Atmospheric room-temperature plasma mutagenesis and heterologous expression of dextranase in Aspergillus niger

Abstract

Dextranase (DEX) is a high-value biocatalyst with substantial potential in oral hygiene and diverse industrial bioprocesses, yet its practical application is frequently constrained by the insufficient yields of wild-type strains. In this study, a high-efficiency DEX production system was established through an integrated strategy comprising Atmospheric and Room Temperature Plasma (ARTP) mutagenesis, molecular mechanism elucidation, and host-specific promoter engineering. A stable mutant, Chaetomium gracile B12, was isolated, exhibiting a 30.6-fold increase in DEX activity (16.23 U/mL) compared to the parental strain. Sequence analysis and multi-scale computational simulations, including AlphaFold2-based modeling and 100 ns molecular dynamics, generated a testable hypothesis that this mutation may contribute this enhancement. This substitution was found to augment structural rigidity and refine the spatial orientation of the substrate within the active pocket, facilitating a more “productive alignment” for catalysis. Furthermore, heterologous expression optimization uncovered a distinct host-specific promoter preference: the heterologous Pcdna1 promoter was most effective in C. gracile, whereas the endogenous inducible PgpdA proved superior in Aspergillus niger. Notably, this pioneering heterologous expression of DEX in A. niger streamlined the production cycle, slashing the fermentation period from 6 days to 48 h. Following systematic optimization of carbon/nitrogen sources and aeration intensity, the engineered A. niger strain H achieved a peak DEX activity of 36.00 U/mL. Biochemical characterization revealed that the recombinant DEX possessed a favorable near-neutral optimal pH (6.0–7.0) and a temperature optimum of 50 °C. Overall, this study reports the first heterologous expression of a dextranase in A. niger, reducing the fermentation period from 6 days to 48 h, and provides a comprehensive strategy for the overproduction of dextranase and offers molecular insights into enzymatic “fine-tuning”, yielding a high-titer, neutral-pH compatible biocatalyst with significant promise for oral care and industrial applications.

Supplementary Information

The online version contains supplementary material available at 10.1186/s13568-026-02074-w.

Introduction

Dextranases (α-1,6-glucan-6-glucanohydrolase, EC 3.2.1.11) are essential biocatalysts that specifically hydrolyze the α-1,6-glycosidic linkages in dextran. These enzymes have garnered significant attention due to their diverse applications in the sugar industry for reducing juice viscosity and, more importantly, in the medical and oral healthcare sectors for the degradation of dental plaque biofilms (Eggleston and Monge 2004). Given that the oral environment maintains a near-neutral pH, there is a growing industrial demand for dextranases that exhibit high catalytic efficiency and stability under neutral conditions (Khalikova et al. 2005; Pleszczyńska et al., 2017).

However, the yield of dextranase from natural microbial sources is generally low, leading to high production costs and limiting its large-scale industrial application. To overcome these bottlenecks and enhance enzyme activity, the breeding of high-producing microbial strains has remained a key research focus (Mir et al. 2023; Muthuvelu et al. 2020). Despite their potential, wild-type isolates often lack the productivity required for commercial viability. In recent years, mutagenesis and directed evolution have emerged as powerful strategies to enhance enzyme activity. Traditional mutagenesis methods, such as ultraviolet (UV) irradiation and chemical agents like EMS, are often limited by high toxicity, low mutation rates, and operational risks (Zhang et al. 2014). As a promising alternative, Atmospheric and Room Temperature Plasma (ARTP) mutagenesis has gained widespread attention. This novel physical technique offers mild operating conditions, rapid mutagenesis, a broad mutation spectrum, and high safety, generating diverse DNA damage through chemically reactive species to produce rich mutant libraries with high mutation efficiency (Zhang et al. 2014). ARTP has been successfully applied across various microorganisms, including bacteria (Ndayisenga et al. 2018), fungi (Chen et al. 2019), and microalgae (Liu et al. 2023), with notable examples such as enhancing cellulase production in Trichoderma reesei (Liu et al. 2019) and improving pullulanase activity in engineered strains (Bouassida et al. 2018). These successful applications highlight ARTP as an effective tool for overcoming the catalytic limitations of natural enzymes and obtaining mutants with desirable traits for industrial application.

In addition to directed evolution, selecting an optimal chassis cell for heterologous expression is critical for industrial enzyme production. Aspergillus niger, a filamentous fungus with a “Generally Recognized as Safe” (GRAS) status, is widely regarded as a superior host for heterologous protein expression. Compared to bacterial hosts like Escherichia coli or yeasts, A. niger possesses unparalleled advantages: it lacks endotoxins, has a robust capacity for secreting large amounts of extracellular proteins, and boasts advanced eukaryotic post-translational modification systems, which are essential for the high specific activity and thermal stability of fungal enzymes like dextranase (Kolb et al. 2005). Moreover, A. niger can utilize inexpensive carbon sources, making it highly suitable for cost-effective, high-density industrial fermentation (Meyer et al. 2011).

To establish an efficient heterologous expression system for the mutant dextranase, the selection of both a robust gene delivery platform and optimal regulatory elements is paramount. Traditional fungal transformation techniques, such as PEG-mediated protoplast transformation and biolistic bombardment, are often constrained by tedious operational procedures, low regeneration rates, and the risk of homology-dependent gene silencing caused by multi-copy tandem integrations (Lubertozzi and Keasling 2009; Michielse et al. 2005; Punt et al. 2002). In contrast, Agrobacterium tumefaciens-mediated transformation (ATMT) has emerged as a superior alternative, offering operational simplicity by bypassing the fragile protoplast stage and predominantly yielding stable, single-copy integration events (de Groot et al. 1998). This genetic stability is crucial for maintaining consistent enzymatic productivity during industrial fermentation. However, beyond stable genomic integration, the ultimate protein titer is heavily dictated by the strategic selection of regulatory elements. Promoter engineering—specifically the comparative evaluation of endogenous inducible promoters (e.g., PgpdA), constitutive promoters (e.g., PgpdA), and heterologous constitutive promoters (e.g., Pcdna1)—represents a critical, host-specific factor in overcoming expression bottlenecks. By integrating the high-fidelity ATMT system with tailored promoter-host combinations, it is possible to synchronize stable gene inheritance with robust transcriptional initiation, thereby maximizing the secretory potential of industrial chassis such as A. niger and C. gracile (Wu et al., 2024).

In this study, we report a comprehensive strategy to enhance dextranase production by integrating ARTP-induced evolution with rational host engineering. A high-yield mutant, C. gracile B12, was first isolated, and its superior performance was investigated through the lens of a novel A289V point mutation. Using multi-scale computational simulations, we explored how this substitution potentially refines the enzyme’s catalytic microenvironment. To further maximize production, the mutant gene was expressed in both C. gracile and A. niger under various promoters, followed by systematic fermentation optimization. To our knowledge, this study reports the first heterologous expression of a dextranase in A. niger, reducing the fermentation period from 6 days to 48 h. Furthermore, it provides a rare integrated example linking ARTP-derived mutation (A289V) to structural dynamics via computational simulation, offering a strategy that combines strain improvement, host engineering, and process optimization. This work not only provides a high-performance dextranase candidate for oral care applications but also offers a detailed roadmap for the synergistic improvement of fungal biocatalysts.

Materials and methods

Strains and materials

C. gracile CGMCC 3.3783 used as the initial strain for ARTP mutagenesis, was purchased from the China General Microbiological Culture Collection Center (CGMCC). A. niger ATCC 20,611 serving as the heterologous expression host, was obtained from the American Type Culture Collection (ATCC). E. coli DH5α was employed for gene cloning, while the plasmid pCAMBIA1300 (stored in our laboratory) provided the foundational framework for constructing the DEX expression cassette. Agrobacterium tumefaciens LBA4404 was used for the genetic transformation of A. niger and C. gracile. The nucleotide sequences of the DEX (A289V)-encoding gene, as well as the promoters (PglaA, PgpdA, Pcdna1) and the terminator (TglaA) utilized for heterologous expression, are provided in the Supplementary Materials. All primers (Table S1) were synthesized by Genesis Biotechnology Co., Ltd. (Nanning, China). Strains and plasmids used or constructed in this study are listed in Tables S2.

Streptomycin B, acetylacetone (AS), rifampicin (Rif), and cefoperazone sodium (Cefo) were procured from Sangon Biotech Co. Ltd. (Shanghai, China). DNA polymerase, FastPure Plasmid Mini Kit, and ClonExpress Ultra One Step Cloning Kitwere all obtained from Vazyme (Nanjing, China). All other chemicals were procured from Aladdin Biochemical Technology Co., Ltd. (Shanghai, China).

ARTP mutagenesis and lethality rate analysis

ARTP mutagenesis was performed using an ARTP-IIS system (Wuxi Yuanqing Tianmu Biotechnology Co., Ltd., China). Single colonies of Chaetomium gracile exhibiting clear hydrolysis halos were inoculated into 10 mL of liquid PDA medium and incubated at 30 °C with shaking at 200 rpm for 72 h. Mycelia were then harvested by centrifugation (4 °C, 12,000 × g, 10 min), washed twice with sterile physiological saline, and resuspended. The spore suspension was adjusted to approximately 1×10⁶–1 × 10⁸ spores/mL using a hemocytometer. An aliquot of the spore preparation was preserved at −80 °C in 1 mL of 5% (v/v) glycerol (Zhang et al. 2014).

For mutagenesis, 10 µL of the spore suspension was spread onto a sterile glass slide and treated with the ARTP instrument under the following parameters: radiofrequency power 100 W, helium gas flow rate 10 standard liters per minute (SLM), distance between the plasma jet nozzle and sample 2 mm, and treatment temperature 25 °C. Based on the preliminary lethality curve, an exposure time of 70 s was selected, with the 0 s treatment serving as the untreated control (Li et al. 2024). After treatment, the slide was transferred into a 1 mL microcentrifuge tube containing sterile water and vortexed for 1 min to elute the treated spores, yielding a mutagenized spore suspension ready for subsequent plating (Yu et al. 2022; Yuan et al. 2024; Zhang et al., 2023a).

Lethality curves were generated by varying exposure durations. To generate lethality curves, different exposure durations were applied. Resuscitated cells were serially diluted and plated on dextran screening medium, followed by incubation at 30 °C for 3 days. The lethality rate was calculated according to the following formula (Zhang et al., 2024):

Strain screening and genetic stability analysis

For primary screening of dextran-degrading strains, the mutagenized spore suspension was spread onto dextran screening medium (crude dextran 10 g/L, glucose 50 g/L, (NH₄)₂SO₄ 2 g/L, CaCl₂ 0.9 g/L, NaH₂PO₄ 0.5 g/L, K₂HPO₄ 0.5 g/L, MgSO₄·7 H₂O 0.2 g/L, and agar 20 g/L) and incubated at 30 °C for 72 h. After incubation, the diameter of each colony and its surrounding hydrolysis halo were measured. The ratio of halo diameter to colony diameter was calculated as the primary screening index. Mutants exhibiting a significantly higher halo-to-colony ratio than the WT strain were selected for secondary screening. The secondary screening was based on enzyme activity of the fermentation supernatant of mutant strains.

Genetic stability of selected strains was evaluated by subculturing them for six consecutive generations in dextran screening medium. DEX activity assay was measured at each generation to confirm phenotypic consistency.

DEX activity assay and protein analysis

The enzyme activity was determined by 3,5-dinitrosalicylic acid (DNS) method. The reaction mixture consisted of an appropriately diluted enzyme solution and 1% (w/v) dextran substrate dissolved in 100 mM citric acid-disodium hydrogen phosphate buffer (pH 7.0) was carried out at 50 °C for 10 min. The reaction was stopped by the addition of DNS followed by heating for 5 min in boiling water. The reaction system was cooled to room temperature, and the resulting solution was centrifuged at 12,000 rpm for 5 min. The absorption of the mixture supernatant was determined at 540 nm. An enzyme activity unit (U) is defined as the amount of enzyme required to release 1 µmol glucose per minute under the above determination conditions. The protein of fermentation supernatant was analyzed using 12% sodium dodecyl-sulfate polyacrylamide gel electrophoresis (SDS-PAGE), and the protein content was determined by BCA method.

Quantitative real-time PCR analysis

To assess gene expression levels, we employed quantitative real-time PCR (qPCR). The total qPCR reaction volume was 10 µL, consisting of SYBR Green, each primer, cDNA template, and nuclease-free water. The amplification conditions were a 30-s pre-denaturation at 95 °C, followed by 40 cycles of 95 °C for 5 s and 60 °C for 30 s, concluding with a melting curve analysis. Each sample was run in triplicate and included a negative control with nuclease-free water instead of cDNA. Relative gene expression levels were computed using the 2^−ΔΔCt method (Pfaffl 2001).

Sequence analysis, molecular docking, and molecular dynamics simulation

The DEX gene was amplified via PCR using cDNA as a template and specific primers (DEX-F/R, Table S1). The resulting amplicons were sequenced and verified through multiple sequence alignment.

Three-dimensional structures of the wild-type (WT) dextranase and the B12 variant (A289V) were modeled using AlphaFold2 (Gómez-Suárez et al. 2023). The structure of isomaltotetraose was retrieved from the Protein Data Bank (PDB ID: 4BFN). Molecular docking was performed via the HDOCK server (Yan et al. 2020), and the optimal enzyme–substrate complexes were visualized and analyzed using PyMOL (v3.1). Molecular dynamics (MD) simulations were conducted using GROMACS. The simulation systems were prepared via the CHARMM-GUI server, employing the AMBER ff99SB-ILDN force field for the protein and GAFF2 with AM1-BCC partial charges for the ligand. Systems were solvated in a cubic box of TIP3P water and neutralized with 50 mM NaCl. Following energy minimization, the systems were equilibrated under NVT and NPT ensembles for 2 ns each at 300 K and 1 atm. A 100 ns production run was then performed under periodic boundary conditions. Post-simulation analyses, including radius of gyration (), hydrogen bonding, and free energy landscape (FEL), were executed to assess structural stability and binding dynamics.

Construction of expression vectors for mutant B12 DEX gene

To construct the expression system, the inducible promoter PglaA and constitutive promoter PgpdA from A. niger, the strong promoter Pcdna1 from T. reesei, and the glaA terminator (TglaA) from A. niger were utilized. The pCAMBIA1300 plasmid served as the backbone for constructing six distinct DEX expression vectors.

Initially, the pCAMBIA1300 backbone was modified to replace the original CaMV 35 S promoter. A linearized vector fragment (8281 bp) excluding the 35 S promoter was amplified using the primer set 1300-gpdA-F/R. Simultaneously, the full-length gpdA promoter (1024 bp) was amplified from A. niger genomic DNA. These fragments were assembled using the ClonExpress II One Step Cloning Kit (Vazyme, Nanjing, China), yielding the intermediate plasmid 1300-gpdA-hygB. To generate a G418-resistant version, the hygB gene was replaced by a G418 resistance cassette (amplified from pPIC9K) via homologous recombination, resulting in the plasmid 1300-gpdA-G418.

For the assembly of DEX expression cassettes, three components—promoter, DEX gene, and terminator—were fused using overlap extension PCR (OE-PCR). Specifically, the glaA and cdna1 promoters were amplified with a 5′ EcoR I site and 3′ homology to the DEX gene. The DEX gene (1828 bp) was amplified from the genomic DNA of the C. gracile mutant using specific primer pairs (D-glaA-DEX-F/R, D-gpdA-DEX-F/R, or D-cdna1-DEX-F/R) to introduce flanking homology to the respective promoters and the TglaA terminator. The TglaA fragment was amplified with a 3′ Hind III site. The resulting ~ 4 kb fusion fragments (glaA-DEX-T, gpdA-DEX-T, and cdna1-DEX-T) were verified for correct assembly (Kirchgaessner et al. 2023).

Finally, these three expression cassettes were individually cloned into the EcoR I/Hind III sites of linearized 1300-gpdA-hygB or 1300-gpdA-G418 vectors using the ClonExpress Ultra One Step Cloning Kit. This yielded six final expression vectors: 1300-gpdA-hygB-(glaA/gpdA/cdna1)-DEX and 1300-gpdA-G418-(glaA/gpdA/cdna1)-DEX.

Transformation of 1300-gpdA- (glaA/gpdA/cdna1)-DEX expression vectors into C. gracile (G418) and A. niger (hygB)

The three dextranase expression vectors, 1300-gpdA-G418-glaA-DEX, 1300-gpdA-G418-gpdA-DEX and 1300-gpdA-G418-cdna1-DEX were, respectively, introduced into the genome of the high-yielding mutant strain B12 (derived from C. gracile via ARTP mutagenesis) using Agrobacterium tumefaciens-mediated transformation (ATMT), following previously established protocols (de Groot et al. 1998). Following antibiotic selection, positive transformants were identified and further evaluated for their enzyme production potential. Representative transformants were spot-inoculated onto differential agar plates containing dextran as the sole carbon source. After incubation at 30 °C for 72 h, the ratio of the hydrolysis zone diameter to the colony diameter (HC ratio) was calculated to facilitate the preliminary selection of candidates with enhanced dextranase-producing capacity.

Similarly, the three dextranase expression vectors, 1300-gpdA-hygB-glaA-DEX, 1300-gpdA-hygB-gpdA-DEX and 1300-gpdA-hygB-cdna1-DEX, were individually integrated into the genome of A. niger ATCC 20,611 via ATMT as previously described (de Groot et al. 1998; Wang et al. 2014; Zhong et al., 2007). Genetically stable transformants were obtained through hygromycin B selection. To assess their enzymatic performance, representative transformants and the parental strain (A. niger ATCC 20611, as a control) were spot-inoculated onto screening plates containing dextran as the sole carbon source. Following 72 h of incubation at 30 °C, the HC ratio was measured to identify transformants exhibiting superior dextranase secretion.

Optimization of dextranase production from A. niger

To maximize the extracellular production of DEX in A. niger ATCC 20,611, the one-factor-at-a-time method was adopted to optimize the culture conditions. This included the selection of carbon and nitrogen sources, their respective concentrations, as well as the culture volume and agitation speed. All experiments were conducted in triplicate.

Effects of temperature and pH on recombinant enzyme activity

The optimum temperature of DEX was determined by measuring its activity at 10 °C intervals within the range of 30–80 °C. For thermal stability, the enzyme solution was incubated at 30–80 °C for 1 h in the absence of substrate. After incubation, the samples were immediately cooled to room temperature, and the residual activity was determined under standard assay conditions. The activity of the untreated (non‑incubated) enzyme was defined as 100%.

The pH dependence of DEX activity was evaluated over a pH range of 3.0–10.0 using two buffer systems, including 100 mM citrate–phosphate buffer (pH 3.0–8.0) and 50 mM glycine–NaOH buffer (pH 9.0–10.0). To determine pH stability, the enzyme was incubated in buffers of different pH values (pH 3.0–10.0) at 4 °C for 1 h, followed by measurement of residual activity under standard assay conditions. For both temperature and pH assays, the highest activity observed under the tested conditions was defined as 100%, and all other activities were expressed as relative activity.

Statistical analysis

Triplicate independent experiments were performed for all assays to guarantee both experimental repeatability and statistical credibility. The obtained data were reported as mean ± standard deviation. Statistical comparisons between two groups (e.g., wild-type vs. mutant in qRT-PCR) were performed using two-tailed Student’s t-test, while comparisons involving more than two groups (e.g., different carbon sources in fermentation optimization) were analyzed by one-way ANOVA followed by Tukey’s post-hoc test. All statistical analyses were executed using Origin 9.0 software (OriginLab Corporation, Massachusetts, USA). A p value < 0.05 was considered statistically significant.

Results

ARTP mutagenesis and screening for high-yielding mutants

The ARTP mutagenesis study on C. gracile spores showed a clear, time-dependent loss of viability with increasing irradiation time (Fig. 1A). In this work, irradiation time was employed as the sole variable parameter, tested at 0, 10, 20, 30, 40, 50, 60, 70, 80, 90, 100, and 120 s. The lethality curve exhibited a clear time-dependent increase across this range, reaching approximately 90% at 70 s. To screen for high-yielding variants, the treated spores were plated on a screening medium containing crude dextran as the sole carbon source. Mutants demonstrating potentially enhanced DEX production were primarily identified based on the diameter of the hydrolysis halos formed around the colonies, a well-established rapid screening method for enzyme producers (Teather and Wood 1982). Out of approximately 1000 surviving mutants, a single isolate designated B12, which produced the largest hydrolysis halo, was selected for detailed analysis (Fig. 1B).

Fig. 1.

Screening for a high-DEX-producing mutant generated by ARTP mutagenesis. A Lethality curve of C. gracile strains following ARTP mutagenesis. B Comparison of hydrolysis halos produced by the wild-type strain and the mutant strain B12 on dextran-containing plates. C DEX activity assay of 29 primary mutants with a hydrolysis zone-to-colony diameter ratio exceeding 1.8. D Assessment of the genetic stability of mutant B12

For the screening, six high-yield DEX-producing strains with activities exceeding 10 U/mL were successfully isolated and designated as B2 (10.73 U/mL), B6 (13.46 U/mL), B12 (16.23 U/mL), B19 (13.7 U/mL), B23 (14 U/mL), and B27 (13.51 U/mL) (Fig. 1C). Among them, mutant strain B12 exhibited the largest hydrolysis halo during primary screening and showed the highest extracellular enzyme activity, representing an approximately 30.6-fold increase compared to the parental strain. In addition, strain B12 demonstrated excellent genetic stability. Although a slight decline in dextranase activity was observed after successive subcultures, the relative activity was maintained at approximately 92.4% (Fig. 1D).

Transcriptional and translational analysis of mutant B12

To elucidate the molecular basis for the enhanced dextranase activity in mutant B12, its gene expression was analyzed at both transcriptional and translational levels. Quantitative real-time PCR (qRT-PCR) analysis revealed that the transcription level of the dextranase gene in mutant B12 was significantly upregulated, showing an approximate 382.6-fold increase compared to the WT strain (p < 0.05) (Fig. 2A). This upregulation was consistently confirmed using two independent primer sets, indicating that the genetic modification successfully enhanced transcriptional activity. Following normalization of total protein concentrations in the fermentation supernatants using a protein assay kit, SDS-PAGE analysis revealed a marked increase in the intensity of the ~ 70 kDa dextranase protein band in strain B12 (Fig. 2B). These combined results indicate that the mutation enhances dextranase production through concurrent upregulation at both the transcriptional and translational levels.

Fig. 2.

A Quantitative fluorescence analysis of dextranase (DEX) gene expression in the WT strain and mutant B12. Expression levels were normalized to the 18 S rRNA gene, and data are presented as fold change relative to the WT strain. B SDS–PAGE analysis of extracellular DEX production in the WT strain and mutant B12 (lane M, protein molecular weight markers; lane 1, WT strain; lane 2, mutant B12)

Identification and structural analysis of the key A289V mutation

Given the pronounced increase in dextranase production in B12, we next investigated whether alterations in the enzyme’s primary sequence and structure might further contribute to the enhanced activity. The full-length dextranase genes from the parental strain and mutant B12 were cloned and sequenced. Comparative analysis revealed a single nonsynonymous point mutation (c.866 C > T) in B12, resulting in an alanine-to-valine substitution at position 289 of the protein (p. Ala289Val, A289V).

To identify the conserved catalytic residues of the dextranase from C. gracile, amino acid sequences of GH49 family dextranases from various fungal sources were retrieved from the NCBI GenBank and CAZy databases. Multiple sequence alignment (MSA) performed with DNAMAN9 revealed highly conserved regions across the family (Fig. 3A). By using the characterized catalytic residues of Penicillium minioluteum dextranase (PDB: 1OGM) as reference anchors, the putative catalytic triad of C. gracile dextranase was successfully mapped to Asp378, Asp397, and Asp398 (numbering based on the C. gracile sequence).

Fig. 3.

Sequence, structural analysis, and molecular dynamics (MD) simulations of C. gracile dextranase. A Multiple sequence alignment of the dextranase from C. gracile and representative GH49 family homologs. Conserved residues are highlighted, with the predicted catalytic triad indicated by red triangles. B Global three-dimensional model of the wild-type (WT) C. gracile dextranase in complex with isomaltotetraose (IM4). The red dashed box is a close-up view of the WT active-site pocket, illustrating key residues and their hydrogen-bonding distances (Å) with the substrate. C Global three-dimensional model of the B12 variant (A289V) in complex with IM4. The red dashed box is a close-up view of the B12 variant active site, highlighting the local substrate rearrangement and optimized interaction network. D RMSD of backbone C atoms of the A289V mutant and wild-type (WT) complexes during the 100 ns trajectory, reflecting global structural convergence and stability. E Per-residue RMSF profiles of the A289V mutant and wild-type (WT) complexes. F Evolution of the total number of hydrogen bonds of the A289V mutant and wild-type (WT) complexes within the protein-substrate complex over time. G Time-dependent distance fluctuations between key catalytic/structural residues of the A289V mutant and wild-type (WT) complexes

To assess the structural impact of this mutation, three-dimensional models of the wild-type (WT) and B12 variant were generated using AlphaFold2, and molecular docking with isomaltotetraose (IM4) was performed. The overall binding mode of the substrate remained consistent between the two enzymes (Fig. 3B and C). However, subtle shifts in key interaction distances were observed in the mutant, notably a “tightening” effect involving residues such as Asp319 and Gln376 (Fig. S1), suggesting a refined spatial orientation of the substrate within the active pocket.

Molecular dynamics (MD) simulations and binding energy

To further evaluate the dynamic consequences of the A289V mutation, 100 ns molecular dynamics (MD) simulations were performed. The A289V variant exhibited a lower and more stable backbone RMSD (~ 0.20 nm) post-equilibration compared to the WT (~ 0.25 nm), indicating enhanced global structural rigidity (Fig. 3D). Notably, the flexibility of the catalytic triad residues (Asp378, Asp397, Asp398) remained consistently low in both systems (RMSF < 0.15 nm), confirming that the mutation does not compromise the integrity of the active site (Fig. 3E).

Analysis of inter-residue distances and hydrogen bonding further supported a mutation-induced conformational tightening (Fig. 4F and G). Consistently, the free energy landscape (FEL) of the A289V variant revealed a narrow, deep energy well representative of a restricted conformational ensemble, in stark contrast to the broad, shallow basin observed for the WT, which is indicative of higher conformational entropy (Fig. S2).

Fig. 4.

Schematic overview of selection marker optimization and vector construction for improved fungal genetic transformation. A Antibiotic sensitivity profiles of A. niger and C. gracile. B Modification of the pCAMBIA1300 plasmid backbone. C Plasmid construction, Agrobacterium tumefaciens-mediated transformation (ATMT), and selection of positive transformants

MM/PBSA binding free energy calculations (Table 1) showed a slight enhancement in total binding affinity for the A289V variant (ΔG_total = -21.64 kcal/mol) compared to the WT (-20.69 kcal/mol). Per-residue energy decomposition revealed that this was not a uniform change, but rather a redistribution of contributions among key residues, with a notable optimization of the interaction energy for Arg430 in the mutant (Fig. S3). These findings are hypothesis-generating rather than conclusive, direct experimental validation will be necessary in the future work.

Table 1.

Binding free energy components calculated by MM/PBSA method for A289V mutant and WT complexes (kcal/mol)

System	MM/PBSA (kcal/mol)
	ΔVDWAALS	ΔEEL	ΔEGB	ΔESURF	ΔGGAS	ΔGSOLV	ΔTOTAL
A289V	− 38.44	0.00	16.99	− 0.19	− 38.44	16.80	− 21.64
WT	− 31.86	0.00	10.32	0.84	− 31.86	11.17	− 20.69

ΔΔG = ΔG(A289V) − ΔG(WT). Negative ΔΔG_total indicates enhanced binding affinity in the mutant

Engineering of expression systems in C. gracile and A. niger

To facilitate the selection of positive transformants, antibiotic resistance systems were optimized for both strains. First, the sensitivity profiles of A. niger ATCC 20,611 and C. gracile B12 toward hygromycin B (hygB) and geneticin (G418) were established, identifying the minimum inhibitory concentrations required for subsequent selection (Fig. S4). Consequently, two recombinant vectors were engineered. To ensure sufficient expression of the selectable marker in A. niger, the original CaMV 35 S promoter in pCAMBIA1300 was substituted with the endogenous PgpdA promoter from A. niger, resulting in pCAMBIA1300-gpdA-hygB (Fig. 4A and B). Given the inherent resistance of C. gracile CGMCC 3.3783 to hygB, the hygB cassette in the aforementioned plasmid was replaced with a G418 resistance marker, producing the customized expression vector pCAMBIA1300- gpdA -G418 (Fig. 4C).

To maximize the heterologous expression of the mutant DEX gene in C. gracile strain B12, three expression cassettes were constructed using distinct promoters: the inducible PglaA and constitutive PgpdA from A niger, and the Pcdna1 promoter from T. reesei. All cassettes were equipped with the A. niger TglaA terminator and integrated into the host genome via Agrobacterium-mediated transformation. Following primary screening, transformants for each promoter—designated as X3 (PglaA), X83 (PgpdA), and XXX12-161 (Pcdna1)—were selected based on their superior hydrolysis halos (Fig. S5A).

Quantitative analysis of the hydrolytic zones revealed that strain XXX12-161 achieved a halo-to-colony diameter ratio of 3.75, which was significantly higher than those of strains B12 (2.38), X3 (2.28), and X83 (2.33) (Fig. 5A). These plate-based results were further validated by shake-flask fermentation, where XXX12-161 reached a peak enzyme activity of 22.81 U/mL, representing a substantial increase over B12 (16.22 U/mL), X3 (17.33 U/mL) and X83 (16.93 U/mL) (Fig. 5B). Such pronounced variations in DEX production underscores the pivotal role of promoter selection in heterologous protein synthesis.

Fig. 5.

Comparison of DEX expression levels in C. gracile and A. niger ATCC 20,611 driven by different heterologous promoters. A The ratio of hydrolysis halo diameter to colony diameter for C. gracile strains B12, X3, X83, and XXX12-161. B Extracellular DEX enzymatic activity of C. gracile transformants post-fermentation. C The ratio of hydrolysis halo diameter to colony diameter for the WT and engineered A. niger strains H, HH-23, and HHH34-7. D DEX enzymatic activity of the WT and A. niger transformants after shake-flask fermentation. Data are presented as mean ± SD from triplicate experiments

To enhance the heterologous expression of mutant DEX in A. niger ATCC 20,611, three expression cassettes were constructed using the inducible PglaA and constitutive PgpdA promoters from A. niger, and the heterologous Pcdna1 promoter from T. reesei, all paired with the TglaA terminator (Fig. 4). These cassettes were integrated into A. niger genome via ATMT. Following transformation, candidates exhibiting the most prominent hydrolysis halos for each promoter were identified and designated as H (PglaA), HH23 (PgpdA), and HHH34-7 (Pcdna1), with the wild-type (WT) strain as the control (Fig. S5B). Quantitative analysis showed that the ratios of halo diameter to colony diameter for the WT, H, HH23, and HHH34-7 strains were 1.47, 7.50, 5.16, and 5.86, respectively (Fig. 5C). Subsequent fermentation confirmed that strain H achieved the highest DEX activity at 26.02 U/mL, significantly outperforming HH23 (14.00 U/mL), HHH34-7 (17.88 U/mL), and the WT (0.64 U/mL) (Fig. 5D). Based on the superior halo-to-colony ratios and enzymatic yields, strain H was selected for further fermentation optimization.

Optimization of fermentation conditions for A. niger strain H

To further enhance the DEX production of the engineered A. niger strain H, systematic optimization of fermentation conditions was performed, including carbon source, nitrogen source, agitation speed, and culture volume.

To evaluate the effect of carbon sources, various dextran types (T5, T40, T100, and T2000) were tested by replacing the original carbon source in the fermentation medium. Screening of the carbon sources indicated that dextran T100 was the most effective for DEX production (Fig. 6A). Subsequently, to investigate optimal concentration, different concentrations of dextran T100 (0.5%, 1.5%, 2%, 2.5%, and 3%, w/v) were supplemented to the medium. The results showed that maximum enzyme activity was achieved at 0.5% (w/v) (Fig. 6B), with higher concentrations leading to a progressive decline in DEX yield.

Fig. 6.

Optimization of fermentation conditions for enhanced dextranase (DEX) production by A. niger strain H. A Effect of different dextran molecular weights on DEX production. Crude dextran was used as the control (100%). B Effect of Dextran T100 concentration (0.5%–3.0%, w/v). The activity at 0.5% was defined as 100%. C Effect of various organic and inorganic nitrogen sources. Ammonium sulfate was set as the reference (100%). D Effect of ammonium sulfate concentration (0.1%–1.0%, w/v). The initial concentration of 0.2% was used as the control (100%). E Effect of agitation speed (160–240 rpm). The initial speed of 220 rpm was used as the control (100%). F Effect of culture volume in 250 mL flasks (25–125 mL). The initial volume of 50 mL was used as the control (100%). Error bars represent the standard deviation of three independent experiments (mean ± SD, n = 3)

To evaluate the effect of nitrogen sources, various organic and inorganic nitrogen sources, including peptone, yeast extract, beef extract, urea, and ammonium sulfate were tested in the carbon source-optimized medium. Among these, ammonium sulfate supported the highest DEX production (Fig. 6C). Further optimization of ammonium sulfate concentration (0.1%, 0.2%, 0.4%, 0.6%, and 0.8%, w/v) showed that enzyme activity peaked at 0.4% (w/v) (Fig. 6D), with deviations above or below this concentration resulting in reduced yields.

Agitation intensity influences oxygen transfer and mycelial morphology, both of which are critical for fungal fermentation (Veiter et al. 2018; Wucherpfennig et al. 2010). Under the optimized carbon and nitrogen source conditions, various rotation speeds (160, 180, 200, 220, and 240 rpm) were tested. The results showed that 180 rpm was optimal for DEX production by A. niger H, representing a significant improvement over the initial control condition of 220 rpm (Fig. 6E).

To further investigate the impact of oxygen supply, different culture volumes (25, 50, 75, 100, and 125 mL) were evaluated in 250 mL shake flasks. The results revealed that DEX production was significantly affected by liquid volume. Maximum enzyme activity was achieved at the highest tested volume of 125 mL (Fig. 6F), while lower volumes resulted in reduced DEX yields.

To evaluate the dynamic relationship between cell growth and enzyme production, samples were collected at 12-hour intervals from 12 to 72 h. Dry cell weight (DCW) was determined by centrifuging the fermentation broth and drying the pellet to constant weight. The results indicated that DEX production was growth-associated, with biomass and enzyme activity peaking at 72 h and 48 h of fermentation, respectively (Fig. 7). A comparison of the fermentation kinetics revealed a distinct decoupling between cell growth and enzyme production under the optimized conditions. Although the final biomass at 72 h in the optimized medium (0.172 g) was slightly lower than that of the original medium (0.225 g), the dextranase activity was significantly enhanced (36.00 U/mL vs. 26.02 U/mL).

Fig. 7.

Time course of DEX activity and biomass (dry cell weight, DCW) of A. niger strain H under both original and optimized fermentation conditions. Samples were collected at 12-hour intervals over a 72-hour period

Biochemical characterization of recombinant dextranase

The thermal profile and thermostability of the recombinant DEX are presented in Fig. 8A and B. The enzyme exhibited robust catalytic activity across a temperature gradient of 30–60 °C, peaking at 50 °C (100% relative activity) (Fig. 8A). Beyond this thermal optimum, a pronounced attenuation in biocatalytic efficiency was observed, with the residual activity diminishing to 35.6% and 11.4% at 60 °C and 70 °C, respectively. Regarding thermal tolerance (Fig. 8B), DEX maintained substantial structural and functional integrity at temperatures up to 50 °C. However, incubation at temperatures exceeding 60 °C triggered rapid thermal inactivation.

Fig. 8.

Biochemical characterization of the recombinant dextranase (DEX). A Temperature-activity profile of DEX. Relative activity was determined across a range of 30–80 °C, with the maximum activity at 50 °C defined as 100%. B Thermal stability of DEX. The enzyme was pre-incubated at various temperatures (30–80 °C) for 1 h, followed by the assessment of residual activity under standard assay conditions. The initial activity prior to incubation was normalized to 100%. C pH-dependency of DEX catalytic activity. The activity was evaluated using 100 mM citrate-phosphate (■) and 50 mM glycine-NaOH (●) buffer systems. The peak activity within the tested range was defined as 100%. D pH stability of DEX. The enzyme was incubated in buffers of varying pH values (3.0–10.0) at 4 °C for 1 h, and the remaining activity was subsequently measured under standard assay conditions. The activity was evaluated using 100 mM citrate-phosphate (■) and 50 mM glycine-NaOH (●) buffer systems. The activity of the enzyme stored at 4 °C in its optimal buffer was used as 100% reference. Data points represent the mean ± standard deviation (SD) of triplicate experiments (n = 3)

The pH-dependency of DEX was evaluated using Citrate-Phosphate and Glycine-NaOH buffer systems (Fig. 8C and D). The enzyme displayed a distinct catalytic preference for near-neutral to slightly acidic environments, with an optimal pH range of 6.0–7.0 (Fig. 8C). Conversely, the enzyme was virtually inactive under alkaline conditions (pH 9.0–10.0). As shown in Fig. 8D, DEX exhibited maximal stability at pH 4.0, retaining over 100% of its initial activity, followed by a fluctuating yet declining trend as the alkalinity increased. Exposure to highly alkaline environments resulted in a precipitous loss of catalytic competence, indicative of irreversible denaturation.

Discussion

The significant improvement in dextranase (DEX) activity achieved through ARTP mutagenesis in this study highlights the efficacy of this technology in generating beneficial genetic diversity. The selection of a 70 s irradiation time (~ 90% lethality) aligns with established mutagenesis theories suggesting that 80–95% lethality optimizes the balance between high mutation frequency and library viability (Zhang et al. 2023b; Peng et al. 2021). The 30.6-fold increase in activity observed in mutant B12 is particularly noteworthy, as it substantially exceeds the typical 2- to 4-fold improvements reported in classical fungal breeding (Peng et al. 2021). This magnitude of enhancement suggests that the C. gracile genome contains considerable latent potential for DEX overproduction that can be unlocked via targeted physical mutagenesis.

Mechanistically, the increased activity in B12 appears to be a result of concurrent upregulation at the transcriptional and translational levels, as evidenced by qRT-PCR and SDS-PAGE. Our qRT-PCR analysis revealed a 382.6-fold increase in DEX transcription in mutant B12 versus the wild type—an unexpectedly high upregulation for a point mutation from ARTP mutagenesis, suggesting possible alterations in cis-regulatory elements or trans-acting factors. However, this dramatic mRNA rise translated to only a 30.6-fold increase in extracellular enzyme activity. This may be the translational limitations, such as ribosomal queuing or errors due to mRNA oversaturation (Uematsu and Qian 2025), endoplasmic reticulum (ER) protein folding and processing bottlenecks, potentially triggering ER stress and unfolded protein response, which reduce synthesis and increase ER-associated degradation (ERAD) (Dabsan et al. 2025; Read and Schröder 2021).

The identification of the A289V mutation suggests that structural modifications may also play a critical role. Our simulations suggest that the A289V mutation may enhance catalytic efficiency by refining the spatial orientation of the substrate, rather than by creating new global hydrogen-bonding networks. This interpretation remains speculative and will be experimentally validated in future research. The “tightening” effect observed in the interaction distances (e.g., O8–Asp319) likely facilitates a more productive alignment for catalysis. Furthermore, MD simulations indicate that the substitution of alanine with the more hydrophobic valine reinforces hydrophobic packing, leading to increased structural rigidity and a more compact global conformation (Pace et al. 2011). The reduction in conformational entropy, visualized by the single global minimum in the free energy landscape, implies that the enzyme is effectively “locked” into a constrained structural ensemble optimized for catalysis (Frauenfelder et al. 1991). The strategic selection of promoters proved pivotal for maximizing DEX production across different hosts. In A. niger ATCC 20,611, the superior performance of transformant H underscores the efficacy of the endogenous inducible PglaA promoter over the constitutive PgpdA and heterologous Pcdna1 alternatives. This superiority likely stems from a high degree of promoter-host compatibility, where PglaA benefits from the optimal recruitment of native transcription factors and carbon-source-specific induction, leading to more robust transcriptional initiation (Meyer et al. 2011; Blumhoff et al. 2013). Conversely, the success of the heterologous T. reesei Pcdna1 promoter in C. gracile suggests that leveraging foreign regulatory elements can be an equally effective strategy to circumvent internal expression bottlenecks in specific fungal systems.

Fermentation optimization further revealed that A. niger strain H is highly sensitive to its microenvironment. The superiority of dextran T100 suggests that its specific molecular weight offers an optimal balance between substrate accessibility and hydrolysis rate, providing a steady supply of inductive oligosaccharides for the PglaA promoter (de Vries and Visser 2001). Notably, the optimal concentration was relatively low (0.5%, w/v), and the reduction in enzyme activity at higher concentrations is likely attributable to carbon catabolite repression (CCR). At elevated dextran concentrations, rapid accumulation of hydrolysis products, such as glucose, may activate the CreA repressor, thereby inhibiting the PglaA promoter (Adnan et al. 2017). This finding emphasizes the importance of precise carbon source control to maintain the derepressed state required for high-level expression (Hondmann and Visser 1994). The preference for ammonium sulfate over complex organic nitrogen sources suggests that the recombinant strain utilizes readily assimilable inorganic nitrogen more efficiently for DEX biosynthesis. Organic nitrogen sources, while rich in nutrients, may contain undefined components that could induce protease secretion, potentially leading to enzyme degradation (van den Hombergh et al. 1997). Furthermore, the optimal concentration of 0.4% (w/v) highlights the importance of maintaining an appropriate carbon-to-nitrogen (C/N) ratio. At concentrations exceeding 0.4%, the excess nitrogen likely shift metabolic flux towards biomass accumulation rather than protein secretion (Papagianni 2004), or may cause detrimental acidification of the medium, ultimately reducing enzyme yield (Nevalainen et al. 2005).

The observation that 180-rpm yielded higher DEX activity than the initial 220-rpm control is noteworthy. While higher agitation generally improves oxygen transfer, it also generates excessive hydrodynamic shear stress (Fazenda et al. 2008). Filamentous fungi such as A. niger are sensitive to shear forces; excessive agitation at 220 rpm likely caused hyphal fragmentation or mechanical damage to the mycelia, thereby impairing metabolic activity (Klöckner et al., 2014). Thus, the optimal speed of 180 rpm appears to strike a crucial balance: it provides sufficient dissolved oxygen for aerobic metabolism while maintaining a gentle enough environment to preserve fungal morphological integrity (Grimm et al. 2005).

The finding that the highest liquid volume (125 mL) supported maximum DEX production is consistent with the observed optimal agitation speed of 180 rpm. Although increasing culture volume typically reduces the oxygen transfer coefficient in shake flasks (Büchs 2001), the superior performance at 125 mL suggests that strain H favors a low-shear environment over high oxygenation. The reduced turbulence in the larger liquid volume likely minimized hydrodynamic stress on the mycelia, preserving their structural integrity and metabolic function (Wucherpfennig et al. 2010). These results indicate that protecting the fungal cells from physical damage is more critical than maximizing oxygen supply for this specific fermentation process (Driouch et al. 2009).

The optimization method used in this study is one-factor-at-a-time, which is a simple and effective strategy for identifying dominant factors, but does not consider possible interactions between variables (such as between carbon concentration and stirring speed, or between nitrogen source and culture volume). Response surface methodology (RSM) would be better suited for capturing such interactions and locating the true global optimum (Bezerra et al. 2008). Therefore, the optimal conditions reported here (0.5% Dextran T100, 0.4% (NH₄)₂SO₄, 180 rpm, 125 mL/250 mL flask) should be considered as an improved operational point rather than an absolute optimum. Encouragingly, the OFAT-guided improvements (a > 38% increase) provide a strong foundation for subsequent RSM-based fine-tuning. Moreover, our primary aim was to establish a functional and efficient heterologous expression system for dextranase in A. niger, which had not been previously reported. From this perspective, the substantial reduction in fermentation time (from 6 days to 48 h) and the significant titer improvement are meaningful achievements irrespective of the optimization method used. Future work will systematically apply RSM to explore interactions among the three most influential factors identified here, with the goal of further maximizing DEX productivity.

Following systematic fermentation optimization, the recombinant A. niger strain reached 36.00 U/mL dextranase activity in only 48 h of submerged fermentation. Compared with representative high-producing recombinant systems in Komagataella phaffii, which report 62.6–97.3 U/mL after 100 h or 69–110 U/mL after 51 h under glucose-based conditions and up to 1614 U/mL in 30 h under methanol induction (Fadel et al. 2020), the volumetric activity of our system is lower in absolute terms but is obtained in a similarly short or shorter fermentation time and without the need for hazardous inducers. Methanol-inducible K. phaffii processes raise concerns related to flammability, toxicity, and increasingly stringent regulations for food and feed enzymes (Fadel et al. 2020; Arnau et al. 2020; Deckers et al. 2020). In contrast, A. niger is a widely used, food-grade filamentous fungus for industrial enzyme production, valued for its strong secretion capacity and history of safe use (Arnau et al. 2020; Deckers et al. 2020). A methanol-free A. niger expression system therefore offers clear advantages for applications in sugar processing and potentially oral-care formulations. The observed decoupling of cell growth and enzyme production under optimized conditions indicates a successful redirection of metabolic flux from vegetative biomass accumulation toward recombinant protein synthesis. By limiting excessive vegetative growth, the optimized parameters likely enabled the fungal cells to maintain a higher specific productivity, thereby maximizing the efficiency of substrate-to-product conversion.

Biochemically, the near-neutral optimal pH (6.0–7.0) of the recombinant DEX distinguishes it from many other fungal dextranases that exhibit more acidic, such as those from C. globosum (pH 5.5) (Yang et al. 2018)d cyclopium (pH 5.0) (Wang et al. 2022). This characteristic, combined with its stability profile, makes it a promising candidate for applications in neutral-pH environments, such as oral hygiene products (Ebaya et al. 2020). Future studies should focus on verifying the genetic diversity of the broader mutant library and exploring the specific interactions between the A289V mutation and the secretion pathway to further maximize industrial yields.

Conclusion

In conclusion, this study successfully developed a high-efficiency production system for a neutral-pH stable dextranase by integrating directed evolution, molecular mechanism elucidation, and host-specific regulatory engineering. ARTP mutagenesis of C. gracile yielded a stable mutant (B12) with a 30.6-fold increase in enzymatic activity. Multi-scale simulations suggest that the A289V mutation may enhance structural rigidity and binding affinity, which requires experimental validation in the future work. Furthermore, strategic use of ATMT enabled stable, single-copy integration of the mutant gene. Promoter optimization revealed a distinct host-specific preference: the heterologous Pcdna1 promoter worked best in C. gracile, while the endogenous inducible PglaA promoter was superior in Aspergillus niger. Notably, this first successful heterologous expression of dextranase in A. niger shortened the fermentation time from 6 days to just 48 h. After systematic optimization, the engineered A. niger strain H reached a peak activity of 36.00 U/mL. The recombinant enzyme has a near-neutral pH optimum (6.0–7.0), making it suitable for oral care and other neutral-pH industrial applications where conventional acidic dextranases perform poorly. This work offers three main advances over previous reports: (1) establishing A. niger as an efficient host for dextranase production; (2) uncovering a host-specific promoter preference relevant to fungal engineering; and (3) providing a computational hypothesis for the A289V mutation’s effect. Although the individual methods are standard, their integrated application—especially the first heterologous expression of dextranase in A. niger—represents the key novelty. Overall, the synergistic strategy of promoter tailoring and fermentation optimization presented here provides both a high-performance biocatalyst and a robust methodological framework for the scalable overproduction of industrial enzymes in filamentous fungi.

Supplementary Information

Below is the link to the electronic supplementary material.

Author contributions

Xinyue Huang: Methodology, Experimental design, Formal analysis, Writing – original draft. Liyan Yang: Methodology, Data curation, Investigation, Writing – original draft. Wenbo Zhang: Methodology. Dengfeng Yang and Hao Guo: Supervision, Funding acquisition. Bin Zhang: Validation. Mingguo Jiang: Conceptualization, Supervision. Lixia Pan: Conceptualization, Funding acquisition, Resources, Methodology, Project administration, Supervision.

Funding

This research was funded by financial support from the Guangxi Science and Technology Challenge Project (Guike JB2504850001), the Science and Technology Major Project of Guangxi (Guike AA24206051), the National Natural Science Foundation of China (Grant No. 32360227, 32360244, and 32460009), Guangxi Science and Technology Program(Gui Ke Fa [2024] 102-4), and the Basic Science and Research Foundation of Guangxi Academy of Sciences (CQ-C-202302).

Data availability

The data is included in the manuscript.

Declarations

Competing interests

The authors declare no competing interests.