- Open Access
A novel sucrose-inducible expression system and its application for production of biomass-degrading enzymes in Aspergillus niger
Biotechnology for Biofuels and Bioproducts volume 16, Article number: 23 (2023)
Filamentous fungi are extensively exploited as important enzyme producers due to the superior secretory capability. However, the complexity of their secretomes greatly impairs the titer and purity of heterologous enzymes. Meanwhile, high-efficient evaluation and production of bulk enzymes, such as biomass-degrading enzymes, necessitate constructing powerful expression systems for bio-refinery applications.
A novel sucrose-inducible expression system based on the host strain Aspergillus niger ATCC 20611 and the β-fructofuranosidase promoter (PfopA) was constructed. A. niger ATCC 20611 preferentially utilized sucrose for rapid growth and β-fructofuranosidase production. Its secretory background was relatively clean because β-fructofuranosidase, the key enzyme responsible for sucrose utilization, was essentially not secreted into the medium and the extracellular protease activity was low. Furthermore, the PfopA promoter showed a sucrose concentration-dependent induction pattern and was not subject to glucose repression. Moreover, the strength of PfopA was 7.68-fold higher than that of the commonly used glyceraldehyde-3-phosphate dehydrogenase promoter (PgpdA) with enhanced green fluorescence protein (EGFP) as a reporter. Thus, A. niger ATCC 20611 coupled with the PfopA promoter was used as an expression system to express a β-glucosidase gene (bgla) from A. niger C112, allowing the production of β-glucosidase at a titer of 17.84 U/mL. The crude β-glucosidase preparation could remarkably improve glucose yield in the saccharification of pretreated corncob residues when added to the cellulase mixture of Trichoderma reesei QM9414. The efficacy of this expression system was further demonstrated by co-expressing the T. reesei-derived chitinase Chi46 and β-N-acetylglucosaminidase Nag1 to obtain an efficient chitin-degrading enzyme cocktail, which could achieve the production of N-acetyl-D-glucosamine from colloidal chitin with a conversion ratio of 91.83%. Besides, the purity of the above-secreted biomass-degrading enzymes in the crude culture supernatant was over 86%.
This PfopA-driven expression system expands the genetic toolbox of A. niger and broadens the application field of the traditional fructo-oligosaccharides-producing strain A. niger ATCC 20611, advancing it to become a high-performing enzyme-producing cell factory. In particular, the sucrose-inducible expression system possessed the capacity to produce biomass-degrading enzymes at a high level and evade endogenous protein interference, providing a potential purification-free enzyme production platform for bio-refinery applications.
Filamentous fungi possess an impressive natural capacity to secrete copious amounts of proteins (mainly hydrolytic enzymes) into the environment, highlighting their intriguing roles as hosts for the industrial-scale production of homologous and heterologous enzymes [1, 2]. In particular, Aspergillus niger has gained immense attention due to its GRAS (generally regarded as safe) status as a preferred, versatile cell factory [2, 3]. Proteome analysis reveals that A. niger can secrete a diverse range of glycoside hydrolases to cope with changing carbon sources, such as glucoamylase in the maltose-grown culture, xylanase in the xylose-grown culture [4, 5]. This property not only makes A. niger a great reservoir of glucoamylase and xylanase but also provides robust and adjustable promoters of glycoside hydrolases for enzyme production [2, 6]. For example, the promoter of the highly expressed glucoamylase-encoding gene (PglaA) has been one of the foremost inducible promoters for driving the expression of various heterologous enzymes, including β-glucosidase and cellobiohydrolase [3, 7, 8]. However, certain highly secreted background proteins, such as alpha-amylases and alpha-glucosidases, are also expressed with target enzyme under inducing conditions, which could restrict the ascension of target enzyme production [9,10,11]. Meanwhile, complex protein background is highly disadvantageous for the production of a specific protein or a relatively pure monocomponent enzyme, as it renders the purification of the target enzyme more tedious and costly [9, 12].
Attempts have been made to express enzymes in the A. niger host strains with low protein backgrounds [9, 13,14,15]. The tannase from A. niger and the Nuclease P1 from Penicillium citrinum have been successfully expressed in a low-background host strain A. niger Bdel4, which was achieved by deleting the major secreted proteins: glucoamylase, acid α-amylase and two putative α-amylases (An05g02100 and An12g06930) [13, 14]. Alternatively, Dong et al.  knocked in two copies of trehalase genes from Myceliophthora thermophila at the loci of two highly expressed extracellular secreted alpha-amylases by the CRISPR/Cas9 tool, resulting in a 793.60% increase in trehalase activity along with a substantial abatement of protein background. Nonetheless, there is a high burden associated with achieving low secretion background hosts by multiple gene deletions, which necessitates the use of multiple screening markers in a single host and efficient gene targeting system [2, 12]. Considering that the expression of amylolytic enzymes is controlled by the transcription activator AmyR, Zhang et al.  constructed an host strain ∆amyR with a low background of protein secretion by deleting amyR in A. niger CICC246. Whereas, downstream promoters of AmyR, such as PglaA, PamyA, and PamyB, reduced their activity in this strain. In this case, constitutive promoters, such as the commonly used glyceraldehyde-3-phosphate dehydrogenase promoter (PgpdA), have the advantage of independent AmyR regulation and can be candidates for expressing the target enzymes. Zhang et al.  adopted a strategy of increasing the copy numbers of gpd box to enhance the transcription efficiency of the PgpdA promoter. However, its transcription efficiency was still limited, which was only 65% of that of PglaA for A. niger CICC2462. Therefore, the establishment of a novel A. niger expression system with high efficiency and low endogenous protein interference is essential for expressing target enzymes and scaling up enzyme production to the industrial level.
Biomass, such as cellulose and chitin, is the most abundant renewable resource in nature and has been devoted to its valorization as a viable alternative to petroleum-based fuels, chemicals, and materials [17,18,19]. Nevertheless, enzymatic degradation of biomass into monomers that can be subsequently utilized by the fuel or chemical industries still presents significant barriers, such as low efficiency and high costs of biomass-degrading enzymes [18, 19]. Thus far, the commercial cellulases for cellulose degradation are predominantly from filamentous fungi, of which, Trichoderma reesei is extensively used as a source of cellulases but has a less abundance of β-glucosidase (BGL) [18, 20]. BGL plays a vital role in lignocellulose degradation by undertaking the rate-limiting final step of hydrolyzing cellobiose, which is an intermediate product and also an inhibitor of cellulase activities, into glucose [21, 22]. As such, supplementing the T. reesei cellulase preparation with exogenous BGLs is of crucial importance for efficient biomass degradation [20, 23, 24]. The addition of purified BGL to the T. reesei Rut-C30 cellulase preparation has been proven to bring about an 80% increase in glucose yield during the saccharification of pretreated cornstalk . In addition, Xia et al.  applied an engineered BGL from Talaromyces leycettanus JCM12802 that collaborated with the commercial T. reesei cellulase Celluclast 1.5L, providing a two-fold improvement in saccharification efficiency. However, the preparation of purified BGL also introduced additional complexity and cost, making it necessary to find a suitable fungal expression system for the production of high-purity BGL . Unlike the enzymatic degradation of cellulose is widely used in industry, the enzymatic degradation of chitin cannot be scaled up because the monomeric subunit N-acetyl-D-glucosamine (GlcNAc) conversion rate is always low [25, 26]. Moreover, the purity of GlcNAc remains challenging . Most crude chitinolytic enzymes from natural microorganisms always produce a mixture of GlcNAc and N-acetyl chitooligosaccharides from the enzymatic hydrolysis of chitin [28,29,30]. In general, the degradation of chitin into GlcNAc involves the synergistic action of chitinase and β-N-acetylglucosaminidase (GlcNAcase) . Thus, heterologous expression, purification, and assembly of chitinase and GlcNAcase in appropriate proportions are currently common strategies to improve the hydrolysis efficiency of colloidal chitin [31,32,33]. For example, under the co-action of purified chitinase SaChiA4 and GlcNAcase SvNag2557 with a mass ratio of 1:2, the final conversion rates of colloidal chitin to GlcNAc reached 80.2% . However, the bioprocess for GlcNAc production still needs to be optimized since chitin-degrading enzyme use for industrial applications is at an unreasonable cost [25, 26]. In addition, although Escherichia coli is the frequently used heterologous system for the expression of chitinolytic enzymes, the expression of fungal chitinases in a bacterial system is restricted by post-translational modifications, and thus filamentous fungi systems can be better alternatives [1, 27, 31, 32]. In this regard, the cost-effective and highly efficient production of biomass-degrading enzymes, such as BGL and chitinolytic enzymes, in low-background A. niger strains, would make an immense contribution to increasing the economic feasibility of large-scale biomass-based bio-refineries.
β-fructofuranosidase (FopA) is the key biocatalyst for commercial sucrose-to-fructooligosaccharides (FOS) biotransformation . Due to the high FopA productivity, A. niger ATCC 20611 has long been used for FOS production in the food industry [34,35,36]. Despite its importance as an industrially relevant organism, strain improvement by genetic engineering is limited until recently, a PEG-mediated transformation method was successfully developed in A. niger ATCC 20611 [36, 37]. Given that A. niger ATCC 20611 can use sucrose as a fermentation carbon source to produce high-level FopA, the promoter of the fopA gene (PfopA) could be a promising candidate for controlling gene expression in A. niger ATCC 20611. Moreover, FopA appears to accumulate mainly in the mycelium cells, since the mycelium cells are always utilized for the sucrose-to-FOS biotransformation [35,36,37]. Thus, the extracellular protein background of A. niger ATCC 20611 may be relatively low compared to the general enzyme-producing A. niger host strains, such as A. niger CICC2462 and CBS 513.88, which preferentially use starch for growth and enzyme production [6, 9]. Therefore, A. niger ATCC 20611 was hypothesized to be an outstanding host strain for the construction of a sucrose-inducible expression platform. To validate the hypothesis, the distribution of FopA and the extracellular protease activity of this strain under sucrose conditions were explored. In parallel, the induction pattern of PfopA was characterized, and the strength of PfopA was assessed using EGFP as a reporter. On this basis, an A. niger sucrose-inducible expression system was constructed. The system allowed high titer production of a β-glucosidase (BGLA) from A. niger C112 and a chitinolytic enzyme cocktail (including the chitinases Chi46 and GlcNAcase Nag1) from Trichoderma reesei. More importantly, these recombinant enzymes were enabled to be purification-free and used directly to degrade biomass materials.
Effect of carbon sources on the mycelial growth and β-fructofuranosidase production
A. niger ATCC 20611 was inoculated in FM with different carbon sources for 84 h to assess its growth pattern, nutritional preference, and FopA production. Figure 1 depicted the time profiles of mycelial growth and FopA production during shake flask fermentation. As shown in Fig. 1A, when fructose, glucose, sucrose, xylose, or maltose were used as the carbon source, mycelia showed excellent growth, and biomass accumulated rapidly. Moreover, the highest biomass concentration (approximately 10.14 mg/mL) was obtained in glucose and fructose after 72 h of cultivation, followed by sucrose (9.42 mg/mL) at 72 h. While growth in starch or lactose was relatively poor, especially the latter exhibiting the lowest growth of about 2.16 mg/mL. Meanwhile, the fungal cultures from different carbon sources were collected for the FopA activity assay (Fig. 1B). It was found that only the sucrose-grown culture possessed high FopA activity, especially, the maximum production of FopA reached 355.67 U/g at 36 h during the exponential growth stage. As for the other carbon source-grown cultures, the enzyme activity was below 10 U/g (Fig. 1B). Thus, sucrose was the preferred carbon source for A. niger ATCC 20611 growth as well as an essential inducer for the β-fructofuranosidase production.
Low background of hydrolase secretion and low level of extracellular protease production in A. niger ATCC 20611
Although FopA from A. niger ATCC 20611 has been characterized and used as a key biocatalyst in the industrial sucrose-to-FOS biotransformation for more than 30 years , little is known about the role that FopA plays in the sucrose utilization of this strain. To determine whether FopA was the key enzyme responsible for mycelial growth under sucrose conditions, the FopA-encoding gene (fopA) was deleted and overexpressed in A. niger ATCC 20611 to construct DF5 and OF55 strains, respectively. Then, the fungal growth and sugar consumption of the resultant mutants in CD media containing sucrose as the sole carbon source were measured. It was found that disrupting fopA resulted in a complete loss of the FopA activity and was accompanied by a severe defect in mycelial growth (Fig. 2A, B). In detail, at the end of the culture period, the fopA deletion strain DF5 displayed only 26.67% of biomass accumulation with respect to the parental strain (Fig. 2B). Correspondingly, the sucrose consumption of DF5 was 67.32% less than that of A. niger ATCC 20611 (Fig. 2B). Instead, compared to the parental strain A. niger ATCC 20611, the fopA overexpression strain OF55 showed a 1.31-fold increase in FopA production, a 48.26% increase in biomass accumulation, and a much shorter time required to consume sucrose (Fig. 2A, B). These data revealed the crucial role of FopA in the sucrose utilization of mycelia. Subsequently, the distribution of FopA activity in mycelium cells and the fermentation broth was investigated. A significant accumulation (approximately 90.23%) of the FopA activity in the mycelial cells was observed in the sucrose-grown culture, indicating A. niger ATCC 20611 was a promising host strain with a low background of hydrolase secretion (Fig. 2A).
Considering that proteolytic degradation by fungal proteases is one of the major bottlenecks interfering with efficient enzyme production [39, 40], extracellular protease produced by A. niger ATCC 20611 under the sucrose condition was determined. Equal amounts of conidia of A. niger MGG029, which is deficient in the expression of several protease genes due to a regulatory mutation, and A. niger ATCC 20611, were inoculated on the skim milk-agar plates for 3 d cultivation. The ratio of halo diameter to colony diameter was calculated to imply protease secretion ability. As depicted in Additional file 1: Figs. S1A, B, a clear halo was formed around the colony of MGG029 and the ratio of the halo diameter to the colony diameter reached 1.50, while the proteolytic halo around ATCC 20611 was invisible. In addition, the proteolytic activities toward azocasein in the fermentation broth supernatants of the fungal strains were given in Additional file 1: Fig. S1C. The extracellular protease activity of ATCC 20611 was significantly lower, only 16.59% of that of MGG029. Taken together, these findings suggest that A. niger ATCC 20611 had the advantage of low endogenous protein interference and was suitable to be developed as a host for enzyme production.
Effect of sucrose concentrations on the β-fructofuranosidase production
As mentioned above, FopA was only expressed in the presence of sucrose, implying that the promoter of the fopA gene (PfopA) was a sucrose-inducible promoter. To further characterize the induction pattern of PfopA, reverse transcription quantitative real-time PCR (RT-qPCR) was performed to monitor the changes in the fopA gene transcript levels in response to different sucrose concentrations (Fig. 3A). fopA transcript levels were enhanced with increasing sucrose concentrations ranging from 0.5 to 20%. For instance, fopA transcript levels were 5.11-fold higher at 20% (w/v) sucrose concentration than at 0.5% (w/v) sucrose concentration. Moreover, the strength of PfopA was compared to the promoter of the key glycolytic enzyme glyceraldehyde-3-phosphate dehydrogenase gene (Pgpd). It was found that the gpd gene was constitutively transcribed in the strain, regardless of the initial sucrose concentration used (Fig. 3A). Moreover, significantly higher transcript levels were observed for fopA than for gpd under all cultivation conditions (Fig. 3A). Even at the minimal sucrose induction concentration (0.5%), fopA transcript level was up to 1.43-fold higher than that of gpd, suggesting that PfopA was a strong promoter. To evaluate whether the increase in the fopA transcript level led to enhanced enzyme production, the FopA activities of mycelial cells grown in FM containing different sucrose concentrations were measured. As depicted in Fig. 3B, the FopA production showed a positive correlation with the fopA transcript levels at the sucrose concentrations used. Moreover, the maximum FopA activity could reach up to 583.23 U/g at a 20% sucrose concentration, which was 3.05-fold and 1.47-fold higher than the FopA activity of mycelial cells grown in FM with 0.5% and 4.0% sucrose as the carbon source, respectively. These data indicated that the sucrose concentration had a significant impact on the ability of PfopA to drive gene expression. However, it should be noted that high sucrose concentrations resulted in increased enzyme production but impaired mycelial cell growth. For example, when 20% (w/v) sucrose was used as a carbon source, there was a 64.45% reduction in biomass accumulation compared to 4% sucrose (data not shown). Thus, considering the optimal balance of FopA production and mycelial cell viability, an initial sucrose concentration of 4% was chosen for subsequent strain fermentation.
Furthermore, A. niger ATCC 20611 was inoculated into FM containing sucrose (S) and glucose (G) as the mixed carbon sources (2% S + 0.5% G, or 2% S + 2% G) to test whether glucose had a negative impact on sucrose induction. As shown in Fig. 4A, the gpd gene was constitutively transcribed in the strain regardless of the carbon source used. fopA was not transcribed when glucose was used as the carbon source but was excessively transcribed when sucrose was present, and the transcription level of fopA of cultures with both sucrose and glucose was similar to that of cultures with sucrose alone (Fig. 4A). The FopA production and sugar consumption of this strain under different culture conditions were further explored. When glucose was used as the sole carbon source (0.5–2% G), glucose was rapidly consumed but no FopA was produced (Fig. 4B, C). When different concentrations of glucose (0.5–2%) were added to 2% sucrose, the strain still could utilize sucrose in the early stages of fermentation (0–18 h) simultaneously with the rapid accumulation of FopA. Moreover, no reduction in FopA activity was detected during the fermentation process compared to ATCC 20611 grown with sucrose as a single carbon source (Figs. 4D–F). These results indicated that PfopA was not regulated by glucose repression, so it was able to mediate the fast production of enzymes after sucrose induction, which could facilitate a highly efficient fermentation process with complex carbon sources for enzyme production.
Comparison of the sucrose-inducible promoter PfopA and the commonly used constitutive promoter PgpdA from A. nidulans for enhanced green fluorescent protein (EGFP) expression
The PgpdA promoter from A. nidulan is the most frequently used promoter for the genetic engineering of Aspergillus spp. to provide high levels of constitutive expression of target genes . Moreover, PgpdA has been successfully used to drive the expression of β-fructofuranosidase variants in A. niger ATCC 20611 [36, 37]. Here, the strength of PfopA was compared with that of PgpdA using enhanced green fluorescent protein (EGFP) as a reporter. The EGFP expression cassettes under the control of PfopA and PgpdA were constructed to generate A. niger strains, EGFPPfopA, and EGFPPgpdA, respectively. The conidia of EGFPPfopA, EGFPPgpdA, and the parental strain ATCC 20611 were spotted on MM-sucrose and MM-glucose plates with coverslips and grown at 30 °C for 48 h. The mycelia attached to the coverslips were analyzed by fluorescence microscopy under blue light (Fig. 5A). The mycelia of ATCC 20611 did not show fluorescence on sucrose or glucose. While the strain EGFPPgpdA displayed strong cytoplasmic EGFP fluorescence on both sucrose and glucose due to the constitutive expression of EGFP. As for EGFPPfopA, a high intensity of fluorescence emission in the mycelia was observed under the sucrose condition, while no fluorescence was shown under the glucose condition. These findings confirmed that PfopA and PgpdA successfully drove egfp expression, additionally, the expression of the egfp gene under the control of PfopA was completely repressed in the absence of sucrose.
The strains were subsequently subjected to flask-level cultivation under different sucrose concentration conditions (0.5, 1, 2, 4, 10, and 20%). The transcription levels of the egfp gene driven by PgpdA and PfopA were quantified by RT-qPCR (Fig. 5B). The egfp gene was constitutively transcribed in the strain EGFPPgpdA regardless of the sucrose concentration, while the transcription levels of egfp in the strain EGFPPfopA were 1.44 to 6.32 times higher than those of the strain EGFPPgpdA after 0.5 to 20% sucrose induction. In agreement with this observation, the intensity of fluorescence of EGFPPgpdA on sucrose did not vary according to the concentrations tested (0.5, 1, 2, 4, 10, and 20%). The fluorescence intensity of EGFPPfopA was dependent on the used sucrose concentration, and the value was gradually elevated in response to 0.5 to 20% sucrose (Fig. 5C). As a result, the fluorescence intensity of EGFPPfopA could reach 7.68-fold higher than that of EGFPPgpdA at 20% (w/v) sucrose concentration. Even at the minimum induced concentration, the fluorescence intensity of EGFPPfopA was still 2.25-fold enhanced than that of EGFPPgpdA. Thus, PfopA-driven egfp expression was obviously dependent on sucrose concentration, and the transcriptional activation capacity of PfopA was significantly higher than that of PgpdA. These results indicated that the sucrose-inducible promoter PfopA appeared to be a powerful promoter to express heterologous enzymes in A. niger ATCC 20611.
Sucrose-inducible PfopA-driven expression of β-glucosidase in A. niger ATCC 20611
Low BGL activity has been recognized as a drawback for efficient cellulose biomass conversion in Trichoderma reesei [21, 22]. Thus, supplementing exogenous BGL is a powerful strategy for formulating an efficient cellulolytic cocktail and maximizing saccharification efficiency [42, 43]. In particular, BGLs from Aspergillus spp. have higher specific activities and more appropriate kinetic parameters in the hydrolysis of cellobiose in comparison to other fungal BGLs [43,44,45]. To facilitate access to available BGL for cost-effective biomass conversion, an expression cassette containing the β-glucosidase gene (bglA) from A. niger C112 under the control of PfopA was constructed and co-transformed with pyrithiamine resistance gene (ptrA)-containing plasmid pME2892 into A. niger ATCC 20611. The candidate transformants were first screened on esculin plates to test the BGL production. The size of the halo around the colony was positively correlated with the BGL activity [20, 21]. Among the transformants, FBL9 and FBL20 showed much larger black halos around the colonies than the parental strain ATCC 20611 (Fig. 6A). Subsequently, the recombinant β-glucosidase (aBGLA) expression strains were further analyzed via Southern blot assay (Fig. 6B). The digested genomic DNA was hybridized by the bgla probe, and two fragments (3.5 and 3.8 kb) were found for FBL20 and three bands (3.5, 4.0, and 4.4 kb) were observed for FBL9, while the band specific for the bgla gene was not detected in the ATCC 20611 genome. These results suggested that two and three copies of bgla were successfully integrated into the genomes of FBL20 and FBL9, respectively.
Furthermore, the two transformants, FBL9 and FBL20, were fermented in FM containing 4% sucrose as the carbon source to investigate their capacity to produce aBGLA. It was found that the relative expression of bgla normalized to the actin gene exhibited high levels in FBL9 and FBL20, while the transcript of bgla was not detected in the parental strain (Fig. 6C). After 72 h of fermentation, the fermentation broths were subjected to the enzyme assay. The secreted proteins from the fungal strains were analyzed by SDS-PAGE without further purification. As shown in Fig. 6D, a clear band of approximately 120 kDa, which was not present in ATCC 20611, was observed in the secreted proteins of FBL9 and FBL20. It was also found that the endogenous protein bands were essentially invisible in the extracellular proteins. MS identification further confirmed that the specific band was the aBGLA expressed by transformants FBL9 and FBL20 (data not shown). These results indicated that the aBGLA was successfully secreted into the culture medium. In accordance with the above result, the BGL activities of FBL9 and FBL20 measured with pNPG as a substrate could reach up to 17.84 U/mL and 15.27 U/mL, respectively (Fig. 6E). Moreover, the extracellular protein concentrations of the fungal strains were also detected. As shown in Fig. 6F, the extracellular protein concentrations of FBL9 and FBL20 (1.68 mg/mL and 1.44 mg/mL, respectively) were significantly increased compared to the parental strain (0.19 mg/mL). It can be estimated that the amount of aBGLA secreted accounted for more than 86.81% of the total amount of extracellular protein. These results revealed that PfopA can stimulate high amounts of high-purity BGL production in ATCC 20611 under the sucrose condition.
Saccharification of the corncob substrates by a combination of the T. reesei cellulase mixture and the β-glucosidase produced from A. niger FBL9
To test whether the heterologously expressed aBGLA could improve the saccharification ability of the T. reesei cellulase mixture, the culture supernatant produced by A. niger FBL9, which had the strongest ability to secret BGL among the transformants, was added into the cellulase mixture of T. reesei QM9414 with the ratio of the FPA to β-glucosidase activity as 1:1 for saccharification of pretreated corncob residues. As shown in Fig. 7A, when the acid-pretreated corncob residue (ACR) was used as the substrate, the final glucose released by T. reesei supplemented with A. niger FBL9 after a 48-h reaction was 10.35 mg/mL (corresponding to 29.65% cellulose conversion), which was 56.11% higher than that released by T. reesei (6.63 mg/mL, corresponding to 18.99% cellulose conversion). In the saccharification of delignified corncob residues (DCR), the released glucose by T. reesei with FBL9 (23.85 mg/mL, corresponding to 68.34% cellulose conversion) was 80.27% higher than the value for T. reesei (13.23 mg/mL, corresponding to 37.91% cellulose conversion) after a total enzymatic reaction of 48 h (Fig. 7B). The equal fermentation broth of the parental strain was also added to the T. reesei cellulase mixture as a control, but it had no significant effect on saccharification efficiency (Fig. 7A, B). Taken together, the aBGLA produced by A. niger FBL9 contributed to optimizing the T. reesei cellulase mixture and exhibited superior performance on the saccharification of differently pretreated corncob substrates. Therefore, it has the potential to be a cost-effective complement to traditional cellulase products for application in cellulose bioconversion.
Sucrose-inducible simultaneous expression of the T. reesei-derived chitinase and β-N-acetylglucosaminidase in A. niger ATCC 20611
To date, the heterologous expression of well-performing chitinolytic enzymes at a high level and in a cost-effective manner is essential to optimizing the enzymatic production of GlcNAc . Ike et al.  reported that the chitinase Chi46 derived from T. reesei could hydrolyze colloidal chitin to yield mainly (GlcNAc)2, and Chen et al.  found that the GlcNAcase NAG1 derived from T. reesei could hydrolyze (GlcNAc)2 to produce GlcNAc. To obtain a highly active chitinolytic enzyme-producing strain, the chi46 and nag1 expression cassettes under the control of PfopA were constructed and co-transformed with the pyrithiamine resistance gene (ptrA)-containing plasmid pME2892 into ATCC 20611 to generate transformants. 31 transformants, which grow rapidly on MM plates containing pyrithiamine, were transferred to the CDCC plates to examine their capacity to secrete chitinolytic enzymes. As shown in Fig. 8A, clear chitin hydrolysis haloes were formed around the colonies of transformants harboring the chi46 and nag1 expression cassettes, while the parental strain ATCC 20611 could not exhibit a chitin hydrolysis halo. In addition, the hydrolysis halo diameters and the colony diameters of the transformants were measured to indicate the chitinolytic enzyme production. It was found that there were considerable differences in the ratios of halo diameter (Dhalo) to colony diameter (Dcolony) of transformants (Fig. 8B). In particular, a transformant, FCN30, showed the highest value (2.85) in comparison to other fungal colonies, implying its superior ability to produce recombinant chitinolytic enzymes.
Furthermore, FCN30 was fermented using 4% sucrose as a carbon source to determine its capacity to produce chitinolytic enzymes. As shown in Fig. 9A, B, high transcription levels of chi46 and nag1 were observed in FCN30, while the transcripts of chi46 and nag1 could not be detected in ATCC 20611. The fermentation broth was used for the enzyme assay. First, SDS-PAGE analysis was applied to detect the secreted proteins from the fungal strain. Two specific protein bands at approximately 46 kDa and 80 kDa, which were in accordance with the molecular weights of Chi46 and Nag1, were observed in the fermentation broth of FCN30, while the bands were nonexistent in ATCC 20611. It was also found that the endogenous protein bands were largely invisible in the extracellular protein (Fig. 9C). MS identification further confirmed that the two specific bands were the recombinant chitinase aChi46 and GlcNAcase aNag1 expressed by FCN30 (data not shown). These results showed that the chitinolytic enzymes were successfully secreted into the culture medium. In addition, the activities of chitinolytic enzymes were quantified. When colloidal chitin was used as the substrate, the effect of temperature and pH on enzyme activity was analyzed, and the optimum temperature and pH for the chitinase were found to be 40 °C and pH 5.0 (data not shown). Under this condition, the chitinase activity of FCN30 was 4.26 U/mL, whereas the parental strain exhibited a relatively low enzyme activity of 0.82 U/mL (Fig. 9D). The activity of GlcNAcase in the fermentation broth was also measured using p-Nitrophenyl-N-acetyl–D-glucosamine (pNP-NAG) as the substrate. The optimum temperature and pH for the GlcNAcase were found to be 60 °C and pH 4.0 (data not shown). As shown in Fig. 9E, the GlcNAcase activity of FCN30 could reach up to 4.58 U/mL, while the parental strain possessed a significantly lower enzyme activity of 0.14 U/mL. In addition, the extracellular proteins were quantified by the Bradford method (Fig. 9F). The concentrations of extracellular proteins of FCN30 (1.55 mg/mL) were significantly higher than those of ATCC 20611 (0.20 mg/mL). It could be speculated that the amount of chitinolytic enzymes accounted for more than 87.11% of the total amount of extracellular proteins in FCN30. Taken together, these results revealed that PfopA could simultaneously express and secrete the T. reesei-derived bienzymatic system of chitinase and GlcNAcase in A. niger ATCC 20611 with high purity.
Bioconversion of colloidal chitin into GlcNAc by the enzyme mixture produced from A. niger FCN30
The ability of the chitinolytic enzymes produced by A. niger FCN30 to hydrolyze colloidal chitin was investigated. It was found that when the fermentation supernatant of FCN30 was used directly to hydrolyze colloidal chitin, insoluble colloidal chitin was converted to soluble components after 24 h of reaction (Fig. 10A). In contrast, the fermentation supernatant of ATCC 20611 did not significantly reduce the amount of insoluble colloidal chitin (Fig. 10A). Furthermore, the hydrolytic products were analyzed at different time intervals by TLC (Fig. 10B). Following a 30-min incubation, colloidal chitin could be hydrolyzed into GlcNAc, and elongation of the reaction time led to rapid accumulation of GlcNAc. Thus, the chitinolytic enzyme mixture produced by FCN30, which contained aChi46 and aNAG1, efficiently converted colloidal chitin to GlcNAc as the end product. However, the enzyme mixture of ATCC 20611 was unable to hydrolyze colloidal chitin into GlcNAc (Fig. 10B). Subsequently, HPLC was used to quantify the accumulation of GlcNAc in the products of the hydrolysis of colloidal chitin by the chitinolytic enzyme mixture produced by FCN30. At the end of the reaction of 24 h, the highest colloidal chitin conversion ratio of 91.83% at a GlcNAc concentration of 9.18 mg/mL was obtained (Fig. 10C). These results implied that the crude chitinolytic enzyme mixture produced by FCN30 could achieve high-purity GlcNAc production without the assistance of other enzymes and have promising potential for industrial application in chitin valorization.
Over the past few decades, the filamentous fungus A. niger has been extensively exploited as a platform organism for the production of heterologous enzymes, benefiting from its outstanding secretory capacity [49, 50]. However, target enzymes expressed in A. niger are commonly secreted with endogenous proteins, especially when carbon source-inducible promoters are used, thus affecting enzyme purity . Moreover, carbon source-inducible promoters are commonly regulated by carbon catabolite repression (CCR), and their application may negatively influence target enzyme synthesis as glucose is produced during the growth phase . On the other hand, biomass-degrading enzymes play a vital role in the conversion of biomass to biofuels and bio-products in sugar platform bio-refineries [18, 52]. Nevertheless, cost-effective access to available pure enzymes is considered one of the main obstacles that hinder biomass valorization. This study aimed to develop an efficient sucrose-inducible expression system in A. niger ATCC 20611 with minimal endogenous protein contamination for the high-level production of high-purity biomass-degrading enzymes.
Production of target enzymes by A. niger expression systems is interfered with by the complex and abundant secretome of the host strains [9, 53]. In particular, the high abundance of hydrolases and proteases in the fungal secretome leads to difficulties in the purification of the target enzymes as well as undesired degradation [4, 5]. Thus, genetic modifications by knocking out the major extracellular proteins, such as glucoamylases, and the proteases, including PepA, PepB, PepD, and PepF, were performed in the A. niger natural strains [13, 54, 55]. However, conventional and laborious deletion of multiple genes in fungi is difficult using an inadequate number of transformants and selection markers [12, 40]. Recently, a reasonable strategy of deletion of transcription factors to suppress the downstream extracellular protein expression is adopted to obtain a host with a lower background for enzyme production [9, 40]. For example, the deletion of PrtT, a transcription factor controlling the expression of extracellular protease, significantly enhanced the yield and stability of heterologous proteins [40, 56]. Nevertheless, high-secretory hydrolytic enzymes in the deletion strain still remain, such as alpha-amylase and alpha-glucosidase, leading to a low purity of target enzymes [40, 57]. Interestingly, deletion of AmyR not only reduced the secretion of amylolytic enzymes but also decreased the extracellular protease activity in the A. niger ∆amyR strain, making ∆amyR a low protein background host strain . Nevertheless, the absence of AmyR resulted in reduced activity of the commonly used PglaA and PamyA promoters . Therefore, the expression hosts constructed by the above strategies have not meet the needs of large-scale production of the target enzymes. In this study, the industrial β-fructofuranosidase-producing strain A. niger ATCC 20611 was used as the host strain. Unlike commonly used industrial glucoamylase-producing strains, such as CBS513.88 , ATCC 20611 grew poorly with starch but efficiently used sucrose to achieve rapid growth and FopA production (Fig. 1). In particular, FopA, the key enzyme responsible for sucrose utilization in ATCC 20611, was basically not secreted into the fermentation medium (Fig. 2). Furthermore, the extracellular protease activity of the strain under sucrose culture conditions was very low, with only 16.59% of A. niger MGG029, a prtT-deletion strain (Fig. 2, Additional file 1: Fig. S1). Thus, the extracellular protein background of A. niger ATCC 20611 was relatively clean, and its endogenous protein interfered minimally with the production of the target enzyme. Compared to other A. niger hosts [9, 13, 40], A. niger ATCC 20611 is an excellent host for enzyme production without disrupting the internal regulatory circuits.
The efficient expression of target enzymes depends largely on a robust promoter [3, 49, 51]. Generally, the application of inducible promoters from glycoside hydrolases, such as glucoamylase, xylanase, and amylase, is an overwhelming strategy to achieve high yields of heterologous enzyme production . However, these promoters remain subject to several limitations that hinder their widespread application, such as leakage under non-inductive conditions, carbon catabolite repression, and endogenous protein interference [9, 49, 50]. This study identified a novel sucrose-inducible promoter of the β-fructofuranosidase gene, PfopA, which was strongly induced only under sucrose conditions and completely repressed under non-inducible conditions. In particular, the existence of a positive correlation between gene relative expression and sucrose concentration made this promoter a promising candidate for fine-tuning gene expression. Moreover, PfopA could be activated by sucrose even in the presence of glucose, indicating the activity of this promoter was not repressed by glucose (Fig. 4). This property allows A. niger ATCC 20611 to use agro-industrial wastes, such as sugarcane bagasse (containing sucrose, glucose, and fructose), as a carbon source instead of pure sucrose for growth and enzyme production, enhancing the economics of applying this promoter. In addition, since FopA was essentially not secreted into the medium, PfopA was enabled to express the target enzyme without interference from endogenous proteins, resulting in highly pure enzyme production (Fig. 3). To our knowledge, some sucrose-inducible promoters have already been reported in bacteria, yeasts, and other A. niger strains for intra- and extracellular protein production [50, 60, 61], such as the promoter of a levansucrase (PsacB) from Bacillus megaterium . However, given that SacB is a secreted protein, the use of PsacB results in the background protein being secreted along with the target enzyme, thus reducing the purity of the target enzyme. And PsacB has basal promoter activity under non-inducing conditions, which is detrimental to the expression of somehow unfavorable or deleterious for the host. As for the promoter of invertase (Psuc2) from Saccharomyces cerevisiae , the Psuc2-regulated genes are tightly repressed by glucose, and thus the Psuc2-driven protein expression requires the use of purified sucrose as a substrate, which is uneconomical for large-scale cultivation of industrial fungi. Especially, a series of expression vectors based on the PsucA promoter have been constructed in A. niger ARAn10 for the intracellular production of recombinant proteins . However, time-consuming procedures about cell disruption, protein extraction, and affinity chromatography purification are required to obtain high-purity recombinant proteins. In contrast to these sucrose-inducible promoters, PfopA has the advantages of low background protein secretion, high efficiency, tight control, and not being subject to catabolite repression, making it a promising genetic tool for implementing dynamic regulatory systems to express enzymes in A. niger.
The economic utilization of cellulose or chitin as the feedstock in the bio-refinery industry represents a profound shift in industrial carbon utilization, in which the biopolymer must first be broken down into constituent monosaccharides for further bioconversions . Hence, there is a rising demand for biomass-degrading enzymes, but cost remains a limiting factor for the widespread use of enzymes. Especially for the acquisition of pure enzymes, common protocols for purifying enzymes usually include fractional precipitation, ion exchange, or gel permeation chromatography, which will increase the cost of enzyme production [31,32,33, 40]. In this regard, the application of expression systems with a low background protein secretion would facilitate the easy purification of the target enzyme or, in some cases, even eliminate the need for additional purification steps. In this study, the sucrose-inducible expression system was successfully utilized for the production of a BGL from A. niger C112, which is the rate and cost-limiting enzyme for efficient utilization of cellulose. The BGL activity of FBL9 could reach 17.84 U/mL, which is higher than the previous report, in which the BGL yield ranged from 5.87 to 8.91 U/mL in different expression systems [63, 64]. Notably, the enzyme samples could be obtained directly from the supernatant after 72 h of cultivation without any purification procedure, and the purity of the secreted aBGLA was estimated to be over 86% (Fig. 6D, E). The ability of a BGL in the synergetic enzymatic saccharification process is undoubtedly a symbolic index for estimating its utility value . The high purity of aBGLA provided an important prerequisite for the direct application of the fermentation supernatant to degraded corncob residues. When the fermentation supernatant was added to the T. reesei cellulase mixture and the ratio of FPA to BGL was increased to 1:1, the saccharifying ability toward ACR and DCR improved by 56.11%, and 80.27%, respectively. In comparison, when the ratio of FPA and purified pBGL1 from Penicillium decumbens C114-2 was 1:1, the saccharifying ability toward ACR and DCR improved only 41.9% and 65.5% respectively. When the ratio of FPA and BGL was 1:1 by adding the commercial enzyme Novozyme NS-50010, the saccharifying ability toward ACR and DCR enhanced by 60.5% and 87.4% respectively. . Thus, aBGLA seems to be more efficient and cost-effective in boosting the saccharification process than pBGL1. aBGLA is not as efficient as Novozyme NS-50010, probably because Novozyme NS-50010 preparation contains other (hemi)cellulase in addition to BGL, which will facilitate the conversion of the substrate . Collectively, the successful production and application of aBGLA not only provide a favorable supplement to reduce the cost of industrial cellulose degradation but also demonstrate that this expression system could be expected to produce other bulk enzymes.
At present, studies on the synergistic action of chitinase and GlcNAcase to hydrolyze colloidal chitin for GlcNAc production have been reported [31,32,33]. In this study, a recombinant A. niger strain FCN30 co-expressing the T. reesei chitinase Chi46 and GlcNAcase NAG1 was constructed using the sucrose-inducible expression system, and the fermentation broth of this strain was further used to convert colloidal chitin. It was found that the enzyme cocktail of FCN30 efficiently hydrolyzed colloidal chitin and generated GlcNAc as the only end product, yielding 9.18 mg/mL with a high conversion ratio of up to 91.83% after 24-h incubation (Fig. 9). The final conversion ratio is much higher than those of previous reports (62.2% ; 79% ; 80.2% ; 83% ; 85% ), and is near to that of the report by Fu et al. (92.6% ) and Cardozo et al. (93% ). It is noteworthy that the reaction time (24 h) in this study is much shorter than those of previous reports (2 d [69, 70]; 5 d ; 7 d ; 10 d ). A further advantage is that the enzyme cocktail of FCN30 is purification-free and can be used directly for chitin degradation to achieve the same conversion rate as using heterologously expressed and purified chitinolytic enzymes. Thus, the recombinant A. niger strain FCN30 could be a good candidate for GlcNAc production. Furthermore, the successful construction and application of FCN30 demonstrate the potential of this sucrose induction system for the simultaneous expression of multiple genes, providing a reference for the development of A. niger ATCC 20611 as a cell factory.
In general, promoter strength can be further enhanced by genetic modification, including deleting repressor binding sequences and adding activator binding sequences . As a consequence, the identification of transcription factors that regulate fopA expression is a prerequisite for modifying the PfopA. As a reference, a Zn(II)2Cys6-type positive-acting transcription factor InuR has been identified in N402, which is essential for the induced expression of genes involved in inulin and sucrose metabolism . However, the InuR binding sequence provided in their study has not been experimentally confirmed. Future studies are needed to focus on the transcription factors that regulate fopA expression and their binding sequences, which will be meaningful for the construction of artificial promoters to further improve enzyme production at the promoter level. In addition, sucrose molasses is considered to be a more promising inducer for bio-refineries than pure sucrose because of its higher cost-effectiveness and rich sugar content . The economic efficiency of industrial enzymes can be further improved by replacing the carbon source of the fermentation medium.
In this study, a sucrose-inducible expression system was developed based on an adjustable promoter, PfopA, and a low background host strain, A. niger ATCC 20611. The system was successfully used to produce the mono-enzyme β-glucosidase and the chitin-degrading enzyme cocktail consisting of chitinase Chi46 and GlcNAcase NAG1 at high levels. The fermentation broths of recombinant strains can be directly applied to the substrate without additional purification procedures, enabling efficient and cost-effective biomass degradation. This study not only facilitates the implementation of the industrial strain A. niger ATCC 20611 as an efficient enzyme-producing cell factory but also provides a purification-free expression platform for bulk enzyme production.
Materials and methods
Strains and culture media
A. niger ATCC 20611, an important industrial fructo-oligosaccharides-producing strain, was used as the parental strain for A. niger transformation. A. niger MGG029 ((prtT, glaA∷fleorpyrG)  was used as a control strain for the protease production assay. The cellulase producer T. reesei QM9414 was used for the saccharification of pretreated corncob residues . Strains were grown on the potato dextrose agar plate (PDA) for conidia production . The spore suspension was obtained by washing the PDA plate with solution A (containing 0.9% NaCl and 0.5% Tween 80). Fermentation medium (FM) was used for A. niger cultivation, FopA production, and sugar consumption assay . Czapek-Dox medium (CD) was used for A. niger cultivation and sugar consumption assay . T. reesei QM9414 was inoculated as described by Qian et al. for cellulase production . Transformation medium was used for protoplast regeneration . A minimal medium (MM; ) supplemented with 2 μg/mL pyrithiamine (Sigma, USA) was applied for assessing the genetic stability of transformants. And the MM agar plate containing sucrose (2%) or glucose (2%) as carbon resources was used to observe the fluorescence of the A. niger strains. The skim milk-agar plate was used to determine the protease secretion ability of strains . The esculin plate was used to screen the transformants with high β-glucosidase activity . The CD agar plate containing colloidal chitin was applied for screening of high chitinolytic enzyme producers.
Fungal growth, β-fructofuranosidase production, extracellular protease production, and sugar consumption
The spore suspension (108 spores/mL) of A. niger ATCC 20611 was inoculated in 200 mL of FM/CD. Following the sampling of these cultures, the mycelia were separated from the culture broth to measure fungal biomass and β-fructofuranosidase activity and the culture broth was used to analyze extracellular protease production and sugar consumption during the fermentation process.
After 10 mL of the cultures were filtered through a Buchner funnel, the collected mycelia were washed with distilled water three times and then dried at 85ºC to constant weight. The dry weight of mycelia was calculated as the difference between the weights of the dried filter paper with and without mycelia.
β-fructofuranosidase activity was determined according to the method described by Zhang et al. . One unit (U) was defined as the amount of enzyme required to produce 1 μmol of reducing sugars per min. Protease activity was detected according to the method described by Sun et al.  with protease K (Genview, USA) as the standard.
SBA-40C biological sensor analyzer (BISAS, Shandong, China) was used to detect the glucose concentration at different time points during the fermentation process. Sucrose Content Assay Kit (Solarbio Science & Technology Co., Ltd., Beijing, China) was used to detect the sucrose concentration at different time points during the fermentation process.
Expression cassette and deletion cassette construction
Phanta® Super-Fidelity DNA Polymerase (Vazyme Biotech Co., Ltd., Nanjing, China) was used for PCR amplification. Primers were designed using the primer premier 5.0 software and were listed in Additional file 2: Table S1.
A 2.2-kb fragment located upstream of the FopA-coding sequence (fopA) from the A. niger ATCC 20611 genome was amplified using primer pair PfopA-UF1/ PfopA-UR1 and used as the PfopA promoter. The fragments of PgpdA and TtrpC were amplified from plasmid pAN7-1 (GenBank accession number Z32698.1) with prime pairs PgpdA-UF/PgpdA-UR and TtrpC-F/TtrpC-R, respectively. The genomic DNA of T. reesei QM6a (GenBank accession number GCA_000167675.2) was used as the template to amplify the terminator Tcbh1 using primer pair Tcbh1-1DF/Tcbh1-1690DR.
For construction of an enhanced green fluorescence protein (EGFP) expression cassette, Plasmid pIG1783  was used as the template to amplify the EGFP encoding gene egfp with the primer pair egfp-F (PfopA)/egfp-R (TtrpC). PfopA/PgpdA, egfp, and TtrpC were fused by Double-joint PCR to get the final EGFPPfopA and EGFPPgpdA expression cassette using the primer pair PfopA-UF1/ TtrpC-R and PgpdA-UF1/ TtrpC-R.
For construction of a β-glucosidase (BGLA) expression cassette, the BGLA coding sequence was cloned from A. niger C112 genome (GenBank accession number KP307454.1) using the primer pair bgla-F (PfopA)/bgla-R (TtrpC). The bgla fragment was fused with PfopA and TtrpC to get the BGLA expression cassette using the primer pair PfopA-UF1/ TtrpC-R.
For construction of a chitinase (Chi46) expression cassette, the Chi46 coding gene (GenBank accession number GCA_000167675.2) was amplified from the genomic DNA of T. reesei QM6a using the specific primers chi46-F (PfopA)/chi46-R (Tcbh1). Then the chi46 fragment was fused with PfopA and Tcbh1 to get the expression cassette using the primer pair PfopA-UF2/ Tcbh1-R.
For construction of a β-N-acetylglucosaminidase (Nag1) expression cassette, the signal peptide of the cellobiohydrolase I gene (cbh1) from Trichoderma reesei QM6a (GenBank accession number GCA_000167675.2) was synthesized and added to the 5ʹ end of the nag1 gene using the primer pair nag1-F (SPcbh1)/nag1-R (TtrpC). Then, the nag1 fragment was fused with PfopA and TtrpC to get the NAG1 expression cassette using primer pair PfopA-UF2/ TtrpC-R1.
To generate the fopA deletion cassette, the 5ʹ and 3ʹ flank fragments of fopA were amplified from the genome of A. niger ATCC 20611 using the prime pairs PfopA-UF1/ PfopA-UR1 and TfopA-F/TfopA-R, respectively. The expression cassette of resistance gene ptrA was amplified from the plasmid of pIG1783 using the prime pair ptrA-F (PfopA)/ptrA-R (TfopA). Then the three purified DNA fragments were fused to amplify the FopA deletion cassette using the prime pair PfopA-UF1/ TfopA-R1.
For fopA overexpression cassette construction, the fopA gene from the A. niger ATCC 20611 genome was amplified using primer pair fopA-F (PgpdA)/fopA-R (TtrpC). Then, the fopA fragment was fused with PgpdA and TtrpC to get the fopA overexpression cassette with the primer pair PgpdA-UF/TtrpC-R1 using Double-joint PCR method .
Transformation of A. niger ATCC 20611
The polyethylene glycol (PEG)/CaCl2-mediated transformation of protoplasts was performed based on the method described by Zhang et al. . Protoplasts of A. niger ATCC 20611 were co-transformed with the expression cassette and the plasmid pME2892 , which carries the selectable marker ptrA gene. All target transformants were purified three times via single-colony isolation on the MM plate containing 2 μg/mL Pyridinium thiamine (Sigma, USA).
Expression of EGFP in A. niger
For detection of EGFP expression, the spores of the A. niger strain were spotted on MM-sucrose/MM-glucose plates with coverslips and grown for 48 h at 30 °C. EGFP expressed in mycelia was visualized using fluorescence and light microscopy (Nikon Eclipse 80i fluorescence microscope).
For quantification of the fluorescence intensity, the A. niger strains were grown in FM for 48 h. Mycelial cells were subjected to fluorescent intensity analyses using a multimode plate reader (Tecan Infinite Pro 200, Switzerland) with excitation and emission wavelengths of 488 and 520 nm, respectively. The parental strain of ATCC 20611 without an integrated EGFP expression cassette was regarded as a control.
β-Glucosidase activity assay
For visualization of BGL expression and secretion, the purified transformants containing BGLA expression cassette were grown onto CMC-esculin plates at 30℃ for 10 h. To further quantify the expression of β-glucosidase, the spores of transformants and the parental strain ATCC 20611 were cultured in FM with 4% sucrose as a carbon source at 30 ℃, 200 rpm for 72 h. The fermentation supernatants were collected by centrifugation at 10,000 rpm for 10 min and filtered by a 0.22-micron membrane. The β-glucosidase activity was measured using p-Nitrophenyl-β-d-glucopyranoside (pNPG) as a substrate . One unit of enzyme activity was defined as the amount of enzyme required that liberated 1 μmol p-nitrophenol per min under the assay condition, with p-nitrophenol as the standard.
Chitinolytic enzyme activity assay
The ability of the strain to produce chitinolytic enzymes was first assessed using colloidal chitin as a substrate. Colloidal chitin was prepared as described by Sandhya et al. . A. niger strains were plated on CD-colloidal chitin (1%) agar plates (CDCC) and grown for 72 h. To further determine the production of chitinolytic enzymes, the spores (106/mL) of transformants FCN30 and the parental strain ATCC 20611 were cultured in FM with 4% sucrose as a carbon source at 30 °C and 200 rpm for 96 h. The fermentation supernatants were collected by a 0.22-micron membrane and subjected to enzyme assays.
The activity of chitinolytic enzymes was determined following the method of Sandhya et al.  using colloidal chitin as a substrate. One unit (U) of chitinase activity was defined as the amount of enzyme required to liberate 1 μmol of reducing sugars per minute under the detection condition.
p-Nitrophenyl-N-acetyl-β-d-glucosaminide (pNP-NAG) was used as the substrate to determine the GlcNAcase activity . One unit of enzyme activity was defined as the amount of enzyme that produced 1 μmol of p-nitrophenol per minute under the assay conditions.
Extracellular protein assay
The protein concentration of the fermentation broth was measured using the Bio-Rad DC Protein Assay kit (Sangon Biotech, Shanghai, China) with bovine serum albumin (BSA) as the standard protein.
For the SDS-PAGE assay, samples were prepared by boiling fermentation supernatants (10 μL) with loading buffer (containing β-Mercaptoethanol) at 100 °C for 10 min. The Unstained Protein Molecular Weight Marker (ThermoScientific, Carlsbad, CA, USA), ranging from 14.4 kDa to 116.0 kDa, was used as the size standard. The protein electrophoresis was performed with a 5% stacking gel and a 12% separating gel, and then the protein bands were visualized with Coomassie Brilliant Blue R-250 staining and acetic-acid/methanol/water (2:3:35, v/v/v) decolorizing. The predicted bands were excised for MALDI-TOF-MS identification.
Saccharification assay of pretreated corncob residues
ACR and DCR were used as substrates in the saccharification process . T. reesei QM9414 was cultured as described by Qian et al. . The fermentation broth was collected by centrifugation at 10,000 rpm for 5 min and subjected to filter paper activity (FPA) and BGL activity assays [21, 22]. 5% corncob residue was used as substrate. The enzyme mixture with equal FPA activity (10 U/g substrates) and BGL activity (10 U/g substrates) was loaded by adding citric acid buffer (pH 4.8) to make up the total volume to 30 mL. The reaction was carried out at 50 °C for 48 h, and the amount of glucose released was measured with an SBA-40C biological sensor analyzer (BISAS, Shandong, China). Cellulose conversion was calculated as described by Qian et al. .
Conversion of colloidal chitin to GlcNAc
The degradation efficiency of chitinolytic enzymes from A. niger was analyzed using TLC with colloidal chitin as the substrate. The reaction mixture containing 1% (w/v) colloidal chitin prepared in 50 mM citrate buffer (pH 5.0) with 1 mL of the fermentation broth was incubated at 50 °C. Aliquots were withdrawn at various time intervals, and the reaction was immediately stopped by boiling for 10 min. After centrifugation (10,000 g, 10 min), the supernatants were spotted onto a TLC Aluminum Silica Gel 60 F254 plate (Merck, Germany). TLC plates were developed with n-butanol/methanol/ammonia/water (5:4:2:1, v/v/v/v) and sprayed with aniline-diphenylamine (DPA) reagent, followed by heating at 105 °C in an oven until spots were visualized. N-acetyl-chitooligosaccharides ranging from GlcNAc to (GlcNAc)6 were used as standards.
The concentration of GlcNAc was quantitatively analyzed by HPLC, using a Shimadzu-LC-20A series instrument (Kyoto, Japan) equipped with a RID-20A detector and a Zorbax carbohydrate column (250 × 4.6 mm, 5 μm, Agilent Technologies). The column temperature was maintained at 45 °C. The isocratic elution condition for N-acetylglucosamine (GlcNAc) was set to acetonitrile/water (80:20, v/v) at a flow rate of 1 mL/min. GlcNAc was used as a standard. The GlcNAc concentration of the reaction was calculated by comparing the peak area to the standard solution. The conversion rate is the percentage of released GlcNAc weight (mg) to initial colloidal chitin weight (mg) .
Total RNA extraction and reverse transcription quantitative real-time PCR (RT-qPCR)
For total RNA extraction, approximately 200 mg of ATCC 20611 mycelia was harvested by a filter, then the wet mycelia were frozen by liquid nitrogen immediately. Total RNA was extracted using Trizol Reagent, according to the manufacturer’s protocol. PrimeScript™ RT reagent kit with gDNA Eraser (TaKaRa, Dalian, Liaoning, China) was used to synthesize cDNA. Primers for RT-qPCR were listed in Additional file 2: Table S1. The SYBR Premix and 1 μl of appropriately diluted cDNA were used with 200 nM of forward and reverse primers in a final volume of 10 μl. RT-qPCR protocols were set following the instructions from the SYBR Premix Ex Taq™ (Tli RNaseH Plus) kit (TaKaRa, Dalian, Liaoning, China) manufacturer. Gene transcription level was analyzed using LightCycler 480 System. Actin was used as the reference gene and the 2−ΔΔCt method was applied for calculating relative expression levels.
Southern blot analysis
The probe of bgla was a fragment amplified through PCR using primer bgla-430F/bgla-1206R to detect the bgla gene. The BgIII/BamHI-digested genomic DNA was hybridized by the bgla probe. The probe-hybridized DNA fragment was detected with the DIG High Prime DNA Labeling and Detection Starter Kit I according to the manufacturer’s protocol (Roche Diagnostics, Mannheim, Germany).
Student’s t test was used to analyze the significant difference. Significant differences were marked at p < 0.05 and p < 0.01 levels.
Availability of data and materials
The data supporting the conclusions of this article are included in this article and its Additional files. Further datasets used and analyzed during the current study are available from the corresponding author upon reasonable request.
Filter paper activity
Acid-pretreated corncob residues
Delignified corncob residues
Lübeck M, Lübeck PS. Fungal cell factories for efficient and sustainable production of proteins and peptides. Microorganisms. 2022;10(4):753.
Madhavan A, Arun KB, Sindhu R, Alphonsa Jose A, Pugazhendhi A, Binod P, et al. Engineering interventions in industrial filamentous fungal cell factories for biomass valorization. Bioresour Technol. 2022;344:126209.
Ntana F, Mortensen UH, Sarazin C, et al. Aspergillus: a powerful protein production platform. Catalysts. 2020;10(9):1064.
Lu X, Sun J, Nimtz M, Wissing J, Zeng AP, Rinas U. The intra- and extracellular proteome of Aspergillus niger growing on defined medium with xylose or maltose as carbon substrate. Microb Cell Fact. 2010;9:23.
de Oliveira JM, van Passel MW, Schaap PJ, de Graaff LH. Proteomic analysis of the secretory response of Aspergillus niger to D-maltose and D-xylose. PLoS ONE. 2011;6(6): e20865.
Cairns TC, Barthel L, Meyer V. Something old, something new: challenges and developments in Aspergillus niger biotechnology. Essays Biochem. 2021;65(2):213–24.
Amaike Campen S, Lynn J, Sibert SJ, Srikrishnan S, Phatale P, Feldman T, et al. Expression of naturally ionic liquid-tolerant thermophilic cellulases in Aspergillus niger. PLoS ONE. 2017;12(12): e0189604.
Reilly MC, Kim J, Lynn J, Simmons BA, Gladden JM, Magnuson JK, et al. Forward genetics screen coupled with whole-genome resequencing identifies novel gene targets for improving heterologous enzyme production in Aspergillus niger. Appl Microbiol Biotechnol. 2018;102(4):1797–807.
Zhang H, Wang S, Zhang XX, Ji W, Song F, Zhao Y, Li J. The amyR-deletion strain of Aspergillus niger CICC2462 is a suitable host strain to express secreted protein with a low background. Microb Cell Fact. 2016;15:68.
Wei SZ, Zhang H, Li J. Homologous expression of asparaginase gene in Aspergillus niger. J Food Sci Biotechnol. 2015;34:554–9.
Zhang YY, Zhang H, Li J. Construction of β-mannanase engineering strain of Aspergillus niger. Biotechnology. 2013;23:31–6.
Zhang Z, Xiang B, Zhao S, Yang L, Chen Y, Hu Y, Hu S. Construction of a novel filamentous fungal protein expression system based on redesigning of regulatory elements. Appl Microbiol Biotechnol. 2022;106(2):647–61.
Liu F, Wang B, Ye Y, Pan L. High level expression and characterization of tannase tan7 using Aspergillus niger SH-2 with low-background endogenous secretory proteins as the host. Protein Expr Purif. 2018;144:71–5.
Chen X, Wang B, Pan L. Heterologous expression and characterization of Penicillium citrinum nuclease P1 in Aspergillus niger and its application in the production of nucleotides. Protein Expr Purif. 2019;156:36–43.
Dong L, Yu D, Lin X, Wang B, Pan L. Improving expression of thermostable trehalase from Myceliophthora sepedonium in Aspergillus niger mediated by the CRISPR/Cas9 tool and its purification, characterization. Protein Expr Purif. 2020;165: 105482.
Zhang H, Yan JN, Zhang H, Liu TQ, Xu Y, Zhang YY, et al. Effect of gpd box copy numbers in the gpdA promoter of Aspergillus nidulans on its transcription efficiency in Aspergillus niger. FEMS Microbiol Lett. 2018;365(15).
Cho EJ, Trinh LTP, Song Y, Lee YG, Bae HJ. Bioconversion of biomass waste into high value chemicals. Bioresour Technol. 2020;298: 122386.
Kumar Saini J, Himanshu H, Kaur A, Mathur A. Strategies to enhance enzymatic hydrolysis of lignocellulosic biomass for biorefinery applications: a review. Bioresour Technol. 2022;1:127517.
Cao S, Liu Y, Shi L, Zhu W, Wang H. N-Acetylglucosamine as a platform chemical produced from renewable resources: opportunity, challenge, and future prospects. Green Chem. 2022;24:2638.
Singhania RR, Patel AK, Raj T, Tsai ML, Chen CW, Dong CD. Advances and challenges in biocatalysts application for high solid-loading of biomass for 2nd ggeneration bio-ethanol production. Catalysts. 2022;12(6):615.
Qian Y, Zhong L, Hou Y, Qu Y, Zhong Y. Characterization and strain improvement of a hypercellulytic variant, Trichoderma reesei SN1, by genetic engineering for optimized cellulase production in biomass conversion improvement. Front Microbiol. 2016;7:1349.
Gao J, Qian Y, Wang Y, Qu Y, Zhong Y. Production of the versatile cellulase for cellulose bioconversion and cellulase inducer synthesis by genetic improvement of Trichoderma reesei. Biotechnol Biofuels. 2017;10:272.
Ma L, Zhang J, Zou G, Wang C, Zhou Z. Improvement of cellulase activity in Trichoderma reesei by heterologous expression of a beta-glucosidase gene from Penicillium decumbens. Enzyme Microb Technol. 2011;49(4):366–71.
Xia W, Xu X, Qian L, Shi P, Bai Y, Luo H, Ma R, Yao B. Engineering a highly active thermophilic β-glucosidase to enhance its pH stability and saccharification performance. Biotechnol Biofuels. 2016;9:147.
Le B, Yang SH. Microbial chitinases: properties, current state and biotechnological applications. World J Microbiol Biotechnol. 2019;35(9):144.
Zhou J, Chen J, Xu N, Zhang A, Chen K, Xin F, et al. The broad-specificity chitinases: their origin, characterization, and potential application. Appl Microbiol Biotechnol. 2019;103(8):3289–95.
Zhang A, Mo X, Zhou N, Wang Y, Wei G, Hao Z, Chen K. Identification of chitinolytic enzymes in Chitinolyticbacter meiyuanensis and mechanism of efficiently hydrolyzing chitin to n-acetyl glucosamine. Front Microbiol. 2020;11: 572053.
Kuk JH, Jung WJ, Jo GH, Kim YC, Kim KY, Park RD. Production of N-acetyl-beta-D-glucosamine from chitin by Aeromonas sp. GJ-18 crude enzyme. Appl Microbiol Biotechnol. 2005;68(3):384–9.
Suresh PV, Anil Kumar PK. Enhanced degradation of α-chitin materials prepared from shrimp processing byproduct and production of N-acetyl-D-glucosamine by thermoactive chitinases from soil mesophilic fungi. Biodegradation. 2012;23(4):597–607.
Kudan S, Eksittikul T, Pichyangkura R, Park RD. Preparation of N-acetyl-D-glucosamine and N, N′-diacetylchitobiose by enzymatic hydrolysis of chitin with crude chitinases. J Biotechnol. 2010;150:89.
Li J, Gao K, Secundo F, Mao X. Biochemical characterization of two β-N-acetylglucosaminidases from Streptomyces violascens for efficient production of N-acetyl-d-glucosamine. Food Chem. 2021;364: 130393.
Fu X, Yan Q, Yang S, Yang X, Guo Y, Jiang Z. An acidic, thermostable exochitinase with β-N-acetylglucosaminidase activity from Paenibacillus barengoltzii converting chitin to N-acetyl glucosamine. Biotechnol Biofuels. 2014;7(1):174.
Jiang S, Jiang H, Zhou Y, Jiang S, Zhang G. High-level expression of β-N-Acetylglucosaminidase BsNagZ in Pichia pastoris to obtain GlcNAc. Bioprocess Biosyst Eng. 2019;42(4):611–9.
Hidaka H, Hirayama M, Sumi N. A fructooligosaccharide-producing enzyme from Aspergillus niger ATCC 20611. Agric Biol Chem. 1988;52(5):1181–7.
Zhang J, Wang L, Luan C, Liu G, Liu J, Zhong Y. Establishment of a rapid and effective plate chromogenic assay for screening of Aspergillus species with high β-fructofuranosidase activity for fructooligosaccharides production. J Microbiol Methods. 2019;166: 105740.
Zhang J, Liu C, Xie Y, Li N, Ning Z, Du N, Huang X, Zhong Y. Enhancing fructooligosaccharides production by genetic improvement of the industrial fungus Aspergillus niger ATCC 20611. J Biotechnol. 2017;249:25–33.
Wang J, Zhang J, Wang L, Liu H, Li N, Zhou H, et al. Continuous production of fructooligosaccharides by recycling of the thermal-stable β-fructofuranosidase produced by Aspergillus niger. Biotechnol Lett. 2021;43(6):1175–82.
Hirayama M, Sumi N, Hidaka H. Purification and properties of a fructooligosaccharide-producing β-fructofuranosidase from Aspergillus niger ATCC 20611. Agric Biol Chem. 1989;53(3):667–73.
Sun Y, Qian Y, Zhang J, Yao C, Wang Y, Liu H, et al. Development of a novel expression platform for heterologous protein production via deleting the p53-like regulator Vib1 in Trichoderma reesei. Enzyme Microb Technol. 2022;155: 109993.
Kamaruddin N, Storms R, Mahadi NM, Illias RM, Bakar FDA, Murad AMA. Reduction of extracellular proteases increased activity and stability of heterologous protein in Aspergillus niger. Arabian J Sci Eng. 2018;43(7):3327–38.
Lu Y, Zheng X, Wang Y, Zhang L, Wang L, Lei Y, et al. Evaluation of Aspergillus niger six constitutive strong promoters by fluorescent-auxotrophic selection coupled with flow cytometry: a case for citric acid production. J Fungi. 2022;8(6):568.
Chen M, Qin Y, Liu Z, Liu K, Wang F, Qu Y. Isolation and characterization of a β-glucosidase from Penicillium decumbens and improving hydrolysis of corncob residue by using it as cellulase supplementation. Enzyme Microb Technol. 2010;46(6):444–9.
Zhao J, Shi D, Yang S, Lin H, Chen H. Identification of an intracellular β-glucosidase in Aspergillus niger with transglycosylation activity. Appl Microbiol Biotechnol. 2020;104(19):8367–80.
Del Pozo MV, Fernández-Arrojo L, Gil-Martínez J, Montesinos A, Chernikova TN, Nechitaylo TY, et al. Microbial β-glucosidases from cow rumen metagenome enhance the saccharification of lignocellulose in combination with commercial cellulase cocktail. Biotechnol Biofuels. 2012;5(1):73.
Dotsenko GS, Gusakov AV, Rozhkova AM, Korotkova OG, Sinitsyn AP. Heterologous β-glucosidase in a fungal cellulase system: comparison of different methods for development of multienzyme cocktails. Process Biochem. 2015;50(8):1258–63.
Qin X, Xin Y, Su X, Wang X, Zhang J, Tu T, et al. Heterologous expression and characterization of thermostable chitinase and β-N-acetylhexosaminidase from Caldicellulosiruptor acetigenus and their synergistic action on the bioconversion of chitin into N-acetyl-d-glucosamine. Int J Biol Macromol. 2021;192:250–7.
Ike M, Nagamatsu K, Shioya A, Nogawa M, Ogasawara W, Okada H, et al. Purification, characterization, and gene cloning of 46 kDa chitinase (Chi46) from Trichoderma reesei PC-3-7 and its expression in Escherichia coli. Appl Microbiol Biotechnol. 2006;71(3):294–303.
Chen F, Chen XZ, Qin LN, Tao Y, Dong ZY. Characterization and homologous overexpression of an N-acetylglucosaminidase Nag1 from Trichoderma reesei. Biochem Biophys Res Commun. 2015;459(2):184–8.
Li C, Zhou J, Du G, Chen J, Takahashi S, Liu S. Developing Aspergillus niger as a cell factory for food enzyme production. Biotechnol Adv. 2020;44: 107630.
Roth AH, Dersch P. A novel expression system for intracellular production and purification of recombinant affinity-tagged proteins in Aspergillus niger. Appl Microbiol Biotechnol. 2010;86(2):659–70.
Zhang T, Liu H, Lv B, Li C. Regulating strategies for producing carbohydrate active enzymes by filamentous fungal cell factories. Front Bioeng Biotechnol. 2020;8:691.
Sethupathy S, Morales GM, Li Y, Wang Y, Jiang J, Sun J, et al. Harnessing microbial wealth for lignocellulose biomass valorization through secretomics: a review. Biotechnol Biofuels. 2021;14(1):154.
Tsang A, Butler G, Powlowski J, Panisko EA, Baker SE. Analytical and computational approaches to define the Aspergillus niger secretome. Fungal Genet Biol. 2009;46(Suppl 1):S153–60.
van den Hombergh JP, Sollewijn Gelpke MD, van de Vondervoort PJ, Buxton FP, Visser J. Disruption of three acid proteases in Aspergillus niger-effects on protease spectrum, intracellular proteolysis, and degradation of target proteins. Eur J Biochem. 1997;247(2):605–13.
Wang Y, Xue W, Sims AH, Zhao C, Wang A, Tang G, et al. Isolation of four pepsin-like protease genes from Aspergillus niger and analysis of the effect of disruptions on heterologous laccase expression. Fungal Genet Biol. 2008;45(1):17–27.
Punt PJ, Schuren FH, Lehmbeck J, Christensen T, Hjort C, van den Hondel CA. Characterization of the Aspergillus niger prtT, a unique regulator of extracellular protease encoding genes. Fungal Genet Biol. 2008;45(12):1591–9.
Levasseur A, Benoit I, Asther M, Asther M, Record E. Homologous expression of the feruloyl esterase B gene from Aspergillus niger and characterization of the recombinant enzyme. Protein Expr Purif. 2004;37(1):126–33.
Pel HJ, de Winde JH, Archer DB, Dyer PS, Hofmann G, Schaap PJ, et al. Genome sequencing and analysis of the versatile cell factory Aspergillus niger CBS 51388. Nat Biotechnol. 2007;25(2):221–31.
Fleissner A, Dersch P. Expression and export: recombinant protein production systems for Aspergillus. Appl Microbiol Biotechnol. 2010;87(4):1255–70.
Biedendieck R, Gamer M, Jaensch L, Meyer S, Rohde M, Deckwer WD, et al. A sucrose-inducible promoter system for the intra- and extracellular protein production in Bacillus megaterium. J Biotechnol. 2007;132(4):426–30.
Williams TC, Espinosa MI, Nielsen LK, Vickers CE. Dynamic regulation of gene expression using sucrose responsive promoters and RNA interference in Saccharomyces cerevisiae. Microb Cell Fact. 2015;14:43.
Sweeney MD, Xu F. Biomass converting enzymes as industrial biocatalysts for fuels and chemicals: recent developments. Catalysts. 2012;2(2):244–63.
Zhan P, Sun J, Wang F, Zhang L, Chen J. Process Optimization of β-glucosidase Production by a Mutant Strain, Aspergillus niger C112. BioResources. 2017;12(4):8937–52.
Zhu YR, Zeng BQ, Zeng L, Liu H. Cloning and bioinformatics analysis of cellobiase gene from Aspergillus niger C112. Biotechnol Bull. 2016;32(2):116.
Jung WJ, Souleimanov A, Park RD, Smith DL. Enzymatic production of N-acetyl chitooligosaccharides by crude enzyme derived from Paenibacillus illioisensis KJA-424. Carbohydr Polym. 2007;67(2):256–9.
Jamialahmadi K, Behravan J, Fathi Najafi M, Tabatabai Yazdi M, Shahverdi AR, Faramarzi MA. Enzymatic production of N-Acetyl-d-Glucosamine from chitin using crude enzyme preparation of Aeromonas sp PTCC1691. Biotechnology. 2011;10(3):292–7.
Pichyangkura R, Kudan S, Kuttiyawong K, Sukwattanasinitt M, Aiba S. Quantitative production of 2-acetamido-2-deoxy-D-glucose from crystalline chitin by bacterial chitinase. Carbohydr Res. 2002;337:557–9.
Cardozo FA, Gonzalez JM, Feitosa VA, Pessoa A, Rivera ING. Bioconversion of α-chitin into N-acetyl-glucosamine using chitinases produced by marine-derived Aeromonas caviae isolates. World J Microbiol Biotechnol. 2017;33(11):201.
Suresh PV. Biodegradation of shrimp processing bio-waste and concomitant production of chitinase enzyme and N-acetyl-D-glucosamine by marine bacteria: production and process optimization. World J Microbiol Biotechnol. 2012;28(10):2945–62.
Suresh PV, Anil Kumar PK. Enhanced degradation of α-chitin materials prepared from shrimp processing byproduct and production of N-acetyl-D-glucosamine by thermoactive chitinases from soil mesophilic fungi. Biodegradation. 2012;23:597–607.
Sashiwa H, Fujishima S, Yamano N, Kawasaki N, Nakayama A, Muraki E, et al. Production of N-acetyl-D-glucosamine from α-chitin by crude enzymes from Aeromonas hydrophila H-2330. Carbohydr Res. 2002;337:761–3.
Yuan XL, Roubos JA, van den Hondel CA, Ram AF. Identification of InuR, a new Zn(II)2Cys6 transcriptional activator involved in the regulation of inulinolytic genes in Aspergillus niger. Mol Genet Genomics. 2008;279(1):11–26.
Xie Y, Zhou H, Liu C, Zhang J, Li N, Zhao Z, Sun G, Zhong Y. A molasses habitat-derived fungus Aspergillus tubingensis XG21 with high β-fructofuranosidase activity and its potential use for fructooligosaccharides production. AMB Express. 2017;7(1):128.
Conesa A, van den Hondel CA, Punt PJ. Studies on the production of fungal peroxidases in Aspergillus niger. Appl Environ Microbiol. 2000;66(7):3016–23.
Penttilä M, Nevalainen H, Rättö M, Salminen E, Knowles J. A versatile transformation system for the cellulolytic flamentous fungus Trichoderma reesei. Gene. 1987;61(2):155–64.
Yu JH, Hamari Z, Han KH, Seo JA, Reyes-Domínguez Y, Scazzocchio C. Double-joint PCR: a PCR-based molecular tool for gene manipulations in flamentous fungi. Fungal Genet Biol. 2004;41(11):973–81.
Sandhya C, Adapa LK, Nampoothiri KM, Binod P, Szakacs G, Pandey A. Extracellular chitinase production by Trichoderma harzianum in submerged fermentation. J Basic Microbiol. 2004;44(1):49–58.
Liu Y, Jiang Z, Ma J, Ma S, Yan Q, Yang S. Biochemical characterization and structural analysis of a β-N-acetylglucosaminidase from Paenibacillus barengoltzii for efficient production of N-Acetyl-d-glucosamine. J Agric Food Chem. 2020;68(20):5648–57.
We would like to thank Jingyao Qu, Zhifeng Li, and Sen Wang of the Core Facilities for Life and Environmental Science, State Key Laboratory of Microbial Technology of Shandong University for help and guidance in RT-qPCR and Fluorescence observation. We would also like to thank Xiangmei Ren of the Core Facilities for Life and Environmental Science, State Key Laboratory of Microbial Technology of Shandong University for help and guidance in Fluorescence intensity measurement and HPLC.
This work was supported by grants from the National Key R&D Program of China (2021YFC2101300, 2018YFA0900503), the National Natural Science Foundation of China (No. 31970070), and the Youth Interdisciplinary Science and Innovative Research Groups of Shandong University (2020QNQT006).
Ethical approval and consent to participate
Consent for publication
All authors consent to the publication of this manuscript.
The authors declare that they have no competing interests.
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Additional file 1: Figure S1.
Detection of extracellular protease production by A. niger ATCC 20611 and A. niger MGG029. (A) Skim milk-agar plate assay for protease activity. Colonies of ATCC 20611 (left) and MGG029 (right) were grown on skim milk-agar plates for 3 d. (B) The ratios of halo diameter (Dhalo) to colony diameter (Dcolony). (C) Detection of protease activity in the supernatant of ATCC 20611 and MGG029 using azoic casein as substrate. Values and error bars of triplicates are presented. Asterisks indicate statistically significant differences (**p < 0.01) as assessed by Student’s t test.
Additional file 2: Table S1.
Primers used in the present study.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.
About this article
Cite this article
Wang, L., Xie, Y., Chang, J. et al. A novel sucrose-inducible expression system and its application for production of biomass-degrading enzymes in Aspergillus niger. Biotechnol Biofuels 16, 23 (2023). https://doi.org/10.1186/s13068-023-02274-7
- Aspergillus niger ATCC 20611