i) the presence of a band around 100 kDa, attributable to CDH and (ii) the absence of 70 kDa band corresponding to laccase. Confirmation by the zymogram technique showed the DCPIP decoloration by the 100 kDa band corresponding to CDH activity ( Figure 1

P. cinnabarinus CDH sequence analysis
Based on P. cinnabarinus ss3 cdh sequence, primers were designed to clone the cdh gene starting from 4-day-old culture induced with cellulose. The cdh sequence of 2310 bp was compared with available cdh sequences. Nucleotide sequence analysis showed 97% identity between cdh of P. cinnabarinus I-937 described by Moukha et al. [8] and the cdh from P. cinnabarinus ss3, a monokaryotic strain isolated from the fruit-like structure of P. cinnabarinus I-937, a wild-type dikaryotic strain. These observed differences in the nucleotide sequence resulted in eight amino acid differences at positions 96 (Ala -Glu), 331 (Arg Ser), 354 (Ala Thr), 357 (Asn Lys), 386 (Tyr Ser), 426 (Tyr Phe) and 495 (Gln Glu). Comparison with T. versicolor CDH and P. chrysosporium CDH resulted in amino acid sequence identities of 77% and 70%, respectively. P. cinnabarinus CDH amino acid sequence exhibited conserved regions with GMC oxidoreductase [42] conserved domain. The linker region rich in Thr-Ser (from position 182 to position 215), the FAD binding site and the Met/His ligands for heme fixation were also identified. Interestingly, the Thr-Ser region was also rich in Pro (28% Thr, 25% Pro, 13% Ser). Analysis of the gene encoding the CDH from P. cinnabarinus has shown high sequence homology with cdh from class I. Indeed, phylogenetic analysis of cdh genes revealed two major classes [42]. The class I cdh genes are found only in basidiomycetes while the class II contain more complex ascomycetes CDHs, that sometimes present a family 1 carbohydrate-binding module (CBM) at the C-terminal position. Emergence of a third class of CDHs in ascomycetes fungi was recently reported [32].

Heterologous expression of CDH in P. pastoris
The coding sequence of cdh was inserted into the P. encoding the yeast α-factor secretion peptide and a (His) 6 tag located at the C terminus. The recombinant gene was then introduced into the Pichia genome under the control of the methanol-inducible promoter. Multicopy transformants were screened to select a clone that exhibited satisfactory levels of production. CDH activity was successfully detected in the supernatant after induction, indicating correct processing of the α-factor signal sequence.
A maximum activity of 1176 U/L was observed after 4 days of induction, and this clone was chosen for this study. To scale up enzyme production, we optimized CDH expression in a 1 liter bioreactor with the best-performing clone of P. pastoris. The recombinant CDH was secreted at high levels, reaching 7800 U/L. Recombinant CDH was purified after 4 days of induction, taking advantage of the (His) 6 tag. Also, only trace amounts of endogenous proteins were present in the culture supernatant of the transformant secreting CDH. The purified enzyme exhibited a specific activity of 22.2 U/mg.

Biochemical characterization of rCDH
Recombinant CDH was purified to homogeneity, i.e. one major band displaying a relative molecular weight around 110 kDa ( . The heme-containing domain was not seen after staining, probably owing to the weak presence of aromatic residues [43]. Binding studies of CDH confirmed the ability of the enzyme to bind cellulose without the presence of a cellulose-binding domain. Dissociation constant (K d ) and binding capacities (B max ) of CDH were determined and were respectively 0.064 μM and 0.2 μmol/g of Avicel ( Figure 3).
When DCPIP is used as electron acceptor, the optimal temperature for CDH is 70°C. The recombinant enzyme displayed activity over a wide range of temperatures, 16% of residual activity at 10°C and 55% of residual activity at 80°C ( Figure 4A).
After incubation of CDH at 45, 50 and 55°C for 33 h, residual enzyme activity was 90%, 80% and 63% respectively. However, CDH was not stable at 65°C, with only 15% of activity remaining after 9 h ( Figure 4C). The optimal pH for recombinant CDH ( Figure 4A) was pH 4.5. The recombinant CDH had V max = 22.2 U/mg and K M = 35.5 μM for cellobiose with DCPIP as electron acceptor. Using cyt c as electron acceptor, we found V max = 3.9 U/mg and K M = 14.7 μM ( Table 2).

Effect of CDH on the saccharification of wheat straw
The range of lignocellulosic enzymes found in the supernatant of P. cinnabarinus makes it a candidate for supplementation of the T. reesei cocktail for saccharification of wheat straw. We thus set out to compare the efficiency of the P. cinnabarinus supernatant with the purified rCDH for supplementation of industrial cocktails. The T. reesei cocktail supplemented with β-glucosidase was used as reference. Addition of P. cinnabarinus secretome (10, 20 and 40 U of CDH) or pure rCDH (10 and 20 U) gave similar results ( Figure 5 and Figure 6). DNS assays were used to measure reducing ends of sugars released after saccharification. Supplementation with purified rCDH or secretome containing CDH showed less response on DNS titration than control with cocktails ( Figure 5). However, overall hydrolysis was increased by addition of CDH, with the production of large amounts of gluconic acid, from 5 to 100 mg per g of wheat straw, compared with control. Also, greater yields of xylose, galactose and arabinose, which increased respectively from 35 to 44, 1.9 to 4 and 9.5 to 13.5 mg/g of wheat straw were observed with addition of 10 U of CDH ( Figure 6A). Production of gluconic acid by CDH can be explained by the formation of cellobionolactone following by its spontaneous hydrolysis in cellobionic acid. This last compound can be cleaved by β-glucosidase into glucose and gluconic acid. Experiments were performed on Dionex (data not shown).
To confirm the strong production of gluconic acid, purified recombinant CDH was used to supplement the T. reesei and A. niger cocktails. The effect on wheat straw was comparable to that obtained with P. cinnabarinus secretome ( Figure 6B). Also, supplementation with 10 U of rCDH did not affect the yield of glucose, increased hemicellulose yield and resulted in the formation of gluconic acid in large amounts.

Discussion
In the last few decades the white-rot fungus P. cinnabarinus has been studied for its ligninolytic system, which is based on phenoloxidases such as laccases, without the presence of peroxidases [44]. This system, and especially laccase, has been used to produce high value compounds [45,46] and applied to the design of biotechnological processes [47]. Here we investigated the cellulolytic and oxidative system of P. cinnabarinus grown in cellulolytic conditions.
In the P. cinnabarinus secretome, we found hemicellulase activities already reported in the literature: αgalactosidase, xylanase or β-galactosidase [48,49,41], together with mannosidase and arabinofuranosidase activities not hitherto described in P. cinnabarinus. Endoglucanase and exoglucanase were identified by zymogram (CMCase) and by hydrolysis of Avicel and CMC. Peroxidase activity assay (manganese peroxidase and lignin peroxidase) was performed on the secretome, but no activity was recovered. P. cinnabarinus is a well-known producer of laccase [50], but in cellulolytic conditions, laccase production seems to be repressed, whereas the zymogram shows activity on ABTS around 50 kDa. Similar results were observed in P. chrysosporium grown in cellulolytic condition with the presence of several laccase bands on the zymogram around 50 kDa confirmed by electron paramagnetic resonance [51]. Production of CDH was previously described [41,25] and its activity was followed in P. cinnabarinus culture.
We cloned and expressed P. cinnabarinus CDH in P. pastoris. CDH of T. versicolor [52], P. chrysosporium [53] and more recently N. crassa [13] were previously expressed in the same host. These results confirm that P. pastoris heterologous expression is an efficient way to produce fungal CDHs at high levels.  Enzymatic characterization of recombinant CDH gave values of kinetic parameters (V max , K M ) in the same range as those observed previously for the native enzyme [25] and more generally for the recombinant CDH cited in the literature [12,52]. However, recombinant CDH of P. cinnabarinus is more thermostable than the other fungal CDHs, with an optimal temperature around 70°C. Optimal pH 4.5 is in close agreement with the literature. Some CDHs produced by ascomycetes and soft-rot fungi contain a carbohydrate binding module (CBM) and are able to bind cellulose. In the case of P. chrysosporium CDH, the ability to bind cellulose seems to be mediated by a specific domain with a structure different from CBM [31]. The ability of the purified enzyme to bind Avicel in the absence of CBM was confirmed experimentally.
CDH is produced simultaneously with cellulase. Its role in the degradation of cellulose was shown by Bao et al., who found that P. chrysosporium CDH increased the sugar yield from cellulose and produced cellobionolactone [39]. In this work, we decided to use CDH to supplement cellulase cocktail on complex substrate such as wheat straw.
In a first set of experiments, we used the P. cinnabarinus secretome containing CDH added directly to cellulase cocktail for the saccharification of wheat straw.
Results on wheat straw showed (i) increased yield in C5 sugars from hemicelluloses, consistent with the lignin degradation effect of the secretome, and (ii) a slight decrease in glucose yield correlated with the formation of large amounts of gluconic acid due to cleavage of cellobionic acid (the main product of the reaction performed by CDH) by β-glucosidase.
Supplementation with purified rCDH gave similar results on wheat straw and even no decrease in glucose yield, but gluconic acid and C5 sugar hemicellulose production was enhanced for 10 U CDH supplementation. Results point to synergy between CDH and cellulases for degradation of raw material. In P. cinnabarinus secretome, β-glucosidase activity was significantly detected (Table 1). However, when no β-glucosidase was added to the saccharification assay, more cellobionic acid was produced instead of gluconic acid by T. reesei cocktail supplemented with P. cinnabarinus secretome (data not shown). It is well established that β-glucosidases are inhibited by gluconolactone and more generally that lactones are inhibitors of many glycosidases [54,55]. Nevertheless, sugar lactones are unstable in aqueous solution, and the rate of spontaneous hydrolysis to the corresponding aldonic acid, i.e. gluconic acid or cellobionic acid, depends on the pH and temperature of the reaction. Aldonolactonase, found in several fungi, catalyzes the hydrolysis of lactones to aldonic acid [56]. This hydrolysis should relieve inhibition of β-glucosidase and glycosidase by lactone, as suggested by Bruchman et al. [57]. β-Glucosidase is able to cleave cellobionic acid into glucose and gluconic acid [58]; cellobionic acid and gluconic acid production decreases the number of reducing ends as shown by the decrease in DNS titration. In the presence of CDH, DNS titration is not a relevant method for monitoring cellulose degradation. The presence of cellobionic acid seems due to a faster reaction rate of CDH than βglucosidase versus cellobiose, as shown by Yoshida et al. [59]. Supplementation with β-glucosidase compensates for the difference in reaction rate, leading to a greater production of gluconic acid. Conversely, as the accumulation of cellobiose induces inhibitory effects on cellulase [60], CDH may decrease the cellobiose concentration in the medium faster and so avert inhibition.

Conclusions
Supplementation of T. reesei secretome by CDH increases the overall degradation of lignocellulose and produces appreciable amounts of gluconic acid. In saccharification processes, the use of gluconic acid should offer a way to improve the profitability of the whole process. Several organisms use gluconic acid through the pentose phosphate pathway. Zymomonas mobilis, for example, is able to produce ethanol from gluconic acid by the Entner-Doudoroff pathway [61]. Alcoholic fermentation from gluconic acid by Saccharomyces bulderi has also been reported [62]. The introduction of such organisms able to use pentose and gluconic acid should increase the overall yield of ethanol by using less fermentable components and should offer a way to design a sustainable process for second generation bioethanol production.

Biological material
P. cinnabarinus ss3 monokaryotic strain BRFM 137 isolated from the fruit-like structure of the P. cinnabarinus  I-937 dikaryotic strain was maintained as previously described [50]. P. pastoris strain X33 is a component of the Pichia Easy Select Expression System and the pPIC-ZαA vector (Invitrogen, Cergy-Pontoise, France).
Media and culture conditions P. cinnabarinus was grown at 30°C on MYA2 plates (maltose: 20 g/L; yeast extract: 1 g/L; agar 16 g/L). After 10 days of incubation, precultures in Roux flasks containing 200 mL of medium were inoculated with five disks of P. cinnabarinus grown in MYA 2 plates. Inoculum was obtained from 10-day-old static precultures incubated at 30°C. We used 10 mL of inoculum suspension obtained from Ultra-Turrax-mixed mycelial mats to inoculate 500  [41]. For the heterologous expression of CDH in Pichia pastoris, all media and protocols are described in the Pichia expression manual (Invitrogen). Cloning procedures were performed using one-shot TOP 10 and DH5α chemically competent Escherichia coli cells (Invitrogen).

Isolation of mRNA and cloning of cdh cDNA gene
Isolation of total RNA was performed on a 4-day-old culture of P. cinnabarinus on cellulose medium using Total RNA Purification from Plant (Macherey-Nagel, Düren, Germany) as prescribed by the manufacturer. Contaminant DNA was digested by Turbo DNase (Ambion Inc., Austin, TX, USA) according to the manufacturer's instructions. First-strand cDNA synthesis was performed using SuperScript reverse transcriptase (Invitrogen) and oligo(dT 18 ) primer following the manufacturer's instructions. The amplification of the full-length cdh cDNA was performed using specific primers (with restriction sites underlined): forward primer cdhF (5' TA GAA TTC CAA GTG GCA GCG CCA TAC 3') and reverse primer cdhR (5' TA TCT AGA CCA GGA CCT CCC GCA AGG GC 3') designed from P. cinnabarinus I-937 cdh gene (NCBI AF081574): 315 ng of cDNA was mixed with 300 pmol of each primer cdhF and cdhR, 200 μM dNTPs, and 0.5U Pfu DNA polymerase (Promega, Madison, WI, USA). The reaction was performed with the following amplification program: 1 cycle at 95°C for 5 min, 30 cycles composed of three steps for each cycle (95°C for 1 min, 65°C for 30 s and 72°C for 4 min), and a final step at 72°C for 10 min. PCR amplicons generated by Pfu DNA polymerase are bluntended. To add an A-tail on these PCR fragments before subcloning into pGEMT-easy vector, Taq DNA polymerase (Promega) was used as described in the pGEMT-easy vector Technical Manual (Promega). The 2.3 kb PCR product was purified using the Qiaquick gel extraction kit (Qiagen, Valencia, CA, USA) and subcloned into pGEMT easy vector.

Construction of pPiCZaA expression vector
The cdh cDNA cloned into pGEMT easy vector was digested using EcoRI and XbaI and purified with a Qiaquick gel extraction kit. In parallel, pPICZαA was linearized using the same restriction enzymes, and cdh cDNA was ligated at the corresponding sites into pPIC-ZαA in frame with both the yeast α-secretion factor and C-term-(His) 6 -tag encoding sequences. Expression vector pPICZαA-cdh was purified by Qiagen Midiprep and sequenced using 3'AOX and 5'AOX primers to confirm the correct sequence insertion.

Transformation and screening
Transformation of competent P. pastoris X33 was performed by electroporation with PmeI linearized pPICZα A-cdh as described in Couturier et al. 2010 [64]. The vector pPICZα without insert was used as a control. Transformants were first screened on YPDS plates with different concentrations of zeocin (100 to 1000 μg/mL). After incubation at 30°C, transformants were picked from minimal dextrose (MD) plates and transferred to minimal methanol plates (MM). Zeocin-resistant P. pastoris transformants were then screened for protein expression in 10 mL of BMGY (in 50 mL tubes) at 30°C in an orbital shaker (200 rpm) for 16 h to an OD 600 of 2-6, and expression was induced by transferring cells into 2 mL of BMMY and growing for a further 3 days. Each day the medium was supplemented with 3% (v/v) methanol. The supernatant was then analyzed by SDS-PAGE to determine which transformant had the best secretion yield.

Recombinant CDH production
The best-producing transformant was grown in 1 liter of BMGY in shaken flasks as described above. The cells were then transferred to 200 mL of BMMY and stirred at 200 rpm at 30°C for 4 days.
Bioreactor production of the best-producing transformant was carried out in a 1-liter bioreactor Tryton (Pierre Guerin, Mauze, France) according to the Pichia Fermentation Process Guidelines (Invitrogen) except for the volume of methanol added in the methanol fed batch, which was changed from 3.6 mL/h/L to 3 mL/h/ L.

Enzyme purification
Culture supernatant was concentrated at least 10 times using Amicon centrifugal units with a 30 kDa cut-off, 4000× g or Amicon vivaflow (Millipore, Bedford, MA, USA) with a 30 kDa cut-off, depending on culture volume. The concentrated supernatant was dialyzed against buffer A (Tris-HCl 50 mM 7.8, NaCl 150 mM and imidazole 10 mM), and loaded on a nickel chelate His-Bind Resin (GE Healthcare, Buc, France) column (0.7 × 5 cm) connected to an Äkta FPLC (GE Healthcare) and equilibrated with buffer A. The His-tagged rCDH was eluted with buffer B (Tris-HCl 50 mM pH 7.7, imidazole 500 mM and NaCl 150 mM). Active fractions were pooled, concentrated and dialyzed against sodium acetate buffer (50 mM, pH 5)

SDS-PAGE, Western blot and zymogram
Polyacrylamide gel electrophoresis (SDS-PAGE) (12%) was prepared as described by Laemmli [65]. Protein bands were stained with Coomassie blue G 250. The molecular mass under denaturating conditions was determined with reference standard proteins (LMW, Amersham Pharmacia Biotech, Orsay, France or unstained protein molecular weight marker, Euromedex, Souffelweyersheim, France).
Enzyme activities were assayed in polyacrylamide gels containing the appropriate substrates. Enzyme preparations were run on an SDS-PAGE gel copolymerized with 0.2% soluble xylan, 0.2% carboxymethylcellulose (CMC) or 0.2% locust bean gum for the analysis of xylanase, CMCase or mannanase activities, respectively. The protein samples were mixed in the loading buffer (3% SDS w/v, 10% glycerol w/v and 30 mM Tris-HCl buffer pH 6.8) without reducing agent, heated at 100°C for 1 min and then separated using a 12% polyacrylamide gel. After electrophoresis, the gel was washed with deionized water and soaked in 2.5% (v/v) Triton X-100. After 1 h incubation at 4°C, the gel was soaked in 100 mM sodium phosphate buffer (pH 5) at 45°C for 2 h for the detection of xylanase and CMCase activity or in 100 mM sodium phosphate buffer (pH 7) for 1 h at 50°C for the detection of mannanase activity. After incubation the gel was stained with 0.1% Congo red solution under gentle shaking for 1 h and destained with 1 M NaCl for 1 h. Protein bands exhibiting xylanase, CMCase and mannanase activity were seen as clear bands on the red background.
For laccase and CDH zymograms, samples were mixed with the same loading buffer as described above without heating; they were incubated at ambient temperature for 15 min and the gel was run. After electrophoresis the gel was soaked in 2.5% Triton X-100 for 1 h at 4°C, rinsed with deionized water and incubated for 2 h at 25°C in 50 mM sodium acetate buffer (pH 5) with 4 mM sodium fluoride for CDH and 50 mM sodium tartrate buffer (pH 4) for laccase. Visualization was performed by adding 5 mM ABTS to stain for laccase and adding 50 mM DCPIP for CDH, staining the gel dark blue. CDH activity was then visualized by adding 100 mM cellobiose. Protein bands exhibiting CDH activity were seen as clear bands on the dark blue background. Western blot analysis was performed as described previously, using the monoclonal anti-polyhistidine alkaline phosphatase conjugate (Sigma) for Western blot analysis of rCDH expressed in P. pastoris. For Western blot analysis, purified rCDH was run on a 12% SDS/polyacrylamide gel and blotted onto a PVDF membrane using the iBlot Dry Blotting System (Invitrogen). Membranes were placed in a Snap Protein Detection System (Millipore, Bedford, MA, USA) used for immunodetection. Following the manufacturer's instructions, the PVDF membrane was incubated in TBS blocking solution (10 mM Tris, 150 mM NaCl and 0.1% Tween 20, pH 8) with addition of 0.1% (w/v) of skimmed milk powder and then washed with TBS. Immunodetection was performed using the monoclonal anti-polyhistidine alkaline phosphatase conjugate (Sigma,). Signal detection was carried out using 60 μL of BCIP (5bromo-4-chloro-3-indolyl-phosphate), 60 μL of NBT (4-nitro blue tetrazolium) (Roche Applied Science, Meylan, France) in 20 mL carbonate buffer 0.05 M pH 9.6 with addition of 5 mM MgCl 2 .
Papain cleavage of the two CDH domains was carried out as described by Henriksson et al. [43]. Deglycosylation was performed using PGNase (New England Biolabs, Saint-Quentin-en-Yvelines, France) to remove rCDH N-linked glycans according to the manufacturer's instructions.

Protein assay
Protein concentration was determined using the Bio-Rad Protein Assay (Bio-Rad, Marnes-la-Coquette, France), based on the Bradford procedure, using bovine serum albumin as standard [66].

Binding studies
Assays were performed with 1 mg/mL of Avicel PH-101 (Sigma) in 50 mM citrate phosphate buffer pH 5 under orbital agitation at room temperature, and rCDH was added in the range 0.02-0.8 μg/L. Two controls were run, one without rCDH and the other with BSA at 1 μg/μL to estimate unspecific fixation of rCDH. Measures were repeated at least three times.

Enzyme assays
To measure enzyme activities in the P. cinnabarinus culture supernatant, each aliquot was centrifuged for 5 min at 3500 rpm and filtered through a 0.45 μm membrane (Millipore, Bedford, MA, USA). CDH activities were determined by monitoring the reduction of 0.2 mM 2,6-dichlorophenol indophenol (DCPIP) in 100 mM sodium acetate buffer (pH 5) containing 2 mM cellobiose and 4 mM of sodium fluoride (sodium fluoride was used as a laccase inhibitor). The decrease in absorption at 520 nm (ε = 6800 M -1 .cm -1 ) was monitored at 30°C for 1 min. Alternatively, CDH activity was determined by monitoring the reduction of 50 μM cytochrome c (cyt c) in 100 mM sodium-acetate buffer (pH 5) containing 2 mM cellobiose. The decrease in absorption at 550 nm (ε = 33,700 M -1 .cm -1 ) was monitored at 30°C for 1 min. Glucose oxidase was measured using the D-gluconic acid / D-glucono-δ-lactone assay (Megazyme). Laccase activity was determined quantitatively by monitoring the oxidation of 5 mM ABTS (2, 2'-azinobis (3-ethylbenzthiazoline-6-sulfonic acid)) at 420 nm (extinction coefficient 36,000 mM. -1 cm. -1 ) in the presence of 50 mM NaK tartrate, pH 4.0. Lignin peroxidase activity was determined spectrophotometrically at 30°C by the method of Tien and Kirk [67]. Manganese peroxidase activity was determined spectrophotometrically at 30°C by the method of Paszczynski et al. [68] using H 2 O 2 and vanillylacetone as substrate. Enzyme activity was expressed in international units (IU). One unit of activity is defined as the quantity of enzyme that transforms 1 μmol of substrate in one minute.
Hydrolysis assays for glycosidases were carried out in 50 mM acetate buffer pH 5 containing 1 mM of substrate in a final volume of 100 μL. Substrates pNP-β-D-glucopyranoside, pNP-β-D-cellobiopyranoside, pNPβ-D-xylopyranoside, α-D-galactopyranoside and pNP-β-D-mannopyranoside were purchased from Sigma. Assays were performed with 0.5 and 1 μg of protein, and incubated for 37°C for 1 h with shaking (300 rpm). To stop the reaction, 130 μL of Na 2 CO 3 1 M was added, and absorbance was read at 410 nm. A control was run with 100 μL of 50 mM acetate buffer pH 5 and references ranging from 0.02 to 0.2 mM of 4-nitrophenyl were measured in parallel. Enzymatic activity was based on colorimetric assay of free pNP present in the reaction after hydrolysis. This activity is expressed in U/mg of proteins.
Hydrolysis assays were carried out in 50 mM acetate buffer pH 5 containing 1% (w/v) of substrates. Carboxymethyl cellulose (CMC, low viscosity) and citrus pectin were purchased from Sigma. Wheat arabinoxylan (low viscosity) and galactomannan (low viscosity) were from Megazyme. Assays were performed with 10 and 30 μg of protein, and incubated at 37°C for 1 h with shaking (150 rpm). Reducing sugars released during hydrolysis were quantified by DNS (3, 5-dinitrosalycylic acid) visualization at 540 nm as described in Navarro et al., 2010 [69]. Controls were run with 50 mM acetate buffer pH 5 and references ranging from 1 to 10 mM of glucose were measured in parallel for each series. Enzymatic activity is expressed in U/mg of proteins. Three controls were performed with the secretome alone to quantify sugars present in culture supernatant. Controls were subtracted from measured values. All assays were performed in triplicate.

Effect of pH and temperature on the activity and stability of rCDH
To determine the optimum pH of the rCDH, the activity was measured with DCPIP using 50 mM citrate phosphate buffer in the pH range 2.5-7 at 30°C. For optimum temperature determination, activity on DCPIP was measured using 50 mM citrate phosphate buffer in the temperature range 10-80°C. Thermal stability of rCDH was determined by incubating enzymes for 33 h at 45, 50 and 55°C and for 10 h at 65°C. Native CDH activity assay was performed in triplicate as described above.

Enzyme kinetics
The kinetic parameters (V max and K m ) were determined for cellobiose oxidation measured at 30°C in 50 mM citrate phosphate buffer pH 4.5 using DCPIP or cytochrome c. The concentration of cellobiose ranged from 10 to 700 μM with both electron acceptors (DCPIP and cytochrome c). Triplicates were run to ensure reliable kinetic parameter determination.
Graphpad prism v.4 (Graphpad Software) was used for the nonlinear regression calculation and kinetic parameter determination.

Carbohydrate determination
Monosaccharides, cellobiose and gluconic acid generated after hydrolysis of wheat straw were quantified by highperformance anion exchange chromatography (HPAEC) coupled with amperometric detection (PAD) (ICS 3000, Dionex, Sunnyvale, CA, USA) equipped with a Carbo-Pac PA-1 analytical column (250 × 4 mm). Enzymatic reactions were stopped by adding 18 mM NaOH before injection (5 μL) into the HPAEC system. Elution (1 mL/ min) was carried out on a sodium acetate gradient (0-250 mM in 25 min). Calibration curves were plotted using galactose, arabinose, glucose, xylose, cellobiose and gluconic acid standards (Sigma-Aldrich), from which response factors were calculated (Chromeleon program, Dionex) and used to estimate the amount of product released in test incubations. All the assays were carried out in triplicate. Reducing sugars released during saccharification assays were quantified by DNS (3, 5dinitrosalycylic acid) method and visualized at 540 nm as described by Navarro et al. [69]. Controls were run with 50 mM acetate buffer pH 5 and references of glucose were ranging from 1 to 10 mM. All assays were performed in triplicate.