Development of an efficient heterologous protein expression platform in Aspergillus niger through genetic modification of a glucoamylase hyperproducing industrial strain

Abstract

Background

Aspergillus niger is widely used in industrial enzyme production due to its strong secretion capacity and the status of generally recognized as safe (GRAS). However, heterologous protein expression in A. niger is frequently constrained by high levels of background endogenous protein secretion, limited access to native high transcription loci, and limitations in the efficiency of the secretory machinery. To address these limitations, this study genetically engineered a chassis strain based on an industrial glucoamylase-producing A. niger strain AnN1 for constructing the improved heterologous protein expression.

Results

In this study, by using CRISPR/Cas9-assisted marker recycling, we deleted 13 of the 20 copies of the heterologous glucoamylase TeGlaA gene and disrupted the major extracellular protease gene PepA, resulting in the low-background strain AnN2. Compared to the parental strain AnN1, AnN2 exhibited 61% less extracellular protein and significantly reduced glucoamylase activity, while retaining multiple transcriptionally active integration loci. Four diverse proteins were integrated into the high-expression loci originally occupied by the TeGlaA gene in the chassis AnN2. These recombinant protein included a homologous glucose oxidase (AnGoxM), a thermostable pectate lyase A (MtPlyA), a bacterial triose phosphate isomerase (TPI), and a medical protein Lingzhi-8 (LZ8). All target proteins were successfully expressed and secreted within 48–72 h, with yields ranging from 110.8 to 416.8 mg/L in 50 mL shake-flasks cultivation. The enzyme activities of AnGoxM, MtPlyA and TPI reached ~ 1276 − 1328 U/mL, ~ 1627. 43 − 2105.69 U/mL, and ~ 1751.02 to 1906.81 U/mg after 48 h, respectively. Additionally, Overexpression of Cvc2, a COPI vesicle trafficking component, further enhanced MtPlyA production by 18%, highlighting the benefit of combining transcriptional and secretory pathway engineering.

Conclusions

Our results demonstrated that the chassis AnN2 served as a robust, modular, and time-efficient platform for heterologous protein expression in A. niger. Through site-specific integration of target genes into native high-expression loci and strategic modulation of the secretory pathway, we successfully enabled the rapid production of functional enzymes and bioactive proteins from diverse origins. This dual-level optimization strategy, which integrates rational genomic engineering with targeted enhancement of the secretory pathway, enabled high-yield expression while minimizing background interference. Together, these findings offer a practical framework for constructing versatile fungal expression systems and highlight the potential of combining genetic and cellular engineering to improve recombinant protein production in filamentous fungi.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12934-025-02786-x.

Background

The use of cell factories for recombinant protein production has revolutionized modern biotechnology, enabling the large-scale synthesis of bioactive proteins for applications in biopharmaceuticals, industrial enzyme production, agriculture, and food additives [1–3]. In the biopharmaceutical industry, recombinant proteins such as insulin, monoclonal antibodies, and vaccines have revolutionized the treatment of diseases like diabetes, cancer, and infectious diseases [4]. For instance, recombinant epidermal growth factor (rEGF) has been successfully produced in mammalian cell, Escherichia coli, Saccharomyces cerevisiae, and Pichia pastoris for clinical and cosmetic applications [5, 6]. In biotechnology and bioenergy sectors, industrial enzymes including glucoamylases, amylases, lipases, and cellulases, are extensively used in biofuel production, detergent formulation, textile processing, and biocatalysts [7–10]. Currently, the global market for biopharmaceutical proteins is approaching $400 billion annually, while the industrial enzyme sector was valued at approximately $7.1 billion in 2023 and is projected to surpass $11 billion by 2028, driven by increasing demand in food processing, biofuels, and pharmaceutical manufacturing [11]. To address the increasing demand for high-yield, cost-effective, and scalable recombinant protein production, microbial expression systems have been extensively developed and optimized by genetic engineering techniques [12, 13]. Among these, prokaryotic hosts such as E. coli and Bacillus species are widely utilized due to their rapid growth, ease of genetic manipulation, and scalability [14, 15]. The eukaryotic expression platforms, including yeast, filamentous fungi, insect, and mammalian cells, are preferred for producing complex proteins that require correct disulfide bond formation for proper folding, N- or O-linked glycosylation to ensure structural stability or bioactivity, and additional post-translational modifications such as phosphorylation or proteolytic processing for full functionality [16–18].

The filamentous fungus Aspergillus niger is widely recognized as a robust industrial host for enzyme production due to its exceptional protein secretion capacity and generally recognized as safe (GRAS) characteristics status [19–23]. Among the commercial enzymes, glucoamylase (GlaA) is one of the most abundantly produced [24], with industrial strains achieving titers reaching 30 g/L [25]. The global glucoamylase market was valued at approximately USD 1.35 billion in 2022 and is projected to reach USD 2.5 billion by 2032, driven by its widespread applications in food processing, biofuel production, and pharmaceuticals. This high productivity is attributed to strong promoters, efficient secretion pathways improvement, morphological engineering, and optimized metabolic networks [26–31]. For example, the introduction of multi-copies of the native or heterologous GlaA gene is useful strategy for generating GlaA overproducer [26, 27]. It is widely accepted that, in filamentous fungi, protein secretion predominantly occurs at the growing hyphal tips. Significant enhancement of glucoamylase secretion was observed in A. niger mutants exhibiting either a hyperbranching phenotype due to the deletion of the Rho-GTPase racA [28] or a short rod-shaped hyphal morphology resulting from the knockout of the tertiary kinase system (bck1-mkkA-mpkA) [29]. These findings suggest that increased accumulation of post-Golgi secretory vesicles at hyphal tips plays a key role in secretion efficiency and that morphological regulation could be leveraged to optimize A. niger for industrial enzyme production. Furthermore, metabolic engineering of the NADPH-regenerating genes gndA and maeA [30], as well as three NADH kinases [31], enhanced the intracellular NADPH pool and significantly improved GlaA production. These findings suggest that optimizing NADPH regeneration represents a promising strategy for enhancing enzyme production in filamentous fungi. Recent advancements in genetic engineering tools and synthetic biology approaches, such as CRISPR-Cas9/Cas12a system and CRISPR-Cas9-mediated multi-copy gene expression, have enabled precise modifications of fungal strains to improve their capacity for producing and secreting both native and heterologous proteins, including the sweet protein monellin (0.284 mg/L), human erythropoietin (73.9 mg/L), alkaline serine protease (10.8 mg/mL), and trehalase (4268.29U/mL) [32–36].

The secretory pathway in A. niger involves coordinated vesicle-mediated transport between the ER and Golgi apparatus [28]. COPII-coated vesicles mediate anterograde transport, directing newly synthesized proteins from the ER to the Golgi, whereas COPI-coated vesicles facilitate retrograde trafficking by recycling ER-resident chaperones and maintaining ER-Golgi homeostasis [37]. These tightly regulated processes are critical for sustaining high-level protein secretion, and disruptions—particularly under strong expression conditions—can compromise secretion efficiency. Despite significant advances in optimizing A. niger for secretory protein production, heterologous protein expression remains a persistent bottleneck, typically yielding titers substantially lower than those of native counterparts such as glucoamylase (GlaA). This limitation stems from multifactorial constraints spanning transcriptional inefficiencies, codon usage bias, misfolding of heterologous polypeptides, and incomplete post-translational modifications [38]. ER stress triggered by the accumulation of misfolded polypeptides can activate the unfolded protein response (UPR) and lead to protein degradation via the ER-associated degradation (ERAD) pathway [39]. Moreover, inefficient vesicular transport and suboptimal cargo sorting further restrict secretion capacity. Extracellular proteolytic degradation and incompatible signal peptides can additionally exacerbate protein loss, together imposing significant barriers to the accumulation of heterologous products [25, 37]. Systematic dissection and resolution of these interconnected biological barriers are imperative to advance A. niger as a robust platform for recombinant protein biosynthesis.

Recently, we developed efficient genetic manipulation tools for two industrial A. niger strains, N1 and O1, including transformation protocols, a marker-free CRISPR/Cas9 technique [40], and a flow cytometry-based plating-free technology [41]. Notably, the A. niger N1 (AnN1), carrying 20 copies of the Talaromyces emersonii GlaA (TeGlaA) genes, is an industrial glucoamylase-producing strain, demonstrating robust transcriptional and secretion machinery and highlighting its potential as a promising host for recombinant protein expression. In this study, we employed the industrial strain AnN1 as a host to establish a heterologous protein expression platform. This was accomplished by deleting13 out of the 20 tandemly integrated TeGlaA gene copies by using the CRISPR/Cas9 System, while subsequently integrating target genes into the corresponding high-transcription loci formerly occupied by TeGlaA. The resulting chassis strain, AnN2, was generated using a CRISPR/Cas9-mediated marker recycling technique, disrupting the high-copy TeGlaA genes and the PepA gene.

To facilitate efficient recombinant protein expression, we developed a modular donor DNA plasmid system incorporating the native AAmy promoter and the AnGlaA terminator as homologous arms for CRISPR/Cas9-mediated integration. The platform was validated using a panel of homologous and heterologous proteins, including A. niger glucose oxidase (AnGoxM), a thermostable pectate lyase from Myceliophthora thermophila (MtPlyA), a bacterial triosephosphate isomerase (TPI), and the immunomodulatory protein LZ-8 from Ganoderma lucidum. These proteins were selected to represent diverse functional classes and phylogenetic origins, covering industrial enzymes of fungal and bacterial origin, and a bioactive pharmaceutical protein, highlighting the versatility and broad applicability of the expression platform. After incubation for 48 h, all target proteins were successfully secreted into the culture supernatant, indicating this platform’s high efficiency and versatility. Furthermore, MtPlyA production was significantly enhanced by overexpressing the COPI component Cvc2, optimizing vesicular trafficking along the secretory pathway. By enabling CRISPR/Cas9-mediated multi-copy integration at endogenous high-expression loci using native promoters, this platform provides a robust and modular approach for high-yield recombinant protein production, with promising applications in industrial enzyme manufacturing and biopharmaceutical development.

Results and discussion

Construction and evaluation of an engineered A. niger host strain AnN2 for heterologous protein expression

In this study, we constructed and validated a robust chassis strain of A. niger, derived from the industrial strain AnN1, to support high-efficiency heterologous protein expression. By targeting and re-utilizing the native high-expression loci originally occupied by the multi-copy industrial glucoamylase gene TeGlaA, and employing the CRISPR/Cas9 marker-recycling multiplex genomic editing developed in our previous reports [39, 41], we established a versatile and low-background expression platform with modular genome editing capacity (Fig. 1). The AnN1 strain harbors 20 copies of the heterologous glucoamylase gene TeGlaA (UniProt: Q9C1V4), one copy of the native glucoamylase gene AnGlaA (UniProt: A2QHE1), and one copy of an acid α-amylase gene (AAmy, UniProt: A2QW02). Both TeGlaA and AnGlaA are expressed under the control of strong native promoters (PAAmy and PGlaA, respectively) and terminated by TGlaA. To characterize the secretome of the parental strain, we conducted SDS-PAGE analysis after 4 and 6 days of shake-flask cultivation (Fig. 1A). The major protein bands were excised and subjected to LC-MS/MS analysis, which confirmed that the most abundant secreted proteins were TeGlaA, AnGlaA, and AAmy. These top LC-MS/MS hits have now been annotated in Fig. 1A for clarity and direct correspondence between electrophoretic bands and protein identities (Additional file 2: Table S2). These features contribute to strong protein secretion, while maintaining a relatively simplified secretome, making AnN1 a suitable chassis candidate. To reduce the background secretion and generate transcriptionally active integration sites, we applied a CRISPR/Cas9-assisted marker recycling system [42] to selectively delete multiple TeGlaA copies (Fig. 1B). To maintain the high secretory capacity of the host strain, we retained the native AnGlaA and AAmy loci in AnN2, which not only contribute to strong endogenous protein secretion but also serve as transcriptionally active sites suitable for future targeted integration of heterologous genes. Two sgRNAs were designed targeting the 5’ and 3’ ends of the TeGlaA ORF, and donor DNA (donor-TeGlaA-TAA) was constructed using the PAAmy promoter and TGlaA terminator as homologous arms. Additional engineered sgRNA recognition sites were introduced to support efficient downstream gene integration at the deleted loci. Simultaneously, the major extracellular protease gene PepA [42] was disrupted to minimize proteolytic degradation of heterologous proteins.

Fig. 1.

Construction of an engineered chassis strain AnN2 by CRISPR/Cas9-mediated deletion of glucoamylase gene loci and generation of transcriptionally active knock-in sites. (A) SDS-PAGE analysis of secreted protein of the AnN1 strains after 4- and 6-days of incubation. The major secreted proteins were identified by LC-MS/MS as TeGlaA, AnGlaA, and AAmy (UniProt: Q9C1V4, A2QHE1, and A2QW02, respectively), and are annotated on the gel accordingly. Each lane was loaded with 10 µL of extracellular protein. (B) Schematic representation of the genome editing strategy used to convert the high-glucoamylase-producing strain AnN1 into the chassis host strain AnN2. A CRISPR/Cas9-assisted system was employed to delete multiple copies of TeGlaA, while retaining its native promoter (PAAmy) and terminator (TGlaA) sequences for future recombinant gene integration. The resulting genome harbors multiple high-transcription loci available as knock-in sites for gene expression

After transformation, ten positive transformants were isolated and verified by diagnostic PCR (Additional file 3: Fig. S1B). To assess the effect of TeGlaA deletion on protein secretion, we performed SDS-PAGE analysis after 6 days of shake-flask cultivation. RT-qPCR analysis was performed to quantify the residual TeGlaA copy numbers in the engineered strains. This revealed a clear correlation between reduced TeGlaA copy number and decreased extracellular protein secretion and enzymatic activity (Fig. 2D). Specifically, strain M3 retained only seven TeGlaA copies—13 fewer than the parental AnN1 strain—indicating successful partial deletion and functional impact. As shown in Fig. 2A, SDS-PAGE analysis clearly illustrated the loss of the hyper-production phenotype in the culture supernatants of the knockout strains following 6 days of shake-flask cultivation. Quantitative analysis of extracellular proteins revealed that the engineered strains secreted approximately 15–61% fewer proteins compared to the parental strain AnN1 (Fig. 2B). Similarly, glucoamylase activity assays showed a significant reduction of approximately 20–59% in the mutant strains relative to AnN1 (Fig. 2C). Among these mutants, strain M3 displayed notably reduced extracellular protein secretion, retaining only about 39% of total protein levels and approximately 41% of glucoamylase activity compared to AnN1. These results were consistent with the markedly reduced levels of TeGlaA-derived peptides observed in the secretome analysis (Additional file 2: Table S2).

Fig. 2.

Reduction of glucoamylase expression and copy number in the engineered chassis mutants. (A) SDS-PAGE analysis of total secreted proteins from the parental strain AnN1 and mutant strains (M1–M10) after 6 days of shake-flask cultivation. Each lane was loaded with 10 µL of extracellular protein. (B) Assays for total extracellular protein concentration in the A. niger strains. (C) Glucoamylase activity in the culture supernatants of the strains, expressed as a percentage of AnN1 activity. (D) Assay of the copy number of the TeGlaA gene in the mutant strains by RT-qPCR analysis. Error bars indicate the SD from three replicates. Different lowercase letters (Fig. 2B and D) denote significant differences as determined by Tukey’s HSD test (P < 0.05)

Further LC-MS/MS analysis of the secretome revealed that the prominent 135–180 kDa band in the mutants was primarily composed of native glucoamylase (AnGlaA), along with a smaller proportion of residual heterologous TeGlaA and low levels of AAmy peptides (Additional file 2: Table S2). The emergence of this band was particularly evident in mutants with reduced TeGlaA burden, suggesting that secretion capacity may have been redistributed toward other endogenous proteins upon partial relief of the secretory load. Interestingly, despite similar TeGlaA copy numbers in M3 and M4, their secreted protein profiles differed, implying that post-editing regulatory variation or chromatin remodeling may influence final secretion outputs. We conducted three independent rounds of CRISPR-mediated transformation. Complete removal of all 20 TeGlaA copies proved challenging, as further deletion consistently led to impaired colony growth or genomic instability, possibly due to extensive genome rearrangement or metabolic stress. The M3 mutant, which retained seven residual copies of TeGlaA, exhibited the most favorable balance between reduced background secretion, robust growth, and genetic stability. Therefore, M3 was selected as the optimized chassis strain and designated as AnN2 for subsequent studies. This moderate reduction preserved host viability and protein secretion machinery, while minimizing interference from endogenous proteins. This delicate balance is crucial in filamentous fungi, where global suppression of secretion pathways often leads to pleiotropic effects such as impaired growth, altered hyphal morphology, and metabolic imbalance [43–45]. A schematic overview of the expression platform and the diverse set of recombinant proteins including AnGoxM, MtPlyA, TPI, and LZ8 expressed in AnN2 is shown in Fig. 3.

Fig. 3.

Schematic overview of the engineered expression platform enabling multi-copy integration at high-expression loci in the engineered chassis strain AnN2. Two expression modules were constructed using native high-expression loci: PAAmy–TGlaA and PGlaA–TtrpC. Target genes were integrated into these loci via CRISPR/Cas9-mediated homologous recombination. Four functionally and taxonomically diverse proteins were selected to demonstrate platform versatility: a thermostable bacterial pectate lyase (MtPlyA), a fungal-origin glucose oxidase (AnGoxM), a bacterial triosephosphate isomerase (TPI), and a pharmaceutical fungal protein (LZ8). This strategy enables robust and tunable heterologous protein expression through modular, multi-copy genome integration at high-expression sites

Expression of a homologous glucose oxidase in the engineered A. niger host strain AnN2

The first recombinant protein tested in the AnN2 system was a thermostable variant of glucose oxidase (AnGoxM). Glucose oxidase (GOX) is a commercially significant enzyme that catalyzes the oxidation of β-D-glucose to gluconic acid and hydrogen peroxide [24, 46–49]. GOX enzymes have extensive applications in various industries, including food processing, clinical diagnostics, pharmaceuticals, and biotechnology [24]. These enzymes are predominantly produced by fungal genera such as Aspergillus and Penicillium [24, 46, 47]. To evaluate the potential of the AnN2 strain as a host for co-expression of recombinant proteins in a reduced-amylolytic background, we introduced the engineered glucose oxidase gene (AnGoxM, previously characterized from A. niger by Jiang et al., 2021 [47]) into the AnN2 strain. The AnGoxM gene was integrated into the transcriptionally active genomic loci originally occupied by the multi-copy TeGlaA and single-copy AnGlaA genes (Figs. 3 and 4A).

Fig. 4.

Multi-locus expression of the glucose oxidase (AnGoxM) in the A. niger chassis strain AnN2. (A) Schematic representation of the CRISPR/Cas9-mediated multi-copy integration strategy for AnGoxM. The gene was inserted into transcriptionally active loci originally occupied by TeGlaA (on chromosomes I, II, and VII) and the single-copy AnGlaA locus (on chromosome VI). (B) SDS-PAGE analysis of the total secreted proteins in the supernatants of RS-AnGoxM after 48 h of shake-flask culture. Each lane was loaded with 10 µL of extracellular protein. AnGoxM expression was confirmed by the appearance of a specific band (~ 75 kDa, blue arrow) in RS-AnGoxM but not in the parental strain AnN2. (C) Western blot analysis of the culture supernatants from the same samples using anti-His antibody confirmed successful secretion of AnGoxM. Each lane was loaded with 5 µL of extracellular protein. (D) Glucose oxidase activity of three RS-AnGoxM strains measured after 48 and 72 h of cultivation. Error bars indicate the SD from three replicates

The donor DNA construct for protein expression was designed by cloning the target gene fused to a 6×His-tag into the donor-TeGlaA-TAA vector, placing it under the control of the strong promoter PAAmy, the AAmy signal peptide, and the TGlaA terminator to facilitate transcription and secretion. For integration at the AnGlaA locus, the donor construct included the AnGoxM gene fused with the AAmy signal peptide at the N-terminus and a 6×His-tag at the C-terminus, controlled by the GlaA promoter and the TrpC terminator (Fig. 4A). Homologous recombination was guided by sequences from the GlaA promoter and the GlaA 3’ flanking region. Following targeted double-strand breaks (DSBs) generated by Cas9 in the presence of locus-specific sgRNAs, repair occurred via homologous recombination, enabling precise integration of the donor DNA constructs into the genome. Diagnostic PCR and qPCR analyses were subsequently performed, leading to the selection of three transformants that each contained 12–13 integrated copies of the AnGoxM gene (Additional file 4: Fig. S2).

The SDS-PAGE and western blotting analyses clearly demonstrated the successful secretion of AnGoxM into the culture supernatants of three RS-AnGoxM strains after 48 h of shake-flask culture (Fig. 4B and C). This phenomenon was consistent with previous observations reported in other expression hosts, such as Yarrowia lipolytica [46], Pichia pastoris [47], and T. reesei [48]. The theoretical molecular weight of AnGoxM is approximately 65 kDa, whereas SDS-PAGE analysis revealed an apparent size of ~ 75 kDa, indicating the presence of post-translational modifications. To explore this possibility, we analyzed the amino acid sequence of AnGoxM using the NetNGlyc 1.0 server, which predicted eight potential N-glycosylation sites matching the consensus Asn-X-Ser/Thr motif. This is consistent with previous studies showing that AnGox is a glycoprotein with 10–16% of its molecular mass attributed to carbohydrate content [49]. Furthermore, Western blot analysis showed an even higher apparent molecular weight (~ 100 kDa), which may reflect enhanced sensitivity of immunodetection to glycosylated forms and reduced electrophoretic mobility. Together, these observations support the hypothesis that the observed size shifts are primarily due to extensive N-linked glycosylation. Notably, no corresponding bands or glucose oxidase activity were detected in the parental host strain AnN2 under the culture conditions, thus the measured activity reflected the production level of AnGoxM in the chassis cells. To assess the genetic stability of the integrated expression cassettes, representative recombinant strains expressing AnGoxM were subjected to five rounds of serial sub-cultivation under selective conditions (G418-containing medium). Enzyme activity was measured at both the second and fifth passages to evaluate whether gene expression remained stable across generations. As shown in Fig S2C, glucose oxidase activity in RS-AnGoxM strains remained consistent between the second and fifth passages, with no significant difference observed, indicating stable expression of AnGoxM. Subsequently, recombinant AnGoxM was purified from culture supernatants using Ni²⁺-NTA affinity chromatography. SDS-PAGE analysis of the purified enzymes revealed a single protein band consistent with the molecular weight observed by western blotting (Fig. 4B and C). The glucose oxidase activities of the RS-AnGoxM strains were measured at ~ 1276 − 1328 U/mL after 48 h and ~ 1304 − 1458 U/mL after 72 h of cultivation (Fig. 4D). Compared to previous reports where A. niger GOX was overexpressed in yeasts P. pastoris and Y. lipolytica with activity levels around 400–600 U/mL under shake culturing [24, 47, 48], the AnN2-based platform exhibited 2–3-fold higher production, indicating the strong secretion capacity of the engineered fungal chassis. The production level of rAnGoxM in recombinant strains reached ~ 364 mg/L after 72 h culture time in the shake flask. The presence of a single, clear band on the SDS-PAGE gels demonstrated that a single-step purification method effectively yielded highly pure recombinant enzymes suitable for further biochemical analyses.

Heterologous expression of bacterial TPI enzyme in the engineered chassis strain AnN2

The applicability of the AnN2 chassis was further explored by expressing heterologous proteins with diverse structural and phylogenetic characteristics. Triose phosphate isomerase (TPI) is an essential enzyme involved in the glycolytic pathway, catalyzing the reversible inter conversion of dihydroxyacetone phosphate and glyceraldehyde-3-phosphate [50, 51]. To evaluate the capability of the engineered strain AnN2 to produce bacterial enzymes, we selected the E. coli-derived TPI as the target protein. Using the CRISPR/Cas9-mediated integration approach described in this work, multiple copies of the optimized tpi gene were successfully integrated into highly active transcriptional loci originally occupied by the TeGlaA and AnGlaA genes in the AnN2 genome (Fig. 5A). Positive transformants (RS-TPI) were were initially verified by colony PCR (Additional file 5: Fig. S3). The TPI gene copy numbers were subsequently quantified by qPCR, revealing successful integration of the 11–12 TPI gene copies into these loci (Fig. 5B).

Fig. 5.

Multi-copy expression and activity analysis of bacterial triose phosphate isomerase (TPI) in the chassis strain AnN2. (A) Schematic diagram showing the CRISPR/Cas9-mediated integration of the TPI gene into multiple native expression loci in the AnN2 genome, including transcriptionally active sites originally occupied by TeGlaA and the AnGlaA locus. (B) Assay of the tpi copy number in the recombinant strains by RT-qPCR. (C) SDS-PAGE analysis of culture supernatants from RS-TPI transformants after 48 h of culture. Each lane was loaded with 10 µL of extracellular protein. The protein band corresponding to TPI (~ 28 kDa, blue arrow) was observed in RS-TPI strains but absent in the control strain AnN2. (D) Western blotting using anti-His antibody confirmed the presence of secreted TPI protein in RS-TPI strains. Each lane was loaded with 5 µL of extracellular rTPI protein. (E) Enzymatic activity of recombinant TPI in three RS-TPI strains at 48 h and 72 h post-inoculation. Error bars represent the standard deviation from three biological replicates

Three verified RS-TPI strains were selected for further expression assessment via shake flask culture. After incubation for 48 h, secretion of TPI was analyzed by SDS-PAGE and western blotting assays. A prominent protein band corresponding to approximately 28 kDa was observed in the culture broth of all three RS-TPI strains but was absent in the parent strain AnN2 (Fig. 5C and D). Additionally, the enzyme activity remained steady across the two passages TPI-expressing strains (Fig. S3B), further demonstrating the robustness of the integration strategy in maintaining expression without activity loss during sub-culturing on selective conditions. The purified recombinant TPI protein displayed a clear single band at approximately 28 kDa on SDS-PAGE gels, consistent with the calculated monomeric molecular weight.

Enzyme activity assays showed that TPI reached peak activities ranging from ~ 1751.02 to 1906.81 U/mg at 48 h. However, enzyme activity significantly decreased by approximately 30% at 72 h post-inoculation (Fig. 5E). This reduction in enzyme activity corresponded to the decreased intensity of the TPI protein band observed in SDS-PAGE and western blot analyses at the later time point, suggesting that the recombinant TPI was susceptible to degradation or instability during extended cultivation. Given that A. niger secretes multiple extracellular proteases under prolonged culture conditions, it is plausible that partial proteolytic degradation contributed to the observed activity loss. The high activity detected at early time points indicates that the bacterial TPI was successfully expressed and secreted in the A. niger system. Nonetheless, the sharp decline at day 3 highlights a potential limitation in protein stability. This instability could result from residual extracellular protease activity or from misfolding-induced degradation via the endoplasmic reticulum-associated degradation (ERAD) pathway. To improve the stability and yield of TPI in future studies, several strategies could be considered. These include engineering host strains with reduced protease secretion, optimizing secretion signal peptides, fusing stabilization domains or tags to the protein, and co-expressing molecular chaperones like BipA and PDI to enhance folding and secretion efficiency. Incorporating these approaches may help maintain enzyme activity during longer cultivation periods and improve overall productivity.

Expression of an immunoregulatory protein derived from a medicinal fungus in the engineered A. niger strain AnN2

Ganoderma lucidum, widely known as Ling Zhi, is a traditional medicinal fungus extensively utilized in Asian medicine for centuries due to its numerous health-promoting properties [52]. Recently, bioactive compounds derived from G. lucidum have gained substantial attention for their potential use in pharmaceutical and cosmetic industries [53]. One of the most notable bioactive proteins isolated from G. lucidum is the immunomodulatory protein Lingzhi-8 (LZ8), first identified in the fungal fruiting body [54]. LZ8 is composed of 111 amino acids, with a theoretical molecular mass of approximately 14 kDa.

To further assess the capabilities of our engineered protein-expression platform in producing bioactive pharmaceutical proteins, LZ8 was selected as a target protein (Fig. 3). Employing the previously described CRISPR/Cas9-mediated genome-editing approach, multiple copies of the LZ8 gene were successfully integrated into transcriptionally active loci originally occupied by TeGlaA and AnGlaA (Fig. 6A). Five recombinant strains (designated RS-LZ8-1 to RS-LZ8-5) were confirmed as positive transformants via colony PCR. The qPCR analysis confirmed that these strains contained approximately 10–13 copies of the integrated lz8 gene (Additional file 6: Fig. S4). This copy number range is consistent with other target genes expressed in the AnN2 platform, including AnGoxM (12–13 copies), MtPlyA (8–14 copies), and TPI (10–12 copies), as determined by qPCR analyses.

Fig. 6.

CRISPR/Cas9 system-mediated multi-copy expression of the immunomodulatory protein LZ8 from G. lucidum in the engineered AnN2 strain. (A) Schematic illustration of the multi-copy integration of the lz8 gene into the TeGlaA-derived high-expression loci and the AnGlaA locus in the AnN2 chassis. (B) SDS-PAGE analysis of culture supernatants from five RS-LZ8 transformants and AnN2 control strain after 48 h culture. Each lane was loaded with 10 µL of extracellular protein. A distinct protein band (~ 14 kDa, red arrow) was observed in RS-LZ8 strains but absent in the AnN2 control. (C) Western blot analysis using anti-His antibody confirmed the secretion of LZ8 protein in all RS-LZ8 strains, with consistent band patterns. Each lane was loaded with 5 µL of extracellular protein. (D) Assays of secreted LZ8 concentrations in five recombinant strains. Data were obtained from three biological replicates, and error bars represent SD. (E) SDS-PAGE of purified rLZ8 from the RS-LZ8-5 strain

Subsequently, the five recombinant strains underwent shake-flask cultivation. Following 48 h of cultivation, secreted recombinant LZ8 was analyzed by SDS-PAGE and Western blotting assays (Fig. 6B and C). A clear and consistent protein band with an apparent molecular weight of around 14 kDa was observed in all RS-LZ8 recombinant strains but not in the host control strain AnN2, confirming successful extracellular secretion. The protein yields of LZ8 in these recombinant strains ranged from approximately 72.7 to 130.5 mg/L after 48 h of culture (Fig. 6D). Additionally, the relatively low levels of background proteins secreted by the host strain AnN2 significantly simplified the downstream purification processes. Consequently, recombinant LZ8 (rLZ8) was efficiently purified from the culture broth using a single-step Ni-NTA affinity chromatography. SDS-PAGE analysis of the purified protein revealed a single homogeneous band at approximately 14 kDa, matching its theoretical molecular weight and verifying its purity (Fig. 6E). LZ8 was successfully secreted and purified as a distinct band with appropriate molecular weight, suggesting that the host system can accommodate small, non-endogenous functional proteins. Notably, the relatively low background of secreted host proteins in AnN2 facilitated downstream purification, which may benefit product quality and reproducibility in follow-up applications.

Collectively, these findings strongly indicate the high efficiency and practicality of the AnN2 engineered chassis platform for rapid and high-yield production of recombinant bioactive proteins, exemplified here by the successful expression and purification of the medicinal fungus-derived LZ8 protein. This highlights the potential of our approach for various pharmaceutical protein production applications. While LZ8 was successfully expressed and secreted in our system, its biological activity has not yet been functionally validated in this study, and further testing will be required to assess its pharmaceutical potential.

Enhanced expression of a thermostable fungal pectinase in the AnN2 by overexpressing the COPI component Cvc2

To examine the production performance of the thermostable fungal protein, the pectate lyases MtPlyA from thermophilic fungus M. thermophila [55] was chosen to express in this engineered platform (Fig. 3). The expression cassettes of MtPlyA were constructed using a similar method that mentioned above (Fig. 7A). By performing the knock-in, transformants were screened by diagnostic PCR and RT-qPCR analysis (Additional file 7: Fig. S5). Then, five positive recombinant strains contained 8, 10, 11, 13, or 14 copies of the M. thermophila MtPlyA gene were generated and renamed as the RS-MtPlyA-1 to 5. The integrated copy numbers varied among independent recombinant strains expressing AnGoxM, TPI, LZ8, and MtPlyA. This variation suggests that while the CRISPR-based multicopy integration strategy is generally effective, some target loci may remain partially unedited or undergo differential recombination during transformation. These observations support the notion that residual knock-in sites can persist following CRISPR-mediated editing, likely due to incomplete disruption of all intended loci or inherent variability in homologous recombination efficiency.

Fig. 7.

Multi-copy heterologous expression and activity analysis of the thermostable pectinase MtPlyA in the chassis strain AnN2. (A) Schematic overview of multi-copy knock-in of the MtPlyA gene into the high-transcription loci originally occupied by TeGlaA and the AnGlaA locus in AnN2 using CRISPR/Cas9. (B) SDS-PAGE analysis of culture supernatants from five recombinant RS-MtPlyA strains compared to the AnN2 host after 72 h cultivation. Each lane was loaded with 10 µL of extracellular protein. A prominent band at ~ 33 kDa (orange arrow) was observed in the recombinant strains but absent in the control. (C) Western blotting using anti-His antibodies confirmed secretion of MtPlyA in all recombinant strains, with a strong signal at the expected molecular weight. Each lane was loaded with 5 µL of extracellular protein. (D) Enzymatic activity of MtPlyA in culture broth was measured at 48 h and 72 h. Data are shown as mean ± SD from three biological replicates

The RS-MtPlyAs and its parent strain AnN2 were also cultivated in the glucose-containing medium of shake flasks for 72 h. As shown in Fig. 7B, the RS-MtPlyA-1 to 5 strains expectedly exhibited a clear strong band corresponding to the expected molecular weights of MtPlyA (~ 30 kDa), respectively. The bands were detected by western blotting using an anti-His antibody, confirming successful production (Fig. 7C). The protein band of MtPlyA was easily visualized on the SDS-PAGE gel on 72 h post incubation, indicating that MtPlyA was expressed to a higher level. Consistent with this high secreted protein production, the pectinase activities in the RS-MtPlyAs were ~ 1627. 43 − 2105.69 U/mL and ~ 2355.43 − 2732.34 U/mL after incubation for 48 h and 72 h, respectively (Fig. 7D). For the MtPlyA-expressing strains RS-MtPlyA-1 to RS-MtPlyA-5, pectate lyase activity was also maintained at comparable levels between the second and fifth passages (Fig. S5C), showing no significant decline in secretion or function. These results collectively confirm the genetic and phenotypic stability of the integrated expression cassettes across multiple generations under selective pressure, supporting the long-term reliability of the engineered platform for recombinant protein production.

To further enhance the heterologous expression of the MtPlyA in this platform, the factor Cvc2 of the secretory pathway was selected for overexpression (Fig. 8A). Cvc2 is a vesicle-associated component involved in COPI-mediated retrograde transport in A. niger between the Golgi apparatus and endoplasmic reticulum (ER), potentially affecting vesicle assembly, cargo sorting, and membrane fusion [56]. The recombinant strain RS-MtPlyA-5 with highest pectinase activity and the 14 copies of MtPlyA, was utilized as the host to enhance the production efficiency of MtPlyA. Subsequently, the Cvc2 was overexpressed in the host RS-MtPlyA-5 and three transformants were selected and named MtPlyA-OECvc2-M1 to M3. The Cvc2 overexpression strains, along with the parent strain RS-MtPlyA, were cultured in shake-flask culture for 72 h. The extracellular protein concentration in the MtPlyA-OECvc2 strains (M1–M3) increased by approximately 6.8–9.1% and 13.0–14.6% at 48 h and 72 h (Fig. 8B), respectively, compared to the parent RS-MtPlyA. As shown in Fig. 8C, Cvc2 overexpression led to a statistically significant increase in enzyme activity at 48 h, the improvement became less pronounced at 72 h when normalized to secreted protein levels. These results were further validated by SDS-PAGE analysis (Fig. 8D), clearly indicating enhanced secretion levels of MtPlyA. Specifically, the crude enzyme activity of MtPlyA in the culture broth reached approximately 3202.19–3243.49 U/mL after 72 h of cultivation. The production of rMtPlyA in the MtPlyA-OEcvc2 strains reached approximately ~ 410.1-423.7 mg/L after 72 h of shake-flask culture. Finally, the purified recombinant enzyme rMtPlyA displayed a single clear band on SDS-PAGE analysis, consistent with its predicted molecular mass (Fig. 8E). The overexpression of Cvc2 likely enhances vesicle formation and cargo trafficking, thereby contributing to improved secretion of recombinant proteins. This enhanced vesicle turnover may also relieve intracellular stress associated with protein misfolding or transport inefficiencies. While the underlying mechanism remains to be fully elucidated, our results reinforce the practical value of combining transcriptional optimization with rational engineering of the secretory pathway.

Fig. 8.

Enhancement of MtPlyA expression and secretion by overexpression of the COPI vesicle component Cvc2. (A) Schematic diagram illustrating the intracellular secretory pathway and overexpression strategy. The MtPlyA gene was integrated into high-expression loci in AnN2 and expressed under the control of native strong promoters (PAAmy or PGlaA), combined with an AAmy signal peptide and a 6×His-tag. To improve secretion efficiency, the COPI β-subunit gene Cvc2 was overexpressed to enhance ER-Golgi vesicle trafficking. (B and C) Comparison of extracellular protein concentration (B) and pectinase activity (C) in the parental RS-MtPlyA strain and three cvc2-overexpressing strains (MtPlyA-OEcvc2-M1 to M3) after 48 h and 72 h of shake-flask culture. Error bars indicate the SD from three replicates. Different lowercase letters denote significant differences as determined by Tukey’s HSD test (P < 0.05). (D) SDS-PAGE analysis of culture supernatants from MtPlyA-OECvc2 strains revealed a stronger band at ~ 33 kDa (orange arrow), corresponding to secreted MtPlyA, compared to the parental strain. Each lane was loaded with 10 µL of extracellular protein. (E) Purified rMtPlyA protein obtained via Ni²⁺-NTA affinity chromatography displayed a single band on SDS-PAGE analysis at the expected molecular mass

Taken together, our results demonstrate that integrating target genes into the multiple original high-expression loci of TeGlaA via the CRISPR/Cas9-mediated marker recycling system significantly enhances both the expression and secretion efficiency of homologous and heterologous proteins in the engineered AnN2 chassis platform. Furthermore, overexpression of the vesicular transport protein Cvc2 markedly improved the secretion and enzymatic activity of the fungal pectinase MtPlyA, highlighting the effectiveness and potential of targeted secretory pathway engineering strategies for optimizing recombinant protein production in A. niger.

Conclusion

In this study, we established and validated a modular, genome-engineered expression platform based on the industrial fungus A. niger. By integrating recombinant genes into transcriptionally active native loci and fine-tuning the secretory pathway, we achieved rapid and functional expression of enzymes and bioactive proteins from diverse sources. These included thermostable enzymes, bacterial metabolic proteins, and pharmaceutically relevant bioactive factors, demonstrating the system’s broad compatibility and adaptability. Importantly, the AnN2 chassis strain featured a simplified secretory background and exhibited high secretion efficiency within a short cultivation period (48–72 h), which facilitated downstream purification and functional recovery of target proteins. Furthermore, the enhancement of protein trafficking via overexpressing vesicular transport component highlights the importance of integrating genetic and cellular engineering to alleviate bottlenecks in protein production.

Overall, this work establishes a practical and adaptable framework for fungal expression system development. The approach may serve as a useful foundation for future applications in enzyme production, recombinant protein expression, and microbial strain engineering within the broader context of fungal synthetic biology.

Materials and methods

Strains and culture conditions

The industrial A. niger strain N1, a high-level glucoamylase-producing strain used in this study, was kindly provided by Longda Biotechnology Inc. (Shandong, China). Both the parental strain N1 and the derived transformants were cultured on Czapek-Dox new (CPZ-new) solid medium at 30 °C for 7 to 10 days to obtain conidia, following the composition described by Liu et al. [40]. Selection of positive transformants was performed by supplementing the medium with G418 or hygromycin as appropriate. For shake-flask cultivation, the culture medium (g/L) consisted of 100 g glucose, 30 g soybean flour, and 30 mL/L corn steep liquor, with the initial pH adjusted to 5.6. Conidial suspensions (1 × 10⁶ conidia/mL) were inoculated into 50 mL of medium in 250-mL Erlenmeyer flasks and incubated at 30 °C with shaking at 240 rpm for 4 days. Culture broth samples were collected after 2, 3, and 4 days of cultivation for extracellular protein quantification, enzyme activity assays, and secretome analysis. The soybean flour was included at 3% (w/v) as an organic nitrogen source. This concentration is commonly used in industrial fungal fermentation and was selected to support biomass accumulation and protein expression. During the cultivation period (48–72 h), most of the soybean-derived nutrients are expected to be metabolized by A. niger. Additionally, prior to SDS-PAGE and LC-MS/MS analysis, all culture supernatants were filtered through 0.22 μm membranes to remove residual solids and prevent interference from particulate medium components. For plasmid construction and selection, Escherichia coli Tran1-T1 was cultured at 37 °C in Luria–Bertani (LB) medium supplemented with kanamycin (50 mg/L) or ampicillin (100 mg/L) as required.

Plasmids construction for genetic engineering and protein expressing

All primers used in this study are listed in Additional file 1: Table S1. PCR fragments were amplified using Super-Fidelity DNA Polymerase (Vazyme Biotech Co., Ltd, China). Vector construction was performed with the Gibson Assembly Cloning Kit (NEB, Beijing, China). The selected recombinant proteins included A. niger glucose oxidase (AnGoxA, NCBI: MK071618.1), M. thermophila pectate lyase (MtPlyA, NCBI: XP_003664579), G. lucidum immunomodulatory protein LZ-8 (NCBI: ACD44335), porcine epidermal growth factor (pEGF, NCBI: MK071618.1), and a bacterial enzyme from E. coli, triose-phosphate isomerase (Tpi, NCBI: WP_087902427). All recombinant genes were codon-optimized for expression in A. niger and synthesized by Life Sciences Research Services (Genewiz, Suzhou, China), with a 6×His-tag inserted at the C-terminus before the stop codon.

To achieve multi-copy integration of heterologous protein genes, two donor constructs were designed for each target gene: one for the TeGlaA loci and another for the AnGlaA locus. For multiple gene knock-out or knock-in of TeGlaA-targeting integration, the 5′ and 3′ flanking sequences of the target TeGlaA gene were amplified from the A. niger N1 genome and ligated into the pUC118 vector using the Gibson Assembly Kit to generate donor-TeGlaA-TAA. The 5′ homology arm was derived from the AAmy (An12g06930) promoter region (641 bp), while the 3′ homology arm was obtained from the GlaA (An03g06550) terminator region (672 bp). A consistent homologous arm size was employed across all recombination events. The open reading frames (ORFs) of the recombinant genes, along with the 641 bp 5′ and the 672 bp 3′ TeglaA flanking regions, were assembled and inserted into the pUC118 vector, generating the following overexpression plasmids: donor-TeGlaA-AnGoxM, donor-TeGlaA-MtPlyA, donor-TeGlaA-LZ8, donor-TeGlaA-pEGF, and donor-TeGlaA-TPI. For gene deletion, the 5′ and 3′ flanking sequences of the protease-encoding gene pepA (An14g04710) and the native AnGlaA gene were amplified using paired primers. The selectable marker fragments PtrpC-neo and PtrpC-hph were amplified by PCR using the plasmids p0380-neo and pPK2-hph-gfp [39] as templates.

For integration into the AnGlaA locus, a separate donor plasmid was constructed using the native AnGlaA promoter as the 5′ regulatory element and the TtrpC terminator as the 3′ element. This promoter-terminator combination was selected to avoid homologous recombination with the PAAmy–TGlaA cassettes used at the TeGlaA sites and to ensure independent and stable expression. The use of distinct regulatory elements also facilitates modular expression tuning across loci. The ORFs of the target genes, along with the 5′ and 3′ AnGlaA homology arms, were assembled into pUC118, generating donor-AnGlaA-AnGoxM, donor-AnGlaA-TPI, donor-AnGlaA-MtPlyA, and donor-AnGlaA-LZ8. The sgRNA expression vector, driven by the A. niger U6 promoter, was designed using the sgRNACas9 tool and constructed by overlapping PCR, followed by cloning into the pJET1.2/blunt vector for sequencing. To overexpress Sect. 31 (An02g01690), the coding sequence of the COPII component Sect. 31 was amplified and inserted into the pAN52-AnPtef-TtrpC vector using Gibson Assembly to generate the plasmid pAN-AnPtef-Sect. 31-TtrpC.

Construction of A. niger strains for recombinant genes expression

A modified version of the previously established protoplast-mediated transformation and CRISPR/Cas9 method [40] was employed for genetic manipulation of A. niger N1. To construct the A. niger chassis strain for recombinant gene expression, multiple copies of the heterologous TeGlaA gene were disrupted via homologous recombination using donor-TeGlaA-TAA and our previously developed CRISPR/Cas9-mediated marker recycling system. To minimize proteolytic degradation during heterologous protein production, the PepA gene was disrupted via homologous recombination. A donor cassette containing the hph gene under control of a constitutive promoter was inserted at the PepA locus, allowing both gene disruption and selection of transformants. Briefly, approximately 10 µg of the Cas9-expression cassette, four sgRNA expression cassettes targeting TeGlaA, the sgRNA cassette targeting PepA, along with the donor DNA fragments donor-TeGlaA-TAA and donor-PepA-hph, were transformed into N1 protoplasts. Transformants were selected on Vogel’s medium supplemented with hygromycin (100 mg/L) and verified by diagnostic PCR and RT-PCR using specific primers. Positive transformants were designated as the AnN2 chassis strain.

For recombinant strains carrying multiple gene knock-ins, four sgRNAs targeting donor-TeGlaA-TAA, two sgRNA targeting native GlaA, and the corresponding donor DNA sequences were integrated into the AnN2 chassis, replacing TeGlaA or GlaA via CRISPR/Cas9-mediated homologous recombination. Additionally, sgRNA expression cassettes and donor DNA fragments of the first marker gene hph and the protease-encoding gene PepA were co-transformed into AnN2 protoplasts. Transformants were selected on G418-containing medium and further validated through diagnostic PCR and Western blot analysis following flask-shaking cultivation.

Stability assay of recombinant strains

All recombinant strains were cultivated in G418-containing medium (200 µg/mL) to maintain selection pressure and prevent cassette loss or reverse recombination. To evaluate the genetic stability of the integrated cassettes, representative recombinant strains were serially passaged under selective conditions for five generations. Cultures were incubated in shake flasks at 30 °C, and samples were collected at the 2nd and 5th passages for enzyme activity assays.

Protein, SDS-PAGE, and enzyme activity assay

The positive recombinant strains as well as their host strain AnN2 were cultured in 50 mL culture medium to further analysis. After incubation in glucose medium, extracellular protein concentrations in the culture supernatants were quantified using the Bio-Rad DC Protein Assay Kit (Bio-Rad, Hercules, CA, USA) with bovine serum albumin (BSA) as the standard. Appropriate volumes of culture supernatants were subjected to SDS-PAGE and enzyme activity assays. For SDS-PAGE, 5 µL of unconcentrated culture supernatant was loaded onto a NuPAGE^® Novex Bis-Tris polyacrylamide gel (Thermo Fisher Scientific, USA). To estimate the concentrations of specific recombinant proteins, SDS-PAGE was performed followed by Coomassie Brilliant Blue staining. Gel images were scanned and analyzed using Image Lab software (Bio-Rad). Densitometric analysis was performed to quantify the intensity of the target protein bands. The relative concentration of each recombinant protein was estimated by calculating the proportion of its band intensity relative to the total lane intensity, thereby allowing for semi-quantitative comparison of expression levels across strains and time points. Glucoamylase activity was measured using the 3,5-dinitrosalicylic acid (DNS) method, as previously described by Guo et al. [57]. One unit of glucoamylase activity was defined as the amount of enzyme catalyzing the release of 1 µmol of glucose per minute from starch hydrolysis, measured at an absorbance of 540 nm. Pectate lyase activity was determined by quantifying reducing sugars released from orange peel pectin in citric acid-phosphate buffer (pH 6.0) at 50 °C for 30 min, using the DNS method [55]. One unit of pectate lyase activity was defined as the amount of enzyme required to produce 1 µmol of galacturonic acid per minute from pectin hydrolysis. The enzyme activities of glucose oxidase (GOX), fructose-1,6-bisphosphatase (FBP), and triose-phosphate isomerase (TPI) in culture supernatants were assessed using commercially available assay kits following the manufacturer’s protocols (Solarbio Life Sciences, Beijing, China), including GOX Assay Kit (#BC0690) and TPI Assay Kit (#BC2260). One unit of glucose oxidase activity was defined as the amount of enzyme catalyzing the production of 1.0 nM of o-dianisidine dihydrochloride (oxidized) per min.

Western blot analysis and protein purification

For Western blot analysis, culture supernatants from recombinant strains grown in glucose medium for 2 days were collected and subjected to SDS-PAGE. Proteins were transferred onto a PVDF membrane and blocked with 5% skimmed milk in TBS buffer for 1 h at room temperature. The membrane was then incubated overnight at 4 °C with anti-His rabbit antibody (Cat. No. AE086, Abclonal, Wuhan, China) diluted 1:5000 in TBS buffer. After washing, the membrane was incubated for 1 h at room temperature with HRP-conjugated goat anti-rabbit IgG antibody (Cat. No. AS014, Abclonal, Wuhan, China) diluted 1:10000. Protein bands were visualized by using SuperSignal™ West Pico PLUS Chemiluminescent Substrate Kit (Thermo Scientific, Cat. No. 34580, USA) and a ChemiDoc MP Imaging System (Bio-Rad, USA). Recombinant C-terminal 6×His-tagged proteins, including AnGoxM, MtPlyA, LZ8, pEGF, and TPI were collected from culture supernatants after 2 days of shake-flask culture. For purification, 100 mL of filtered culture supernatant was first concentrated approximately 2-fold using binding buffer (20 mM sodium phosphate, pH 7.4, 500 mM NaCl, and 5 mM imidazole) through ultrafiltration with a Vivaflow 20 ultrafiltration membrane (3 kDa cutoff; Sartorius Stedim Biotech, Germany). The concentrated protein sample was then loaded onto a 5 mL HisPur™ Ni-NTA chromatography cartridge (Thermo Fisher Scientific, USA) connected to an ÄKTA Purifier 10 system (GE Healthcare, USA). Recombinant proteins were eluted using a linear imidazole gradient (5–500 mM) in the same binding buffer, and eluted fractions were collected for further SDS-PAGE analysis.

Quantitative real-time PCR and secretome analysis

Fungal mycelia were harvested by vacuum filtration, immediately frozen in liquid nitrogen, and homogenized prior to total RNA extraction using Trizol reagent (Invitrogen, Carlsbad, CA, USA). Quantitative real-time PCR (qRT-PCR) was performed following a previously described method [41]. To quantify the integrated gene copy number in A. niger N1, genomic DNA (gDNA) was extracted and used as a template for quantitative real-time PCR (RT-qPCR). The assay was performed using SYBR Green Real-time PCR Master Mix (TOYOBO, Japan) on a CFX96 Real-Time PCR Detection System (Bio-Rad, USA) by described protocol [41]. The actin gene (ANI_1_106134) served as an internal reference for normalization. Amplification efficiency was evaluated by constructing standard curves from serially diluted genomic DNA, and all reactions were performed in triplicate. Primer sequences used in this study are listed in Table S2. For secretome analysis via liquid chromatography-mass spectrometry (LC-MS/MS), the culture supernatants were collected after 4 days of growth, filtered through 0.22 μm PES membranes (Millipore), and subjected to protein quantification using the Bio-Rad protein assay kit. The supernatants from 4-day cultures were analyzed by SDS-PAGE, and subsequent LC-MS/MS analyses for protein identification were performed following the method previously described by Guo et al. [57].

Statistical analysis

All of the experiments were carried out in three independent repeated assays. Differences among experimental groups were analyzed via one-way ANOVA, with significance defined as P < 0.05.

Electronic supplementary material

Below is the link to the electronic supplementary material.

Abbreviations

CRISPR

Clustered regularly interspaced short palindromic repeats

DSBs

Double-strand breaks

AnN1

A. niger N1 strain

AnN2

A. niger N2 strain

GlaA

Glucoamylase

TeGlaA

T. emersonii glucoamylase

AnGlaA

A. niger glucoamylase

AAmy

Acid α-amylase

GOX

Glucose oxidase

MtPlyA

M. thermophila pectate lyase A

pEGF

Porcine epidermal growth factor

LZ8

Lingzhi-8

TPI

Triose-phosphate isomerase

RS-AnGoxM

Recombinant strain for AnGoxM

RS-TPI

Recombinant strain for TPI

RS-LZ8

Recombinant strain for LZ8

RS-MtPlyA

Recombinant strain for MtPlyA

Author contributions

QL and CT designed the project. FG, DL, and CG carried out the experiments. QL, FG, DL, KW, XW and YC analyzed the data and analyzed the results. QL and CT wrote the manuscript. QL, CT and YC substantively revised manuscript. All authors read and approved the final manuscript.

Funding

This study was financially supported by the National Key Research & Developmental Program of China (2023YFC3402300), the Strategic Priority Research Program of the Chinese Academy of Sciences (XDA0510300), and the National Natural Science Foundation of China (U22A20441).

Data availability

No datasets were generated or analysed during the current study.

Declarations

Ethics approval and consent to participate

Not applicable.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.