Two structurally discrete GH7-cellobiohydrolases compete for the same cellulosic substrate fiber
- Fernando Segato1, 4,
- André R L Damasio1, 5,
- Thiago Augusto Gonçalves4,
- Mario T Murakami3,
- Fabio M Squina4,
- MariadeLourdesTM Polizeli6,
- Andrew J Mort2 and
- Rolf A Prade1, 4Email author
© Segato et al.; licensee BioMed Central Ltd. 2012
Received: 14 December 2011
Accepted: 30 March 2012
Published: 11 April 2012
Cellulose consisting of arrays of linear beta-1,4 linked glucans, is the most abundant carbon-containing polymer present in biomass. Recalcitrance of crystalline cellulose towards enzymatic degradation is widely reported and is the result of intra- and inter-molecular hydrogen bonds within and among the linear glucans. Cellobiohydrolases are enzymes that attack crystalline cellulose. Here we report on two forms of glycosyl hydrolase family 7 cellobiohydrolases common to all Aspergillii that attack Avicel, cotton cellulose and other forms of crystalline cellulose.
Cellobiohydrolases Cbh1 and CelD have similar catalytic domains but only Cbh1 contains a carbohydrate-binding domain (CBD) that binds to cellulose. Structural superpositioning of Cbh1 and CelD on the Talaromyces emersonii Cel7A 3-dimensional structure, identifies the typical tunnel-like catalytic active site while Cbh1 shows an additional loop that partially obstructs the substrate-fitting channel. CelD does not have a CBD and shows a four amino acid residue deletion on the tunnel-obstructing loop providing a continuous opening in the absence of a CBD. Cbh1 and CelD are catalytically functional and while specific activity against Avicel is 7.7 and 0.5 U.mg prot-1, respectively specific activity on pNPC is virtually identical. Cbh1 is slightly more stable to thermal inactivation compared to CelD and is much less sensitive to glucose inhibition suggesting that an open tunnel configuration, or absence of a CBD, alters the way the catalytic domain interacts with the substrate. Cbh1 and CelD enzyme mixtures on crystalline cellulosic substrates show a strong combinatorial effort response for mixtures where Cbh1 is present in 2:1 or 4:1 molar excess. When CelD was overrepresented the combinatorial effort could only be partially overcome. CelD appears to bind and hydrolyze only loose cellulosic chains while Cbh1 is capable of opening new cellulosic substrate molecules away from the cellulosic fiber.
Cellobiohydrolases both with and without a CBD occur in most fungal genomes where both enzymes are secreted, and likely participate in cellulose degradation. The fact that only Cbh1 binds to the substrate and in combination with CelD exhibits strong synergy only when Cbh1 is present in excess, suggests that Cbh1 unties enough chains from cellulose fibers, thus enabling processive access of CelD.
KeywordsCellobiohydrolase Cellobiohydrolase I Cellobiohydrolase D Aspergillus niveus Aspergillus fumigatus Crystalline cellulose breakdown Biofuels Cellulases Biomass decomposition
Carbohydrate-Active enZYmes Database (http://www.cazy.org/)
CaZy GH7 (Cel7A) cellobiohydrolase 1
CaZy GH7 (Cel7B) cellobiohydrolase D
Degree of synergism
Whatman #3 filter paper
Minimal Aspergillus medium
Phosphoric acid swollen (Avicel) cellulose
Sodium dodecyl sulfate polyacrylamide gel electrophoresis.
Biofuel generation from sources such as cornstarch, sugarcane or sweet sorghum syrups, produces large amounts of biomass waste products. For example, over 90% of the plant is unused in the case of ethanol production from cornstarch [1–3]. Current commercial enterprises produce ethanol from cornstarch, sugar cane or sweet sorghum syrups and large amounts of waste biomass accumulate alongside refineries and most of them are not recycled. Biofuels production would therefore be significantly more efficiently utilized if this biomass could be converted into fermentable sugars.
Biomass polysaccharides consist of cellulose, hemicellulose and pectin. Cellulose is a linear, crystalline self-assembled nanofiber formed from the linear polymer containing exclusively glucose monomers linked through β-1,4-glycosydic bonds [1–4]. Recalcitrance of cellulose towards enzymatic degradation is a widely reported phenomenon [1, 5, 6] and is the result of the wide range of possible intra- and inter-molecular hydrogen bonds within and among linear cellulose molecules assuming crystalline or amorphous cellulosic nanofiber structures. Cell walls from different plants contain various amounts of crystalline/amorphous cellulose fibers , reflected by the relative crystalinity index (RCI).
Because cellulose is structurally complex and crystalline in nature, it is recalcitrant towards microbial or enzymatic attack. Unfortunately, current pretreatment methods – e.g., acid hydrolysis [8, 9] or pyrolysis [10, 11], generate compounds that inhibit subsequent fermentation. Enzymatic hydrolysis of cellulose results in glucose the universal carbon source for all organisms to drive oxidative metabolism including the production of biofuels, chemicals and pharmaceuticals.
A complete enzymatic cellulose degrading system consists of at least three related partially redundant biochemical reactions. Endo-glucanases (EC 184.108.40.206) randomly hydrolyze internal glycosydic bonds to decrease the length of the cellulose chain and multiply polymer ends [12–14]. Exo-glucanases (EC 220.127.116.11) split-off cellobiose from cellulose termini [15–17] and β-glucosidases (EC 18.104.22.168) hydrolyze cellobiose and oligomers to render glucose [18–20]. All three types of enzymes have similar catalytic domains all splitting the β-1,4-glycosidc bond between glucose molecules, however they differ in their binding and substrate interaction domains resulting in cooperation and synergism in releasing glucose [21–23].
Exo-glucanases hydrolyze cellulose chains by removing cellobiose either from the non-reducing end (GH6, EC 22.214.171.124) or reducing end (GH7, EC 126.96.36.199), which in both cases results in the release of reducing sugars (cellobiose) but little polymer length reduction . In fungi, exo-glucanases are commonly known as 1,4-β-D-glucan cellobiohydrolases . The main characteristic of a cellobiohydrolase is its processive action on individual cellulose chains by reiterated release of cellobiose [24, 26–32].
The catalytic domain (CD) of a typical cellobiohydrolase forms a channel-shaped cavity topped by several flexible loops resulting in a tunnel like structure [33–39], which is frequently connected through a linker to a carbohydrate binding domain (CBD) [40, 41]. CBDs are thought to bind to the crystalline cellulosic fiber sliding down the cellulosic surface [6, 42] and carrying the CD with it. Three tyrosine residues on the CBD hydrophobic surface align with the cellulose chain adjacent to the reducing end and a fourth tyrosine residue moves from its internal position to form van der Waals interactions with the cellulose surface resulting in an induced change near the surface allowing the CBD to progress [16, 40].
From a comprehensive bioinformatics survey of seven Aspergillii completely sequenced genomes, we found between three and five genes encoding cellobiohydrolases, typically two in each of the CAZy families GH6 and GH7, respectively. Remarkably, for each GH family 6 or 7 fungal cellobiohydrolase examined, one enzyme was linked to a CBD and a second was not.
Here we describe two GH7-cellobiohydrolases, Cbh1 and CelD from Aspergillus niveus. The structural differences between these two enzymes were the lack of a CBD in CelD and a conserved loop that obstructed the catalytic tunnel in enzymes that were linked to a CBD, but missing in enzymes that did not link to a CBD, ensuring that the tunnel was always open. These structural differences reflected directly on kinetic properties, thermostability, inhibition by glucose and cellobiose and the way these proteins interacted with crystalline substrate molecules. We hypothesize that the cellobiohydrolase linked to a CBD is competent to release cellulosic chains from the crystalline nanofiber and the cellobiohydrolase without a CBD can only use loose ends, without the aid of a CBD.
Aspergillii hold precisely two GH7-cellobiohydrolases, one with and another without a cellulose-binding domain
Fungi and bacteria produce enzymes that at least partially degrade plant cell wall polysaccharides [43, 44]. More often than not, cellulase-encoding genes appear as multiple copies in the genome of sequenced microorganisms and enzymes occur as functionally redundant with overlapping biochemical functions.
Genome wide distribution of gh7-cellobiohydrolases
Prot ID Locus tag
Prot ID Locus tag
A. clavatus NRRL1
A. fumigatus Af293
XP_750600 AFUA 6G07070
XP_751044 AFUA 6G11610
A. nidulans FGSC A4
A. niger CBS 513.88
A. terreus NIH 2624
A. oryzae RIB40
A. flavus NRRL-3357
P. chrysogenum W54-1255
Neurospora crassa OR 74A
CAA38275 Pccbh 1-2
Trichoderma viride (Hypocrea rufa)
A. niveus a filamentous fungus analogous to A. fumigatus (the genomic DNA sequence is up to 97% identical) grew at high temperatures, up to 50°C, on medium containing complex carbon sources such as cellulose and hemicellulose. As expected, the genome of A. niveus encoded two GH7-cellobiohydrolases: Cbh1-like 1,4-beta-D-glucan cellobiohydrolase with a linker and CBD and a CelD-like 1,4-beta-D-glucan cellobiohydrolase with no linker and CBD. Genes, cbh1 and celD were expressed and proteins secreted into the medium when the fungus was grown on cellulose (e.g., Avicel) as the carbon source (data not shown).
The Cbh1 and CelD catalytic domains share 72% identity and 82% similarity in their amino acid sequences. The main difference between these proteins is the presence of a cellulose-binding domain in Cbh1 and the absence of such a domain in CelD. The objective of this study was to investigate the mechanistic differences between these two enzymes.
Minor structural differences in Cbh1 and CelD are catalytically relevant
Cbh1 and CelD catalytic domains (CD) were modeled based on the 3D-structural template derived from Talaromyces emersonii cellobiohydrolase 1 (PDBID: 1Q9H, ). Modeled structures were subjected to energy minimization and final models presented high-quality local and overall stereochemistry with approximately 98% of amino acid residues lying in allowed regions of the Ramachandran plot (Figure 1B-E).
The Cbh1 and CelD catalytic core comprises a typical CaZy GH7 family β-sandwich surrounded by numerous surface loops, which outline the substrate-binding channel (Figure 1B-E). The native T. emersonii structure presented a disordered loop encompassing the amino-acid segment 190–200. This sequence was fully conserved among all other Cbh1 proteins (Figure 1F), and likely became structured upon substrate binding similarly, to what has been reported for other glycosyl hydrolases . Structural superposition resulted in a RMSD value of 0.15 Cα for Cα atoms indicating high conservation of Cbh1 and CelD three-dimensional structures. The catalytically relevant residues and the substrate channel are fully conserved including Trp38, Tyr168, Asp170, Glu209, Asp211, Glu214, Trp371 and Trp380 (Figure 1C). The only significant difference was at the entrance of the catalytic tunnel formed by loops 43–63, 94–103 and 190–200 that partially blocked the catalytic channel in Cbh1, due to a four-residue insertion in the 94–103 loop (Figure 1C and 1F). In all CelD proteins, this four amino acid insertion is missing (Figure 1F) and CelD proteins do not have a CBD thus suggesting that in the absence of a CBD, the catalytic domain only functions if the substrate channel is open.
Cbh1 and CelD are catalytically functional
Cbh1 and CelD kinetic properties on crystalline cellulosic substrates
U mg prot-1*
Only Cbh1 binds to cellulose
Thermal inactivation leads to differential protein unfolding
Differential glucose inhibition implies two differentially adapted catalytic domains
Cbh1 and CelD combine efforts to hydrolyze the same substrate molecules
Cellobiohydrolase activity (U) was determined by mixing 20 nM (2:1) and 40 nM (4:1) of Cbh1 with 10 nM of CelD or 20 nM (1:2) and 40 nM (1:4) of CelD with 10 nM of Cbh1, combined with various forms of crystalline (Figure 5A) and partially hydrated (Figure 5B) cellulosic substrates and the degree of synergy (DS = ab/a + b) was determined .
Equimolar Cbh1/CelD mixtures had no combinatorial effect (DS ≤1) on cellulose breakdown. Excess of Cbh1 over CelD resulted in a combinatorial effort (up to 345%) to attack crystalline (Figure 5A) forms of cellulose while less crystalline forms remained unchanged (Figure 5B). Excess of CelD over Cbh1 resulted in little or no gain in synergy (DS up to 1.91) for crystalline substrates (except 1:4 FP) and activity was inhibited with less crystalline substrates. For details on activity and degree of synergy, see Additional file 3: Figure S 2 and Methods.
Thus, molar excess of Cbh1 improved the ability to hydrolyze cellulose chains while excess of CelD only partially improved the activity, which could be correlated to the crystalinity (or available loose cellulose chain ends) of the substrate. When the substrate was PASC, an Avicel artificially swollen with phosphoric acid, there was no apparent combinatorial effort because of the excess number of untied cellulose chains. When the substrate was CMC, a soluble but substituted form of cellulose, there was no consequence when Cbh1 was present in excess or equimolar conditions, however activity was severely affected by the excess of CelD. Interestingly, when incubated alone, Cbh1 was far more active in CMC than was CelD (Figure 1A).
It is a widespread feature of sequenced fungal genomes to contain multiple loci that encode similar plant cell wall degrading enzymes. In Aspergillii, cellobiohydrolase genes are one such example. In A. fumigatus (or the closely related A. niveus), one GH6 cellobiohydrolase with a CBD and two GH7 cellobiohydrolases, one with and one without a CBD are present. Here we investigated the functionality of the two GH7 cellobiohydrolases and focus on whether the CBD domain is an essential domain for typical cellobiohydrolase function.
Fungi growing in the presence of cellulose (Avicel or cotton cellulose) expressed and secreted both enzymes to the extracellular medium. Thus, we transferred both genes cbh1 and celD to a controlled high-yield expression/secretion system to recover enriched and purified Cbh1 and CelD enzyme preparations useful to study their biochemical properties.
Initially we investigated enzyme thermal tolerance (Figure 3). Both enzymes were thermolabile at temperatures of 60°C and above. However Cbh1 appeared to be significantly more thermostable at lower temperatures, 40 and 50°C compared to CelD, thus indicating that the presence of a CBD provided some thermo-protection [49, 50]. We then investigated the inhibition effect of cellobiose and glucose on both enzymes (Figure 4). Both enzymes were severely inhibited by cellobiose, however Cbh1 was much less sensitive to the presence of glucose (Figure 4B). It remains unclear how the presence of a CBM could affect the inhibition by glucose. However, the opened catalytic channel in the CelD catalytic channel, could explain the high sensitivity towards the presence of glucose. Interestingly, Bu and cols  while probing absolute binding free energies for cellobiose and glucose on T. reesei GH7-cellobiohydrolase show that glucose is less stable in the catalytic channel.
We compared specific activity and other enzyme kinetic parameters using crystalline cellulosic substrates and the artificial substrate, p-nitrophenyl cellobiose (Table 2 and Additional file 3: Figure S 2). Cbh1 specific activity was between 16 and 4 fold more active on Avicel and cotton cellulose respectively, while CelD and Cbh1 exhibited identical specific activities when assayed with pNPC (Table 2). Nearly indistinguishable specific activities with pNPC highlights the fact that both enzymes have almost identical catalytic domains and the differences in activity on crystalline substrates emphasizes involvement of other structural binding features such as the CBD in Cbh1 and a lid-type open-loop structure in CelD.
Cbh1 kcat substrate turnover rates (Table 2) were 8.47 fold higher than CelD in Avicel but only 6.05 fold higher in cotton cellulose indicating that crystalinity of the substrate had a direct effect on catalytic efficiency (kcat/km), 1.22 versus 0.06 with Avicel and 0.98 versus 0.18 with cotton cellulose. The difference in substrate turnover rates was likely related to the presence of a CBD in Cbh1, which allowed the untying of cellulose chains from the original hydrogen bonded nanofiber while the lid-like open-loop feature on the CelD catalytic channel allowed binding and catalysis of already untied cellulose chains (Figure 1).
Thus, it seemed reasonable to assume that the amount of enzyme in the presence of a constant amount of crystalline substrate should have a synergistic effect for the protein that was capable of binding to the substrate. Indeed that was precisely what Figure 5A showed, where increasing molar amounts of Cbh1 favors specific activity over CelD, which performs at a lesser level. Therefore, protein concentration as well as substrate crystalinity differentially affects Cbh1 and CelD activity. Moreover, this could suggest some sort of interaction at the one-on-one substrate molecule level. Kurasin and Väljamäe measured Cel7A processivity and found cellulose hydrolysis was more than an order of magnitude lower than the values of the ratio of catalytic and dissociation rate constants, suggesting that the length of the obstacle-free path available for a processive run on a cellulose chain limits the processivity of cellobiohydrolase . Igarashi and cols found that the sliding velocity of Cel7A on crystalline cellulose was 3.5 nm/s, and interestingly, the catalytic domain without a cellulose-binding domain moved at similar rates to that of the intact enzyme . Moreover, Cel7A molecules slide along the crystalline cellulose surface and at a given point undergo collective halting [54, 55].
In our experiment, when both enzymes were mixed at equal molecular amounts, Cbh1 and CelD probably occupied all untied cellulose chains and overall activity was reduced because of the poor performance of CelD on cellulose chains that were not loose. Excess of Cbh1 molecules rescued activity, because many Cbh1 molecules initiated fresh untying cellulosic fibers, halted and changed to a new strand allowing CelD to initiate a processive run. Thus, when both enzymes are mixed together they progress into a combinatorial effort whereas Cbh1 unties substrates chains and CelD hydrolyzes these cellulosic chains. When CelD was present in excess, the effect could only be partially overcome while CelD could not initiate new loose cellulosic chains only used the ones that were already available. Hence, the recovery of excessive CelD was dependent on the crystalinity of the substrate, less pronounced on CC than AV and FP showing little to no combinatorial effort effect with PASC, an artificially swollen Avicel. Activity of cellobiohydrolases on soluble (CMC) and partially hydrated substrates has been reported .
The two cellobiohydrolases investigated in this study are similar in amino acid sequence differing mainly by the presence of a cellulose-binding domain in Cbh1 which makes this enzyme a substrate bound and CelD a soluble substrate unbound enzyme. Both enzymes have similar catalytic properties however differ in thermostability, inhibition by glucose and protein concentration dependent specific activity. The fact that Cbh1 binds to its substrate and specific activity is dependent on protein concentration suggests that both enzymes employ a combinatorial effort in attacking the crystalline forms of cellulose.
Cellulosic and hemicellulosic substrates were purchased from the best source possible, Sigma Aldrich, MO and Megazyme, UK. For synthesis of APTS-labeled cellopentaose 1 mg of cellopentaose, β-D-Glc-[1 → 4])4-D-Glc, D(+)-cellopentaose (Sigma Aldrich, MO) was mixed with 10 μl of 10 mg APTS (8-aminopyrene-1,3,6-trisulfonic acid trisodium salt) in 200 μl of 25% acetic acid and 10 μl of 1 M sodium cyanoborohydride in DMSO, heated at 80°C for 60 min and purified as described in . The cellulosic substrates used throughout were carboxymethylcellulose (CMC), cotton linters, SigmaCel50 (CC) crystalinity index (CI) of 91.2, Avicel PH-101 91.7 CI as determined by the x-ray diffraction method , phosphoric acid swollen-Avicel (PASC) and filter paper, Whatman #3 (FP). Proteins were quantified by the Bradford method , validated for purity by SDS-PAGE  and used for biochemical studies.
Construction of pEXPYR-client protein plasmids
The pEXPYR Aspergillus “shuttle” expression plasmid for expression and secretion of client proteins was used . PCR-amplified gene-fragments (for primers of gene models see Additional file 3: Figure S 2) were digested with NotI and XbaI, isolated by gel excision of a thin-slice from a 0.8% agarose electrophoresis gel, purified with QIAquick Gel Extraction kit (Quiagen), ligated onto NotI/XbaI digested pEXPYR plasmid with T4-fast ligase (Promega, WI) and transformed into Ca+ competent Escherichia coli TOP 10 F’ competent cells (Invitrogen, CA). Random ampicillin-resistant colonies were selected and grown in 5 ml LB-ampicillin broth, plasmids purified , restricted with NotI/XbaI and insert size verified by 1% agarose gel electrophoresis . Plasmids with the correct insert size DNA were fully sequenced at the Oklahoma State University Core Facility and clones with the correct DNA sequence used for transformation. Recombinant pEXPYR-Cbh1 or pEXPYR-CelD plasmid was introduced through integrative transformation into the A. nidulans strain FGSC A773 (pyrG pyroA) genome  and recombinants selected on MM supplemented with 1 mM pyridoxine and 100 μg/ml of zeocin. Five pyrG + , zeocin resistant transformants were grown on 10 ml MM, pyridoxine and 5% maltose containing plates for 48 h at 37°C. Accumulation of Cbh1 and CelD in the medium was analyzed by SDS-PAGE and one transformant for each enzyme was used for further investigation.
Production and secretion of client proteins
107-108 spores/ml were inoculated in liquid minimal medium supplemented with 0.5 to 15% of maltose, distributed onto dishes (20 ml in 150 mm Petri-dishes and 500 ml onto cafeteria trays) and incubated (stationary) at 37°C for 2–3 days. The mycelial mat was lifted with a spatula and discarded and the medium collected by filtration, centrifuged at 10,000xg for 10 min prior to concentration by ultra-filtration (10,000 kDa cutoff Amicon), quantified by the Bradford method , validated for purity by SDS-PAGE  and used for biochemical studies.
For biochemical studies, the recombinant Cbh1 and CelD proteins were produced by heavy inoculation of fresh conidia onto Petri dishes containing MM supplemented with pyridoxine and 5% maltose and incubated at 37°C for 48 h. Cbh1 and CelD were routinely recovered with this stationary incubation method and proteolysis avoided due to the 2 day incubation period. The medium was harvested by filtration, centrifuged at 10,000xg and concentrated by ultra filtration on Amicon 10 kDa cut-off micro columns. The majority of the protein content recovered from culture filtrates was Cbh1 or CelD.
Crude ultra filtrated protein extracts were resolved by SDS-PAGE, Comassie-blue stained (CB) and the SDS removed by successive washes with 25% isopropanol solution. After transferred to 50 mM ammonium acetate buffer pH 5.0 a 1% carboxymethylcellulose (CMC) solution was infused into the SDS-free polyacrylamide gel, incubated at 37°C for 120 min, and stained with Congo red [65–68].
Cbh1 and CelD were purified by two steps. The concentrated and dialyzed protein samples (500 μl aliquots) were applied to ion exchange Resource Q® column equilibrated with 20 mM sodium phosphate buffer, pH 7.4 and proteins eluted with a linear 0 to 1 M sodium chloride gradient (Äkta Purifier, GE). Fractions active on pNPC were collected and loaded onto a Superdex G-75® (10x30 mm) gel filtration column, equilibrated with 50 mM ammonium acetate buffer, pH 5.0 and eluted fractions showing enzymatic activity were analyzed by SDS-PAGE. Single band containing fractions were combined, concentrated and used for further biochemical analysis. The flow rate used for both chromatographic steps was 0.5 ml min-1. Purified Cbh1 and CelD fractions were validated by SDS-PAGE (Additional file 1: Table S 1), and pNPC, pNPG activity measurement comparisons (Additional file 3: Figure S 2).
Optimal pH, temperature and thermostability
Optimal pH was measured at 50°C in the presence of 18 mM pNPC with the pH ranging from pH 3.0 to pH 8.0 using 50 mM phosphate/citrate buffer. Optimal enzyme operating temperatures for Cbh1 and CelD were measured at their optimal pH, 5.0, with temperatures ranging from 30°C to 80°C. Thermal stability of Cbh1 and CelD was tested at optimal pH and exposure for various times (up to one hour). Purified enzyme in 50 mM ammonium acetate (without substrate) was incubated at temperatures ranging from 40 to 70°C. Samples were drawn from a master mix and residual activity assayed with 18 mM pNPC.
Cellulose specific CelD or Cbh1, CBD-dependent binding
To reveal the functionality and specificity of the predicted CBM1 domain, the binding of Cbh1 or CelD was evaluated by a pull-down assay. 20 μg of Cbh1 or CelD was incubated at 4 C for 30 min in a rotary shaker, with 200 μl of 30% cotton cellulose or Avicel slurry in 50 mM, pH 5.0, ammonium acetate buffer. The reaction was centrifuged at 14,000xg for 15 min, supernatant (free fraction) collected and concentrated in an Amicon, 10 kDa cutoff ultra filtration column mixed with 2X Laemmli buffer and subjected to SDS-PAGE. The bound fraction was released from the Avicel or cotton linter or Avicel slurry by addition of 40 μl of 2X Laemmli buffer, vigorous agitation and boiling for 10 min. The SDS protein-solubilized slurry was centrifuged and supernatant (bound fraction) subjected to SDS-PAGE.
Cellobiohydrolase activities, substrate- and protein-dependent kinetics
Substrate specificity of cellobiohydrolases was determined by incubating 1 μg of Cbh1 or CelD with a, 1% slurry of cotton cellulose (Sigmacell 50), Avicel PH-101, PASC or 1% solution of CMC or 18 mM pNPCellobiose, incubated for 120 min or as indicated at 50°C and the release of reducing sugars determined with the DNS method . Specific activity was defined as U per mg protein at 50°C whereas U was the amount of enzyme that produced 1 μmole of reducing sugar (glucose or cellobiose) per minute. Activity towards starch, polygalacturonic acid, wheat arabinoxylan, arabinan from sugar beet, xylan birchwood, xylan beechwood and xyloglucan from tamarind could not be detected and is not shown.
Michaelis-Menten kinetic constants were determined from Lineweaver–Burk plots. Reaction rates were measured using Avicel and cotton cellulose ranging from 0 to 100 mg/ml of insoluble substrate suspended in 50 mM ammonium acetate buffer pH 5.0. Reactions were carried out over a 120-min period at 50°C, boiled, and the released reducing sugars determined by DNS assay . The raw substrate activity plots are shown in Additional file 4: Figure S 3.
The degree of synergy (D.S.) was calculated by dividing the activity when Cbh1 and CelD were incubated together (ab) by the sum of the activity when Cbh1 and CelD were incubated separately (a + b) as proposed by . Enzyme mixtures with dissimilar molar amounts (e.g., 4xCbh1:1xCelD), the individual activities were adjusted correspondingly (4a + 1b). A value of >1 indicated synergy and a value below 1 indicated antagonist competition for the same substrate molecules.
Homology molecular modeling
The atomic coordinates of the cellobiohydrolase I (CBH IB) from Talaromyces emersonii (PDBID: 1Q9H, ) was used as template for generating structural models of both CBHI and CelD by restraint-based modeling as implemented in the program MODELLER . To guarantee sufficient conformational sampling, an ensemble of 50 models was built, from which the best final model was selected based on evaluation of stereo chemical values from MOLPROBITY , the objective function from MODELLER (DOPE score) and by visual inspection. Those models were then minimized using the steepest descent minimization algorithm as implemented in the UCSF chimera software . Incomplete side-chains were replaced using the Dunbrack rotamer library .
Cellobiose and glucose Inhibition
To analyze Cbh1 and CelD inhibition, 1 μg enzyme was incubated in 18 mM pNPC dissolved in 50 mM ammonium acetate buffer pH 5.0 with glucose or cellobiose added in a range from 0 to 100 mM at 50°C. After 15 min, enzyme activity was stopped by adding 100 μl of a 2% Na2CO3. The pNPC chromophore release was spectrophotometrically quantified at 410 nm with a Multimode Infinte M200 Reader (Tecan, SC).
We are indebted for insightful discussions with Dr. Babu Fathepure and Dr. Richard J Ward, the expert enzyme kinetic analysis prepared by Junio Cota, at the Brazilian Bioethanol Science and Technology Laboratory and generous funding from the Oklahoma Bioenergy Center and Department of Energy, awards 06103-OKL and ZDJ-7-77608-01 and FAPESP award 2010/18198-3.
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