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. 2026 Aug 26. Online ahead of print. doi: 10.1039/d6sc05697k

A portable pathway specific transcription factor-promoter circuit enables heterologous production of fungal nonribosomal peptides

Peng-Lin Wei a,b,†, Yuanyuan Li a,c,†, Zedong Ren a, Shengquan Zhang a,c, Jinwei Ren a, Jie Fan a,b,✉, Wen-Bing Yin a,c,✉
PMCID: PMC13532921  PMID: 42683484

Abstract

Most fungal nonribosomal peptide synthetase (NRPS) biosynthetic gene clusters remain transcriptionally silent, severely restricting the discovery of bioactive peptide metabolites. Standard constitutive promoters fail to support robust expression of large multidomain NRPS enzymes. Here, we identify a conserved subgroup of Zn(ii)2Cys6 transcription factors (TFs) encoded within fungal NRPS clusters and functionally characterize TdaN, which triggers transcription from its cognate promoter PtdaA by recognizing paired CGG-type cis-regulatory motifs. The TdaN–PtdaA pair is repurposed as a portable transcriptional circuit capable of activating five out of seven heterologous NRPS/NRPS-like genes in Trichoderma hypoxylon, notably the recalcitrant silent synthetase Afpes1 that previously demanded screening over 90 distinct promoters for trace product yields. Circuit-driven overexpression of ftmA yields abundant cyclo-l-Trp-l-Pro (cWP) and seven structurally diversified cWP derivatives. This study reveals a conserved regulatory paradigm for fungal nonribosomal peptide biosynthesis and delivers a versatile transcriptional module to expand cryptic peptide chemical diversity.

A native fungal PSTF-promoter circuit is repurposed as a portable transcriptional tool box to activate silent NRPS pathways, enabling heterologous production of diverse nonribosomal peptides and expanding fungal natural product diversity.graphic file with name d6sc05697k-ga.webp

Introduction

Fungal nonribosomal peptide synthetases (NRPSs) assemble structurally diverse peptide natural products (NPs) with important biological and pharmaceutical activities, including antimicrobial, cytotoxic, immunosuppressive and ecological functions.1 Genome sequencing has revealed a large number of fungal NRPS biosynthetic gene clusters (BGCs), yet only a few of their products have been characterized. A major obstacle is that many fungal NRPS genes are transcriptionally silent or expressed at very low levels under standard laboratory conditions.2 In addition, fungal NRPS genes are often large, multidomain and sensitive to the transcriptional and metabolic context of the host.3–5

Heterologous expression has become a powerful strategy for linking fungal BGCs to their products and for exploring biosynthetic logics.6,7 Refactoring fungal BGCs with strong constitutive or inducible promoters has enabled the discovery of many cryptic metabolites. For example, quantitative assessment of more than 90 promoters enabled the identification of strong promoter candidates, which were subsequently used to activate the silent NRPS gene Afpes1 from Aspergillus fumigatus, leading to the discovery of the cyclic tetrapeptides fumiganins A and B.8 However, strong promoters do not always guarantee efficient expression of large fungal NRPS genes. Successful activation may also depend on promoter architecture, host compatibility, chromatin environment, precursor supply and downstream tailoring capacity.9 These challenges highlight the need for more pathway oriented regulatory tools that can drive fungal NRPS expression.10,11

Native fungal BGCs offer a potential solution because they contain not only biosynthetic genes but also dedicated regulatory elements that coordinate pathway expression.12 Many fungal BGCs encode pathway specific transcription factors (PSTFs), which regulate cluster genes by recognizing cognate regulatory motifs within target promoters.4,13,14 Overexpression of PSTFs has been widely used to activate silent BGCs or enhance metabolite production.15,16 Nevertheless, PSTF overexpression alone is not always sufficient or predictable because transcriptional activation depends on both TF and its responsive promoter. Thus, a native PSTF and its cognate promoter should be considered as a coupled regulatory unit rather than as two independent genetic parts.

Compared with direct expression from a conventional strong promoter, such a PSTF-promoter circuit preserves native transcriptional recognition while allowing flexible reconstruction of biosynthetic pathways. For example, coexpression of carotenoid biosynthetic genes under the control of stc promoters (PstcA, PstcB and PstcM) together with the PSTF aflR/S under the control of nitrate inducible promoters enabled carotenoid production in a heterologous host.17 Quantitative characterization of two PSTF–promoter regulatory circuits from fumagillin and sterigmatocystin BGCs using a microfluidic platform enabled the predictable discovery of novel NPs.18 However, whether such circuits can be used as practical modules to drive large fungal NRPS and NRPS-like genes, particularly those that are silent or difficult to express, remains insufficiently explored.

In this study, we surveyed fungal NRPS-associated BGCs and identified a conserved subgroup of single-domain Zn(ii)2Cys6 PSTFs. Using TdaN and its responsive promoter PtdaA as a representative pair, we established a portable PSTF-promoter circuit for heterologous nonribosomal peptide (NRP) production. By placing fungal NRPS or NRPS-like genes downstream PtdaA and supplying TdaN as the cognate activator, this circuit enabled the production of diverse metabolites in Trichoderma hypoxylon, including cyclodipeptides, a cyclic tetrapeptide, NRPS-like enzyme-derived products and structurally diversified cyclo-l-Trp-l-Pro (cWP) derivatives. These findings demonstrate that fungal PSTF-promoter circuits can be repurposed as compact transcriptional modules to enhance heterologous NRPS expression and expand fungal peptide chemical diversity.

Results and discussion

Survey of fungal BGCs identifies a conserved single-domain Zn(ii)2Cys6 TF subgroup

To investigate TF architectures associated with fungal secondary metabolism, we systematically analysed 537 fungal BGCs, including 536 documented BGCs retrieved from the MIBiG database and the tda cluster identified in our previous study.19,28 Among these clusters, 201 contain putative TFs, and 58 of them are associated with NRPS encoding genes. Notably, 45 of the 58 NRPS BGCs encode a total number of 48 Zn(ii)2Cys6 TFs, indicating that this TF family is the dominant regulatory class governing fungal NRP biosynthetic pathways (Fig. 1A). Although our study focuses on NRPS-associated regulatory systems, Zn(ii)2Cys6 TFs are also widely distributed among other major fungal secondary metabolite (SM) classes. 137 out of 146 PKS-associated BGCs and 15 out of 25 terpene cyclase-associated BGCs encode Zn(ii)2Cys6 TFs (Table S1).

Fig. 1. TdaN represents a PF00172 single-domain Zn(ii)2Cys6 TF associated with fungal NRPS BGCs. (A) PSTF prediction of tda BGC and 536 fungal BGCs from the MIBiG database.19 201 of the 537 fungal BGCs contain putative TFs, including 58 NRPS BGCs. Of these NRPS BGCs, 45 encode Zn(ii)2Cys6 TFs. (B) Pfam-based classification of the 48 Zn(ii)2Cys6 TFs encoded by 45 NRPS BGCs. (C) Comparison of domain architectures of Zn(ii)2Cys6 TFs associated with fungal NRPS BGCs and the known TFs. Gal4, Pho7 and Leu3 are canonical Zn(ii)2Cys6 TFs reported in previous studies.20–22 Fungal Zn(ii)2Cys6 binuclear cluster domain PF00172, fungal specific TF domain PF04082, CcsR C-terminal domain PF27599 and leucine zipper domain IPR046347 are shown in different colors.23–26 (D) Structural model of TdaN predicted by AlphaFold3.27 The Zn(ii)2Cys6 DBD spanning residues 14–54 is shown in orange and forms a compact zinc cluster DNA-binding module. The disordered region spanning residues 58–112 is shown in light purple. Cys21, Cys24, Cys31, Cys38, Cys41 and Cys48 are predicted to coordinate two zinc ions, conforming to the canonical Cys-X2-Cys-X6-Cys-X6-Cys-X2-Cys-X6-Cys motif. (E) Coiled-coil prediction of TdaN and the representative fungal Zn(ii)2Cys6 TFs using CoCoNat.

Fig. 1

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We next analysed the domain architectures of Zn(ii)2Cys6 TFs. All 48 PSTFs from fungal NRPS BGCs share the Zn(ii)2Cys6 binuclear cluster domain PF00172. Based on additional domain compositions, they were classified into three groups (Fig. 1B and Table S2). Group 1 comprises 18 PSTFs containing only the PF00172 domain, such as TdaN from pretrichodermamide BGC, GliZ from gliotoxin BGC, CBF73428 from aspercryptin BGC and ACI30651 from beauvericin BGC (Fig. 1C).28–31 Group 2 comprises 15 PSTFs containing both PF00172 and fungal specific TF domain PF04082, such as NotL from notoamide A BGC, KKP04592 from harzianopyridone BGC and EPS34233 from atpenin B BGC.32,33 Group 3 comprises 15 PSTFs containing PF00172 together with a CcsR C-terminal domain PF27599, such as EAU38972 from dihydroisoflavipucin BGC, ImqK from imizoquin A BGC and OJJ98493 from acurin A BGC.34–36 This classification indicates that the single-domain PF00172 represents a prevalent and conserved feature among Zn(ii)2Cys6 regulators in fungal NRPS BGCs.

We previously have identified the pretrichodermamide biosynthetic pathway in T. hypoxylon, which is encoded by a 22-gene BGC.37,38 This cluster contains 21 biosynthetic genes and the putative PSTF TdaN. Considering that TdaN belongs to the PF00172-only subgroup and is encoded in an experimentally characterized NRPS BGC, it was then selected as a representative for structural analysis. The AlphaFold3-predicted model suggests that TdaN contains a conserved N-terminal Zn(ii)2Cys6 DNA-binding domain (DBD) spanning residues 14–54, forming a compact zinc-cluster fold. Six conserved cysteines, including Cys21, Cys24, Cys31, Cys38, Cys41 and Cys48, coordinate two zinc ions, consistent with the canonical Cys-X2-Cys-X6-Cys-X6-Cys-X2-Cys-X6-Cys pattern (Fig. 1D). In contrast to the well-characterized Zn(ii)2Cys6 TFs, such as Gal4, Ppr1, Put3, Hap1 and Leu3, TdaN is predicted to lack a clearly defined coiled-coil or leucine zipper-like dimerization region adjacent to the DBD (Fig. 1E).20,22,24,39,40 An intrinsically disordered region was shown spanning residues 58–112, which is enriched in disorder-promoting residues (K/R, S/G/P) and not predicted to form a stable secondary structure (Fig. 1D). These features suggest that group 1 Zn(ii)2Cys6 TF, exemplified by TdaN, represents a minimal architecture characterized by a compact zinc-cluster DBD coupled with a flexible disordered region that may contribute to regulatory specificity.

TdaN acts as a pathway activator of the pretrichodermamide BGC

To investigate the regulatory role of TdaN in the native tda BGC, we deleted tdaN in T. hypoxylon from the Δtri5 background, which lacks the dominant trichothecene metabolites (Fig. 2A).28,41,42 LC-MS analysis of ΔtdaNΔtri5 mutant showed the complete abolishment of pretrichodermamides, including pretrichodermamide (1), gliovirin (2) and the desulfurized derivative trichodermamide A (3) (Fig. 2B). To determine whether metabolite loss was associated with transcriptional regulation, we then examined tda gene expression by semiquantitative PCR and transcriptomic analysis. Most tda biosynthetic transcripts were readily detected in Δtri5 but were strongly reduced or undetectable in the ΔtdaNΔtri5 mutant (Fig. 2C and D). These genes include tdaA, which encodes the NRPS responsible for initial cyclodipeptide formation, as well as multiple tailoring enzymes such as cytochrome P450 monooxygenases TdaB, TdaG, TdaI, TdaQ and TdaS.37 These results indicate that TdaN is required for the biosynthesis of pretrichodermamides and functions as a pathway-level positive regulator.

Fig. 2. TdaN specifically regulates the biosynthesis of pretrichodermamides 1–3. (A) Schematic representation of tda gene cluster containing the PSTF tdaN and 21 biosynthetic genes. (B) LC-MS analysis of extracts from T. hypoxylon strains. Δtri5 and ΔtdaNΔtri5 were compared to evaluate the role of TdaN in tda cluster expression. To construct the strain carrying PgpdA::tdaA, tdaA was reintroduced via integration at the native tda locus under the control of the constitutive promoter PgpdA in the ΔtdaA–VΔtri5 background. The strain carrying PgpdA::tdaN::PtdaA::tdaA retained native PtdaA-driven expression of tdaA together with PgpdA-driven expression of tdaN in the ΔtdaA–VΔtri5 background. Extracted ion chromatograms (EICs) refer to [M + H]+ or [M + Na]+ of 1–4 with a tolerance range of ± 0.25. (C and D) Expression analyses of tda genes in Δtri5 and ΔtdaNΔtri5 by semiquantitative PCR (C) and transcriptomic analysis (D), respectively.

Fig. 2

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To further validate the functional relationship between TdaN and the tda promoter, we constructed a synthetic expression system TYLYY4 (ΔtdaA–VΔtri5), in which the entire 49.6-kb tda cluster and tri5 has been deleted to eliminate production of 1–3 and trichothecene, as well as the related intermediates.37,41 In this system, tdaA was reintroduced at the native tda locus under the control of the constitutive promoter PgpdA (PgpdA::tdaA) (Fig. 2B). However, no detectable product was observed. In contrast, coexpression of tdaN under the control of PgpdA together with tdaA driven by its native promoter PtdaA (PgpdA::tdaN::PtdaA::tdaA) resulted in the accumulation of the TdaA-derived cyclodipeptide cyclo-l-Phe-l-Phe (cFF, 4). These results defined TdaN and tda promoters, represented by PtdaA, as a functional PSTF-promoter circuit.

To explore the molecular basis of TdaN-dependent transcriptional activation, we analysed promoter regions of TdaN-responsive genes and identified a conserved cis-regulatory motif, 5′-(A/C/T)CGGNNNCCGA-3′ (Fig. 3A and S3). This motif contains paired CGG-like half-sites, a common feature recognized by fungal Zn(ii)2Cys6 TF, suggesting that it may serve as the core TdaN-binding element. To test the functional contribution of these half-sites, three PtdaA variants were constructed in which the CGG half-site, the CCGA half-site, or both were replaced with TTT/TTTT substitutions while maintaining native spacing (Fig. 3B). Structural modelling of TdaN bound to the wild-type promoter fragment predicted that the Zn(ii)2Cys6 DBD domain contacts the paired motif region. His25 and Lys28 are located near the CGG half-site and its flanking nucleotides, whereas Arg18, His25, Ser27 and Lys28 are positioned in proximity to the CCGA half-site and adjacent bases, suggesting that these residues may contribute to sequence recognition and local DNA stabilization (Fig. S4). In contrast, mutations in either half-site disrupted the predicted protein–DNA interaction network, consistent with reduced binding affinity in silico (Fig. 3C). Together, these results support a promoter-centred recognition mechanism in which TdaN specifically engages paired CGG-like elements within tda promoters to activate transcription of the corresponding genes. Therefore, we consider the TdaN–PtdaA regulatory pair as a portable expression circuit for heterologous gene expression.

Fig. 3. Identification of TdaN–PtdaA regulatory circuit. (A) Motif analysis identified a conserved 5′-(A/C/T)CGGNNNCCGA-3′ candidate cis-regulatory motif in TdaN-regulated promoters. (B) Three PtdaA variants were designed by replacing the CGG half-site, the CCGA half-site or both half-sites with TTT/TTTT sequences. (C) AlphaFold3-predicted models of TdaN in complex with the wild-type or PtdaA variants are shown. Mutated nucleotides are highlighted in green.

Fig. 3

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The TdaN–PtdaA circuit enables heterologous production of NRPS-derived metabolites

To test the utility of TdaN–PtdaA regulatory circuit, we constructed a TdaN–PtdaA-based NRPS expression cassette for individual fungal NRPS or NRPS-like genes (Fig. 4A). In this cassette, tdaN was expressed under the control of the constitutive promoter PgpdA, whereas the selected heterologous NRPS gene was placed downstream the native promoter PtdaA. In this configuration, TdaN serves as the transcriptional activator and PtdaA functions as the output promoter to drive expression of the inserted NRPS gene. Then, the cassette, together with the AfpyrG selectable marker, was introduced by homologous recombination into the original tda locus of the engineered T. hypoxylon host TYLYY4.37 Using this system, we successfully constructed seven mutants, respectively carrying ataP from Aspergillus terreus, Afpes1, gliP, ftmA and hasD from A. fumigatus, as well as micA and asqK from A. nidulans (Fig. S2 and Table S3).

Fig. 4. Heterologous production of diverse NRPS-derived metabolites in T. hypoxylon using the TdaN–PtdaA regulatory circuit. (A) Design and construction of the TdaN–PtdaA::nrps activation cassette. TdaN was expressed under the control of the constitutive promoter PgpdA, and heterologous NRPS genes were placed downstream the native promoter PtdaA. (B) LC-MS analysis of engineered strains expressing NRPS genes, including ataP, Afpes1, micA, gliP and ftmA. The accumulation of 5–8 and 8′ were identified by comparison with previously characterized compounds from our laboratory. EICs refer to [M + H]+ or [M + Na]+ of 5–8 with a tolerance range of ± 0.25. (C) Structures of 5–8 and 8′.

Fig. 4

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LC-MS analysis showed that five of the seven NRPS expression strains produced detectable NRPS-derived metabolites (Fig. 4B). Product assignments were supported by LC-MS analysis and the comparison with authentic standards where available. Three of the tested genes, ataP, gliP and ftmA, have previously been shown to catalyse cyclodipeptide formation. Consistent with their assigned functions, expression of ataP, a tdaA homolog from the acetylaranotin BGC of A. terreus, led to the production of 4.43 Expression of gliP from the gliotoxin BGC produced cyclo-l-Phe-l-Ser (5).44–51 Expression of ftmA from the fumitremorgin BGC generated cyclo-l-Trp-l-Pro (cWP, 6), also known as brevianamide F.52

Afpes1 is a silent NRPS gene from the human pathogen A. fumigatus and encodes a multimodular synthetase with an A–T–E–C–A–C–A–C–A–T–E–C–T–C–T domain architecture. In our previous study, activation of Afpes1 required screening more than 90 promoters, of which only two enabled detectable product formation.8 In the present system, we achieved the activation of Afpes1 using the TdaN–PtdaA circuit, yielding the cyclic tetrapeptide fumiganin A (7). Similarly, micA, an NRPS-like gene from A. nidulans encoding an A–T–TE domain architecture, has previously been activated using the inducible alcohol dehydrogenase promoter PalcA or CRISPR-mediated transcriptional activation.53,54 Here, the micA expression strain produced two microperfuranone epimers, 8 and 8′. In contrast, no ions corresponding to the calculated masses of the predicted metabolites, nor any additional differential metabolites associated with hasD or asqK expression, were detected in the engineered strains carrying hasD or asqK (Fig. S5). This lack of detectable products may reflect differences in gene expression, enzyme folding, substrate availability or the requirement for additional pathway components. Overall, these results indicate that the TdaN–PtdaA circuit can provide an efficient toolbox for expressing selected silent or poorly accessible fungal NRPS and NRPS-like genes and discovering the derived peptide products.

NRPS replacement using the circuit generates diketopiperazine derivatives

In the native pretrichodermamide A pathway, TdaA catalyses the formation of 4 as the initial diketopiperazine (DKP) scaffold, which is subsequently diversified by tailoring enzymes encoded in the tda cluster.37,38 To redirect this pathway toward new DKP derivatives, we replaced the native tdaA with ftmA due to the high yield of 6 in ftmA expression strain with the TdaN–PtdaA regulatory circuit. Briefly, tdaA was first deleted from T. hypoxylon wild-type strain using the hygromycin resistance marker.28 To improve heterologous recombination efficiency, the non-homologous end-joining gene thlig4 was replaced with a cas9 expression cassette. Using this CRISPR/Cas9-enabled system, ftmA was integrated downstream PtdaA while the endogenous tda tailoring genes were retained (Fig. 5A).

Fig. 5. Heterologous expression of ftmA in ΔtdaA mutant generates cWP derivatives. (A) Design and construction of the ftmA expression strain in ΔtdaA background. (B) LC-MS analysis of crude extracts from different T. hypoxylon strains, including TYZD3 (ΔtdaA), TYZD6 (ΔtdaA::ftmA) and TYZD10 (OE::tdaN, ΔtdaA::ftmA). (C) Structures of 9–13, 9′ and 13′. EICs refer to [M + H]+ or [M + Na]+ of 9–13 with a tolerance range of ± 0.25.

Fig. 5

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LC-MS analysis showed that the ΔtdaA::ftmA mutant accumulated compound 6 as the major product, together with several minor product peaks (Fig. 5B). To further enhance production of the new metabolites, we introduced a tdaN overexpression cassette into the ΔtdaA::ftmA mutant. Compared with the ΔtdaA::ftmA strain, the resulting strain exhibited a 13.6% increase in the production of compound 6 based on HPLC peak area analysis. More importantly, overexpression of tdaN increased the accumulation of low-abundance metabolites that were barely detectable in the parental strain, enabling their isolation and structural elucidation. This improved production profile facilitated large-scale cultivation, chromatographic purification, and structural elucidation of compounds 9–13, 9′, and 13′ (Fig. S6–S19 and Tables S4–S19).

Compounds 9 and 9′ were individually purified and characterized by NMR analyses, which established their identical planar structures as the known pyrroloindoline-containing DKP bearing a C3-hydroxy group (Fig. S6, S7 and Tables S5, S6).55 Referring to the reported biosynthesis of brevianamide derivatives, we proposed that H-11 and H-14 were cofacial.56 No diagnostic correlation between H-2 and 3-OH was observed. The two compounds also exhibited nearly identical experimental ECD spectra. Comparison with calculated ECD spectra for the stereochemical candidates narrowed the possible assignments to 2S,3S,11S,14S and 2S,3R,11S,14S configurations (Fig. S13 and Tables S12–S15). The two epimers were subsequently differentiated by DP4+ analyses based on calculated and experimental NMR chemical shifts. Taken together, the ROESY, ECD, and DP4+ results support the assignments of (2S,3S,11S,14S)-9 and (2S,3R,11S,14S)-9′ respectively (Fig. 5C and S14–S17).

Compound 10, which lacks the pyrroloindoline ring system and instead contains a 3-hydroxyl-2-indolone moiety, was identified as the previously reported compound, asperochramide D (Figs. S8 and S18, Table S7).57,58 The formation of 9, 9′ and 10 could be rationalized by divergent transformations involving a putative 2,3-epoxide intermediate, analogous to the reported biosynthesis of brevianamides.56 Although a homolog of BvnB, the enzyme responsible for the corresponding epoxidation in brevianamide biosynthesis, was not identified in T. hypoxylon, other oxygenases may catalyze this transformation. For example, the cytochrome P450 oxygenase TdaB involved in pretrichodermamide biosynthesis catalyses an aromatic epoxidation reaction, suggesting that TdaB may potentially participate in the formation of the proposed 2,3-epoxide intermediate.37 Intramolecular cyclization followed by epoxide ring opening would generate the pyrroloindoline scaffold of 9/9′, whereas alternative hydrolytic opening of the proposed epoxide intermediate followed by oxidation would afford the 3-hydroxyl-2-indolone scaffold of 10 (Fig. S20A).

Compound 11 lacks an intact indole and instead contains an anthraniloyl-type aromatic ketone, matching the previously reported compound, gallaecimonamide A (Fig. 5C and S9, Table S8).59 Compared with 11, compound 12 exhibited an [M + Na]+ ion at m/z 338.1111, corresponding to a mass increase of 28 Da. This difference is consistent with the presence of an additional formyl group. Accordingly, the NMR data of 12 revealed a characteristic N-formyl signal (δH 8.49 and δC 161.2), supported by the diagnostic HMBC correlation from the formyl proton (δH 8.49) to C13 (δC 137.9) (Fig. S10 and Table S9). Thus, compound 12 was identified as an N-formylkynurenine-derived proline DKP, representing the N-formylated analogue of 11. Based on the structure features, compounds 11 and 12 may arise from oxidative cleavage of the indole ring in 6 by the indole-2,3-dioxygenase-like enzyme T_hypo_3384.60 Alternatively, direct activation and condensation of l-kynurenine or N-formyl-l-kynurenine with l-Pro by a promiscuous NRPS module cannot be excluded (Fig. S20B).

Compounds 13 and 13' were obtained as yellow oils and assigned the same molecular formula C27H28N4O4 based on HR-ESI-MS analysis, which showed [M + Na]+ ions at m/z 495.2002 and 495.2004, respectively (calcd 495.2003). The 1H and 13C NMR spectra of both compounds revealed characteristic signals for the cWP core and a second indole unit (Fig. 5C and S11–S12, Tables S10–S11). As for 13, this assignment was supported by the downfield 24-NH signal (δH 10.92) and its HMBC correlations to C21 (δC 114.9), C22 (δC 126.6), C23 (δC 135.8) and C25 (δC 122.9) (Fig. S11 and Table S10). The remaining oxygenated aliphatic fragment was assigned as C20–C30–C31 based on the correlations of H30 (δH 4.26–4.34) with H20 (δH 4.71) and H31 (δH 3.20–3.50). Key HMBC correlations from H20 to C2, C3, C21, C22, C25, C30 and C31, together with correlations from 30-OH to C21, C30 and C31 and from H31 to C20 and C30, established the connection of this diol-containing side chain to both indole units. Accordingly, compounds 13 and 13′ were identified as a pair of stereoisomers with an identical planar structure, corresponding to a bisindole-containing cWP derivative bearing an oxygenated C20–C30–C31 side chain. For 13′, a clear ROESY correlation between H-11 and H-14 supported a cofacial relationship between these protons. Because 13 contained minor impurities, reliable ROESY data could not be obtained. The relative configuration of its diketopiperazine core was therefore proposed due to the similar biosynthesis to 13′. Comparison of the experimental ECD spectra of 13 and 13′ with the calculated spectra of the relevant stereoisomeric candidates supported the assignments of (11S,14S,20S,30S)-13 and (11S,14S,20S,30R)-13′, respectively (Fig. 5C and S19).

The formation of 13 and 13′ may involve a three-component condensation of cWP, indole and glyceraldehyde.61 In the proposed pathway, glyceraldehyde would be first activated by nonenzymatic catalysis or an unidentified enzyme-assisted process, allowing nucleophilic attack of free indole at its C3 position to form an indolyl carbinol intermediate. Subsequent proton transfer and dehydration would generate an indoleninium-like electrophile, which could be captured by the C2 position of the DKP-derived indole in compound 6. Final deprotonation and rearomatization would afford the bisindole DKP scaffold of 13 (Fig. S20C). The enzymes responsible for this proposed transformation remain unknown and will require future biochemical investigation.

Collectively, by placing ftmA under the control of the native PtdaA promoter and further enhancing pathway activation through tdaN overexpression, the engineered strain produced high levels of cWP and accumulated a series of cWP derivatives. The resulting compounds revealed multiple diversification routes, including the putative indole epoxidation, oxidative indole-ring cleavage and bisindole–aldehyde condensation. These findings highlight the utility of the TdaN–PtdaA circuit for boosting heterologous NRP production and expanding structural diversity. However, the broader applicability of the TdaN–PtdaA circuit will likely depend on the compatibility between the regulatory components and heterologous hosts. Successful implementation in other fungal hosts, such as Aspergillus nidulans or A. oryzae, cannot be assumed solely by introducing the PSTF and promoter elements. Additional factors, including higher-order regulatory mechanisms, chromatin accessibility and host-specific transcriptional networks, may influence circuit performance. Thus, the TdaN–PtdaA circuit provides a promising framework for fungal pathway activation, while future studies are required to systematically evaluate its portability across diverse fungal hosts.

Conclusions

In this study, we identified a conserved group of single-domain Zn(ii)2Cys6 TFs associated with fungal NRPS BGCs and established TdaN as a representative pathway specific activator. Genetic deletion, transcriptional analysis and promoter reconstruction demonstrated that TdaN positively regulates the native pretrichodermamide BGC through the responsive promoters, represented by PtdaA. Motif analysis and structural modelling further suggest that TdaN recognizes paired CGG-like cis-elements within target promoters, defining a functional PSTF–promoter regulatory circuit.

By repurposing this native TdaN–PtdaA circuit, we developed a transcriptional activation module for heterologous expression of fungal NRPS and NRPS-like genes in T. hypoxylon. This system enabled the production of diverse NRPS-derived metabolites and provided an alternative strategy for activating heterologous biosynthetic genes. Further application of the circuit to enhance ftmA expression promoted production of cWP and enabled the discovery of multiple cWP derivatives generated through indole epoxidation, oxidative indole-ring cleavage and bisindole–aldehyde condensation.

Overall, our work demonstrates that fungal PSTF–promoter pairs can serve not only as regulatory elements of native biosynthetic pathways but also as portable expression circuits for NP discovery. The TdaN–PtdaA circuit provides a practical strategy for boosting heterologous NRP production and expanding the chemical diversity.

Author contributions

W.-B. Y. and J. F. conceived the study, acquired funding and supervised the research. P.-L. W., Y. L., Z. R and S. Z. performed experiments including mutant construction and NP identification. Z. R. and J. R. identified the structures. P.-L. W. performed computational analyses. P.-L. W. and J. F. drafted the manuscript. All authors read and approved the final manuscript.

Conflicts of interest

There are no conflicts to declare.

Supplementary Material

SC-OLF-D6SC05697K-s001

Acknowledgments

We thank Dr Wenzhao Wang from the Institute of Microbiology, Chinese Academy of Sciences for HR-ESI-MS analysis. This work was partially supported by Beijing Natural Science Foundation-Chaoyang Innovation Joint Fund (L259025 to W.-B. Y.) and Tianjin Natural Science Foundation Project (25JCQNJC01450 to J. F.).

Data availability

The data supporting this article, have been included as part of the supplementary information (SI). Supplementary information: materials and methods, Fig. S1–S20 and Tables S1–S22. See DOI: https://doi.org/10.1039/d6sc05697k.

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Supplementary Materials

SC-OLF-D6SC05697K-s001
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Data Availability Statement

The data supporting this article, have been included as part of the supplementary information (SI). Supplementary information: materials and methods, Fig. S1–S20 and Tables S1–S22. See DOI: https://doi.org/10.1039/d6sc05697k.

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