Production of four Neurospora crassa lytic polysaccharide monooxygenases in Pichia pastoris monitored by a fluorimetric assay
© Kittl et al.; licensee BioMed Central Ltd. 2012
Received: 31 July 2012
Accepted: 22 October 2012
Published: 26 October 2012
Recent studies demonstrate that enzymes from the glycosyl hydrolase family 61 (GH61) show lytic polysaccharide monooxygenase (PMO) activity. Together with cellobiose dehydrogenase (CDH) an enzymatic system capable of oxidative cellulose cleavage is formed, which increases the efficiency of cellulases and put PMOs at focus of biofuel research. Large amounts of purified PMOs, which are difficult to obtain from the native fungal producers, are needed to study their reaction kinetics, structure and industrial application. In addition, a fast and robust enzymatic assay is necessary to monitor enzyme production and purification.
Four pmo genes from Neurospora crassa were expressed in P. pastoris under control of the AOX1 promoter. High yields were obtained for the glycosylated gene products PMO-01867, PMO-02916 and PMO-08760 (>300 mg L-1), whereas the yield of non-glycosylated PMO-03328 was moderate (~45 mg L-1). The production and purification of all four enzymes was specifically followed by a newly developed, fast assay based on a side reaction of PMO: the production of H2O2 in the presence of reductants. While ascorbate is a suitable reductant for homogeneous PMO preparations, fermentation samples require the specific electron donor CDH.
P. pastoris is a high performing expression host for N. crassa PMOs. The pmo genes under control of the native signal sequence are correctly processed and active. The novel CDH-based enzyme assay allows fast determination of PMO activity in fermentation samples and is robust against interfering matrix components.
Hydrolysis is still a major cost factor in the production of second-generation biofuels from lignocellulosic biomass, considering the costs of currently available enzyme mixtures. Until recently, only hydrolytic enzymes were thought to play a role in the degradation of recalcitrant cellulose and hemicelluloses to fermentable sugars. The finding that enzymes from glycosyl hydrolase family 61 (GH61) in combination with the flavocytochrome cellobiose dehydrogenase (CDH) enhance the action of hydrolytic enzymes added a new dimension to the classical concept of cellulose degradation, as recently reviewed by Horn et al.. These copper-dependent enzymes were shown to cleave cellulose by an oxidative mechanism provided that reduction equivalents from CDH or low molecular weight reducing agents (e.g. ascorbate) are available[2–4]. GH61 enzymes with this confirmed activity have been termed polysaccharide monooxygenases (PMOs) or, to indicate the reaction mechanism more specificly, lytic PMOs. Recently, it was shown that enzymes of the structurally similar CBM33 family are also capable of cleaving cellulose. Unlike lytic PMOs they are mainly found in bacteria, but also in other eukaryotes.
The CDH/PMO system was shown to improve the degradation of cellulose in combination with cellulases in several studies[2–4, 6, 7]. In the proposed reaction mechanism CDH donates an electron via its cytochrome domain to the type-2 copper in the PMO active site. There, oxygen is partially reduced and attacks the pyranose ring of the glucose moieties at the C-1 (class-1 PMOs) or C-4 (class-2 PMOs) position, thereby destabilizing the adjacent glycosidic bond and breaking it by an elimination reaction[3, 8]. The occurrence of gh61 (pmo) genes has been confirmed in many cellulolytic fungi. In some genomes, gh61 genes even outnumber cellulase genes[9–11]. It remains to be elucidated whether all of these encoded enzymes have PMO activity, but their large number emphasizes the importance of oxidative cellulose cleavage. In the cellulolytic ascomycete Neurospora crassa, 2 cdh genes and 14 pmo genes are present. When the fungus is grown on Miscanthus one cdh gene and 8 pmo genes are upregulated. In our study the expression of four of these pmo genes in Pichia pastoris is investigated. Only few examples of heterologous expression of pmo genes are hitherto known. A PMO from Thermoascus aurantiacus was expressed in Trichoderma reesei by Novozymes, and three PMOs (from Phanerochaete chrysosporium, Sporotrichum thermophile and Aspergillus kawachii) were produced in P. pastoris with the α-factor signal sequence for secretion[7, 12, 13]. Although the expressions and purifications were successful, none of these studies reported production or purification yields. One reason for that is certainly the lack of a fast and robust assay for PMO activity. The assays currently used to determine PMO activity are based on a lengthy incubation of the PMO together with CDH or a reductant such as ascorbate and the polymeric cellulosic substrates Avicel, phosphoric acid swollen cellulose (PASC), carboxymethyl cellulose (CMC) or nano-fibrillated cellulose at elevated temperatures (37°C – 50°C) for several hours or up to three days. Subsequently, the reaction products are analyzed by MALDI-TOF/MS, HPAEC or LC/MS[3, 5, 6, 12–14]. While these time-consuming and labor-intensive assays are a good choice for investigating the reaction mechanism and substrate specificity, they are of little use for monitoring fermentation processes or the progress of a purification procedure. The proposed reaction mechanism points to the possibility that hydrogen peroxide is generated as a by-product of a futile side reaction, which might occur in the absence of cellulose as a substrate. Li et al. found that peroxide can be modeled well in an electron density at the coordination site of the type-2 copper in N. crassa PMO-3 [GenBank: NCU07898]. In this work, we measure the oxygen reducing side reactivity of PMOs by quantifying the formation of hydrogen peroxide. Based on this observation a fast and robust enzymatic assay for PMO activity was developed and tested on the expression and purification of four PMOs from N. crassa.
Results and discussion
Polysaccharide monooxygenase activity assay
Contribution of background activities of CDH IIA and ascorbic acid, respectively, to the total activity measured with various concentrations of PMO (PMO-02916)
5.39 ± 0.23
3.61 ± 0.22
2.34 ± 0.05
1.05 ± 0.01
0.63 ± 0.02
0.49 ± 0.02
6.21 ± 0.22
2.57 ± 0.11
1.77 ± 0.11
0.94 ± 0.05
0.74 ± 0.05
0.70 ± 0.04
We employed CDH IIA from N. crassa for this assay, which is the major CDH in the N. crassa secretome and which was recombinantly produced in P. pastoris. A possible direct interaction of PMO with cellobiose, the electron donor of CDH, was tested by replacing cellobiose by lactose. Lactose interacts almost equally well with Nc CDH IIA and did not alter the peroxide formation by PMOs.
PMO activity in fermentation samples can only be determined with the CDH-driven assay, since the strong reducing agent ascorbate interacts unspecifically with other oxidases present in the P. pastoris culture supernatant (e.g. alcohol oxidase 1) and media components, generating a strong background signal. To exclude the effects of media components (e.g. copper ions) on the CDH-based assay, sample aliquots were additionally centrifuged in mini-spin columns with 10-kDa cut-off, and the blank activity of the permeate was subtracted. A possible interference by proteins in the P. pastoris culture supernatant on the assay was examined by measuring background activities of culture supernatant and its permeate (10 kDa cut-off) of two similar fermentations of a P. pastoris strain without PMO. The media components in the permeate increase the blank activity significantly (1.6 times). The increase of the background H2O2 production by the culture supernatant is 1.7 fold (Additional file2). This shows that compounds with a molecular mass above 10 kDa (proteins) have also a limited effect. The estimated error for using the permeate as blank experiment instead of the full sample matrix is 8-10%, which seems acceptable for the monitoring of PMO activity in fermentations.
Recombinant production of PMO-01867, PMO-02916, PMO-03328 and PMO-08760
Fermentation of PMO enzymes
Purification of recombinant PMOs
Purification scheme of recombinant PMO enzymes
Total activity [U]
Specific Activity [U g-1]
Culture supernatant (2.5 L)
Culture supernatant (2.1 L)
Culture supernatant (2.9 L)
Culture supernatant (2.3 L)
The specific activities of the pure enzymes differ according to the method used to determine the protein concentration. When using the absorbance at 280 nm and the calculated molecular absorption coefficient, the specific activities are lower by a factor of 1.54 (PMO-01867), 2.1 (PMO-02916), 0.93 (PMO-03328) and 1.99 (PMO-08760) compared to the Bradford method for protein quantification with a BSA standard curve. This obviously reflects the inaccuracy of the Bradford assay for glycosylated proteins.
Molecular masses and glycosylation of PMO enzymes
Putative glycosyl. sites
39.6 ± 0.7
57.5 ± 0.8
23.7 ± 0.6
41.4 ± 0.4
Spectral properties of investigated PMO enzymes
Specific activities of PMO enzymes
Reductant/Specific activity [U g-1]
1.07 ± 0.13
4.00 ± 0.18
1.82 ± 0.04
4.47 ± 0.15
15.46 ± 0.97
12.08 ± 0.67
1.53 ± 0.19
4.84 ± 0.13
P. pastoris was found to be a suitable expression host for glycosylated and non-glycosylated N. crassa PMO enzymes. A further advantage of this host is the correct processing of the N. crassa signal sequences of PMOs. This feature is especially important when considering that the active site of PMOs contains the N-terminal histidine. The PMO enzymes expressed in this study reduce oxygen and show typical type-2 copper absorption spectra, which supports the finding that these enzymes are in fact copper-containing PMOs. The newly developed CDH-driven activity assay allows fast PMO activity measurements in fermentation samples and is robust against interfering matrix components. With an easy assay for the target protein available, optimization of fermentation and purification conditions will be greatly simplified and should substantially improve the yields. Additionally, it allows the establishment of a high-throughput screening for improved activity enabling for example a screening for better producing P. pastoris clones. The second assay using ascorbate as reducing agent provides a tool to measure PMO activity independently from the interaction between PMO and its electron donor CDH, but it is too prone to interference to be used in crude enzyme samples such as fermentation media. Recombinant production of PMO enzymes should enable the formulation of more efficient, cost-effective enzyme solutions for biofuel production from lignocellulosic biomass.
Chemicals and microorganisms
All chemicals were of the highest purity grade available and were purchased from Sigma-Aldrich unless stated otherwise. Amplex Red (10-acetyl-3,7-dihydroxyphenoxazine) was purchased from VWR. Restriction endonucleases and T4 DNA ligase were obtained from Fermentas and were used as recommended by the manufacturer. Nucleic acid amplifications were done employing GoTAQ DNA Polymerase (Promega), dNTP mix, oligonucleotide primers (HVD Life Sciences, Vienna, Austria) and a C-1000 thermocycler (Bio-Rad Laboratories). E. coli strain DH5α (Invitrogen) was used for subcloning. The gene coding for a PMO enzyme from N. crassa (GenBank:NCU02916, gh61-3) has been previously expressed and purified by Sygmund et al.. To simplify the denomination and also prevent confusion caused by the newly discovered polysaccharide monooxygenase function, we suggest to name this enzyme PMO-02916 on the basis of its GenBank number, and the other PMOs accordingly PMO-01867 (GenBank:NCU01867, gh61-10, PMO-03328 (GenBank:NCU03328, gh61-6), and PMO-08760 (GenBank:NCU08760, gh61-5). The N. crassa genes encoding the PMO enzymes were codon-optimized for expression in P. pastoris (Additional file5) and commercially synthesized by Invitrogen including their native signal sequences. P. pastoris strain X-33 and the vector pPICZα A are components of the Pichia Easy Select Expression System from Invitrogen. CDH IIA was produced and purified as previously reported.
Amplex Red/horseradish peroxidase assay
The oxygen reactivity of PMOs was measured by a time resolved quantification of H2O2 formation in 96-well plates (total volume of 200 μL) using a Perkin Elmer EnSpire Multimode plate reader. All reactions were performed in 100 mM sodium phosphate buffer, pH 6.0 at 22°C. Based on preliminary studies ascorbate and CDH were used in concentrations of 30 μM and 0.3 μM (0.025 mg mL-1), respectively to prevent a limitation in the PMO reduction step. As electron donor for CDH 500 μM cellobiose was used. When ascorbate was used as reductant, it was added to a final concentration of 30 μM and enzyme assays were started by mixing 20 μL of the respective PMO with 180 μL of the ready-made assay solution containing 30 μM ascorbate, 50 μM Amplex Red and 7.14 U mL-1 peroxidase in 96-well plate wells. In reference experiments without PMO the background signal (H2O2 production by CDH) was measured and subtracted from the assays. When CDH was used as reductant, the PMO assays were started by mixing 20 μL of sample solution and 20 μL CDH solution with 160 μL of the reaction mix containing cellobiose, Amplex Red and peroxidase. Initial fluorescence scans of resorufin showed highest signal intensities and lowest interference with matrix compounds when using an excitation wavelength of 569 nm and an emission wavelength of 585 nm for the selected conditions. The stoichiometry of H2O2 conversion to resorufin formation is 1:1. By using a high concentration of Amplex Red (50 μM) the linearity of the detection reaction was ensured and the molar ratio of Amplex Red:H2O2 was always greater than 50:1. The H2O2 concentration in the assays was far below 1 μM, which leads to a linear concentration/activity response of horseradish peroxidase, which has a KM for H2O2 of 1.55 μM. The high final activity of horseradish peroxidase (7.14 U mL-1) assures a fast conversion of the formed H2O2 and prevents the final reaction to be rate limiting. Additionally, it prevents the accumulation of H2O2, which could have detrimental effects on enzyme stability in the assay. The stability of resorufin fluorescence under these conditions was tested by addition of varying concentrations of hydrogen peroxide (0.1 – 5 μM) to the assay. A stable signal that remained constant throughout the measured period of 45 minutes was observed and maximum signal intensity was reached already during the mixing period before starting the assay. A linear relation between fluorescence and H2O2 concentrations in the range of 0.1 – 2 μM H2O2 was observed and the slope (28450 counts μmol-1) was used for the calculation of an enzyme factor to convert the fluorimeters readout (counts min-1), into enzyme activity. PMO activity was defined as one μmol H2O2 generated per minute under the defined assay conditions.
Construction of pmo expression vectors for P. pastoris
The synthetic pmo genes were digested with the restriction enzymes Bst BI and Xba I and ligated into the equally treated vector pPICZα A using the Rapid DNA Ligation Kit from Fermentas. The procedures resulted in plasmids carrying genes encoding proteins with their native signal sequences cloned under the control of the methanol inducible AOX1 promoter. C-terminal tags for purification or antibody detection were omitted. The plasmids were linearized with the restriction enzyme Sac I and transformed into electro-competent P. pastoris cells. Transformants were grown on YPD plates (10 g L-1 yeast extract, 20 g L-1 peptone, 10 g L-1 glucose and 15 g L-1 agar) containing 100 mg L-1 zeocin.
Enzyme production and purification
The enzymes were produced in a 7-L bioreactor (MBR) with a starting volume of 3 L Basal Salts Medium (21 mL L-1 H3PO4 (85%); 0.93 g L-1 CaSO4·2H2O; 14.9 g L-1 MgSO4·7H2O; 18.2 g L-1 K2SO4; 4.13 g L-1 KOH; 4% (v/v) glycerol) supplemented with 4.35 mL L-1 PTM1 trace salts (Invitrogen), 1 mL Antifoam 204 (Sigma) and 0.1 mM CuSO4. After sterilization, the pH of the medium was adjusted to pH 5.0 with 28% ammonium hydroxide and maintained at this level throughout the entire fermentation process. The fermentations were started by adding 400 mL preculture grown on YPD medium in 1-L baffled shaking flasks at 125 rpm and 30°C overnight. The cultivations were performed according to the Pichia Fermentation Process Guidelines of Invitrogen with some alterations. After depletion of glycerol in the batch medium the fed-batch phase was started with a constant feed of 36 mL h-1 50% glycerol containing 12 mL L-1 PTM1 trace salts overnight to increase biomass. For induction the feed was switched to 100% methanol containing 12 mL L-1 PTM1 trace salts at an initial feed rate of 12 mL h-1 until the culture was fully adapted to methanol. Subsequently the feed rate was adjusted to keep the dissolved oxygen saturation constant at 4% at a constant air supply of 6 L min-1 and a stirrer speed of 800 rpm. After induction the cultivation temperature was reduced to 25°C. Samples were taken regularly and wet biomass, protein concentration and PMO activity were measured.
The fermentation broth was centrifuged at 6400 × g and 4°C for 30 min, and ammonium sulphate was slowly added to the clear culture supernatant to give a 30% saturated solution. Precipitate was removed by centrifugation (6400 × g; 20 min at 4°C) and the clear supernatant was loaded onto a 600-mL PHE-Sepharose Fast Flow column (chromatographic equipment and materials from GE Healthcare Biosciences) equilibrated with a 25 mM sodium acetate buffer, pH 5.0, containing 30% ammonium sulphate saturation. Proteins were eluted within a linear gradient from 30 to 0% ammonium sulphate within 3 column volumes and fractions were collected. Fractions containing the respective PMO were pooled and diafiltered with a hollow fiber cross-flow module (Microza UF module SLP-1053, 10 kDa cut-off, Pall Corporation). The diafiltered pools (conductivity < 1.4 mS cm-1) were applied to a 20-mL column packed with Q15-Source equilibrated with 20 mM TRIS/HCl buffer, pH 8.0. The flow-throughs were collected and contained the PMO enzymes. The solutions were concentrated, first by diafiltration with a Vivaflow cross-flow module (Millipore, cut-off 10 kDa) and subsequently by centrifugation in Amicon centrifugation tubes (Millipore, cut-off 10 kDa, 3200 x g, 15 min at 4°C). Size exclusion chromatography was done with a Superdex 75 column (Pharmacia Biotech) equilibrated with 20 mM TRIS/HCl, pH 8.0 buffer. Fractions containing pure PMO enzyme were pooled and stored at 4°C.
Measurement of cellulolytic activity of PMOs
Microcrystalline cellulose was incubated with the PMOs and analyzed for soluble cello-oligosaccharides. The cellulose concentration in 50 mM sodium phosphate buffer, pH 6.0 was 25 mg mL-1 and PMOs were employed at a concentration of 5 mg g-1 cellulose. CDH IIA was used as reductant for PMO at a concentration of 2 mg g-1; cellulose and lactose (200 μM) were used as electron donors for CDH. A total volume of 400 μL was incubated at 30°C for 72 hours on a shaking incubator operated at 320 rpm and 90% humidity. The reaction tubes were sealed with an air permeable membrane to ensure continuous oxygen supply. Negative controls without CDH IIA and PMOs were performed under the same conditions. After incubation the samples were heated to 95°C for 10 min and protein precipitate and remaining cellulose was removed by centrifugation. Released cellooligosaccharides were detected by HPLC using a CarboPac PA100 Carbohydrate Column equipped with a guard column and an ED40 electrochemical detector (all equipment from DIONEX). The column was equilibrated with 150 mM NaOH (solution A) and reaction products were eluted by a linear gradient from 0 to 30% solution B (500 mM sodium carbonate in solution A) within 25 minutes at a flow rate of 0.5 mL min-1. The column was washed with 100% solution A for 5 minutes and re-equilibrated for 13 minutes with solution B before starting the next run.
Protein contents of crude preparations or partially purified fractions were determined by the dye-binding method of Bradford using a pre-fabricated assay (Biorad Laboratories) and BSA as calibration standard. Protein concentrations of purified samples were measured based on their extinction coefficient at 280 nm (PMO) or 420 nm (CDH IIA).
Spectra of homogenously purified PMO enzymes were recorded with a Hitachi U-3000 spectrophotometer at room temperature. Appropriately diluted enzymes (Abs ~1 at 280 nm) were measured from 250 to 800 nm at a scan speed of 60 nm min-1. To assess the extinction coefficients of the type-2 copper sites, spectra of concentrated enzymes were recorded from 450 to 800 nm at a scan speed of 30 nm min-1. Spectra of reduced species were obtained by addition of an excess of ascorbate to the cuvette. The molar absorption coefficients at 280 nm for all enzymes were calculated using the mature amino acid sequence and the program ProtParam (http://web.expasy.org/protparam/).
PMO enzymes (1.5 mg mL-1, 20 μL) were treated with PNGase F under denaturing conditions. The enzymes were mixed with 2 μL of glycoprotein denaturing buffer (80.5% sodium dodecylsulfate, 1% mercaptoethanol) and incubated at 99°C for 10 min. Then, 4 μL of 0.5 M sodium phosphate buffer, pH 7.5, 4 μL of nonylphenol (40%) and 1 μL of PNGase F were added to the reaction mix and incubated at 37°C for 1 hour. Glycosylation sequons were predicted using the servers NetNGlyc 1.0 (http://www.cbs.dtu.dk/services/NetNGlyc/) and NetOGlyc 3.1 (http://www.cbs.dtu.dk/services/NetOGlyc/).
Mini-PROTEAN TGX precast gels (Bio-Rad Laboratories) with a gradient from 4 – 20% were used for SDS-PAGE analysis of purified enzymes and fermentation supernatants. Protein bands were visualized by staining with Bio-Safe Coomassie, and unstained Precision Plus Protein Standard was used for mass determination. All procedures were done according to the manufacturer’s recommendations (Bio-Rad Laboratories). Three independent gels were used for molecular mass determination of PMO enzymes.
Differential scanning calorimetry
Thermodynamic stabilities of PMO enzymes were measured by differential scanning calorimetry (DSC) using a MicroCal VP-DSC instrument with an autosampler (MicroCal, Northampton, MA). Enzyme concentrations were adjusted to 1 mg mL-1 based on their molar extinction coefficients at 280 nm. All measurements were carried out in 100 mM phosphate buffer, pH 6.0. A linear temperature ramp from 20 to 100°C was applied at a scan rate of 1 K per min. Prior to all measurements, samples were degassed by continuous stirring in vacuo for 15 min. Obtained sample thermograms were corrected for the calorimeter baseline by subtracting a buffer blank that was scanned in the reference chamber. Transition midpoint temperatures (TM) of the enzymes were determined from the peak maximum of transition using the Origin 7.5 software (Origin Lab Corporation, Northampton, MA).
Family 61 glycoside hydrolase
Phosphoric acid swollen cellulose
This work has been supported by the doctoral program “BioTop – Biomolecular Technology of Proteins” (FWF W1224) of the Austrian Science Fund FWF.
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