Introduction
Hemicellulose, one of the main components of plant cell walls together with cellulose and lignin, is the second most abundant renewable carbon source on Earth (Geng et al. 2022). It refers to a group of heterogeneous polysaccharides, with xylans being the predominant form, that differ in their backbone, branches, and type and distribution of glycosidic linkages (Méndez-Líter et al. 2021a, 2021b). Several products of industrial interest can be obtained from hemicellulose such as oligosaccharides with prebiotic effect and fermentable sugars (mostly xylose) from which ethanol and xylitol, among others, can be synthesised (Méndez-Líter et al. 2021b; Geng et al. 2022). Moreover, in plant biomass saccharification processes, the removal of hemicellulose is of great importance to increase cellulose accessibility (Meng and Ragauskas 2014).
Hemicellulose degradation can be conducted in several ways; however, the enzymatic approach is preferred over the chemical one because it is more environmentally friendly and does not generate undesirable side compounds (Ostadjoo et al. 2019). Hemicellulases are a class of glycosyl hydrolases that act on hemicellulose, and they are extensively produced by filamentous fungi. The variable composition of hemicellulose requires the concerted action of multiple hemicellulases for its degradation, being the endo-β-1,4-xylanases and β-xylosidases the most important ones (Méndez-Líter et al. 2021b).
There is a tight regulation that controls the production of hemicellulases and typically relies on the presence of their corresponding substrate in the medium (de Eugenio et al. 2017; di Cologna et al. 2018). The most accepted model suggests that the large size of polysaccharides prevents their direct import into the cells and, in consequence, filamentous fungi need to constitutively express and/or express under carbon starvation certain hemicellulases (Benocci et al. 2017; Méndez-Líter et al. 2021a, 2021b). These enzymes, although produced at low levels, initiate hydrolysis releasing disaccharides and monosaccharides, which enter the intracellular environment and act as inducers of the entire hemicellulolytic system via substrate-specific transcription factors. In this sense, these factors not only control the expression of hemicellulases and other glycosyl hydrolases, but also regulate the expression levels of genes coding for specific transporters and those belonging to catabolic pathways (Benocci et al. 2017; Alazi and Ram 2018).
The ascomycete Talaromyces amestolkiae stands out for its remarkable production of glycosyl hydrolases (Prieto et al. 2021). Almost 200 genes encoding these enzymes have been predicted in its genome, which is a number significantly higher than those found in other fungi involved in cellulose and hemicellulose degradation, such as Trichoderma reesei, Penicillium oxalicum, or Aspergillus niger (de Eugenio et al. 2017). Several glycosyl hydrolases of this fungus, including 3 β-glucosidases, 2 endo-β-1,4-glucanases, 1 β-xylosidase, 2 endo-β-1,4-xylanases, and 2 α-L-arabinofuranosidases, have been characterised and used in saccharification processes, oligosaccharide synthesis, and glycosylation of bioactive compounds (Prieto et al. 2021). In addition, T. amestolkiae strains are an attractive source of natural red colourants (de Oliveira et al. 2022) and other secondary metabolites, such as meroterpenoids, isocoumarins, and benzofurans (Li et al. 2022).
Enhancing the production of hemicellulases in filamentous fungi is a very attractive approach to improve the valorisation of the hemicellulosic fraction of plant biomass. In this regard, an increase in glycosyl hydrolase production by optimising the fermentation process has recently been reported in T. amestolkiae (de Eugenio et al. 2024). Nonetheless, these types of fungal fermentations are often limited by the use of insoluble substrates acting as inducers and the mycelial nature of the fungus (Mattam et al. 2022). In order to further enhance hemicellulase production in this fungus, it is convenient to develop genetic manipulation tools.
Several transformation systems are available for filamentous fungi based on two types of selection markers, which are antibiotic-resistance genes and auxotrophic complementation. Since the use of the first approach involves safety concerns about the spread of antibiotic resistant strains and there are also legal requirements from industries handling food-related products, auxotrophic complementation is preferred (Martín 2015; Mei et al. 2019). Among this type of selection markers, pyrimidine auxotrophy is the most widely used strategy and has already been adapted successfully to some Talaromyces spp. (Borneman et al. 2001; Inoue et al. 2013; Delmas et al. 2014; Zhang et al. 2022). The de novo pathway for pyrimidine biosynthesis comprises six biochemical steps and ends up producing uridine monophosphate (UMP) (Figure S1). pyrF (Aleksenko et al. 1999) and pyrG (Oakley et al. 1987) genes, encoding orotate phosphoribosyltransferase (OPRTase) and orotidine 5′-monophosphate decarboxylase (OMPdecase), respectively, play an essential role in this pathway. pyrF− and pyrG− mutants can be easily obtained by selecting for resistance to the antimetabolite 5-fluoroorotic acid (5-FOA), which is transformed by OPRTase and OMPdecase to the toxic 5-fluoro-UMP. These pyrimidine auxotrophs will then need the addition of uridine and/or uracil as nutritional requirements to be able to grow (Díez et al. 1987).
On the other hand, the most promising approach to improve hemicellulase production in filamentous fungi is manipulating, through genetic engineering, the transcriptional factors that regulate their expression. The main transcriptional activator of hemicellulose degradation and xylose utilisation is XlnR/XYR1/XLR1 (Van Peij et al. 1998b). This transcriptional factor contains a Zn2Cys6-binuclear cluster domain necessary for DNA-binding and a middle homology region with regulatory functions. The binding efficiency and the set of genes regulated by XlnR/XYR1/XLR1, as well as its role in cellulose degradation, differ between species (Benocci et al. 2017; Alazi and Ram 2018).
Several attempts have been made to enhance hemicellulase production in filamentous fungi by modulating the transcriptional regulation of XlnR/XYR1/XLR1. The first set of strategies involved the addition of extra copies of this transcriptional activator (van Peij et al. 1998a; Llanos et al. 2019) or its overexpression with constitutive strong promoters such as the ones of gpdA (Tamayo-Ramos and Orejas 2014; Li et al. 2015), tef1 (Marui et al. 2002), and pdc1 (Wang et al. 2013) genes, which led to an improved hemicellulolytic activity. However, better results were obtained when single residue substitutions were introduced in the C-terminal region of XlnR/XYR1/XLR1. In P. oxalicum, T. reseei, and Neurospora crassa, Ala871Val, Ala824Val, and Ala828Val, respectively, increased to a higher extent the production of hemicellulases (Derntl et al. 2013; Craig et al. 2015; Gao et al. 2017). In
This work aims to increase hemicellulase production in T. amestolkiae by using an approach based on the forced expression and engineering of its transcriptional factor XlnR. To achieve this goal, it was necessary to first develop a genetic modification method, which will also enable the use of this fungus in other biotechnological applications.
Experimental Procedures
Strains, Media and Growth Conditions
The fungus T. amestolkiae (hereinafter Tam_WT) was isolated from cereal wastes and deposited in the culture collection of Center for Biological Research Margarita Salas (CIB, Madrid, Spain), with reference A795. It was maintained at 28 °C in plates of potato dextrose agar (PDA, Becton Dickinson) and Aspergillus complete medium (MCA) (Cove 1966), which contained 1% (w/v) glucose as a carbon source and 5 mM diammonium tartrate as a nitrogen source. Cultures for T. amestolkiae transformation and genome sequencing were grown from conidiospores for 2–3 days in MCA liquid medium. To assess enzymatic activities and obtain crudes for saccharification studies, Tam_WT and its mutant strains (Table 1) were first cultivated on CSS medium (de Eugenio et al. 2017) from conidiospores (4.8 × 104 sp/mL). After 4 days of growth, 2.5 mL of these inocula were used to start cultures on Mandels medium (de Eugenio et al. 2017) supplemented with different substrates at 1% (w/v): beechwood xylan (Megazyme), glucose (Merck) and glycerol (ITW Reagents). All liquid experiments were conducted in 250 mL flasks containing 50 mL of medium, at 28 °C and 180 rpm. 5 mM uridine and 5 mM uracil were added in plates and liquid cultures when required to complement pyrimidine auxotrophy. The conidiospores of Tam_WT and its mutant strains were preserved at −80 °C in 15% (v/v) glycerol, after determining their concentration using a Neubauer chamber.
TABLE 1
| Description, Reference | |
| Fungal strain | |
| Tam_WT | Wild-type |
| Tam_pyrG14 | Pyrimidine auxotroph resistant to 5-FOA, this study |
| Tam_p1439a | Transformed with pFNO3 (p1439), this study |
| Tam_p1393 (colony #1)a | Transformed with pRG3-AMA1-NotI (p1393), this study |
| Tam_XlnR (colony #7)a | Transformed with p1393-PgpdA-XlnR-TtrpC, this study |
| Tam_XlnRA788V (colony #10)a | Transformed with p1393-PgpdA-XlnRA788V-TtrpC, this study |
| Tam_XlnRV785F (colony #12)a | Transformed with p1393-PgpdA-XlnRV785F-TtrpC, this study |
| Plasmid | |
| pFNO3 (p1439) | pPV472 backbone, ampR, kanR, |
| pRG3-AMA1-NotI (p1393) | pUC19 backbone, ampR, |
| pUC18-PgpdA-XlnR-TtrpC | pUC18 backbone, ampR, PgpdA::xlnR::TtrpC cassette, this study, for cloning steps |
| p1393-PgpdA-XlnR-TtrpC | P1393 derivative, PgpdA::xlnR::TtrpC cassette, this study |
| pUC18-PgpdA-XlnRA788V-TtrpC | pUC18 backbone, ampR, PgpdA::xlnRA788V::TtrpC cassette, this study, for cloning steps |
| p1393-PgpdA-XlnRA788V-TtrpC | P1393 derivative, PgpdA::xlnRA788V::TtrpC cassette, this study |
| pUC18-PgpdA-XlnRV785F-TtrpC | pUC18 backbone, ampR, PgpdA::xlnRV785F::TtrpC cassette, with this study, for cloning steps |
| p1393-PgpdA-XlnRV785F-TtrpC | P1393 derivative, PgpdA::xlnRV785F::TtrpC cassette, this study |
Construction of Plasmids
All plasmids used in this work can be found in Table 1. The first attempt to transform T. amestolkiae was performed with the integrative plasmid pFNO3 (p1439 in our collection) (Yang et al. 2004), which contains an ATG-less chimera composed of a repetition of 5 Gly-Ala residues (GA5x) fused in frame to the sGFP, and the
Next, to obtain maximum expression levels, we used the autonomously replicating plasmid pRG3-AMA1-NotI (p1393 in our collection) (Osherov et al. 2000) carrying the AMA1 replicator of Aspergillus nidulans and
Another cassette, harbouring xlnR gene of Tam_WT (GeneBank Accesion Number BHQ10_006915, Figure S2A,B) under the control of
On the other hand, pUC18-PgpdA-XlnR-TtrpC was used as a template to develop two variants of the transcriptional activator by directed mutagenesis. The alanine at position 788 was substituted by a valine (Ala788Val) in one variant, and the valine at position 785 was replaced by a phenylalanine (Val785Phe) in the other variant. For the mutagenic PCR, the ExpandLong Template PCR System (Roche) was applied using the primers of Table S1. The PCR product was digested by DpnI (New England Biolabs) to hydrolyse the parental methylated DNA. After sequencing the new cassettes, the cloning procedure continued as explained before to generate p1393-PgpdA-XlnRA788V-TtrpC and p1393-PgpdA-XlnRV785F-TtrpC plasmids.
Extraction of
In order to analyse the genotype of Tam_WT and its mutant strains, gDNA was extracted from their conidiospores. Briefly, sterile wooden toothpicks were used to collect conidiospores of fresh fungal cultures grown on PDA or MCA plates. Then, conidiospores were placed on 200 μL breaking buffer containing 2% (v/v) Triton X-100, 1% (v/v) SDS, 100 mM NaCl, 10 mM Tris–HCl pH 8 and 1 mM EDTA pH 8. Approximately 150 mg of glass beads (0.4–0.6 mm, Merck) were added to help disrupt the conidiospores. Samples were vortexed for 30 s and subsequently incubated at 70°C for 30 min. Three more vortexing steps of 30 s were performed while incubating. After that, 200 μL phenol/chloroform/isoamyl alcohol (25:24:1 v/v, Meck) were added and samples were vortexed again for 5 min. Disrupted samples were finally centrifuged at 14,000 rpm for 5 min, and 50 μL of the aqueous solution carrying the dissolved gDNA was transferred to a clean tube.
Isolation of 5-
The generation of 5-FOA-resistant mutants from Tam_WT was conducted in MCA plates with 2 mg/mL 5-FOA (Apollo Scientific) and 5 mM uridine and uracil. A toothpick was heavily loaded with fresh conidiospores and used to inoculate a plate at 7 evenly distributed spots. As the toothpick was used repeatedly without reloading, the conidiospores concentration decreased in each spot, which ensures reaching the optimal conditions for the generation of 5-FOA-resistant sectors. 16 plates were prepared following this procedure, resulting in 112 inoculation spots, and incubated at 28 °C for up to 12 days. 5-FOA-resistant mutants were then purified through one selective pass on MCA plates containing the same concentrations of 5-FOA, uracil, and uridine. Singles colonies of each mutant were subsequently tested for pyrimidine auxotrophy on MCA plates with and without uridine and uracil, since the mutants will need these two compounds to grow. To check the genotype of the mutants, gDNA was extracted as described before and used as a template for the amplification of pyrF and pyrG genes involved in pyrimidine synthesis. The PCRs were performed in 50 μL reaction mixtures that contained the PrimeSTAR GXL DNA Polymerase (Takara Bio) and specific primers for both genes (Table S1). The amplified products were purified using the QIAquick PCR purification kit (Qiagen) and finally sequenced with the EZ-Seq service of Macrogen. Alignments of the orthologous pyrF and pyrG genes with the wild-type ones (GeneBank accesion numbers BHQ10_006112 and BHQ10_007154, respectively) were performed to determine the position and type of acquired mutations.
Fungal Transformation and Selection of Transformants
The auxotroph mutant Tam_pyrG14 was transformed as described for
Then, protoplasts were transformed with the circular plasmids of Table 1 by applying CaCl2 and polyethylene glycol (PEG). In the case of plasmid p1439, 2.5 μg of DNA were used, whereas 600 ng of DNA were employed for the autonomously replicating plasmid p1393 and its derivatives. The transformation procedure was carried out in 50 mL tubes by mixing 50 μL of solution E (600 g/L PEG6000, 10 mM Tris–HCl pH 7.5, 0.12 M CaCl2), 40 μL of solution D, the circular plasmids dissolved in 10 μL of sterile water, and 200 μL of the protoplast suspension. A control without DNA was also prepared. The tubes were gently mixed and incubated on ice for 20 min. Subsequently, 1 mL of solution E was added to each tube, and the mixture was incubated for 5 min at RT. Finally, the volume was adjusted to 5 mL with solution D and completed to 20 mL with recovery medium (MCA with 1 M sucrose as osmotic stabiliser and without uracil and uridine) containing 0.75% (w/v) agar. The tubes were briefly mixed by inversion to embed the protoplasts, and their content distributed across 4 recovery medium plates, with1.5% (w/v) agar and without uracil and uridine, to select transformed colonies. The no-DNA control was plated onto 2 selective recovery medium plates (negative control) and 2 plates supplemented with uridine and uracil (positive control for protoplast regeneration). Plates were incubated at 28 °C for at least 7 days.
Potential transformants were further isolated through one selective pass on MCA plates without uracil and uridine. Then, their gDNA was extracted and used to check by PCR the presence of targeted genes of the plasmids. In the case of mutants transformed with p1393 and its derivatives, an enzymatic activity screening to select the best performing mutant of each transformation was carried out.
Whole Genome Sequencing of Tam_XlnRV785F Recombinant Strain
For sequencing the genome of Tam_XlnRV785F, cultures were grown on MCA liquid medium, mycelium was harvested by centrifugation, and gDNA was extracted using the NZY plant/fungi gDNA isolation kit (NZYtech). Paired-end sequencing was then performed on an Illumina PE150 system using the services of the company Novogene, which was also in charge of removing low-quality reads and reads that maintained adaptors using Fastp v.0.20.0 software (Chen et al. 2018). A high quality (Q30 of 92.07%) DNA library composed of 17,434,018 clean reads of 150 bp in length was received. Finally, the Bioinformatics and Biostatistics facility of the CIB analysed the sequencing data to determine the behaviour of p1393-PgpdA-XlnRV785F-TtrpC plasmid in Tam_XlnRV785F. With this purpose, the clean reads were mapped using the BWA/0.7.17 aligner (Li and Durbin 2009) against the scaffolds of T. amestolkiae genome (GenBank Accession Number ASM189636v1) and the sequence of the plasmid. Further studies were conducted with SAMtools/1.14 (Danecek et al. 2021) and R/4.3.2 () softwares.
Enzyme Activity and Protein Assays
To evaluate the enzymatic activities and extracellular protein concentration of Tam_WT, Tam_pyrG14, Tam_p1393, Tam_XlnR, Tam_XlnR,A788V and Tam_XlnRV785F, they were grown on Mandels medium supplemented with 1% (w/v) xylan, glucose, or glycerol. Samples were taken daily and centrifuged for 10 min at max. speed and 4 °C to retrieve the supernatants.
β-xylosidase activity was determined spectrophotometrically at 410 nm by the release of 4-nitrophenol (pNP) (ε410 = 15,200 M−1 cm−1) using 0.1% (w/v) p-nitrophenyl-β-d-xylopyranoside (pNPX, Sigma-Aldrich) in 50 mM sodium acetate buffer pH 5. Reactions (200 μL) were incubated for 10 min at 50 °C and 1200 rpm and stopped by changing the pH through the addition of 500 μL of 2% (w/v) Na2CO3. Endoxylanase activity was calculated from the hydrolysis of 3% (w/v) beechwood xylan (Megazyme) dissolved in 50 mM sodium acetate buffer pH 5. Reactions (500 μL) were incubated for 10 min at 50 °C and 1200 rpm and stopped by heating at 100 °C for 5 min. After centrifuging for 5 min at 20,000 × g and RT, the release of reducing sugars was measured spectrophotometrically at 540 nm using the Somogyi-Nelson method (Somogyi 1945). One unit of activity was defined as the amount of enzyme capable of releasing 1 μmol of pNP or reducing sugars per minute. In the initial screenings conducted to select the best performing transformants, endoxylanase activity was determined using Azo-Xylan Birchwood Powder (Megazyme) following the manufacturer's protocol.
Extracellular protein concentration was quantified utilising the Bradford dye reagent (Bio-Rad), based on the original Bradford method (Bradford 1976), and Bovine Serum Albumin (BSA, Thermo Fisher Scientific) as the standard. Measurements were performed with a spectrophotometer at 595 nm.
For SDS-PAGE and zymograms, enzymatic crudes were concentrated 10× and dialysed against 100 mM sodium acetate buffer pH 5 in Amicon Ultra—0.5 mL 3 kDa devices (Merck). Crudes (14 μg of extracellular proteins) were heated up at 98 °C for 10 min in a buffer containing SDS and β-mercaptoethanol before being loaded into 10% polyacrylamide gels. Precision Plus Protein Dual Colour Standard (Bio-Rad) was the molecular weight marker employed, and proteins were stained with Coomassie Brilliant Blue R-250 (Bio-Rad). For endoxylanase zymograms, 0.2% (w/v) beechwood xylan was incorporated into SDS-PAGE 10% polyacrylamide gels, and 1.5 μg of extracellular proteins from the crudes were loaded. After electrophoresis, gels were washed in 5% (v/v) Triton X-100 solution for 1 h to remove SDS and incubated in distilled water for 30 min at 30 °C to set the conditions for endoxylanase activity. Then, gels were stained with 0.1% (w/v) Congo Red for 30 min and washed with 1 M NaCl until activity bands became visible. Gels were finally immersed in 5% (v/v) acetic acid to turn the background black. For β-xylosidase zymograms, isoelectric focusing was carried out on 5% polyacrylamide gels using pH 3–10 ampholytes (GE Healthcare), with 1 M H3PO4 and 1 M NaOH as anode and cathode buffers, respectively. 1 μg of extracellular proteins were loaded for each crude. The pH gradient was directly measured on the gels with a contact electrode (Crison). β-Xylosidase activity was detected by incubating the gels with 2 mM p-methylumbelliferyl-β-d-xylopyranoside (Sigma-Aldrich) for 10 min at 30°C, and fluorescence was visualised under UV light using a Gel Doc XR+ system (Bio-Rad).
Biomass Determination
Tam_WT, Tam_p1393, Tam_XlnR, Tam_XlnRA788V and Tam_XlnRV785F growth in liquid culture was compared by estimating ergosterol content, which is a specific sterol of fungal cell walls (Ng et al. 2008). 2 mL samples were centrifuged (10 min at max. speed and 4 °C) and lyophilized. The pelleted biomass was transferred to 8 mL KIMAX tubes (DWK Life Sciences) and dissolved in 200 μL petroleum ether (Merck) and 800 μL 10% (w/v) KOH (Merck) solubilised in methanol (Merck). Then, samples were sonicated for 15 min in an ultrasound bath and, after 45 min at RT, incubated at 70 °C for 90 min. Once cooled down, 0.4 g/L cholesterol (Merck) was added as an internal standard, together with 200 μL water and 400 μL petroleum ether for ergosterol and cholesterol extraction. Samples were mixed with a 30 s vortex step and let them stand until the phases were separated. The organic phase, containing petroleum ether together with ergosterol and cholesterol, was placed in a 2 mL tube and dried by evaporation at 25 °C to repeat one more time the extraction. Finally, the extracted samples were resuspended in 400 μL methanol, mixed and filtered through 0.22 μm PTFE syringe filters (Agilent).
The detection and quantification of ergosterol and cholesterol was carried out by HPLC in an Agilent 1200 series LC instrument equipped with a reverse phase ZORBAX Eclipse plus C18 column (Agilent). The mobile phase was a mixture of methanol and acetonitrile (80:20, v/v), at a flow rate of 1 mL/min, and the volume of sample injected was 10 μL. Ergosterol and cholesterol peaks were detected at 254 and 204 nm, respectively, and quantified by comparing their areas with the calibration curves of each compound prepared in methanol. Ultimately, fungal biomass was estimated from the ergosterol content using a calibration curve plotting ergosterol content per gram of lyophilized Tam_WT biomass.
Saccharification Assays
Tam_WT and Tam_XlnRV785F cultures, grown on Mandels medium supplemented with xylan, glucose and glycerol, were centrifuged for 10 min at maximum speed and 4 °C when highest levels of enzymatic activities were reached. Then, the supernatants were sequentially vacuum filtered through 0.8, 0.45 and 0.22 μm nitrocellulose membrane discs (Millipore) to obtain the enzymatic crudes.
Saccharification reactions were performed in 2 mL tubes. Each reaction was composed of 450 uL of filtered enzymatic crudes mixed with 50 mM sodium acetate buffer pH 5 containing 5% (w/v) beechwood xylan (Megazyme) in a final volume of 1 mL. The reactions were incubated at 50 °C and 1200 rpm for 24 h. Samples were taken periodically, and the xylose released was immediately quantified using the D-Xylose Assay Kit (Megazyme).
Bioinformatic Analyses of
The nucleotide sequences of T. amestolkiae pyrG, pyrF, and xlnR were retrieved from the genomic information available under the GenBank Accession Number ASM189636v1. The alignment of wild-type pyrF and pyrG with their corresponding mutated homologues was performed with Snapgene software (). The identification of A788 and V785 residues in XlnR was conducted by alignment using Clustal Omega (Madeira et al. 2024) with the orthologous genes of P. oxalicum (Gao et al. 2017),
Statistical Tests
Activity measurements to compare Tam_WT and its mutant strains were carried out employing biological triplicates and technical duplicates. Statistical differences were determined with the Student's t-test (n = 3, *p < 0.05 and **p < 0.01).
Results and Discussion
Isolation of
Genetic transformation of T. amestolkiae strains had not been reported yet. A widely used method in filamentous fungi, based on the complementation of pyrimidine auxotrophs (Martín 2015; Mei et al. 2019), was chosen for this work. For that, auxotrophic mutations in the de novo pyrimidine biosynthetic pathway were selected using resistance to 5-FOA (Díez et al. 1987). The optimal 5-FOA concentration preventing growth of the fungus was determined to be 2 mg/mL on MCA. Sectors resistant to 5-FOA were obtained by incubating wild-type colonies for 5 to 12 days at 28 °C. 14 putative pyr− sectors were selected and the corresponding strains purified by plating isolated colonies derived from conidiospores. These strains were further confirmed to be uridine/uracil auxotrophs (Figure S3A). Their genotypes were determined after PCR amplification and Sanger sequencing of the coding regions of pyrF (785 bp long) and pyrG (887 bp long), which encode the enzymes catalysing the two final steps of the de novo pyrimidine biosynthetic pathway (Figure S1). In all cases, convincing causative mutations in either pyrG or pyrF were detected. These mutations are listed in Table 2.
TABLE 2 List of
| Mutant | Mutation type | Nucleotide change | Protein change |
| pyrF1 | Nonsense | c.C606T | Early stop codon after E183 |
| pyrF2 | Frameshift | c.Δ(A421 → C458) | Truncation after N122 |
| pyrG3 | Frameshift | c.Δ(G720 → A741) | Truncation after D223 |
| pyrF4 | Deletion of 3 bp | c.Δ(C506 → T508) | Deletion of V151 |
| pyrG5 | Deletion of 18 bp | c.Δ(A558 → T575) | Deletion of T170-D175 |
| pyrF6 | Frameshift | c.ΔA549 | Truncation after I164 |
| pyrF7 | Missense | c.G3A | Met1Ile, loss start codona |
| pyrF8 | Missense | c.T2C | Met1Thr, loss start codona |
| pyrF10 | Frameshift | c.117insT | Truncation after F40 |
| pyrG11 | Deletion of 12 bp | c.Δ(G822 → G833) | Deletion of A258-Q261 |
| pyrF13 | Missense | c.T2C | Met1Thr, loss start codona |
| pyrG14 | Frameshift | c.Δ(C144 → G154) | Truncation after L48 |
| pyrF15 | Frameshift | c.136insT | Truncation after L46 |
| pyrG16 | Frameshift | c.279insA | Truncation after T76 |
The most frequent mutations are deletions, while substitutions and insertions are less represented. Except in three strains, these mutations caused premature truncations of the protein or resulted in early stop codons or loss of the start Met codon. Therefore, these data strongly indicate that these mutations lead to a complete loss-of-function. Indeed, all the mutants were strictly dependent on supplementation with uridine and uracil for their growth. T. amestolkiae pyrG14 (Figure S3B, hereinafter Tam_pyrG14), resulting in the earliest frameshift in pyrG (removing the 231 C-terminal residues of the 278 full length OMPdecase), was chosen for subsequent work. The deleted residues form part of an α-helix located upstream of the main catalytic residues of the enzyme (Traut and Temple 2000) (Figure S4), strongly suggesting that pyrG14 is a null allele. Deletion of 11 pb of the coding region near the codons of catalytic site residues would help prevent the isolation of spontaneous pyrG+ revertants during selection of transformants. These results, in combination with previous reports on T. marneffei (Borneman et al. 2001), highlight the utility of 5-FOA to select pyrimidine auxotrophs in Talaromyces species.
Transformation of
Once a pyrimidine auxotroph of T. amestolkiae was obtained, we used PEG/calcium-mediated transformation of protoplasts following the protocol described for
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Forced Expression of the Transcriptional Activator
The establishment of a transformation protocol for T. amestolkiae enabled the engineering of hemicellulase production through genetic manipulation. We introduced the autonomously replicating plasmid pRG3-AMA1-NotI (p1393 in our collection) (Osherov et al. 2000) carrying a transgene in which T. amestolkiae's transcriptional activator XlnR was expressed under the control of the constitutive
In addition to wild-type XlnR, two mutant versions of the regulator were expressed, based on reports that these mutants increased expression of hemicellulolytic genes in P. oxalicum,
Tam_pyrG14 was transformed with these three plasmids and 6–8 colonies that complemented the pyrimidine auxotrophy were isolated from each transformation. In addition, the empty p1393 vector was also transformed to study the effect that this autonomously replicating plasmid has on fungal growth and to discard the unlikely possibility that the vector on its own was responsible for increasing hemicellulase production. Tam_XlnR, Tam_XlnRA788V, Tam_XlnR,V785F and Tam_p1393 transformants were analysed on 50 mL Mandels medium supplemented with 1% (w/v) beechwood xylan as inducer, to select the best β-xylosidase and endoxylanase producer (Figures S5–S8) using Tam_WT as control, and Tam_pyrG14 to confirm the lack of effect of the pyrimidine auxotrophy. The transformants displaying the highest activity values, which were Tam_XlnR #7, Tam_XlnRA788V #12, Tam_XlnRV785F #10, and Tam_p1393 #1, were chosen for further studies.
These transformants and Tam_WT were cultured on 50 mL Mandels medium with 1% (w/v) beechwood xylan. β-xylosidase and endoxylanase activities, extracellular protein concentration, and fungal growth were measured daily. Figure 2A,B, reveals that Tam_XlnR, Tam_XlnR,A788V and Tam_XlnRV785F mutants increased both β-xylosidase and endoxylanase activities compared to Tam_WT and Tam_p1393, but to a different extent. When calculating the fold change in enzymatic activity of the mutants relative to Tam_WT on day 4 of cultivation, it became evident that the improvement in Tam_XlnR is not very pronounced, showing 1.4-fold and 2.0-fold increments in β-xylosidase and endoxylanase activities, respectively. This suggests that forced expression of xlnR does not lead to a major increase in β-xylosidase and endoxylanase activities.
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While similar results have been reported with T. reesei (Mach-Aigner et al. 2008; Wang et al. 2013), P. oxalicum (Li et al. 2015), Penicillium canescens (Serebryanyi et al. 2006) and Fusarium oxysporum (Calero-Nieto et al. 2007), other reports concluded that xlnR overexpression in
Tam_XlnRA788V transformant showed an increase of 1.5-fold for β-xylosidase activity and of 2.4-fold for endoxylanase activity on day 4 of cultivation, while in Tam_XlnRV785F β-xylosidase activity increased 3.3-fold and endoxylanase activity 3.9-fold. In addition, measurements of extracellular protein concentration revealed that Tam_XlnRA788V and especially Tam_XlnRV785F secreted more protein than the rest of the strains (Figure S9A). Therefore, introducing single residue substitutions in specific regions of XlnR is a more effective approach to improve hemicellulase production than the sole forced expression of this transcriptional activator. In this regard, in T. amestolkiae, Val785Phe has a larger effect than Ala788Val.
Comparison between a wild-type strain and these three XlnR transformants in a single experiment has not been reported yet. In P. oxalicum, xlnR overexpression during growth on wheat bran showed minimal increases in β-xylosidase and endoxylanase activities and in extracellular protein concentration compared to the significant improvement observed with the XlnRA871V mutant. In line with the data from Tam_XlnRV785F, this enhancement was particularly high in the late phases of cultivation (Gao et al. 2017). Nevertheless, the alternative mutation XlnRV868F has not been studied in P. oxalicum. In the case of T. reesei, equivalents of these three strains have been assayed under varying conditions, which makes direct comparison between them unreliable (Derntl et al. 2013; Ellilä et al. 2017; Lv et al. 2023; Zhao et al. 2023).
In order to determine if this improvement of β-xylosidase and endoxylanase activities alters the pattern of proteins secreted by T. amestolkiae, a SDS-PAGE analysis was carried out using equal extracellular protein concentrations of crudes at day 4 of cultivation (Figure 2C). Results revealed a very similar protein pattern across all strains. Given the complexity of T. amestolkiae crudes, which contain a diverse mixture of proteins, directly identifying β-xylosidases and endoxylanases from the gel is challenging. To overcome this limitation, zymograms were conducted for these enzymatic activities. The endoxylanase zymogram was performed employing a SDS-PAGE gel and displayed four bands, with the two larger ones being more intense in Tam_XlnR, Tam_XlnR,A788V and Tam_XlnRV785F mutants than in the wild-type and p1393 control (Figure 2D). The β-xylosidase zymogram was carried out using an isoelectric focusing gel, since BxTW1, a characterised enzyme from T. amestolkiae with this activity (Nieto-Domínguez et al. 2015), is a dimeric protein that would be difficult to renature after SDS-PAGE. This zymogram showed a single band of higher intensity in Tam_XlnRV785F compared to those in the other strains (Figure 2E). These findings suggest that the enhancement of hemicellulase activities results from the differential abundance of specific β-xylosidases and endoxylanases, rather than an overall distinct protein secretion pattern.
Fungal growth was monitored by measuring ergosterol content and correlating it with biomass concentration. Figure 3A shows that Tam_XlnR, Tam_XlnR,A788V and Tam_XlnRV785F grow slower than Tam_WT and Tam_p1393, with these differences being more pronounced on day 1 of cultivation and becoming less noticeable as time progresses. This observation is important because it demonstrates that more hemicellulases are being produced despite diminished biomass. The negative effect on growth caused by forced expression of xlnR and its mutant variants is likely due to a metabolic burden associated with increased synthesis of this transcriptional activator. The higher hemicellulase production observed in the Tam_XlnRV785F recombinant strain does not seem to worsen this metabolic burden since no substantial differences in growth behaviour can be seen when compared with Tam_XlnR. Forced expression of xlnR and its derivatives also changes mycelial morphology, as depicted in Figure 3B: Tam_WT and Tam_p1393 grow in a dense filamentous form with homogeneously dispersed hyphae, while with Tam_XlnR, Tam_XlnR,A788V and Tam_XlnRV785F spherical pellets resulting from hyphal aggregation are observed. These differences in fungal morphology would need to be taken into account when scaling up hemicellulase production because they can greatly affect the performance of the process (Veiter et al. 2018).
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Lastly, there were no major disparities between Tam_p1393 and Tam_WT in any of the parameters measured. Thus, the autonomously replicating plasmid itself is not responsible for any enhancement in hemicellulase activity or for differences in growth among the XlnR transformants. This is in agreement with findings in
In light of all these results, it was decided to further characterise the Tam_XlnRV785F recombinant strain. By whole genome sequencing, we confirmed that the plasmid was autonomously replicating since no integration events were detected. Thus, all the phenotypic effects previously described should be attributed to the transgenes amplified with the vector, and not to any disturbance in T. amestolkiae genome.
Partial Deregulation of Hemicellulase Production in Tam_XlnRV785F Transformant When Using Glucose and Glycerol as Carbon Sources
To evaluate the production of hemicellulases by Tam_XlnR and Tam_XlnRV785F under non-inducing and carbon repressing conditions, they were cultured together with Tam_WT on 50 mL Mandels medium supplemented with 1% (w/v) glucose. Figure 4A,B shows that Tam_XlnRV785F is able to secrete high amounts of β-xylosidases and endoxylanases while barely any activity is detected in Tam_WT. Specifically, a 16.9-fold improvement in β-xylosidase activity on day 5 of cultivation and a 31.9-fold increase in endoxylanase activity on day 3 of growth were observed. This reveals an important partial or complete deregulation of the hemicellulolytic system in this recombinant strain, which is capable of circumventing the strong inhibition that glucose exerts in the wild-type by carbon catabolite repression. Regarding extracellular protein concentration, Tam_XlnRV785F maintained it below that of Tam_WT at the beginning of cultivation, until it surpassed it on day 4 (Figure S9B). However, this final increase in extracellular protein concentration was markedly reduced in Tam_XlnR, as well as the enhancement of β-xylosidase and endoxylanase activities, reaching fold increases of only 1.8 and 1.7 at days 5 and 3 of cultivation, respectively. The modest improvement in hemicellulase production by Tam_XlnR, compared to Tam_XlnRV785F, strongly suggests that the deregulation is not attributable to forced expression of the gene encoding the transcriptional activator XlnR, but rather to the Val785Phe substitution.
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Comparable outcomes were obtained with the sole xyr1 overexpression in T. reesei when grown on glucose and glycerol (Derntl et al. 2019). In agreement with our results, Ala824Val and Val821Phe substitutions in T. reesei XYR1 and Val756Phe in
On the other hand, the SDS-PAGE analysis shows larger differences in the intensities of low molecular weight bands between Tam_XlnRV785F and both Tam_XlnR and Tam_WT, compared to the xylan crudes (Figure 4C). This is in line with the observed results in the zymograms (Figure 4D,E), where the β-xylosidase band is barely detectable in Tam_XlnR and Tam_WT, and the four endoxylanase bands are not visible at all. Consistent with the enzymatic activity data, these observations suggest that the Val785Phe substitution in XlnR enables high β-xylosidase and endoxylanase production on glucose, since the corresponding bands are present in the mutant. However, there are also variations in the intensities of Tam_XlnRV785F bands in the endoxylanase zymograms between the xylan (Figure 2D) and glucose (Figure 4D) crudes. Specifically, the enzymes corresponding to the two lower molecular weight bands are more difficult to detect in the glucose crudes, with this disparity being more evident for the 20 kDa band. This finding indicates that the regulation of hemicellulase production is not completely overridden with the Val785Phe mutation. The most likely explanation for this phenomenon is that, besides XlnR, several other transcription factors are involved in the regulation of the hemicellulolytic genes, and they may act differently depending on the substrate used (Tani et al. 2014; Llanos et al. 2019). For example, the transcriptional activators TctA in Talaromyces cellulolyticus, and ACE2 and ACE3 in T. reesei, have been shown to control hemicellulase production (Benocci et al. 2017). There are other transcription factors, such as CreA/Cre1/Cre-1 in most fungi, ACE1 and XPP1 in T. reesei and HCR-1 in
T. amestolkiae has been demonstrated to be an excellent producer of robust and efficient glycosyl hydrolase cocktails on media with different carbon sources, including low-cost and renewable substrates (Méndez-Líter et al. 2021a; Prieto et al. 2021). In this sense, glycerol is an abundant side-product generated from biodiesel production that could be used to obtain hemicellulases (Monteiro et al. 2018). To test this, Tam_XlnRV785F and Tam_WT were grown on 50 mL Mandels medium supplemented with 1% (w/v) glycerol. The recombinant strain showed a notably high β-xylosidase activity with a 13.8-fold increment on day 6 of cultivation compared to Tam_WT as well as a markedly elevated endoxylanase activity, with a 22.7-fold increase at day 4 of growth. In contrast, barely any of these activities were detected in Tam_WT (Figure 5A,B). Moreover, the measurements of extracellular protein concentration revealed that Tam_XlnRV785F secretes more proteins than Tam_WT throughout the entire cultivation time (Figure S9C). Regarding the SDS-PAGE and zymogram analyses (Figure 5C–E), similar results to those obtained with the glucose crudes were observed, although the differences in β-xylosidase and endoxylanase abundance between Tam_XlnRV785F and Tam_WT are not that pronounced in this case. As explained before, there are also variations in the band intensity pattern of Tam_XlnRV785F depending on the substrate used, demonstrating that some level of regulation still remains. In the case of glycerol, the most intense bands in the endoxylanase zymogram (Figure 5D) are those around 50 and 20 kDa, while the 37 kDa one (the most abundant band in the xylan and glucose crudes) is barely detectable.
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Enhanced Xylan Saccharification Using the Enzymatic Cocktails of Tam_XlnRV785F Transformant
To assess the performance of the improved enzymatic cocktails produced by Tam_XlnRV785F compared to those of Tam_WT, they were applied for the saccharification of beechwood xylan. Thus, Tam_WT and Tam_XlnRV785F crudes were harvested from the cultures using different substrates when maximal hemicellulase activities were observed: after 4 days of growth in the case of xylan and glucose, and on day 6 of cultivation for glycerol. β-xylosidase and endoxylanase activities, as well as extracellular protein concentrations of each crude used for the saccharification are displayed in Table 3. Since the improvement of hemicellulase activities is likely due to an increased concentration of the enzymes, the assays were conducted with equal volumes of the crudes.
TABLE 3 β-xylosidase and endoxylanase activities, and extracellular protein concentrations of the enzymic cocktails from
| Substrate | Strain | β-xylosidase activity (U/L) | Endoxylanase activity (U/mL) | Extracellular protein concentration (mg/L) |
| Xylan t = 4 days | Tam_WT | 998.42 ± 32.78 | 60.77 ± 6.84 | 198.97 ± 5.94 |
| Tam_XlnRV785F | 3292.76 ± 26.38 | 236.41 ± 8.31 | 258.46 ± 1.94 | |
| Glucose t = 4 days | Tam_WT | 44.67 ± 2.66 | 3.69 ± 0.67 | 112.46 ± 4.77 |
| Tam_XlnRV785F | 876.84 ± 20.53 | 126.30 ± 4.8 | 156.68 ± 6.86 | |
| Glycerol t = 6 days | Tam_WT | 68.70 ± 3.33 | 2.47 ± 0.18 | 136.74 ± 1.78 |
| Tam_XlnRV785F | 945.25 ± 6.18 | 119.91 ± 4.99 | 158.62 ± 8.74 |
Saccharifications were performed with 50 g/L xylan, applying a high agitation to ensure the homogeneity of the reactions, and they were monitored by measuring the release of xylose. The results of xylan saccharification with the different crudes of Tam_XlnRV785F and Tam_WT can be found in Figure 6. For all the substrates used to obtain the crudes, Tam_XlnRV785F reaches higher xylose concentrations than Tam_WT, demonstrating greater efficiency in producing hemicellulolytic enzymes, which leads to a more economically viable saccharification of plant biomass residues. This enhancement is even more pronounced for non-inducing substrates, enabling the utilisation of low-cost and renewable materials, such as glycerol, to produce these enzymatic cocktails, an outcome that would not be achievable with the wild-type strain. Specifically, the saccharification yields after 24 h of Tam_XlnRV785F and Tam_WT, respectively, were 51.7% and 40.4% for xylan crudes, 38.5% and 1.2% for glucose supernatants and 36.8% and 1.0% for glycerol cocktails.
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Conclusions
A successful transformation procedure for the genetic manipulation of T. amestolkiae was developed, enabling the directed improvement of its biotechnological potential. Indeed, with this tool we demonstrated that forced expression of a Val785Phe mutant of the transcriptional activator XlnR is a promising strategy to increase hemicellulase production in this fungus, enhancing the efficiency of its hemicellulolytic cocktails for plant biomass valorisation. Furthermore, the Val785Phe substitution of XlnR in T. amestolkiae also leads to a partial deregulation of β-xylosidase and endoxylanase expression under non-inducing conditions. These results show the potential of genetic manipulation tools to design fungal strains that expand the biotechnological applications of T. amestolkiae.
Author Contributions
Ana Pozo-Rodríguez: conceptualization, methodology, writing – original draft, data curation. Miguel Ángel Peñalva: conceptualization, writing – review and editing. Jorge Barriuso: conceptualization, writing – review and editing. Eduardo A. Espeso: conceptualization, writing – review and editing, funding acquisition, supervision. María Jesús Martínez: conceptualization, funding acquisition, writing – review and editing, supervision.
Acknowledgements
The authors thank the Bioinformatics and Biostatistics facility of the Center for Biological Research Margarita Salas (CIB-CSIC) for analysing the Whole Genome Sequencing data of Tam_XlnRV785F transformant. The authors acknowledge the support toward the publication fee by the CSIC Open Access Publication Support Initiative through its Unit of Information Resources for Research (URICI). This work has been funded by H2V2021-05-003, as a part of the CSIC programme for the Spanish Recovery, Transformation and Resilience Plan funded by the Recovery and Resilience Facility of the European Union, established by the Regulation (EU) 2020/2094, as well as the RTI2018-093683-Β-I00 (MICIU/AEI/FEDER) and TED2021-129607B-I00 (MICIU/AEI/NGEU/PRTR) projects. Ana Pozo-Rodríguez thanks MICIU for her FPU19/04192 fellowship.
Conflicts of Interest
The authors declare no conflicts of interest.
Data Availability Statement
Data are provided within the manuscript and S1 files.
Alazi, E., and A. F. J. Ram. 2018. “Modulating Transcriptional Regulation of Plant Biomass Degrading Enzyme Networks for Rational Design of Industrial Fungal Strains.” Frontiers in Bioengineering and Biotechnology 6: 133.
Aleksenko, A., and A. J. Clutterbuck. 1997. “Autonomous Plasmid Replication in Aspergillus nidulans: AMA1 and MATE Elements.” Fungal Genetics and Biology 21: 373–387.
Aleksenko, A., W. Liu, Z. Gojkovic, J. Nielsen, and J. Piskur. 1999. “Structural and Transcriptional Analysis of the pyrABCN, pyrD and pyrF Genes in Aspergillus nidulans and the Evolutionary Origin of Fungal Dihydroorotases.” Molecular Microbiology 33: 599–611.
Arai, T., S. Ichinose, N. Shibata, et al. 2022. “Inducer‐Free Cellulase Production System Based on the Constitutive Expression of Mutated XYR1 and ACE3 in the Industrial Fungus Trichoderma reesei.” Scientific Reports 12: 19445.
Ballmann, P., J. Lightfoot, M. Müller, S. Dröge, and R. Prade. 2019. “Redesigning the Aspergillus nidulans Xylanase Regulatory Pathway to Enhance Cellulase Production With Xylose as the Carbon and Inducer Source.” Microbial Cell Factories 18: 193.
Benocci, T., M. V. Aguilar‐Pontes, M. Zhou, B. Seiboth, and R. P. de Vries. 2017. “Regulators of Plant Biomass Degradation in Ascomycetous Fungi.” Biotechnology for Biofuels 10: 152.
Bertani, G. 1951. “Studies on Lysogenesis I.” Journal of Bacteriology 62: 293–300.
Borneman, A. R., M. J. Hynes, and A. Andrianopoulos. 2001. “STE12 Homolog From the Asexual, Dimorphic Fungus Penicillium marneffei Complements the Defect in Sexual Development of an Aspergillus nidulans steA Mutant.” Genetics 157: 1003–1014.
Bradford, M. M. 1976. “A Rapid and Sensitive Method for the Quantitation of Microgram Quantities of Protein Utilizing the Principle of Protein‐Dye Binding.” Analytical Biochemistry 72: 248–254.
Calero‐Nieto, F., A. Di Pietro, M. I. G. Roncero, and C. Hera. 2007. “Role of the Transcriptional Activator XlnR of Fusarium oxysporum in Regulation of Xylanase Genes and Virulence.” Molecular Plant‐Microbe Interactions 20: 977–985.
Chen, S., Y. Zhou, Y. Chen, and J. Gu. 2018. “Fastp: An Ultra‐Fast All‐in‐One FASTQ Preprocessor.” Bioinformatics 34: i884.
Cove, D. J. 1966. “The Induction and Repression of Nitrate Reductase in the Fungus Aspergillus nidulans.” Biochimica et Biophysica Acta (BBA)—Enzymology and Biological Oxidation 113: 51–56.
Craig, J. P., S. T. Coradetti, T. L. Starr, and N. L. Glass. 2015. “Direct Target Network of the Neurospora Crassa Plant Cell Wall Deconstruction Regulators CLR‐1, CLR‐2, and XLR‐1.” MBio 6: e01452‐15.
Danecek, P., J. K. Bonfield, J. Liddle, et al. 2021. “Twelve Years of SAMtools and BCFtools.” GigaScience 10: giab008.
de Eugenio, L. I., I. de la Torre, F. de Salas, et al. 2024. “Fungal Enzymes for Saccharification of Gamma‐Valerolactone‐Pretreated White Birch Wood: Optimization of the Production of Talaromyces amestolkiae Cellulolytic Cocktail.” Engineering in Life Sciences 24: e202400029.
de Eugenio, L. I., J. A. Méndez‐Líter, M. Nieto‐Domínguez, et al. 2017. “Differential β‐Glucosidase Expression as a Function of Carbon Source Availability in Talaromyces amestolkiae: A Genomic and Proteomic Approach.” Biotechnology for Biofuels 10: 161.
de Oliveira, F., I. L. D. Rocha, D. Cláudia Gouveia Alves Pinto, et al. 2022. “Identification of Azaphilone Derivatives of Monascus Colorants From Talaromyces amestolkiae and Their Halochromic Properties.” Food Chemistry 372: 131214.
Delmas, S., A. Llanos, J.‐L. Parrou, et al. 2014. “Development of an Unmarked Gene Deletion System for the Filamentous Fungi Aspergillus niger and Talaromyces versatilis.” Applied and Environmental Microbiology 80: 3484–3487.
d'Enfert, C. 1996. “Selection of Multiple Disruption Events in Aspergillus fumigatus Using the Orotidine‐5′‐Decarboxylase Gene, pyrG, as a Unique Transformation Marker.” Current Genetics 30: 76–82.
Derntl, C., L. Gudynaite‐Savitch, S. Calixte, T. White, R. L. Mach, and A. R. Mach‐Aigner. 2013. “Mutation of the Xylanase Regulator 1 Causes a Glucose Blind Hydrolase Expressing Phenotype in Industrially Used Trichoderma Strains.” Biotechnology for Biofuels 6: 62.
Derntl, C., R. L. Mach, and A. R. Mach‐Aigner. 2019. “Fusion Transcription Factors for Strong, Constitutive Expression of Cellulases and Xylanases in Trichoderma reesei.” Biotechnology for Biofuels 12: 231.
di Cologna, N. d. M., D. P. Gómez‐Mendoza, F. F. Zanoelo, et al. 2018. “Exploring Trichoderma and Aspergillus Secretomes: Proteomics Approaches for the Identification of Enzymes of Biotechnological Interest.” Enzyme and Microbial Technology 109: 1–10.
Díez, B., E. Alvarez, J. M. Cantoral, J. L. Barredo, and J. F. Martín. 1987. “Selection and Characterization of pyrG Mutants of Penicillium chrysogenum Lacking Orotidine‐5′‐Phosphate Decarboxylase and Complementation by the pyr4 Gene of Neurospora Crassaa.” Current Genetics 12: 277–282.
Ellilä, S., L. Fonseca, C. Uchima, et al. 2017. “Development of a Low‐Cost Cellulase Production Process Using Trichoderma reesei for Brazilian Biorefineries.” Biotechnology for Biofuels 10: 30.
Gao, L., Z. Li, C. Xia, et al. 2017. “Combining Manipulation of Transcription Factors and Overexpression of the Target Genes to Enhance Lignocellulolytic Enzyme Production in Penicillium oxalicum.” Biotechnology for Biofuels 10: 100.
Geng, B., X. Jia, X. Peng, and Y. Han. 2022. “Biosynthesis of Value‐Added Bioproducts From Hemicellulose of Biomass Through Microbial Metabolic Engineering.” Metabolic Engineering Communications 15: e00211.
Gutiérrez, S., A. T. Marcos, J. Casqueiro, et al. 1999. “Transcription of the pcbAB, pcbC and penDE Genes of Penicillium Chrysogenum AS‐P‐78 is Repressed by Glucose and the Repression Is Not Reversed by Alkaline pHs|Microbiology Society.” Microbiology 145: 317–324.
Hasper, A. A., L. M. Trindade, D. van der Veen, A. J. J. van Ooyen, and L. H. de Graaff. 2004. “Functional Analysis of the Transcriptional Activator XlnR From Aspergillus niger.” Microbiology 150: 1367–1375.
Inoue, H., T. Fujii, M. Yoshimi, et al. 2013. “Construction of a Starch‐Inducible Homologous Expression System to Produce Cellulolytic Enzymes From Acremonium cellulolyticus.” Journal of Industrial Microbiology and Biotechnology 40: 823–830.
Jones, P., D. Binns, H.‐Y. Chang, et al. 2014. “InterProScan 5: Genome‐Scale Protein Function Classification.” Bioinformatics 30: 1236–1240.
Jumper, J., R. Evans, A. Pritzel, et al. 2021. “Highly Accurate Protein Structure Prediction With AlphaFold.” Nature 596: 583–589.
Kun, R. S., J. Meng, S. Salazar‐Cerezo, M. R. Mäkelä, R. P. de Vries, and S. Garrigues. 2020. “CRISPR/Cas9 Facilitates Rapid Generation of Constitutive Forms of Transcription Factors in Aspergillus niger Through Specific On‐Site Genomic Mutations Resulting in Increased Saccharification of Plant Biomass.” Enzyme and Microbial Technology 136: 109508.
Li, H., and R. Durbin. 2009. “Fast and Accurate Short Read Alignment With Burrows–Wheeler Transform.” Bioinformatics 25: 1754–1760.
Li, K.‐Y., Q.‐F. Zhu, J.‐L. Ao, et al. 2022. “New Meroterpenoid and Isocoumarins From the Fungus Talaromyces Amestolkiae MST1‐15 Collected From Coal Area.” Molecules 27: 8223.
Li, Z., G. Yao, R. Wu, et al. 2015. “Synergistic and Dose‐Controlled Regulation of Cellulase Gene Expression in Penicillium oxalicum.” PLoS Genetics 11: e1005509.
Llanos, A., S. Déjean, V. Neugnot‐Roux, J. M. François, and J.‐L. Parrou. 2019. “Carbon Sources and XlnR‐Dependent Transcriptional Landscape of CAZymes in the Industrial Fungus Talaromyces versatilis: When Exception Seems to Be the Rule.” Microbial Cell Factories 18: 14.
Lv, D., W. Zhang, X. Meng, and W. Liu. 2023. “Single Mutation in Transcriptional Activator Xyr1 Enhances Cellulase and Xylanase Production in Trichoderma reesei on Glucose.” Journal of Agricultural and Food Chemistry 71: 11993–12003.
Mach‐Aigner, A. R., M. E. Pucher, M. G. Steiger, G. E. Bauer, S. J. Preis, and R. L. Mach. 2008. “Transcriptional Regulation of xyr1, Encoding the Main Regulator of the Xylanolytic and Cellulolytic Enzyme System in Hypocrea jecorina.” Applied and Environmental Microbiology 74: 6554–6652.
Madeira, F., N. Madhusoodanan, J. Lee, et al. 2024. “The EMBL‐EBI Job Dispatcher Sequence Analysis Tools Framework in 2024.” Nucleic Acids Research 52: W521–W525.
Martín, J. F. 2015. “Fungal Transformation: From Protoplasts to Targeted Recombination Systems.” In Genetic Transformation Systems in Fungi, edited by M. A. van den Berg and K. Maruthachalam, vol. 1, 3–18. Springer International Publishing.
Marui, J., N. Kitamoto, M. Kato, T. Kobayashi, and N. Tsukagoshi. 2002. “Transcriptional Activator, AoXlnR, Mediates Cellulose‐Inductive Expression of the Xylanolytic and Cellulolytic Genes in Aspergillus oryzae.” FEBS Letters 528: 279–282.
Mattam, A. J., Y. B. Chaudhari, and H. R. Velankar. 2022. “Factors Regulating Cellulolytic Gene Expression in Filamentous Fungi: An Overview.” Microbial Cell Factories 21: 44.
Mei, Y.‐Z., Y.‐L. Zhu, P.‐W. Huang, Q. Yang, and C.‐C. Dai. 2019. “Strategies for Gene Disruption and Expression in Filamentous Fungi.” Applied Microbiology and Biotechnology 103: 6041–6059.
Méndez‐Líter, J. A., L. I. de Eugenio, N. L. S. Hakalin, A. Prieto, and M. J. Martínez. 2021a. “Production of a β‐Glucosidase‐Rich Cocktail From Talaromyces amestolkiae Using Raw Glycerol: Its Role for Lignocellulose Waste Valorization.” Journal of Fungi 7: 363.
Méndez‐Líter, J. A., L. I. de Eugenio, M. Nieto‐Domínguez, A. Prieto, and M. J. Martínez. 2021b. “Hemicellulases From Penicillium and Talaromyces for Lignocellulosic Biomass Valorization: A Review.” Bioresource Technology 324: 124623.
Meng, X., and A. J. Ragauskas. 2014. “Recent Advances in Understanding the Role of Cellulose Accessibility in Enzymatic Hydrolysis of Lignocellulosic Substrates.” Current Opinion in Biotechnology 27: 150–158.
Monteiro, M. R., C. L. Kugelmeier, R. S. Pinheiro, M. O. Batalha, and A. da Silva César. 2018. “Glycerol From Biodiesel Production: Technological Paths for Sustainability.” Renewable and Sustainable Energy Reviews 88: 109–122.
Ng, H.‐E., S. S. A. Raj, S. H. Wong, D. Tey, and H.‐M. Tan. 2008. “Estimation of Fungal Growth Using the Ergosterol Assay: A Rapid Tool in Assessing the Microbiological Status of Grains and Feeds.” Letters in Applied Microbiology 46: 113–118.
Nieto‐Domínguez, M., L. I. de Eugenio, J. Barriuso, et al. 2015. “Novel pH‐Stable Glycoside Hydrolase Family 3 β‐Xylosidase From Talaromyces Amestolkiae: An Enzyme Displaying Regioselective Transxylosylation.” Applied and Environmental Microbiology 81: 6380–6392.
Noguchi, Y., M. Sano, K. Kanamaru, et al. 2009. “Genes Regulated by AoXlnR, the Xylanolytic and Cellulolytic Transcriptional Regulator, in Aspergillus Oryzae.” Applied Microbiology and Biotechnology 85: 141–154.
Oakley, B. R., J. E. Rinehart, B. L. Mitchell, et al. 1987. “Cloning, Mapping and Molecular Analysis of the pyrG (Orotidine‐5′‐Phosphate Decarboxylase) Gene of Aspergillus nidulans.” Gene 61: 385–399.
Osherov, N., J. Mathew, and G. S. May. 2000. “Polarity‐Defective Mutants of Aspergillus nidulans.” Fungal Genetics and Biology 31: 181–188.
Ostadjoo, S., F. Hammerer, K. Dietrich, M.‐J. Dumont, T. Friscic, and K. Auclair. 2019. “Efficient Enzymatic Hydrolysis of Biomass Hemicellulose in the Absence of Bulk Water.” Molecules 24: 4206.
Pettersen, E. F., T. D. Goddard, C. C. Huang, et al. 2004. “UCSF Chimera—A Visualization System for Exploratory Research and Analysis.” Journal of Computational Chemistry 25: 1605–1612.
Prieto, A., L. de Eugenio, J. A. Méndez‐Líter, et al. 2021. “Fungal Glycosyl Hydrolases for Sustainable Plant Biomass Valorization: Talaromyces amestolkiae as a Model Fungus.” International Microbiology 24: 545–558.
Serebryanyi, V. A., O. A. Sinitsyna, E. A. Fedorova, et al. 2006. “Synthesis of Xylanolytic Enzymes and Other Carbohydrases by the Fungus Penicillium canescens: Transformants With an Altered Copy Number of the Gene xlnR and an Encoding Transcriptional Activator.” Applied Biochemistry and Microbiology 42: 584–591.
Somogyi, M. 1945. “A New Reagent for the Determination of Sugars.” Journal of Biological Chemistry 160: 61–68.
Tamayo‐Ramos, J. A., and M. Orejas. 2014. “Enhanced Glycosyl Hydrolase Production in Aspergillus nidulans Using Transcription Factor Engineering Approaches.” Biotechnology for Biofuels 7: 103.
Tani, S., T. Kawaguchi, and T. Kobayashi. 2014. “Complex Regulation of Hydrolytic Enzyme Genes for Cellulosic Biomass Degradation in Filamentous Fungi.” Applied Microbiology and Biotechnology 98: 4829–4837.
Tilburn, J., C. Scazzocchio, G. G. Taylor, J. H. Zabicky‐Zissman, R. A. Lockington, and R. W. Davies. 1983. “Transformation by Integration in Aspergillus nidulans.” Gene 26: 205–221.
Traut, T. W., and B. R. S. Temple. 2000. “The Chemistry of the Reaction Determines the Invariant Amino Acids During the Evolution and Divergence of Orotidine 5′‐Monophosphate Decarboxylase.” Journal of Biological Chemistry 275: 28675–28681.
van Peij, N. N., M. M. Gielkens, R. P. de Vries, J. Visser, and L. H. de Graaff. 1998a. “The Transcriptional Activator XlnR Regulates Both Xylanolytic and Endoglucanase Gene Expression in Aspergillus niger.” Applied and Environmental Microbiology 64: 3615–3619.
Van Peij, N. N. M. E., J. Visser, and L. H. De Graaff. 1998b. “Isolation and Analysis of xlnR, Encoding a Transcriptional Activator Co‐Ordinating Xylanolytic Expression in Aspergillus Niger.” Molecular Microbiology 27: 131–142.
Veiter, L., V. Rajamanickam, and C. Herwig. 2018. “The Filamentous Fungal Pellet—Relationship Between Morphology and Productivity.” Applied Microbiology and Biotechnology 102: 2997–3006.
Villarino, M., E. A. Espeso, P. Melgarejo, and I. Larena. 2018. “Transformation of Penicillium Rubens 212 and Expression of GFP and DsRED Coding Genes for Visualization of Plant‐Biocontrol Agent Interaction.” Frontiers in Microbiology 9: 1653.
Wang, J., Y. Wu, Y. Gong, S. Yu, and G. Liu. 2015. “Enhancing Xylanase Production in the Thermophilic Fungus Myceliophthora Thermophila by Homologous Overexpression of Mtxyr1.” Journal of Industrial Microbiology and Biotechnology 42: 1233–1241.
Wang, S., H. Chen, X. Tang, H. Zhang, W. Chen, and Y. Q. Chen. 2017. “Molecular Tools for Gene Manipulation in Filamentous Fungi.” Applied Microbiology and Biotechnology 101: 8063–8075.
Wang, S., G. Liu, J. Wang, J. Yu, B. Huang, and M. Xing. 2013. “Enhancing Cellulase Production in Trichoderma reesei RUT C30 Through Combined Manipulation of Activating and Repressing Genes.” Journal of Industrial Microbiology and Biotechnology 40: 633–641.
Xia, C., L. Gao, Z. Li, G. Liu, and X. Song. 2022. “Functional Analysis of the Transcriptional Activator XlnR of Penicillium oxalicum.” Journal of Applied Microbiology 132: 1112–1120.
Yang, L., L. Ukil, A. Osmani, et al. 2004. “Rapid Production of Gene Replacement Constructs and Generation of a Green Fluorescent Protein‐Tagged Centromeric Marker in Aspergillus nidulans.” Eukaryotic Cell 3: 1359–1362.
Zhang, T., Y. Sun, Z. Ma, J. Zhang, B. Lv, and C. Li. 2022. “Developing Iterative and Quantified Transgenic Manipulations of Non‐Conventional Filamentous Fungus Talaromyces pinophilus Li‐93.” Biochemical Engineering Journal 179: 108317.
Zhao, Q., Z. Yang, Z. Xiao, et al. 2023. “Structure‐Guided Engineering of Transcriptional Activator XYR1 for Inducer‐Free Production of Lignocellulolytic Enzymes in Trichoderma reesei.” Synthetic and Systems Biotechnology 8: 732–740.
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Abstract
ABSTRACT
The ascomycete
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; Peñalva, Miguel Ángel 2 ; Barriuso, Jorge 1
; Espeso, Eduardo A. 2 ; Martínez, María Jesús 1
1 Microbial Systems and Protein Engineering Group, Department of Biotechnology, Center for Biological Research Margarita Salas, Spanish National Research Council (CIB‐CSIC), Madrid, Spain
2 Aspergillus Cell Biology Group, Department of Cellular and Molecular Biosciences, Center for Biological Research Margarita Salas, Spanish National Research Council (CIB‐CSIC), Madrid, Spain