The transcription factor STE12 influences growth on several carbon sources and production of dehydroacetic acid (DHAA) in Trichoderma reesei – Scientific Reports

The domain structure of STE12 in filamentous fungi is conserved in T. reesei

In order to integrate STE12 in the context of current network knowledge, we searched for known and predicted interactors using the STRING database (Fig. 1A). The protein interaction network of STE12 in T. reesei revealed a connection to the mating related MAPkinase pathway and TMK1, as well as numerous genes involved in chromatin modification (Fig. 1B). Since ste12 homologues were previously reported to be subject to alternative splicing^26,27, we screened available transcriptome data for coverage of the ste12 gene model, which contains two introns. For evaluation of the gene model used in our analysis for ste12, we checked data from growth on cellulose or glucose in constant light or darkness (Fig. 1C). We found that the predicted introns are clearly present and that ste12 has a relatively long 5′ UTR of roughly 700 bp, which comprises an upstream in-frame stop codon at position − 24. In this UTR region, neither an additional intron nor an upstream open reading frame (uORF;²⁸) was detected, which might interfere with efficient ste12-translation. No indications for alternative splicing were detected.

Characteristics of ste12/TR_36543/TrA1391C and its encoded protein. (A) STRING network of known and predicted interaction partners. The network for interactors of STE12 was drawn using the online STRING search function (https://string-db.org/) in version 12⁹¹ (B) Annotations of predicted interaction partners of STE12. (C) Evaluation of the protein model of STE12 by analysis of aligned reads from available transcriptome data for growth on cellulose (CEL) or glucose (GLU) in constant light (LL) or constant darkness (DD).

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The fungal STE homeodomain is highly divergent, however, in this domain also a conserved stretch of KQKVFFWFSVA resides²⁹. Indeed, a related sequence is also present in T. reesei, albeit with three amino acid alterations: KQKVFYWYSVP. Accordingly, T. reesei STE12 comprises a STE like transcription factor domain (pfam02200; p-value 1.17e-78).

STE12 positively influences cellulase gene transcription

Interestingly, ste12 shows mutations in the T. reesei cellulase-mutant strains NG14 and RutC30^30,31, suggesting a potential contribution to the efficient production of cellulose degrading enzymes in these strains. We asked whether STE12 has a function in regulation of cellulase formation upon growth on cellulose. Therefore, we first tested whether biomass formation on this carbon source would be altered. In darkness, Δste12 showed similar growth as the wild-type, while in light biomass formation was significantly increased by 20% (Fig. 2A). However, while specific cellulase activity in darkness was unaltered and hence consistent with growth data, activity in light remained below detection levels (Fig. 2B). Analysis of cbh1 transcript abundance showed a positive effect of STE12 in light, causing a decrease of cbh1 transcript by 60% upon ste12 deletion but no effect in darkness (Fig. 2C, D). A similar effect was detected for the carbon catabolite repressor gene cre1, with a 40% decrease in transcript abundance for Δste12 in light (Fig. 2E, F). For the cellulase transcription factor gene xyr1, no significant regulation by STE12 was found, although in darkness a negative trend of transcript levels was apparent (p-value 0.071) (Fig. 2G, H). The important regulatory gene vel1, which is required for cellulase induction³² and impacts secondary metabolism also in T. reesei^33,34 is not significantly regulated by STE12 (Fig. 2I, J). Pks4, the gene encoding the polyketide synthase responsible for the green pigment in spores of T. reesei³⁵ shows a trend towards increased abundance in the mutant in constant light, albeit the respective p-value (0.092) is below our threshold for significance set at 0.05 (Fig. 2K, L). The same upregulation is observed for gene expression analysis by RNA-sequencing, which resulted in a significant five-fold upregulation of pks4 in constant light for Δste12, confirming the validity of the sequencing results.

Relevance of STE12 for biomass formation, cellulase activity and gene regulation upon growth on 1% cellulose. (A) Biomass formation relative to wild-type (WT). (B) Specific cellulase activity. (C, D) Transcript levels of cbh1 (C) in constant darkness and (D) constant light. (E, F) Transcript levels of cre1 (E) in constant darkness and (F) constant light. (G, H) Transcript levels of xyr1 (G) in constant darkness and (H) constant light. (I, J) Transcript levels of vel1 (I) in constant darkness and (J) constant light. (K, L) Transcript levels of pks4 (K) in constant darkness and (L) constant light. (M) Transcript levels of ste12 in MAPKinase deletion mutants in constant darkness (DD) and light (LL).

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Of the five MAPkinase cascades of S. cerevisiae, two, Fus3 and Kss1 target Ste12¹² to transmit the pheromone signal. Since STE12 is subject to regulation by MAPkinases also in other fungi^16,36,37, although predominantly in terms of phosphorylation and stability, we asked whether in T. reesei also effects on the transcriptional level are present. Our analysis showed that T. reesei ste12 is not subject to transcriptional regulation by MAPkinases upon growth on cellulose (Fig. 2M).

Growth on different carbon sources is altered in Δste12

A more general role of STE12 in regulation of growth and hence metabolism on diverse carbon sources was investigated using the BIOLOG phenotype microarrays³⁸. We monitored growth patterns from 72 to 144 h after inoculation in constant darkness (supplementary file 1). If two consecutive time points showed statistically significant differences (p-value < 0.05, t-test) in biomass formation as analyzed by turbidimetry at 750 nm, we considered STE12 to be relevant for regulation of growth on this carbon source.

Interestingly, the differences we found for Δste12 were all positive in terms of elevated growth of the mutant strain compared to the wild-type strain (Fig. 3A–E). In many cases, these differences occurred at 120 and 144 h after inoculation, when the mutant strain obviously kept growing, whilst the parental strain did not. Better growth on glycerol and glycogen suggests utilization of these carbon sources instead of storage.

Analysis of carbon source utilization using the BIOLOG phenotype microassay. (A) Schematic representation of carbon sources on which Δste12 grows better than the wild-type along with conversion pathways as deduced from KEGG pathways for T. reesei. (B, C) Growth data of the ste12-deletion strain as represented by turbidimetric analysis of biomass accumulation at 750 nm and compared to the wild-type strain QM6a. The analysis was done in biological triplicates with growth in darkness (DD). Statistical significance was determined by the T-test; * = p-value < 0.05, ** = p-value < 0.01.

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Moreover, sugars including lactose, lactulose, melibiose, maltose and melezitose enable longer growth of the mutant strain, as do γ-hydroxy butyric acid, p-hydroxyphenylacetic acid and α-keto-glutaric acid (Fig. 3A–E).

STE12 impacts gene regulation

We investigated the regulatory impact of STE12 on gene regulation upon growth on cellulose as carbon source in constant light and constant darkness. In total, we found 203 genes to be more than twofold significantly (p-adj < 0.05) regulated directly or indirectly by STE12 (Supplementary file 2). Functional category analysis (supplementary file 2) of these genes revealed a significant enrichment (p-value < 0.05) of genes involved in transport facilities, particularly calcium-, iron- and zinc-ion transport, carbohydrate metabolic process as well as secondary metabolism (amine- and proline catabolic process). Gene ontology (GO) analysis supported these results (Fig. 4).

Gene ontology analysis of genes regulated by STE12. GO enrichment of up- and down-regulated genes for Δste12 in constant darkness and light, visualized with rrvgo in R.

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Of the STE12 targets, 84 genes were up-regulated in light, including nine CAZyme encoding genes, for example a candidate chitinase (TrA0008W/TR_59791, 28.8-fold) and a subgroup beta-chitinase (TrE0823C/TR_43873, 8.8-fold), a candidate glycoside hydrolase (TrC0858W/TR_55886) and a beta-xylosidase, bxl1 (TrC1552C/TR_121127). Other upregulated genes include the conidiation specific con-10, TrD0147C/TR_5084 (11.7-fold), the protein kinase gene gin4 (TrD1202W/TR_64125), which positively influences trichodimerol biosynthesis³⁹ a candidate cutinase transcription factor (TrA0431C/TR_106259, 26.3-fold). TR_106259 is also strongly up-regulated in a deletion mutant of the secondary metabolite regulator of the SOR-cluster, YPR2^8,40,41, corroborating an indirect effect of STE12 on secondary metabolism. The same applies for a second strongly up-regulated transcription factor gene, TrF0487C/TR_112643 (12.7-fold in darkness), which is also strongly up-regulated in Δypr2⁴¹. Further up-regulated genes include the gene encoding the glucose transporter HXT1 (TrE0206W/TR_22912), the predicted sugar transporter gene TrD0036W/TR_50894, which was shown not to be required for growth on lactose⁴² along with several other transporter genes as well as two genes encoding proteins predicted to be involved in plant surface sensing⁴³, the effector protein encoding TrA1330W/TR_72907 and the PTH11 type G-protein coupled receptor gene TrG0742C/TR_45573. Among the up-regulated genes in light, four genes belong to the cytochrome p450 superfamily, where TrF0040C/TR_65036 and TrA1084W/TR_75713 are potential homologues of Aspergillus nidulans alkane hydroxylases, catalyzing the oxidation of alkanes. The other two cytochrome p450 encoding genes are TrE0324C/TR_66453 homolog of N. crassa ci-1, an ent-kaurene oxidase, involved in the biosynthesis of gibberellins⁴⁴ and TrA0963W/TR_67377. Additionally, two polyketide synthase genes, pks4 (TrD1440W/TR_82208), responsible for the green pigmentation of T. reesei conidia³⁵ and pks2 (TrD0448W/TR_65891) are up-regulated (5- and 2.8-fold) in light.

The 23 genes up-regulated in darkness comprise the conidiation associated glucose repressible gene grg-1, TrE0533C/TR_73516, a family 5 carbohydrate esterase, the xylanase gene xyn3 (TrF0312W/TR_54219 and TrC0667W/TR_120229), the non-ribosomal peptide synthase (NRPS) encoding tex2 (TrB1256C/TR_123786) responsible for paracelsin biosynthesis. Furthermore, two mitochondrial transporters TrC0706C/TR_103853 and TrF1000W/TR_121743 and a small cysteine-rich protein encoding gene TrC1533/TR_121135 (90.3-fold).

The 86 genes of the gene set down-regulated in light comprises four CAZyme encoding genes including cbh1/cel7a (4.1-fold), egl3/cel12a (23-fold), which is limiting for high efficiency plant cell wall degradation⁴⁵, the beta-glucosidase bgl1/cel3a (45.2-fold) and a GH 99 gene, TrC1527C, TR_121136 (21.9-fold). Additionally, among the down-regulated genes in darkness are the GprK-like RGS domain containing heterotrimeric G-protein coupled receptor gene TrG0214W/TR_81383 and three transcription factor genes (TrA0076W/TR_3605, TrG1015C/TR_120363 and TrD0324W/TR_80139). The 10 down-regulated genes in darkness include a predicted oligonucleotide transporter gene related to sexual differentiation process protein ISP4 (TrA1796W/TR_124002), and a predicted MFS permease (TrB1842C/TR_68990).

Of all STE12 targets, five genes contain mutations in the high cellulase producer RutC30 (TrB1256C/TR_123786, TrG0579W/TR_56726, TrF0040C/TR_65036, TrF0049W/TR_65039 and TrC0660W/TR_120231).

Plant cell wall degradation specific phosphorylation was detected previously⁴⁶ for six STE12-regulated genes including an amino acid transporter (TrB0212C/TR_123718), grg-1 and a putative methyltransferase gene (TrD1044C/TR_108914).

Regulation by STE12 in both light and darkness

Eight genes show light independent regulation by STE12. Up-regulation in both, light and darkness, was observed for a potential amino acid transporter gene (TrB0212C/TR_123718), the polyketide synthase gene pks2 (TrD0448W/TR_65891), a potential carnitine O-acyltransferase encoding gene (TrC0399W/TR_122240) and TrE0645C/TR_54352. The putative exonuclease protein TrA1281W/TR_57424, a siderophore transporter TrG0054C/TR_82017 and TrA1279C/TR_57823 (PRE containing) were down-regulated in light and darkness. One gene, TrD0165W/TR_50793, encoding a putative homologue of QIP, a putative exonuclease protein involved in quelling with contrasting regulation in light and darkness by STE12 was found.

STE12 influences genes involved in iron homeostasis

Interestingly, several genes involved in iron homeostasis are targeted by STE12: The genes encoding the multicopper peroxidase Fet3b (TrD0040C/TR_5119) and the high affinity iron permease Ftr1b (TrD0041W/TR_80639), both belonging to the reductive iron uptake system⁴⁷, are up-regulated in light in Δste12. Moreover, a gene encoding a predicted, Fet5 related ferroxidase (TrD1438C/TR_124079) as well as a predicted siderophore transporter (TrD0541W/TR_67026) are upregulated in light. In contrast another siderophore transporter gene (TrG0054C/TR_82017) is downregulated in darkness. Additionally, a predicted iron transporter (TrD0323C/TR_38812) is 11-fold down-regulated in light. These findings suggest a contribution of STE12 to light modulated regulation of iron homeostasis.

Presence of the pheromone response element (PRE) in STE12 target promotors

The target sequence motif of STE12 was determined in S. cerevisiae and is called pheromone response element (PRE): 5′ (A)TGAAACA 3′^29,48. Multimerization of S. cerevisiae Ste12 appears to enhance binding to pheromone response elements (PREs) and several adjacent PREs occur in strongly pheromone induced genes^49,50, although a clear correlation was not found and pheromone responsive genes without PREs also exist^51,52.

This sequence is also essential for Ste12 binding in C. neoformans⁵³ and in Colletotrichum lindemuthianum²⁷. Screening the genes regulated by STE12 in T. reesei on cellulose, we found PREs in the promotors of five target genes (TrE0645C/TR_54352, TrA1206C/TR_104816, TrF0872C/TR_107349, TrA0569C/TR_108586 and TrA0485W /TR_121285). The reverse sequence 5’ TGTTTCA 3’ was present in 14 of the T. reesei STE12 target genes (supplementary file 2) CAZyme encoding genes, grg-1 and a putative amino acid transporter. However, in none of these promotors we found more than one motif or a combination of forward and reverse motifs.

STE12 regulates production of dehydroacetic acid and trichodimerol

Functional category analysis of genes regulated by STE12 upon growth on cellulose revealed a significant enrichment of genes associated with secondary metabolism among its targets. Moreover, regulation of development is among the primary functions of STE12 in fungi^13,37,54, which is accompanied with clear alterations in secreted metabolites in T. reesei³³. Consequently, we asked whether STE12 is required for proper chemical communication under conditions favoring sexual development.

Bisorbibutenolide, which was recently shown to be produced by T. reesei and dependent on the presence of the MAPkinase TMK3²⁴, is not regulated by STE12 (Fig. 5A, highlighted in orange (D)). However, STE12 is involved in regulation of dehydroacetic acid (highlighted in green (B, C)) and also trichodimerol (highlighted in yellow (E)) in Fig. 5A.

HPLC analysis of secondary metabolite production and identification of dehydroacetic acid. (A) HPLC–DAD chromatograms of QM6a and Δste12 at 230 nm. QM6a profile is shown in grey for better comparison. Three biological replicates are shown. Strongly regulated peaks are indicated by asterisks. Dehydroacetic acid (B,C) is highlighted in green, (21S)-bisorbibutenolide (D) in orange and trichodimerol (E) in yellow²⁴. (B) UV-spectrum and (C) chemical structure of dehydroacetic acid.

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Preparative column chromatography fractions obtained from T. reesei crude extracts were subjected to NMR and MS analysis and resulted in the identification of dehydroacetic acid (Fig. 5B, C). It was identified in a mixture together with the steroid ergosterol (sample A), in a further purified sample (B) and finally by comparison to a commercially available standard.

The NMR spectroscopic analysis of sample A revealed a content of approx. 90% (mol/mol) ergosterol (Fig. S1 in supplementary file 3). These NMR spectroscopic data of ergosterol are in agreement with those of a commercial reference sample as well as with previously published data of ergosterol⁵⁵. In addition, approximately 7% (mol/mol) of the target compound could be identified from the mixture in sample A. Further purification of this smaller amount in sample A by prep TLC using silica gel 60 glass plates (Merck) yielded 0.6 mg of the target compound (sample B). It was identified as dehydroacetic acid (3-acetyl-6-methyl-3,4-dihydro-2H-pyran-2,4-dione, DHAA).

HR-ESI-TOF–MS in negative ionization of sample A (Fig. S2 in supplementary file 3) shows a deprotonated molecular ion [M-H]^– of m/z 167.0343, which correlates quite well with the calculated [M-H]^– of m/z 167.0350 of the molecular formula C₈H₈O₄. The HR-ESI-TOF–MS of sample B (Fig. S3, S4 in supplementary file 3) shows a deprotonated molecular ion [M-H]⁻ of m/z 167.0349 in the negative ionization mode as well as a [M + Na]⁺ of m/z 191.0309 and a [M + H]⁺ of m/z 169.0489 in positive ionization mode. The isotopic patterns in these spectra of sample B show a weak entry of deuterium into the molecule, because it was previously dissolved in CD₃OD. However, all recorded monoisotopic masses fit well with calculated [M-H]^– of m/z 167.0350, [M + Na]⁺ of m/z 191.0315 and a [M + H]⁺ of m/z 169.0495 of the molecular formula C₈H₈O₄. Further co-chromatographic comparison using commercially available dehydroacetic acid (Thermo Scientific, Waltham, MA; CAS Nr. 520-45-6) as standard confirmed the identity of this compound in sample B (Fig. 5).

1D and 2D NMR measurements of the 7% (mol/mol) dehydroacetic acid in sample A further confirmed the structure of the target compound (Fig. S5 in supplementary file 3). The spectra led to a total number of two methyl-, zero methylen-, one methine groups and five quaternary carbon atoms, resulting in one additional non carbon bound proton. The ¹H NMR signal of the methyl group at pos. 8 (δ_H 2.58 ppm/δ_C 30.7 ppm) shows in HMBC a ²J_H-C coupling to the keto function at C-7 (δ_C 206.7 ppm) and a ³J_H-C coupling to the quaternary C-3 (δ_C 100.9 ppm). Furthermore, the ¹H NMR signal of the methyl group in pos. 9 (δ_H 2.28 ppm/δ_C 21.2 ppm) shows a ²J_H-C to C-6 (δ_C 171.7 ppm) and a ³J_H-C on the of the methylene group at C-5 (δ_H 6.14 ppm/δ_C 102.2 ppm). The corresponding H-5 shows a further ²J_H-C to C-4 (δ_C 180.4 ppm), while the ¹³C NMR signal from C-2 cannot be determined in HMBC and is assumed to be as weak signal at 162.4 ppm. All these chemical shifts and couplings are in good agreement with those reported earlier^56,57. Numbering of protons and carbons as well all chemical shifts and couplings are shown in Fig. S6 in supplementary file 3.

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• Source: https://www.nature.com/articles/s41598-024-59511-8