This project employs Pichia pastoris as a host system to optimize the expression of Plectasin NZ2114 through strategic combinations of promoters and signal peptides. Four promoters (AOX1, FDH1, CAT1, AOX7₁₃) and four signal peptides (α-factor, SP4, SP14, 0030) were systematically combined and evaluated to assess their impact on the expression levels of the target protein.

Guided by predefined design objectives, we constructed a versatile yeast cell factory platform and performed comprehensive expression profiling and functional validation. By comparing the performance of each engineered strain, we identified the optimal promoter-signal peptide combination, thereby establishing a robust foundation for scaling up the industrial production of Plectasin NZ2114.

Beyond its applied significance, this study enhances the mechanistic understanding of how promoters and signal peptides regulate recombinant protein expression in Pichia pastoris and provides a scalable framework for the efficient production of natural antimicrobial peptides.

To evaluate the gene expression driven by different promoters, we performed genetic construct development based on the pPIC9K plasmid vector. The coding sequence of the target gene Plectasin NZ2114 and the sequences of various promoters were obtained via gene synthesis, followed by PCR amplification to generate the desired DNA fragments, as shown in Figure 1.

Fig. 1 DNA electrophoresis of PCR product of promoter fragment and plasmid backbone fragment.

Electrophoretic analysis revealed DNA bands of the expected size for the target gene fragment at the theoretical position. These bands were documented by imaging and analyzed for accuracy. The target bands were then excised from the gel and subjected to DNA recovery and purification for downstream applications. Notably, the AOX713 promoter shares a homologous sequence segment with the AOX1 promoter. Therefore, we constructed an expression plasmid for NZ2114 driven by AOX713 using the AOX1 plasmid as the backbone.

Fig. 2 Transformation of Escherichia coli competent cells with promoter plasmids and verification of colony monoclonal clones

Following purification of the target gene fragment, we employed homologous recombination to assemble the fragment with the pPIC9K plasmid backbone. The ligation mixture was transformed into competent Escherichia coli cells, and transformants were selected on solid LB agar plates supplemented with ampicillin. As shown in Figures 2A and 2B, single colonies were successfully obtained. These colonies were subjected to double digestion analysis and colony PCR to identify clones harboring correctly assembled plasmids. Positive clones were then scaled up for culture, and plasmid DNA was extracted from the bacterial cultures and sent to a sequencing service provider for Sanger sequencing to confirm the accuracy of the inserted sequence.

Fig. 3 Sanger sequencing was used to identify the constructed plasmid DNA

Sequencing results confirmed that 3A corresponds to pPIC9K-AOX1-NZ2114, 3B to pPIC9K-CAT1-NZ2114, 3C to pPIC9K-FDH1-NZ2114, and 3D to pPIC9K-AOX713-NZ2114. The sequencing traces of the constructed plasmid vectors exhibited clean single peaks and were fully consistent with the theoretically designed sequences, confirming accurate assembly of all recombinant plasmids. This successful validation provides a solid foundation for the subsequent transformation of GS115 competent cells.

Fig. 4 Construction and identification of engineered yeast strains for promoters

For the successfully constructed plasmid DNA, linearization was performed followed by purification of the digested products. The linearized fragments were then transformed into GS115 chemically competent cells. Transformants were selected by plating on YPD agar supplemented with G418 and incubated at 30 °C until single colonies appeared. Individual colonies were subjected to colony PCR using a dedicated microbial lysis buffer for cell lysis, and the resulting lysates were used directly as templates. PCR products were analyzed by agarose gel electrophoresis. As shown in Figure 4, distinct bands corresponding to the expected inserts were detected for all promoter-containing plasmids. Positive yeast monoclonal strains were subsequently expanded in liquid culture to generate seed cultures for downstream experiments.

First, based on the construction of the expression plasmid with the promoter, we chose pPIC9K-AOX1-NZ2114 as the backbone plasmid for constructing different signal peptide expression plasmids. We obtained gene fragments of the 0030, SP4, and SP14 signal peptide sequences through PCR amplification. The fragments were analyzed by electrophoresis and matched the theoretical size. Additionally, for the backbone plasmid, we obtained the linearized pPIC9K backbone plasmid DNA fragment through reverse PCR with primers.

Fig. 5 Gel electrophoresis of agarose nucleic acid for identifying signal peptide sequence fragments and plasmid backbone fragments

The electrophoresis results show that the gene fragment sequences of the signal peptides are generally small, around 250 bp, and the band positions match the theoretical sizes. We then proceeded with gel cutting and DNA recovery and purification. Afterward, we obtained purified gene fragment DNA.

Fig. 6 Transformation of signal peptide plasmids into Escherichia coli competent cells and colony identification

Based on the purified target gene product, we conducted homologous recombination ligation reactions according to the corresponding concentrations. We then performed transformation operations on E. coli DH5α competent cells, plated the cells, and successfully obtained monoclonal cells under cultivation at 37 degrees Celsius. At the same time, we verified the monoclonal cells through colony PCR, obtaining positive clones in all three plasmid-transformed monoclonal strains. Subsequently, Sanger sequencing confirmed the successful construction of the plasmid DNA, as shown in the results of Figure 7.

Fig. 7 Sanger sequencing for identifying the plasmid containing the signal peptide and its construction. Figure 7A shows pPIC9K-AOX1-0030-NZ2114, Figure 7B shows pPIC9K-AOX1-SP4-NZ2114, and Figure 7C shows pPIC9K-AOX1-SP14-NZ2114. The sequencing signal peaks are stable, and no interference peaks are observed.

Prior to yeast cell transformation, SalI was used to linearize the constructed plasmid in order to improve transformation efficiency. The digested fragments were purified and subsequently employed as the linearized DNA for transformation into yeast competent cells.

Fig. 8 Construction and identification of signal peptide engineered yeast strains

As shown in Figure 8, using the successfully constructed plasmid DNA, we transformed chemically competent GS115 cells. Following selection on YPD solid medium supplemented with G418, positive colonies were obtained and verified by colony PCR. This resulted in the successful generation of engineered yeast strains harboring distinct signal peptide expression plasmids.

Building upon engineered yeast strains harboring different promoter-signal peptide combinations, we performed a preliminary screening. First, the strains were scaled up for cultivation, followed by fermentation and collection of culture supernatants. The levels of secreted Plectasin NZ2114 in the supernatants were then analyzed by protein electrophoresis. Given the small molecular weight of the NZ2114 antimicrobial peptide—only 4.4 kDa—Tris-Tricine SDS-PAGE was employed for analysis and identification. This system replaces glycine with tricine as the trailing ion, thereby optimizing the buffer composition and significantly enhancing resolution for low-molecular-weight proteins. By effectively modulating the migration rate of small proteins, Tris-Tricine electrophoresis prevents their rapid migration through the gel matrix, which can otherwise lead to poor resolution. Consequently, it has become the method of choice for the separation and detection of proteins and peptides in the 1–30 kDa range.

Fig. 9 Expression of plectasin NZ2114 driven by different promoter and verification by Tris-Tricine PAGE Electrophoresis

As shown in Figure 9, protein expression driven by different promoters resulted in distinct bands at the theoretical molecular weight of 4.4 kDa for NZ2114. Clear and well-defined bands were observed with minimal background from non-specific proteins, highlighting the efficiency and purity advantages of secretory expression in Pichia pastoris. Comparative analysis among promoter groups revealed that the AOX1 promoter consistently drove high-level and stable expression across multiple clones. The FDH1 and CAT1 promoters achieved protein expression levels comparable to AOX1, whereas the AOX713 promoter exhibited significantly lower expression. Collectively, these results demonstrate that AOX1, FDH1, and CAT1 are markedly more effective than AOX713 in mediating recombinant NZ2114 production.

Fig. 10 Expression of Plectasin NZ2114 driven by different signal peptide and Tris-Tricine PAGE Electrophoresis verification

We performed small-scale cultivation and induction of various Pichia pastoris strains expressing NZ2114 under the control of different signal peptides. Culture supernatants were collected and analyzed by SDS-PAGE to assess protein secretion efficiency. As shown in Figure 10, the NZ2114 band intensity was markedly higher in the ⍺-factor group compared to those with SP4, SP14, or 0030 signal peptides. These results indicate that the ⍺-factor signal peptide is the most effective in mediating the extracellular secretion of NZ2114 in Pichia pastoris.

Fig. 11 The inhibition zone experiment was used to detect the fermentation products of different promoter/signal peptide strain strains.

Following the evaluation of regulatory element effects on NZ2114 expression via protein electrophoresis, we observed that the promoters AOX1, FDH1, and CAT1 exhibited comparable expression efficiencies. In contrast, among the signal peptides tested, α-factor demonstrated markedly superior performance in mediating protein secretion. To further assess the functional impact of these elements, we evaluated the antimicrobial activity of harvested yeast fermentation supernatants using a phenotypic assay. As shown in the Oxford cup diffusion assay (Figure 11), the inhibitory activity correlated well with the expression results: promoter strength followed the order AOX1 ≈ FDH1 ≈ CAT1 > AOX713, while signal peptide efficacy was ranked as α-Factor > SP4 ≈ SP14 > 0030. Based on these findings and considering both the broad applicability of the yeast expression system and the reliability of methanol-induced expression, we selected the AOX1 promoter combined with the α-factor signal peptide as the optimal regulatory pair for high-level production of Plectasin NZ2114 in Pichia pastoris. This combination was subsequently employed in large-scale fermentation and functional characterization experiments.

Fig. 12 Determination of the inhibitory growth curve of the culture supernatant of engineered yeast strains: 2h, 4h, 6h, 24h, 30h

For the selected promoter and signal peptide combination, we performed 120-hour fed-batch fermentation of the engineered strain GS115-pPIC9K-AOX1-α Factor-NZ2114 and the control strain GS115-pPIC9K carrying the empty vector. Methanol was supplemented daily at 1% (v/v) to serve as both an inducer for AOX1-driven expression and a carbon source. Upon completion of fermentation, the culture supernatants were collected by centrifugation and stored at -80°C for subsequent use. Fresh cultures of Escherichia coli DH5α and Bacillus subtilis were prepared as indicator strains for antimicrobial activity assays. The antibacterial assay was carried out in a total volume of 2 mL, with inoculation of 1% (v/v) pre-grown seed culture. The yeast supernatant was added at varying proportions: 100% (1000 μL), 50% (500 μL), 25% (250 μL), 10% (100 μL), and 0% (0 μL). OD600 was monitored at regular intervals throughout the incubation period. As shown in Figures 12A and 12B, the E. coli control group (treated with empty vector supernatant) exhibited normal growth, reaching an OD600 of approximately 1. In contrast, the NZ2114-containing supernatant significantly suppressed bacterial growth in a dose-dependent manner: 10% and 25% supplementation resulted in partial inhibition, while 50% and 100% supplementation completely abolished the growth of E. coli DH5α. Similarly, as depicted in Figures 12C and 12D, B. subtilis in the control group grew robustly, whereas all NZ2114 treatment groups—except the 0% (no supernatant) control—completely inhibited bacterial growth, indicating potent and broad-spectrum antimicrobial activity of the expressed peptide.

Fig.13 Luminescence detection of control and NZ2114 treated DH5 alpha and B.subtilis in a 50% ratio of fermentation culture medium.

Based on the antibacterial growth curve analysis, we performed a cell viability assay on bacterial cells treated with 50% fermentation supernatant. Cell viability was assessed using the CellTiter-Glo luminescent assay, which involves cell lysis followed by a chemiluminescent reaction. Luminescence signals were quantified using a microplate reader, and the results are presented in Figure 13. Compared to the control group treated with supernatant from empty vector-carrying GS115 yeast, the supernatant from the NZ2114-expressing strain significantly reduced the viability of both Escherichia coli and Bacillus subtilis at the 50% concentration. Notably, the inhibitory effect on Bacillus subtilis was more pronounced. These findings further confirm that our engineered Pichia pastoris GS115 strain successfully produces functional Plectasin NZ2114, leading to effective suppression of bacterial cell viability and growth.

Given that this study aims to address the clinical challenge of drug-resistant Staphylococcus aureus, Bacillus subtilis was selected as the primary model organism for scanning electron microscopy (SEM) analysis to evaluate the bactericidal efficacy of NZ21114. Both Bacillus subtilis and Staphylococcus aureus are Gram-positive bacteria, yet they differ morphologically—rod-shaped (bacilli) versus spherical (cocci). In contrast, Escherichia coli, a Gram-negative bacterium, exhibits more pronounced differences in cell wall structure and overall biological characteristics compared to Staphylococcus aureus. Therefore, Bacillus subtilis serves as a phylogenetically and structurally relevant surrogate that allows for meaningful assessment of antimicrobial effects while maintaining experimental relevance to the target pathogen.

Fig. 14 SEM electron microscope image of Bacillus subtilis morphology at a 1-micron scale, control group (A, B, C), NZ2114 treatment group (D, E, F)

As shown in Fig. 14, SEM observations at a 1-micron scale were performed across multiple fields of view. In the control group (Fig. 14 A, B, C), Bacillus subtilis cells and spores exhibited smooth and intact surface morphologies. In contrast, NZ2114-treated Bacillus subtilis (Fig. 14 D, E, F) displayed clear morphological alterations, including cell surface wrinkling, collapse, and irregularity. These structural changes are consistent with the known mechanism of antimicrobial peptides, which exert their bactericidal effects by disrupting the bacterial membrane. The observed damage provides direct visual evidence that Plectasin NZ2114 compromises membrane integrity in Bacillus subtilis.

Furthermore, high-resolution SEM imaging at a 200-nanometer scale (Fig. 15) revealed more pronounced ultrastructural damage in the treatment group. Compared to the uniformly smooth and intact surfaces of control bacteria, NZ2114-treated cells not only exhibited severe wrinkling but also developed crack-like fissures on their surfaces. These findings further substantiate the membrane-disrupting mode of action of NZ2114 and confirm its potent bactericidal effect against Bacillus subtilis.

Fig. 15 SEM electron microscope image of Bacillus subtilis morphology at a 200-nanometer scale, control group (A, B), NZ2114 treatment group (D, E)