Synergistic Action of a Lytic Polysaccharide Monooxygenase and a Cellobiohydrolase from Penicillium funiculosum in Cellulose Saccharification under High-Level Substrate Loading

The enzymatic hydrolysis of cellulosic biomass by cellulases continues to be a significant bottleneck in the development of second-generation biobased industries. While increasing efforts are being made to obtain indigenous cellulases for biomass hydrolysis, the high production cost of this enzyme remains a crucial challenge affecting its wide availability for the efficient utilization of cellulosic materials. This is because it is challenging to obtain an enzymatic cocktail with balanced activity from a single host. This report describes the annotation and structural analysis of an uncharacterized lytic polysaccharide monooxygenase (LPMO) gene in Penicillium funiculosum and its impact on biomass deconstruction upon overexpression in a catabolite-derepressed strain of P. funiculosum. Cellobiohydrolase I (CBH1), which is the most important enzyme produced by many cellulolytic fungi for the saccharification of crystalline cellulose, was further overexpressed simultaneously with LPMO. The resulting secretome was analyzed for enhanced LPMO and exocellulase activities and the corresponding improvement in saccharification performance (by ∼20%) under high-level substrate loading using a minimal amount of protein.

ABSTRACT

Lytic polysaccharide monooxygenases (LPMOs) are crucial industrial enzymes required in the biorefinery industry as well as in the natural carbon cycle. These enzymes, known to catalyze the oxidative cleavage of glycosidic bonds, are produced by numerous bacterial and fungal species to assist in the degradation of cellulosic biomass. In this study, we annotated and performed structural analysis of an uncharacterized LPMO from Penicillium funiculosum (PfLPMO9) based on computational methods in an attempt to understand the behavior of this enzyme in biomass degradation. PfLPMO9 exhibited 75% and 36% sequence identities with LPMOs from Thermoascus aurantiacus (TaLPMO9A) and Lentinus similis (LsLPMO9A), respectively. Furthermore, multiple fungal genetic manipulation tools were employed to simultaneously overexpress LPMO and cellobiohydrolase I (CBH1) in a catabolite-derepressed strain of Penicillium funiculosum, PfMig1⁸⁸ (an engineered variant of P. funiculosum), to improve its saccharification performance toward acid-pretreated wheat straw (PWS) at 20% substrate loading. The resulting transformants showed improved LPMO and CBH1 expression at both the transcriptional and translational levels, with ∼200% and ∼66% increases in ascorbate-induced LPMO and Avicelase activities, respectively. While the secretome of PfMig⁸⁸ overexpressing LPMO or CBH1 increased the saccharification of PWS by 6% or 13%, respectively, over the secretome of PfMig1⁸⁸ at the same protein concentration, the simultaneous overexpression of these two genes led to a 20% increase in saccharification efficiency over that observed with PfMig1⁸⁸, which accounted for 82% saccharification of PWS under 20% substrate loading.

IMPORTANCE The enzymatic hydrolysis of cellulosic biomass by cellulases continues to be a significant bottleneck in the development of second-generation biobased industries. While increasing efforts are being made to obtain indigenous cellulases for biomass hydrolysis, the high production cost of this enzyme remains a crucial challenge affecting its wide availability for the efficient utilization of cellulosic materials. This is because it is challenging to obtain an enzymatic cocktail with balanced activity from a single host. This report describes the annotation and structural analysis of an uncharacterized lytic polysaccharide monooxygenase (LPMO) gene in Penicillium funiculosum and its impact on biomass deconstruction upon overexpression in a catabolite-derepressed strain of P. funiculosum. Cellobiohydrolase I (CBH1), which is the most important enzyme produced by many cellulolytic fungi for the saccharification of crystalline cellulose, was further overexpressed simultaneously with LPMO. The resulting secretome was analyzed for enhanced LPMO and exocellulase activities and the corresponding improvement in saccharification performance (by ∼20%) under high-level substrate loading using a minimal amount of protein.

INTRODUCTION

Lignocellulosic biomass continues to attract attention worldwide as an inexpensive source of energy owing to its relatively high abundance, renewable nature, and wide availability in the environment as a low-cost substrate (1, 2). Cellulases and hemicellulases are vital enzymes for breaking down the glycosidic linkages present in the carbohydrate portion of lignocellulosic biomass and making it available for the production of various biofuels and biochemicals (3). These enzymes are mainly produced by filamentous fungi and perform a central role in the global carbon cycle by degrading insoluble cellulose to soluble sugars (4–6). Over time, the secretomes of many fungal genera, such as Trichoderma, Aspergillus, Neurospora, and Penicillium, have been studied for their ability to produce cellulolytic enzymes. Although the secretome of Trichoderma reesei, which serves as a model fungus for cellulase production, is widely used in commercial cocktails, complete saccharification of cellulosic biomass by this secretome is usually not achievable. This is because the secretome of T. reesei under cellulase-inducing conditions contains mainly cellobiohydrolase I (CBH1) (EC 3.2.1.176) and cellobiohydrolase II (EC 3.2.1.91), which constitute approximately 80% of the total secreted proteins (7), while other cellulase components, such as β-glucosidase (BGL) (EC 3.2.1.21) and endoglucanase (EG) (EC 3.2.1.4), which participate in biomass hydrolysis, are secreted at lower levels and have to be produced by other fungal sources to complement the cellobiohydrolases in commercial cocktails (7–9).

Furthermore, while increasing attention is being paid to the fortification of the secretomes of many cellulase-producing strains for improved biomass hydrolysis, recent efforts have identified the significant roles of some synergistic or auxiliary proteins, such as expansins, swollenin, and hydrophobins, in promoting cellulase activity (10–12). Most notably, lytic polysaccharide monooxygenase (LPMO), an enzyme that is auxiliary to cellulases, has been reported to significantly reduce the total cellulase loading needed to achieve the ∼80 to 90% cellulose conversion required by industries for bioethanol production (13–15). LPMOs (EC 1.14.99.53 to -56), a family of recently discovered auxiliary proteins, are unique in terms of their ability to catalyze the cleavage of glycosidic bonds of polysaccharides via an oxidative mechanism rather than hydrolytic means. The mechanism of cellulose cleavage by LPMOs involves the reduction of Cu²⁺ at the active site by some redox partners, including synthetic reducing agents such as ascorbic acid, gallate, and reduced glutathione (16–18); lignin, which naturally exists in lignocellulose (19, 20); or enzymes such as cellobiose dehydrogenase (CDH) and glucose methanol-choline oxidase/dehydrogenase (20–23). This is followed by the binding of a cosubstrate such as O₂ or H₂O₂, which introduces an oxygen atom that consequently attacks the pyranose ring of the glucose moieties at the C-1 or C-4 position, thereby destabilizing the adjacent glycosidic bond and breaking it by an elimination reaction (17, 23, 24). These LPMOs target the crystalline regions of the cellulose surface, which are typically highly recalcitrant to cellulase action and introduce nicks on the substrate surface, thereby providing extra chain ends for glycoside hydrolases (GHs) to act on (25). This action subsequently enhances enzymatic synergy with other cellulases during the saccharification of cellulosic substrates (25–27). However, the critical drawback of not having a single strain that can secrete all the cellulase components (both hydrolytic and accessory) in a balanced ratio subsequently affects the overall enzyme cost required for biomass saccharification, especially at high substrate concentrations. This challenge has therefore prompted many kinds of research to identify better strains that could secrete, to a large extent, all the key enzyme components required for effective biomass hydrolysis (28–30).

In our search for a native producer of cellulolytic enzymes, we identified Penicillium funiculosum (NCIM1228), a filamentous fungus with a secretome that exhibited outstanding potential for hydrolyzing cellulosic biomass (31). Proteomic analysis of its secretome revealed that approximately 58% of 195 proteins detected in the secretome of P. funiculosum under cellulase-inducing conditions were carbohydrate-active enzymes (CAZymes). When the proportion of the cellobiohydrolases (CBH1 and -2), which are the most important and abundant proteins present in the secretome of any cellulase-producing strain, was assessed, it was found that the proportion of CBHs in the secretome of P. funiculosum was only 15% of the total secreted proteins, which is contrary to the secretome of T. reesei, in which CBHs represented 80% of the total secreted proteins (7, 31). Furthermore, during the structure-function characterization of the CBH1 enzyme of P. funiculosum, it was found that CBH1 of P. funiculosum can hydrolyze crystalline biomass with an ∼5-fold-higher efficiency than its counterpart from the well-known industrial host T. reesei (32, 33). Interestingly, further proteomic analysis of the secretome under cellulase-inducing conditions also revealed the detection of an LPMO belonging to auxiliary activity family 9 (AA9) (31, 34), which could have contributed to the high level of saccharification of the secretome of this fungus compared with cellulases from other fungal hosts. (35–37).

Therefore, to further improve this strain as an industrial workhorse for the production of lignocellulolytic enzymes, the regulatory pathway was first manipulated genetically for the deregulation of cellulolytic genes in its genome (38). This deregulation was performed by deleting a homolog of the carbon catabolite repressor (CCR) Mig1 in P. funiculosum NCIM1228, thereby yielding a catabolite-derepressed strain, P. funiculosum Mig1⁸⁸ (PfMig1⁸⁸), which showed a >2-fold improvement in enzyme activity and protein titer in the secretome (38). When the saccharification potential of the PfMig1⁸⁸ secretome toward different chemically pretreated cellulosic biomasses was evaluated, it was found that this secretome efficiently hydrolyzed cellulose to glucose from nitric acid- and ammonium hydroxide-pretreated biomasses, but the conversion efficiencies were much lower when either nitric acid-pretreated biomass (containing high lignin levels) or ammonium hydroxide-pretreated biomass (which contains more hemicellulose) was used (39). This drawback thus necessitated the exploration of better ways of increasing the cellulase content of the secretome.

Based on the above-mentioned factors, this study was designed to further improve the efficiency of the secretome of the PfMig1⁸⁸ strain to enhance its saccharification performance toward highly recalcitrant biomass at higher-level substrate loading with a minimal amount of protein. This was done by fortifying the secretome of PfMig1⁸⁸ through engineering of key hydrolytic and oxidative enzymes, i.e., CBH1 and LPMO, which participate in the saccharification of crystalline cellulosic biomass. Considering that the structure of LPMO from P. funiculosum has never been reported, we first performed in silico structural modeling to identify important residues in the active site of this enzyme and compared them with those in other known LPMOs. We then overexpressed the two selected genes in the catabolite-derepressed strain of P. funiculosum independently and simultaneously. The secretome from each of the resulting transformants was then assessed for improved production of cellulolytic enzymes as well as their saccharification efficiency toward acid-pretreated wheat straw (PTS).

RESULTS AND DISCUSSION

Phylogenetic distribution and computational analysis of the LPMO gene in P. funiculosum.

From different studies and reports, it is known that the number of LPMO genes in fungi varies and can range from a single gene to 47 or more depending on the organism (20, 40–42). Therefore, to ascertain the number of LPMO genes present in P. funiculosum, we first performed whole-genome annotation for this organism and searched for all possible LPMO genes that have been reported across different auxiliary activity (AA) families, i.e., AA9, -10, -11, -13, -14, -15, and -16, using hmmscan software (HMMER3, version 3.0) and the dbCAN database. The annotation indicated the presence of a single gene for LPMO in the genome (Fig. 1A; see also the supplemental material), which belongs to the AA9 family in the CAZy database and is thus designated PfLPMO9. A preliminary analysis of the PfLPMO9 nucleotide and encoded protein sequences showed that the nucleotide sequence consists of 1,020 bp, including two introns, which encode 310 amino acids with a theoretical molecular weight of 32 kDa (Fig. 1A). Since there have been limited reports on LPMO from P. funiculosum, we proceeded to perform phylogenetic analysis with the LPMO sequences available in the NCBI database to assess its neighbors. We found that the LPMO orthologs from Talaromyces pinophilus and Penicillium occitanis in the tree that was the closest shared the same branch, while those from well-studied fungi such as T. reesei and Neurospora crassa were in different branches (Fig. 1B). The phylogenetic analysis placed PfLPMO9 in a cluster together with C-1/C-4-oxidizing LPMOs, which are active toward cellulosic substrates, suggesting that this LPMO could mediate the oxidative cleavage of cellulosic biomass at either the reducing (C-1) or nonreducing (C-4) position (36).

FIG 1.

Schematic representation of P. funiculosum LPMO (A) and phylogenetic tree of LPMO orthologs in fungi (B). Molecular phylogenetic analysis was performed using the maximum likelihood method and the Jones-Taylor-Thornton (JTT) matrix-based model. The tree with the highest log likelihood (−17,992.93) is shown. Initial trees for the heuristic search were obtained automatically by applying neighbor-joining and BIONJ (BIO neighbor-joining) algorithms to a matrix of pairwise distances estimated using the maximum composite likelihood (MCL) approach and then selecting the topology with a superior log-likelihood value. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The bootstrap support corresponding to the numbers on the tree branches was calculated per 1,000 bootstrap replicates (69). This analysis involved the catalytic domains of 29 LPMOs, most of which had been functionally characterized. Evolutionary analyses were conducted using MEGA X software.

To assess putative structural differences (if any) between the C-1/C-4-oxidizing LPMOs, a structural model of PfLPMO9 was built and used for comparison with previously characterized LPMOs. Since there are no three-dimensional (3-D) structures available for LPMO from P. funiculosum in public databases, we first retrieved from the PDB the full-length amino acid sequences corresponding to the catalytic domains of the C-1/C-4-oxidizing LPMOs with 3-D structures that were available and subjected them to multiple-sequence alignment (MSA) alongside the catalytic domain of PfLPMO9. A reasonable degree of amino acid conservation was found among the various LPMO catalytic domains retrieved and analyzed (Fig. 2). The three residues His22, His107, and Tyr196, which are involved in copper coordination and the binding of cellooligosaccharide chains, were conserved among all the LPMOs analyzed. The alignment showed that PfLPMO9 shared 75% identity with the protein from Thermoascus aurantiacus (TaLPMO9A), while 53%, 50%, and 36% identities were observed with LPMOs from Neurospora crassa (NCU07760), T. reesei (HjLPMO9B), and Lentinus similis (LsLPMO9A), respectively.

FIG 2.

Multiple-sequence alignment of PfLPMO9 with LPMOs in auxiliary activity family 9 (AA9) with known C-1/C-4-oxidizing activity. The proteins included are as follows: NCU07760 (UniProtKB accession no. Q7S111), PaLPMO9A (UniProtKB accession no. B2B629), TaLPMO9A (UniProtKB accession no. G3XAP7), FgLPMO9A (UniProtKB accession no. I1REU9), HjLPMO9B (UniProtKB accession no. O14405), GtLPMO9B (UniProtKB accession no. Q7SA19), and LsLPMO9A (UniProt accession no. A0A0S2GKZ1). Fully conserved residues are shown in white on a red background. Blue triangles indicate residues coordinating the copper ion at the active site. The positions of the four cysteine residues with potential disulfide bridge formation are indicated by green dashed lines. Blue frames indicate that more than 70% of the residues in the corresponding columns exhibit similar physicochemical properties (indicated as red residues on a white background). Black boxes indicate variable regions in the LPMO9 family with the corresponding names L2, L3, LC, and LS, which contribute to shaping the substrate-binding surface (70).

A homology model of PfLPMO9 was further constructed using the reference proteins mentioned in Materials and Methods (Fig. 3A). The modeled structure was evaluated for accuracy by Ramachandran plot validation using the PROCHECK module of the PDBSum server (Fig. 3B). The Ramachandran statistics revealed that 90.5% of the amino acid residues from the modeled structure were incorporated in the favored regions (A, B, and L) of the plot, while 8.9% of the residues were in the allowed regions (a, b, l, and p) of the plot, which suggests that the model is reasonable and reliable for homologous structural analysis. Images of the superimposition of the obtained PfLPMO9 structural model with the TaLPMO9A structure as well as the LsLPMO9A structure are shown in Fig. S1 in the supplemental material. We found that the overall folds of PfLPMO9 and TaLPMO9A were very similar. Importantly, the model indicated two possible disulfide bonds in PfLPMO9 (Cys118-Cys122 and Cys77-Cys199), similar to those in TaLPMO9A (Cys97-Cys101 and Cys56-Cys178), which might help in the stabilization of its tertiary structure (Fig. 2). On the other hand, only one disulfide bond was observed in LsLPMO9A (Cys41-Cys167) (43).

FIG 3.

Structural model of PfLPMO9. (A) Best model of the catalytic domain of PfLPMO9, highlighting the three amino acids that make up the copper site. The loop regions that potentially contribute to functional variation among LPMOs, named L2, L3, LS, and LC, are marked with dark blue, light blue, green, and purple, respectively. (B) Ramachandran plot validation of the modeled structure evaluated by PROCHECK. An array of Phi (Φ) and Psi (Ψ) distributions of the nonglycine, nonproline residues is summarized on the plot.

Regulation and functionality of the PfLPMO9 gene product in P. funiculosum.

To investigate the functional role of PfLPMO9 in the degradation of cellulosic biomass, the fungus was grown on cellulosic and hemicellulosic biomasses, and its transcriptional regulation was evaluated by comparing its RNA level with that of fungus grown on glucose as a carbon source. Transcriptional analysis of the PfLPMO9 gene was performed using transcriptome sequencing (RNAseq) data available for this fungus grown on various carbon sources in our laboratory. LPMO was found to be upregulated when the fungus was cultivated on crystalline cellulose (Avicel) and hemicellulose (wheat bran) as well as on alkali-pretreated wheat straw compared to that when the fungus was grown on glucose, a readily metabolizable carbon source (Table 1). It was very striking to see that the LPMO was most highly upregulated in the presence of Avicel, with a log₂-fold change of 7.73 with respect to the level obtained with growth on glucose, strongly suggesting the possible unique role of LPMO in the degradation of crystalline cellulose compared to other biomasses tested (Table 1). The extent of upregulation of the PfLPMO9 gene in response to crystalline cellulose also closely matched that of the CBH1 gene (Table 1), the product of which, cellobiohydrolase I, is well known to act on crystalline cellulose. The results from the transcriptomic analysis indicate that improving the abundance of PfLPMO9 in the secretome may facilitate an improvement in its saccharification efficiency on highly recalcitrant and crystalline biomass, as suggested by our previous proteomic studies (31, 34).

TABLE 1.

Transcriptional regulation of genes coding for PfLPMO9 and PfCBH1 in response to diverse carbon sources

Gene	Fold upregulation in response to carbon source(s)a
	Avicel	Wheat bran	Avicel + wheat bran	Pretreated wheat straw
PfLPMO9	7.73	1.73	3.09	3.53
PfCBH1	8.35	3.58	6.34	6.41

^aThe normalized transcript read counts after 60 h of growth on Avicel, wheat bran, Avicel plus wheat bran, and pretreated wheat straw were analyzed and compared to the normalized read counts obtained for glucose, which was taken as a control. Values presented are the log₂-fold changes obtained on different polymeric carbon sources with respect to glucose.

In addition to monitoring the transcript level of PfLPMO9 in the presence of crystalline cellulose, we also evaluated the functionality of the LPMO in the crude secretome of P. funiculosum (PfMig1⁸⁸) in cellulose hydrolysis. For this, we adopted the aldonic acid/LPMO quantitation method described previously by Dixit et al. (37), by which we were able to analyze the hydrolysates with the help of an extrinsic LPMO electron donor (ascorbate). The actual effect of the LPMO activity on the fungal secretome of PfMig1⁸⁸ was estimated using Avicel as a substrate in the presence and absence of ascorbic acid in the enzymatic hydrolysis reaction mixture. The differential gluconic acid concentration was quantified after the exclusion of background gluconic acid present in the enzymatic hydrolysis reaction setup without ascorbic acid (negative control) from that in the reaction mixture setup with ascorbic acid, as shown in Fig. 4. The differential amounts of gluconic acid produced by the PfMig1⁸⁸ secretome were 0.24 and 0.43 g/liter at 24 and 48 h, respectively (Fig. 4). Interestingly, this amount was found to be larger than those reported for the variant strain P. funiculosum DBT-IOC-P-11 (0.13 g/liter) and commercial Cellic Ctec2 (0.17 g/liter) after 48 h (37), which may be due to the higher LPMO content in the engineered PfMig1⁸⁸ secretome, as discussed in subsequent sections.

FIG 4.

Gluconic acid production in Avicel hydrolysates. The measured gluconic acid concentrations were produced in the absence and presence of 2 mM ascorbic acid (AA). Avicel hydrolysis was carried out with 7 FPU/g of the PfMig1⁸⁸ secretome. The differential gluconic acid [d(Gluconic acid)] concentration represents the actual concentration of gluconic acid produced during the enzymatic hydrolysis of Avicel in the presence of ascorbic acid. This was calculated by subtracting the background level of gluconic acid produced in the reaction mixture without ascorbic acid from the amount of gluconic acid produced in the presence of ascorbic acid.

Recombinant overexpression of PfLPMO9 in the PfMig1⁸⁸ strain to improve LPMO activity.

After ascertaining the functionality of the LPMO in P. funiculosum in the degradation of crystalline cellulose at both the transcript and protein levels, we decided to insert an additional copy of the PfLPMO9 gene in the background of the Mig1-repressor-deleted strain of P. funiculosum, i.e., PfMig1⁸⁸, as it was shown to produce a larger amount of cellulolytic enzymes in our previous study (38). To overexpress the PfLPMO9 gene in the PfMig1⁸⁸ strain, an expression vector containing the endogenous gene along with its promoter and terminator was constructed using the backbone of the pBIF binary vector. The 3.0-kb region containing the LPMO9 gene (Fig. 5A) was amplified from the genome of the parent NCIM1228 strain and cloned into the pBIF vector to generate the pOAO1 plasmid. The pOAO1 recombinant plasmid was confirmed by restriction digestion before transformation into the PfMig1⁸⁸ strain (Fig. S2A) using the agrobacterium-mediated transformation method (AMTM). The hygromycin-resistant transformants obtained were subsequently analyzed (Fig. 5B). The transformants were first confirmed by PCR (Fig. S2B), and Southern blotting was then carried out for five PCR-positive transformants to further check the integration and copy number. For this, genomic DNA (gDNA) from each strain was digested with XhoI and HindIII and probed with a 621-bp LPMO gene fragment (Fig. 5C). The results showed that all the transformants possessed more than one band, in contrast to the control (PfMig1⁸⁸), which had only a single band, further verifying the insertion of an additional copy of the LPMO cassette in the genome of PfMig1⁸⁸.

FIG 5.

Construction of the LPMO expression cassette and its overexpression in PfMig1⁸⁸. (A) Schematic diagram showing the assembly of the LPMO cassette from the P. funiculosum NCIM1228 genome. SP, signal peptide. (B) Transformants of pOAO1 after AMTM transformation in PfMig1⁸⁸. Transformants were selected on 100 μg/ml hygromycin. (C) Southern blotting of transformants confirmed by PCR. Lane M is the HindIII lambda DNA size marker used, lane C indicates the genomic DNA of the nontransformed parental strain, and lanes 1 to 5 represent the genomic DNA from transformants with LPMO cassette integration. (D) LPMO activity expressed as micromoles of H₂O₂ released per milliliter per minute and FPase activity in the fermentation broth of PfMig1⁸⁸ and five transformants.

To evaluate the expression level of the LPMO gene in the LPMO-overexpressing transformants, the five transformants confirmed by Southern hybridization were cultivated in cellulase-inducing medium (CIM) for 5 days. The transformants were then screened based on the H₂O₂ production capability of their culture supernatant via the Amplex red assay, and the results were compared with those for the parent strain, PfMig1⁸⁸ (Fig. 5D). Due to the potential presence of diverse H₂O₂-producing oxidases and H₂O₂-consuming peroxidases in crude enzyme preparations, the Amplex red assay is not specific for determining LPMO activity in the P. funiculosum secretome (44). Nevertheless, this method was used here to allow fast and simple initial screening for LPMO-like activity that results in ascorbic acid-dependent H₂O₂ production. The results showed a significant increase in ascorbate-induced H₂O₂ accumulation, in the range of 155 to 203%, in all the transformants compared to the parent strain, with the maximum H₂O₂ production obtained with transformant T4 (Fig. 5D). The variation in enzyme activity resulting in H₂O₂ production across the transformants as determined by the Amplex red assay might be due to the difference in the integration loci of the expression cassette in the PfMig1⁸⁸ genome, as previously reported (45, 46). However, when the total cellulase (FPase) activity of the transformants was evaluated using the filter paper assay to see if there would be any corresponding impact on the overall cellulase activity, no significant difference in the FPase activity of the transformants was observed compared to that of the parent strain (Fig. 5D). This lack of difference could be due to the absence of reducing agents that serve as electron donors required by the LPMO enzyme to act on pure cellulose in the standard filter paper assay reaction mixture (36). Nevertheless, the marked increase in LPMO activity based on the H₂O₂ production recorded in all the transformants over that in the parent strain indicates that the gene was successfully overexpressed and that the LPMO amount in the PfMig1⁸⁸ cellulase system was significantly improved.

Simultaneous overexpression of PfLPMO9 and PfCBH1 in the P. funiculosum Mig1⁸⁸ strain to improve the cellulase system in its secretome.

Since CBH1 is one of the vital hydrolytic enzymes of particular importance participating in crystalline cellulose deconstruction, the impact of the overexpression of CBH1 on the quality of the total cellulase system of the LPMO transformant was next examined.

In addition to the construction of the dual transformant, a fungal strain was also engineered in which only the CBH1 gene, along with its promoter and terminator, was integrated into the genome of the PfMig1⁸⁸ strain to assess the impact of CBH1 overexpression alone on the cellulase system (Fig. S3A to E and supplemental results and discussion). An approximately 57 to 62% increase in Avicelase activity was found across the five transformants over that of PfMig1⁸⁸. The results also showed up to a 25% increase in the filter paper units (FPU) of the transformants, where the maximum FPase activity achieved was 5.1 FPU/ml for CBH1 transformant T3 (Fig. S3F).

To engineer the fungal strain for the overexpression of both LPMO and CBH1, a systematic approach was utilized for the construction of the desired pOAO5 vector containing the LPMO/CBH1 expression cassette (Fig. 6A), as described in Materials and Methods. For fungal transformation, the pOAO5 plasmid was confirmed by restriction digestion before transformation into the PfMig1⁸⁸ strain (Fig. S2C). The hygromycin-resistant transformants obtained were analyzed by PCR to confirm the integration of the LPMO/CBH1 cassette into the genome (Fig. 6B; Fig. S2D). Similarly, to further confirm the integration and copy number, Southern blotting was carried out for all selected transformants (Fig. 6C). The results showed that all the transformants possessed more than one band, in contrast to the control (PfMig1⁸⁸) strain, which had a single prominent band, further verifying the insertion of an additional copy of the LPMO/CBH1 cassette in the genome of PfMig1⁸⁸. Transformant T5 showed two faint bands at similar locations, which may be due to experimental error leading to lower-level loading of digested genomic DNA. Four out of the five transformants had only one additional band, suggesting a single-copy insertion of the LPMO/CBH1 cassette in the genome, while one of the transformants (T1) had two copies of the integrated cassette (Fig. 6C).

FIG 6.

Simultaneous overexpression of LPMO and CBH1 in PfMig1⁸⁸. (A) Schematic diagram showing the construction cassette for the dual overexpression of the LPMO and CBH1 genes from P. funiculosum NCIM1228. Pro, promoter; SP, signal peptide; Ter, terminator. (B) Transformants of pOAO5 after AMTM transformation in PfMig1⁸⁸. Transformants were selected on 100 μg/ml hygromycin. (C) Southern blotting of transformants confirmed by PCR. Lane M is the HindIII lambda DNA size marker used, lane C indicates the genomic DNA of the nontransformed parental strain, and lanes 1 to 5 represent the genomic DNA from transformants with LPMO/CBH1 cassette integration. (D) Enzymatic profile and H₂O₂ production of the overexpressed enzymes in the fermentation broth of PfMig1⁸⁸ and six transformants.

To further assess the effect of this simultaneous overexpression on the expression level of cellulolytic enzymes by the transformants, the six transformants that were confirmed to be positive by PCR for the integration of the LPMO/CBH1 expression cassette were cultivated in cellulase-inducing medium for 5 days. The resulting secretome from the strains was thereafter recovered and used for enzyme assays. The LPMO, cellobiohydrolase, and total cellulase activities of the transformants were measured and compared with those of the parent PfMig1⁸⁸ strain. As expected, based on the results from the specific overexpression of the enzymes, all the transformants screened exhibited enhancements in all the activities compared with the parent strain, as presented in Fig. 6D. The results showed an increment in H₂O₂ production, likely as a result of the ascorbate-induced action of LPMO over that of PfMig1⁸⁸ across the transformants, which was also confirmed by an increase in the LPMO mRNA level, as discussed below. Similarly, we found an approximately 40 to 66% increase in Avicelase activity and a 14 to 20% increase in filter paper units compared with PfMig1⁸⁸ (Fig. 6D). Although there was variation in the enzyme activities tested across all the transformants, the highest levels of production of all the enzymes as well as H₂O₂ were found with transformant T6, in which the Avicelase and FPase activities were 3.51 U/ml and 5.2 U/ml, respectively, while the level of ascorbate-induced H₂O₂ production was 33.2 μmol/ml (Fig. 6D). It was surprising that there was no difference in enzyme activities between the transformants with single-copy integration and transformant T1, which had two additional copies of the LPMO/CBH1 cassette. This suggested that the integration locus in addition to the gene copy numbers might have also affected the expression of the cellulolytic enzymes (30).

Comparative transcriptional and translational analyses of cellulolytic enzymes in NCIM1228, PfMig1⁸⁸, and the engineered strains.

To gain a further understanding of the overall regulation of the cellulase system due to the overexpression of these key cellulolytic enzymes, the transcript abundance of each overexpressed gene at the mRNA level was next examined. Furthermore, to understand whether the overexpression of LPMO and/or CBH1 had any impact on the expression of other key cellulolytic enzymes, we also checked the transcript levels of some of the major cellulases/hemicellulases known to be produced by NCIM1228 under derepressing conditions (Table 2) (34). For this, cultures of NCIM1228, PfMig1⁸⁸, and the best-performing transformants, PfOAO1 (overexpressing LPMO in the background of PfMig1⁸⁸), PfOAO2 (overexpressing CBH1 in the background of PfMig1⁸⁸), and PfOAO3 (overexpressing LPMO as well as CBH1 in the background of PfMig1⁸⁸), were grown in 4% Avicel for 48 h to obtain good mycelial growth. The transcript levels of all five strains under derepressing conditions were determined by real-time PCR (RT-PCR) with tubulin as a control, and the relative fold change was normalized to the level in NCIM1228 since it was the original strain from which all other mutants were generated. Under LPMO overexpression, we observed a 5-fold increase in LPMO transcripts for PfMig1⁸⁸, while there were 18- and 26-fold increases in transcript levels for the PfOAO1 and PfOAO3 strains, respectively, indicating the reason for the increase in H₂O₂ production, as seen in Fig. 5D and Fig. 6D. As expected, there was no difference in the LPMO transcript level for the PfOAO2 transformant compared to that for PfMig1⁸⁸ (Fig. 7A). Likewise, for the CBH1-overexpressing strains, approximately 4-, 30-, and 39-fold increases in CBH1 transcript levels were observed for the PfMig1⁸⁸, PfOAO2, and PfOAO3 strains, respectively, while there was no change in the CBH1 transcripts of PfOAO1 (Fig. 7A). When the transcript level for the β-glucosidase (BGL) gene in all the strains was checked, the results showed a 5-fold increase in BGL expression for PfMig1⁸⁸ and PfOAO1, while there were 10- and 11-fold increases recorded for the PfOAO2 and PfOAO3 mutants, respectively. The increase in the expression level of BGL in the two CBH1 transformants could be a result of the increase in the cellobiohydrolase level in the transformants. The increased cellobiohydrolase level in the medium may have led to increased cellobiose levels, thereby providing a signal to the cells to produce more BGL, which is required to hydrolyze this cellobiose to glucose. However, no significant difference between the parent PfMig1⁸⁸ strain and all the mutants was observed in terms of the expression levels of endoglucanase (EG) and xylanase (XYL), as shown in Fig. 7A.

TABLE 2.

CAZymes whose transcript levels were monitored in P. funiculosum NCIM1228, PfMig1⁸⁸, and the resulting transformants^a

Enzyme class	Functional classification	CAZY family
AA	LPMO	AA9
Exoglucanase	CBH1	GH7-CBM1
Endoglucanase	EG	GH5-CBM1
β-Glucosidase	BG	GH3
Xylanase	β-1,4-Xylanase	GH10-CBM1

^aAA, auxiliary activity; LPMO, lytic polysaccharide monooxygenase; CBH1, cellobiohydrolase I; EG, endoglucanase; BGL, β-glucosidase.

FIG 7.

Determination of cellulase expression and activities in P. funiculosum NCIM1228, PfMig1⁸⁸, and all the engineered strains. (A) The transcriptional expression of LPMO, cellobiohydrolase, endoglucanase, β-glucosidase, and xylanase in NCIM1228, PfMig1⁸⁸, and all the engineered strains was measured by quantitative real-time PCR after growing the strains for 48 h in the presence of 4% Avicel. The expression levels were normalized to those in NCIM1228 and plotted. (B) CBH1 activity determined using Avicel as the substrate. (C) H₂O₂ production by PfLPMO expressed as micromoles of H₂O₂ released per milliliter determined using Amplex red. (D) β-Glucosidase activity determined using p-nitrophenyl-β-d-glucoside (pNPG). (E) Overall cellulase activity on filter paper. (F) Endoglucanase activity determined using CMC as the substrate. (G) Xylanase activity measured using beechwood xylan as the substrate. (H) Total secreted proteins in all the strains. The data are presented as the means from three independent experiments, and error bars express the standard deviations.

To examine and compare the influences of LPMO and CBH1 overexpression on the overall cellulase system, cultures of NCIM1228, PfMig1⁸⁸, and the best-performing transformants of all the engineered strains were grown in CIM. The resulting supernatants containing the secreted enzymes were recovered from the mycelia and used for enzyme assays. Individual cellulase activities of LPMO, CBH, EG, BGL, and XYL and FPU for all strains were assayed and compared and are presented in Fig. 7B to G. In accordance with the mRNA levels and data from a previous report (38), the PfMig1⁸⁸ strain showed a 2-fold increase in all enzyme activities evaluated and in the total extracellular protein level. It was found that PfOAO2 and PfOAO3 showed ∼65% and 27% increases in Avicelase (Fig. 7B) and FPase (Fig. 7E) activities, respectively, compared to PfMig1⁸⁸, while PfOAO1 showed a nonsignificant change. On the other hand, PfOAO1 and PfOAO3 exhibited greater H₂O₂ production capabilities, while PfAOA2 did not show any significant change (Fig. 7C). In addition to an increase in Avicelase activity, PfOAO2 and PfOAO3 also showed an ∼18% increase in BGL activity and an ∼9% increase in total secreted protein compared with PfMig1⁸⁸ (Fig. 7D and H), as indicated by transcript data. Furthermore, the increase in BGL production by PfOAO2 and PfOAO3 transformants relative to PfMig1⁸⁸ was further confirmed by zymography analysis (Fig. S4). Little change in carboxymethyl cellulase (CMCase) and xylanase activities was observed in the transformants compared to PfMig1⁸⁸ (Fig. 7F and G).

In addition, we investigated whether the specific activities of the overexpressed enzymes also increased in addition to the volumetric activities. For this, the total protein in the secretome of each strain was determined and used to calculate specific enzyme activities as well as H₂O₂ production (Table S1). The results showed a significant increase in the specific H₂O₂ generation activity of the PfOAO1 and PfOAO3 strains relative to that of PfMig1⁸⁸. Similarly, there was a significant increase in the specific activities of CBH1, BGL, and FPase in PfOAO2- and PfOAO3-overexpressing strains. However, there was no significant difference in the specific activities of EG and XYL across all the strains. The increase in the specific activities of the key hydrolytic and oxidative enzymes seen in this study indicates that the performance of the enzyme mixture produced by the overexpressed strains was enhanced compared with that of PfMig1⁸⁸ and may help in reducing the enzyme load required for biomass saccharification.

Hydrolytic efficiency of the cellulase enzyme complex with LPMO and CBH1 overexpression against acid-treated biomass under high-level substrate loading.

To assess the relevant industrial application of the overexpression of these two key enzymes, the secretomes of all the engineered strains as well as the parent strains were used to investigate their saccharification performance on acid-pretreated wheat straw (PWS). The saccharification reaction mixture was set up at 20% substrate loading of PWS using the secretomes of NCIM1228, PfMig1⁸⁸, PfOAO1, PfOAO2, and PfOAO3 at the same protein loading of 30 mg/g dry biomass weight (DBW) and incubated at 50°C for 96 h. Samples were collected every 24 h and analyzed for the production of monomeric sugars (Fig. 8A and B). From the results, linear increases in the concentration of monomeric sugars and hydrolysis efficiency were observed in all the strains over time for 72 h, after which no appreciable increase in the sugar concentration was observed. At the 72-h time point, approximately 27% holocellulose (cellulose plus hemicellulose) conversion was obtained with the NCIM1228 secretome, whereas more than 64% of the pretreated biomass was hydrolyzed by the PfMig1⁸⁸ secretome (Fig. 8A); the total monomeric sugar (glucose plus xylose) concentrations recorded were 37.5 g/liter and 90 g/liter, respectively (Fig. 8B). The results showed a saccharification efficiency similar to that previously reported when the secretome of PfMig1⁸⁸ was used for the saccharification of acid-treated sugarcane bagasse and rice straw (39). The marked increase in the biomass hydrolysis capacity of the PfMig1⁸⁸ secretome could be attributed to a large extent to increased proportions of CBH, EG, and BGL being secreted due to the absence of a functional CCR in PfMig1⁸⁸, as recently reported (39). The 64% hydrolysis achieved with the secretome of PfMig1⁸⁸ at 20% solid loading was highly significant, as it has previously been reported that most saccharification experiments carried out with high-level substrate loading may not proceed beyond 60% even after physical optimization of the saccharification process (47–49).

FIG 8.

Time course of the saccharification of nitric acid-pretreated wheat straw by the secretomes of NCIM1228, PfMig1⁸⁸, and all the engineered strains under 20% solid loading and a protein concentration of 30 mg/g biomass. (A) Percent sugar release measured at 24-h intervals over the 96-h saccharification period; (B) total fermentable sugar obtained at the 72-h saccharification time point.

When the hydrolytic ability of the cellulases produced by the LPMO-overexpressing PfOAO1 strain on PWS relative to that of the enzymes produced by PfMig1⁸⁸ was evaluated, a considerable effect was seen, with increases in the concentration of monomeric sugars and cellulose conversion (Fig. 8A and B). The considerable effect seen with overexpressed LPMO in the secretome is consistent with previous reports of a boost in the performance of LPMO in the bioconversion of lignocellulosic biomass due to its synergistic action with other cellulases, especially CBH1 (47, 50). The concentration of total monomeric sugars released by the secretome of the CBH1-overexpressing PfOAO2 strain was 107 g/liter, which corresponded to 77% holocellulose conversion and a 12% increase over the value obtained with PfMig1⁸⁸. The enhanced saccharification obtained with the PfOAO2 strain could be not only due to the increase in the cellobiohydrolase level in the system but also linked to the enhancement in the production of β-glucosidase, as seen previously (Fig. 7D). This must have facilitated the enhanced enzymatic degradation of PWS since BGL has the capacity to hydrolyze cellobiose to glucose in the final step and alleviate the feedback inhibition effect of cellobiose on the activities of CBH and EG (51). As expected, there was no significant increase in the hydrolysis of hemicellulose in all the engineered strains compared with that in PfMig1⁸⁸ (Table S2).

Interestingly, when both LPMO and CBH1 were co-overexpressed, the concentration of total reducing sugars released by the secretome of the strain increased to 112 g/liter, corresponding to 82% holocellulose conversion (Fig. 8A and B). This shows that the simultaneous overexpression of LPMO and CBH1 led to a highly significant increase in the saccharification efficiency compared to those of NCIM1228 (∼199% increase), PfMig1⁸⁸ (∼20% increase), and the individual strains (∼13% and ∼6% increases for LPMO- and CBH1-overexpressing strains, respectively). The results therefore suggest that the co-overexpression of major cellulase components and the accessory enzymes could facilitate the creation of a more efficient cellulolytic system for the optimal hydrolysis of cellulosic biomass under high-level substrate loading, as previously reported (51, 52). The positive effect of LPMO seen in these experiments with pretreated wheat straw, without the addition of an electron donor such as ascorbate, may be due to the presence of reducing agents in the form of phenolic compounds in lignin, which could have possibly initiated the LPMO activity (20, 53).

In conclusion, multiple fungal genetic tools were utilized to construct a more versatile cellulase system for improving the saccharification performance of a catabolite-derepressed strain of P. funiculosum via the overexpression of key oxidative and hydrolytic enzymes in its genome. The combined overexpression of LPMO and CBH1 provided a significantly more efficient cellulase cocktail with an enhanced saccharification performance on PWS under high-level substrate loading. The engineered LPMO/CBH1 cellulase system exhibited significantly enhanced cellulose conversion after 72 h of enzymatic saccharification of PWS at 20% loading compared to that obtained with PfMig1⁸⁸. The results from this study showed that the engineered strains developed could potentially be used as promising bioresources needed for the production of richer and more balanced cellulase cocktails required for the low-cost production of lignocellulose-based biofuels.

MATERIALS AND METHODS

Plasmids and microbial strains.

All the strains and plasmids used in this study are listed in Table 3, while the protein sequences for LPMO and CBH1 for the study are provided in the supplemental material. Escherichia coli DH5α was used for plasmid propagation throughout the experiments. The Agrobacterium tumefaciens LBA4404 strain used for fungal transformation was maintained on low-sodium LB medium (10 g/liter tryptone, 5 g/liter yeast extract, 5 g/liter sodium chloride) containing 100 μg/ml kanamycin and 30 μg/ml rifampicin. The pBIF vector (54), which was used as the backbone vector for fungal transformation, contains hygromycin and kanamycin resistance genes as selective markers for the selection of transformants. The pBluescript SK(+) (pBSK+) vector containing the ampicillin resistance gene was used as a shuttle vector for the simultaneous cloning of genes for LPMO and CBH1. P. funiculosum NCIM1228 and its derivative P. funiculosum Mig1⁸⁸ (PfMig1⁸⁸), the fungal strains used for this study, were routinely cultivated on petri dishes containing low-malt extract–peptone (LMP) agar for approximately 14 days until full sporulation. For the selection and maintenance of fungal transformants, LMP medium supplemented with hygromycin B at 100 μg/ml was used.

TABLE 3.

Plasmids and strains used in the study

Strain or plasmid	Descriptiona	Source or reference
Strains
Escherichia coli DH5α	F− ϕ80lacZΔM15 Δ(lacZYA-argF)U169 recA1 endA1 hsdR17	Invitrogen
Agrobacterium tumefaciens LBA4404	Ach5 with pAch5; Δtra Δocc ΔTL ΔTR Rifr	38
Penicillium funiculosum NCIM1228	Cellulase-producing fungus obtained from the NCIM	31
PfMig188	Catabolite-derepressed strain of Penicillium funiculosum NCIM1228	38
PfOAO1	LPMO overexpression in PfMig188 with the help of the pOAO1 vector	This study
PfOAO2	CBH1 overexpression in PfMig188 with the help of the pOAO2 vector	This study
PfOAO3	LPMO/CBH1 co-overexpression in PfMig188 with the help of the pOAO5 vector	This study
Plasmids
pBIF	Kanamycin-resistant vector containing EGFP	54
pBluescript(SK+)	Standard cloning vector containing the ampicillin resistance gene	Agilent
pOAO1	pBIF vector containing the LPMO gene along with its native promoter and terminator	This study
pOAO2	pBIF vector containing the CBH1 gene along with its native promoter and terminator	This study
pOAO3	pBluescript SK(+) vector containing the LPMO gene along with its native promoter and terminator	This study
pOAO4	pOAO3 vector containing the CBH1 gene along with its native promoter and terminator	This study
pOAO5	pBIF vector containing the LPMO and CBH1 genes along with their native promoters and terminators	This study

^aNCIM, National Culture Collection Centre, Pune, India; EGFP, enhanced green fluorescent protein.

Cellulosic substrate and pretreatment.

Nitric acid-treated wheat straw was used as the cellulosic substrate in this study. Wheat straw biomass was comminuted in a cutting mill and sieved through a 1.5-mm mesh to obtain a uniform size before chemical pretreatment. Pretreatment was carried out using 0.5% nitric acid in a reactor for 30 min at 120°C at 1.8 × 10⁶ Pa. The pretreated straw was then repeatedly washed until the pH became neutral. Compositional analysis conducted according to National Renewable Energy Laboratory (NREL) procedure TP510-42618 (55) on the pretreated wheat straw yielded a cellulose content of 61.3%, a hemicellulose content of 6.1%, a lignin content of 15.6%, and an ash content of 6%.

Computational analysis of LPMO from P. funiculosum (PfLPMO9).

The LPMO sequence was retrieved from the draft genome sequence of P. funiculosum NCIM1228 available in our laboratory (see the supplemental material) (19). The amino acid sequences of biochemically characterized LPMOs from other species were retrieved from the NCBI database, and molecular phylogenetic analysis by the maximum likelihood method and the Tamura-Nei model was conducted using MEGA X software. Multiple-sequence alignment of PfLPMO9 with C-1/C-4-oxidizing LPMOs obtained from the phylogenetic analysis was performed using the Clustal Omega multiple-sequence alignment program (56). The alignment was visualized using ESPript 3.0 software (57). To build the structural model of PfLPMO9, the LPMO sequence was first compared with all the available PDB structures using psi blast (56). The top six protein structures from the database, namely, those under PDB accession no. 6HA5 (58), 2YET (16), 6H1Z (58), 2VTC (59), 5ACF (43), and 5NLT (60), were selected as references for modeling. The LPMO structure was modeled using Modeller version 9.17, while UCSF Chimera software version 1.14rc was used for molecular graphics and visualization of the protein structure (61).

Engineering of P. funiculosum Mig1⁸⁸ for overexpression of LPMO and CBH1.

All vectors for fungal expression described in this study were constructed on the backbone of the pBIF vector that was previously constructed using the binary vector pCAMBIA1300 as a backbone (54). Binary vectors for the overexpression of the LPMO and CBH1 genes from P. funiculosum were constructed as follows. The endogenous gene encoding LPMO was amplified from the genome of P. funiculosum NCIM1228 using the primers PfLPMO-F and PfLPMO-R (Table 4). The primers were designed according to the LPMO sequence containing both its native promoter and terminator, which was obtained from the draft genome sequence of the strain. The PCR product obtained was digested with the restriction enzymes SbfI and SpeI before being ligated into the pBIF vector previously digested with the enzymes PstI and SpeI to generate the pBIF/LPMO (pOAO1) vector. For the overexpression of CBH1, the CBH1 gene (21) was amplified from the genome of P. funiculosum NCIM1228 using the primers PfCBH1-F1 and PfCBH1-R1 spanning its native promoter and terminator (Table 4). The PCR product containing the SacI and BamHI restriction sites was digested and ligated into pBIF at the corresponding sites to create the pBIF/CBH1 (pOAO2) vector. The ligated products of pOAO1 and pOAO2 were then transformed into E. coli DH5α cells and selected on 50 μg/ml kanamycin. The resulting colonies were screened for positive transformants by colony PCR, followed by restriction digestion of the corresponding plasmids. To create a strain for the combined overexpression of both the LPMO and CBH1 genes, sequential cloning of the two genes was performed using pBSK+ as a shuttle vector. First, the LPMO gene that had previously been amplified from the genome was cloned into the pBSK vector between the PstI and SpeI sites in its multiple-cloning site (MCS) to obtain the plasmid pBSK/LPMO (pOAO3). Next, the CBH1 gene with its native promoter and terminator was further amplified from the genome of P. funiculosum NCIM1228 using the primers PfCBH1-F2 and PfCBH1-R2 containing the AflII and SpeI restriction sites, respectively. The PCR product obtained was digested with the AflII and SpeI restriction enzymes before being ligated into the corresponding sites of the pOAO3 vector to obtain the pOAO4 vector. The resulting SbfI/SpeI fragment obtained in the pOAO4 plasmid was excised and cloned between the PstI and SpeI sites of the pBIF vector, thereby generating the pBIF/LPMO/CBH1 (pOAO5) vector. The verified pOAO1, pOAO2, and pOAO5 plasmids were transformed into the PfMig1⁸⁸ fungal strain according to the agrobacterium-mediated transformation method (AMTM) as previously described (62).

TABLE 4.

Primers used in the study

Purpose	Primer name (sequence)a
LPMO and CBH1 plasmid construction cassette	PfLPMO-F (5′-TCATCGATATCACCTGCAGGTCTGCGCCTGGTCC-3′)
	PfLPMO-R (5′-TACGGTACTAGTCAAGCTGTGGATCGGTTCTC-3′)
	PfCBH1-F1 (5′-TATACCCGAGCTCTATCCCAACAGGTCGATCC-3′)
	PfCBH1-R1 (5′-TATGGATCCGCTAGATGGTGCG-3′)
	PfCBH1-F2 (5′-AGACTTCTTAAGTATCCCAACAGGTCG-3′)
	PfCBH1-R2 (5′-AGTTCAGACTAGTGCTAGATGGTGCGAAAG-3′)
	PgpdA-F (5′-GAATTCTGTACAGTGACCGGTGACTC-3′)
	TrpC-R (5′-GCCAAGCTTCCTCTAAACAAGTGTACCTGTGC-3′)
	PgpdA IR-F (5′-TGCGTCAGTCCAACATTTGT-3′)
	PfCBH1 IR R (5′-GGTACCGGCGTAACGATCAGAGCT-3′)
Southern blotting	LPMO probe F (5′-AGTAGCAGAACCAGTGGCAACAGCG-3′)
	LPMO probe R (5′-TGGGCAACAACTGCAACCGACCTG-3′)
	CBH1 probe F (5′-TACACTGGCAACACTTGGAATAGCGCC-3′)
	CBH1 probe R (5′-GGTACCGGCGTAACGATCAGAGCT-3′)
RT-PCR	LPMO RT F (5′-GCAGAAGCAGCGGCAGC-3′)
	LPMO RT R (5′-GAAACGCAGACGGTGCCCAAAA-3′)
	CBH1 F (5′-GCAAACACGAAGCTGGTATGG-3′)
	CBH1 R (5′-GGTGACATAGGAGCTGCCGG-3′)
	EG-GH5 F (5′-GCAACCATTGGTGAATTCATCAGTCAG-3′)
	EG-GH5 R (5′-CTGCCATTGTACCTTCCATAATTGTGAG-3′)
	BGL-GH3 F (5′-CTGGCGAAGGGTACATAACAGTCG-3′)
	BGL-GH3 R (5′-GCCAGCCCATACTACGGC-3′)
	Xyl (GH10-CBMI) F (5′-GCAATGAAATGGCAACCCACCG-3′)
	Xyl (GH10-CBMI) R (5′-CCTTCAAGGCAGCAATAAGGGTTG-3′)
	Tubulin F (5′-ATTGCTCAGGTTGTCTCCTCCATC-3′)
	Tubulin R (5′-CATGGTGATCTCGTTGACAGAGTTGG-3′)

^aUnderlining indicates the site of the restriction enzyme used for cloning.

Molecular analysis of transformants.

After transformation, the resulting hygromycin-resistant transformants were screened according to the method described previously by Fang and Xia (63). The transformants were verified by both PCR and Southern blot hybridization for the integration of the LPMO and CBH1 gene expression cassettes. For PCR analysis, the primers PgpdA-F and TrpC-R were used to screen for LPMO and CBH1 integration, while transformants of LPMO/CBH1 were screened using the primer set PgpdA IR-F and PfCBH1-IR R (Table 4). Southern hybridization was performed using standard procedures as described previously (64). Genomic DNAs (8 μg) of PfMig1⁸⁸ and the LPMO, CBH1, and LPMO/CBH1 transformants were digested with the XhoI, HindIII, and ApaI restriction enzymes. The digested gDNAs were then size fractionated by electrophoresis on a 0.8% agarose gel in 1× TAE (Tris-acetate-EDTA) buffer. After depurination using 250 mM HCl, denaturation (1.5 M NaCl and 0.5 M NaOH), and neutralization (1.5 M NaCl and 1.0 M Tris-HCl [pH 8.0]), the gel was capillary blotted onto positively charged Hybond-N⁺ membranes (Amersham Biosciences, USA). The 621-bp fragment of the LPMO gene was PCR amplified with the primers LPMO probe F and LPMO probe R and labeled as a probe to detect LPMO integration. Similarly, a fragment of the CBH1 gene (605 bp long) was amplified using the primers CBH1 probe F and CBH1 probe R and used to confirm the insertion of the CBH1 expression cassette in both the CBH1 and LPMO/CBH1 strains (Table 4). The LPMO and CBH1 amplicons for detection were radiolabeled with [α-³²P]dCTP using the NEBlot kit (New England BioLabs [NEB], USA) according to the manufacturer’s instructions and used as probes to determine the copy numbers of transfer DNA (T-DNA) integrations in all the transformants.

Expression analysis of cellulolytic enzymes via real-time PCR.

For real-time PCR experiments, cultures of all strains were grown in minimal Mandel’s medium containing 4% Avicel for 48 h (65). Mycelia were harvested by filtration and frozen in liquid nitrogen. RNA was extracted using an RNeasy kit (Qiagen) according to the manufacturer’s instructions. RNA was treated with DNase (Invitrogen) before cDNA synthesis. One microgram of RNA was used as the template for each quantitative real-time PCR (qRT-PCR). A cDNA synthesis control was performed to ensure the absence of DNA contamination. qRT-PCR was carried out using iTaq universal SYBR green supermix (Bio-Rad) and a Bio-Rad CFX96 qPCR detection system. Primers for transcripts to be tested were designed using the boundary sequence of two exons to avoid any amplification from contaminant genomic DNA. qRT-PCR was performed in biological triplicates with tubulin as the endogenous control. Relative expression levels were normalized to the level of tubulin, and the fold changes in RNA levels were calculated as the ratios of the relative expression levels in PfMig1⁸⁸ and the corresponding transformants of LPMO and CBH1 to that in NCIM1228 under cellulase-inducing conditions (66).

Cellulolytic secretome preparation.

Penicillium funiculosum NCIM1228, PfMig1⁸⁸, and the resulting LPMO, CBH1, and LPMO/CBH1 transformants were cultivated on petri dishes containing low-malt extract agar until full sporulation. After 14 days of incubation, spores were recovered with sterile water, filtered through sterile Mira cloth, and quantified using a hemocytometer. The primary culture of each strain was prepared by culturing 10⁷ conidiophores in potato dextrose broth (PDB) for 36 h. Primary cultures of the strains were added to cellulase-inducing medium in Erlenmeyer flasks at a final concentration of 10%. Cellulase-inducing medium contained soya peptone (24 g/liter), wheat bran (21.4 g/liter), microcrystalline cellulose (MCC) (24 g/liter), KH₂PO₄ (12.4 g/liter), K₂HPO₄ (2.68 g/liter), (NH₄)₂SO₄ (0.28 g/liter), CaCO₃ (2.5 g/liter), corn steep liquor (1%), urea (0.52 g/liter), and yeast extract (0.05 g/liter), with the final pH adjusted to 5.0. The flasks were kept at 28°C for 5 days with orbital shaking at 150 rpm (Innova 44; Eppendorf AG, Germany). Induced cultures were centrifuged at 9,000 rpm for 10 min at 4°C, and the cellulolytic supernatants were collected and stored at 4°C until use.

Determination of the enzyme activities of the engineered secretome.

All enzymatic activity assays in this study were performed according to standard assay procedures. Cellobiohydrolase activity was determined by incubating the appropriate dilution of the enzyme with 1% Avicel PH-101 (Sigma) for 120 min. Endoglucanase, xylanase, and β-glucosidase activities were determined by incubating appropriate dilutions of the enzyme with 2% carboxymethyl cellulose (CMC) (Sigma), 2% beechwood xylan (HiMedia), and p-nitrophenyl-β-d-glucopyranoside (Sigma), respectively, for 30 min, after which the amount of reducing sugars released was measured as previously reported (38). One unit of CMCase, Avicelase, and xylanase activity was defined as the amount of enzyme that released 1 μmol of reducing sugars per min, while 1 U of β-glucosidase activity was defined as the amount of protein that released 1 μmol of p-nitrophenol per min. LPMO activity was assessed by monitoring the amount of H₂O₂ accumulation using the Amplex red assay, as described previously (67). The reaction mixture was composed of 20 μl of the LPMO source (enzyme) and 180 μl of assay solution, which comprised 18 μl of 300 μM ascorbic acid, 18 μl of 500 μM Amplex red, 18 μl of 71.4 U/ml horseradish peroxidase (HRP), 18 μl of 1 M sodium phosphate buffer (pH 6.0), and 108 μl of high-performance liquid chromatography (HPLC)-grade water. Resorufin fluorescence was measured at an excitation wavelength of 530 nm and an emission wavelength of 580 nm after 10 min of incubation at 22°C using a multimode plate reader (Spectra Max M3). In reference experiments without LPMO, the background signal was measured and subtracted from the assay values. A standard curve obtained with various dilutions of H₂O₂ was used to calculate the amount of H₂O₂ liberated (micromoles per milliliter) due to the ascorbate-induced action of LPMO in the fungal secretome. The total cellulase (FPase) activity in the secretome was measured in terms of filter paper units (FPU) per milliliter of the original (undiluted) enzyme solution. The assay requires a fixed degree of conversion of the substrate from 50 mg of filter paper within 60 min at 50°C. An FPU is defined as the amount of enzyme required to produce 2 mg of glucose from 50 mg of filter paper within 60 min of incubation. The total protein content of each secretome was estimated by a bicinchoninic acid (BCA) assay using bovine serum albumin (BSA) as a standard.

SDS-PAGE and zymogram analysis.

Sodium dodecyl sulfate (SDS)-polyacrylamide gels (12%) were prepared, and proteins obtained following culture supernatant preparation were separated via SDS-polyacrylamide gel electrophoresis (PAGE) according to a previously described method (68). A Mini-Protean Tetra cell (Bio-Rad) with a gel size of 8.6 by 6.7 cm² was used for protein separation. After protein separation, the gels were first washed with 1× phosphate-buffered saline (PBS) to remove the SDS. Zymogram analysis was carried out using 5 mM 4-methylumbelliferyl β-d-glucopyranoside (MUG) as the substrate according to standard procedures in 50 mM citrate phosphate buffer (pH 4.0) for 10 min before visualization under UV light. Proteins in the gel were then stained with Coomassie blue R-250 (Sigma-Aldrich, USA), and the molecular mass of the proteins was determined by reference to standard proteins (Thermo Scientific, USA).

Avicel hydrolysis and product analysis of gluconic acid by HPLC-ELSD.

To quantitate the oxidative action of PfLPMO9 in the deconstruction of crystalline cellulose, an enzymatic hydrolysis reaction was set up using Avicel as the substrate under 10% solid loading in 50 mM sodium citrate phosphate buffer (pH 4.0) and 7 FPU/g of the PfMig1⁸⁸ secretome for 48 h at 50°C at 200 rpm in the presence and absence of 2 mM ascorbic acid as described previously by Dixit et al. (37). Samples were withdrawn at various intervals, boiled for 10 min to stop the reaction, and filtered through a 0.45-μm filter before quantification of the hydrolysis product (gluconic acid). The hydrolysates were analyzed using an Agilent 1260 series HPLC instrument coupled with a 1290 Infinity evaporative light scattering detector (ELSD). The oxidized products were estimated using a Rezex RSO-oligosaccharide Ag⁺ (4%) column (200 by 10 mm) with a Rezex RSO-oligosaccharide Ag⁺ (4%) guard column (60 by 10 mm) (Phenomenex), with a mobile phase of 100% water. Throughout the analysis, the column was kept at 80°C, and the mobile phase flow rate was maintained at 0.3 ml/min under isocratic conditions for 65 min. The chromatographic run was initiated by first equilibrating the column for 5 min followed by injecting 20 μl of the sample solution into the column. The ELSD was maintained at 45°C throughout. The nebulizer (nitrogen) gas pressure was set at 2.8 × 10⁵ Pa, and the detector gain was set at 9 × 10⁵ Pa. The concentration of gluconic acid was calculated from calibration curves of external standards (gluconic acid) purchased from Megazyme. To quantitate LPMO-driven gluconic acid production, differential gluconic acid (d-GlcA) measurements were performed by measuring the difference between the gluconic acid levels measured in the enzymatic hydrolysis reaction of Avicel in the presence and those in the absence of ascorbic acid (37).

Biomass saccharification and quantification of fermentable sugars.

The saccharification efficiency of the secretomes of all the strains used in the study with pretreated wheat straw was determined according to the method described previously by Ogunmolu et al. (31), with some modifications. The performance of the secretomes toward nitric acid-treated wheat straw was evaluated at 20% dry weight of biomass using an enzyme concentration of 30 mg/g biomass. Saccharification was performed in 50-ml screw-cap Falcon tubes in an incubator shaker at 50°C for 96 h. The reaction mixture included nitric acid-treated wheat straw under 20% dry weight loading in a 5-ml final reaction volume. The total protein content in the secretomes of all the fungal strains tested was measured, and the appropriate volume of the desired protein concentration (30 mg/g DBW) was added to the reaction mixture. The reactions were set up in 100 mM citrate phosphate buffer (pH 4.0), and the mixtures were incubated at 50°C with constant shaking at 300 rpm for 96 h. Samples were collected every 24 h and analyzed for the production of fermentable sugars. Control experiments were carried out under the same conditions, using substrates without enzymes (enzyme blank) and enzymes without substrates (substrate blank); a substrate-free negative control was set up by filling the Falcon tubes with 100 mM citrate phosphate buffer (pH 4.0), and the background of soluble sugars present in the respective biomass was determined by incubating each biomass in the absence of the enzyme. Following the completion of hydrolysis at each time point, the Falcon tubes were centrifuged at 3,500 rpm for 10 min in a swinging-bucket centrifuge (Eppendorf, Germany) to separate the solid residue from the digested biomass. Supernatants that were recovered after enzymatic hydrolysis of the pretreated wheat straw were analyzed by high-performance liquid chromatography with an Aminex HPX-87H anion-exchange column (Bio-Rad, USA) and a refractive index (RI) detector to analyze the released monosaccharides (glucose and xylose) by anion-exchange chromatography. The filtered mobile phase (4 mM H₂SO₄) was used at a constant rate of 0.3 ml/min with the column, and the RI detector temperature was maintained at 35°C. The concentration of each monosaccharide was calculated from calibration curves of external standards (xylose and glucose) purchased from Absolute Standards Inc. The following equations provided in NREL laboratory analytical procedure (LAP) TP-510-43630 were then used to determine the theoretical conversion of cellulose and hemicellulose (in percentages) to monomeric sugars:

$$\text{glucose yield}\left(\text{\%}\right)=\frac{\text{[glucose] × 0.9 × 100}}{f\text{× [pretreated biomass]}}$$

$$\text{xylose yield}\left(\text{\%}\right)=\frac{\text{[xylose] × 0.88 × 100}}{f\text{× [pretreated biomass]}}$$

where [glucose] is the glucose concentration (grams per liter), [xylose] is the xylose concentration (grams per liter) obtained after hydrolysis, [pretreated biomass] is the dry pretreated biomass concentration at the beginning of enzymatic hydrolysis (grams per liter), and f is the cellulose or hemicellulose fraction in the dry pretreated biomass (grams per gram) (39).

Data and statistical analysis.

All experiments were performed in triplicate, and the results are presented as the means and standard deviations. The data were compiled in a Microsoft Excel spreadsheet, where the averages and standard errors of the means were determined. All graphs were created using GraphPad Prism 8.0 software. The data were further evaluated by one-way analysis of variance (ANOVA) and multiple t tests using GraphPad Prism 8.0 software where appropriate.