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. 2024 Nov 2;17(11):e70038. doi: 10.1111/1751-7915.70038

Metabolic engineering of Streptomyces roseosporus for increased production of clinically important antibiotic daptomycin

Xingwang Li 1, Ziwei Sang 1, Xuejin Zhao 2,✉, Ying Wen 1,✉
PMCID: PMC11530997  PMID: 39487765

Abstract

Daptomycin (DAP), a novel cyclic lipopeptide antibiotic produced by Streptomyces roseosporus, is clinically important for treatment of infections caused by multidrug‐resistant Gram‐positive pathogens, but the low yield hampers its large‐scale industrial production. Here, we describe a combination metabolic engineering strategy for constructing a DAP high‐yielding strain. Initially, we enhanced aspartate (Asp) precursor supply in S. roseosporus wild‐type (WT) strain by separately inhibiting Asp degradation and competitive pathway genes using CRISPRi and overexpressing Asp synthetic pathway genes using strong promoter kasOp*. The resulting strains all showed increased DAP titre. Combined inhibition of acsA4, pta, pyrB, and pyrC increased DAP titre to 167.4 μg/mL (73.5% higher than WT value). Co‐overexpression of aspC, gdhA, ppc, and ecaA led to DAP titre 168 μg/mL (75.7% higher than WT value). Concurrently, we constructed a chassis strain favourable for DAP production by abolishing by‐product production (i.e., deleting a 21.1 kb region of the red pigment biosynthetic gene cluster (BGC)) and engineering the DAP BGC (i.e., replacing its native dptEp with kasOp*). Titre for the resulting chassis strain reached 185.8 μg/mL. Application of our Asp precursor supply strategies to the chassis strain further increased DAP titre to 302 μg/mL (2.1‐fold higher than WT value). Subsequently, we cloned the engineered DAP BGC and duplicated it in the chassis strain, leading to DAP titre 274.6 μg/mL. The above strategies, in combination, resulted in maximal DAP titre 350.7 μg/mL (2.6‐fold higher than WT value), representing the highest reported DAP titre in shake‐flask fermentation. These findings provide an efficient combination strategy for increasing DAP production and can also be readily applied in the overproduction of other Asp‐related antibiotics.

Our findings provide an efficient combination strategy for increasing clinically important daptomycin production by enhancing aspartate precursor supply, abolishing by‐product red pigment production, engineering daptomycin biosynthetic gene cluster (BGC) by promoter replacement, and duplicating the engineered daptomycin BGC in Streptomyces roseosporus wild‐type strain. We finally obtained a maximal DAP titre of 350.7 μg/mL, representing the highest reported titre in shake‐flask fermentation. Such strategies can be readily applied in the overproduction of other antibiotics using aspartate as a precursor.

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INTRODUCTION

The Gram‐positive Streptomyces species are an industrially important group characterized by the production of a large variety of secondary metabolites such as antibiotics, anticancer drugs, immunosuppressants, and insecticides (Li et al., 2021). Antibiotics are the most important secondary metabolites that display a wide range of biological activities and therapeutic spectra and are widely applied in human medicine, agriculture, and animal husbandry. More than half of the known antibiotics are sourced from Streptomyces. In addition, genome sequencing and mining have revealed numerous uncharacterized antibiotic biosynthetic gene clusters (BGCs) in Streptomyces, highlighting their valuable potential for producing novel antibiotics (Blin et al., 2021; Gavriilidou et al., 2022; Lucas et al., 2013). However, large‐scale industrial production of antibiotics using Streptomyces is generally limited by low yields, as antibiotic biosynthesis is tightly controlled by complex regulatory networks and the precursors of antibiotics are typically generated by primary metabolism, which are also required for cell growth (Cao et al., 2020). Therefore, antibiotic biosynthesis competes with primary metabolic pathways for common precursors and insufficient precursor supply is an obstacle to improving antibiotic production.

In recent years, there has been a growing focus on improving antibiotic production by enhancing precursor supply. Wang et al. (2020) found that intracellular triacylglycerol (TAG) pool can be degraded during stationary growth phase, providing necessary acyl‐CoA precursors for polyketide antibiotic biosynthesis. To balance TAG distribution between cell growth and antibiotic biosynthesis, they developed a dynamic degradation of TAG (ddTAG) strategy based on the cumate inducible system to regulate TAG pool, redirecting more carbon flux towards antibiotic biosynthesis without reducing cell growth. Using this strategy, the yields of polyketide antibiotics actinorhodin, oxytetracycline, jadomycin B, and avermectin B1a were significantly increased in four different Streptomyces species. An et al. (2021) increased acyl‐CoA precursor supply by overexpressing acetyl‐CoA carboxylase, propionyl‐CoA carboxylase, and acetyl‐CoA synthetase genes in heterologous spinosyn‐producer Streptomyces albus J1074, which led to increased spinosad production. Our laboratory increased acyl‐CoA precursor supply by overexpressing Streptomyces avermitilis native β‐oxidation pathway genes fadAB and fadD, or/and heterologous cyanobacterial CO2‐concentrating mechanism genes bicA and ecaA using the native temporal promoter pkn5p (active mainly in middle‐to‐late fermentation stage), thereby increasing avermectin B1a production in an industrial strain (Hao et al., 2022). These findings indicate the importance of enhancing precursor supply for increasing antibiotic production. However, the current precursor supply strategies in Streptomyces are mainly focused on enhancing acetyl‐CoA precursor supply for polyketide antibiotic production.

Antibiotic BGCs are tightly controlled by multiple levels of transcriptional regulators, resulting in a complex regulatory network (Liu et al., 2013; Urem et al., 2016). However, the regulatory mechanisms involved in antibiotic biosynthesis in Streptomyces are insufficiently understood. Thus, promoter engineering offers an efficient strategy for rapid and effective BGC reconstruction by using well‐characterized promoters for transcriptional activation or optimization of BGC expression (Bu et al., 2021; Ji et al., 2018, 2022; Zhao et al., 2024). In addition, increasing copy number of BGC is also an effective approach for overexpressing biosynthetic genes, thereby enhancing antibiotic production (Li et al., 2019; Li, Gao, et al., 2022; Li, Pan, et al., 2022). However, there have been no reports on combining promoter engineering and multiple copies of BGC to enhance antibiotic production.

Daptomycin (DAP), produced by Streptomyces roseosporus, is a novel calcium‐dependent cyclic lipopeptide antibiotic that is clinically important for treatment of infections caused by Gram‐positive bacteria, including methicillin‐resistant Staphylococcus aureus and vancomycin‐resistant Enterococci (Baltz, 2009; Gonzalez‐Ruiz et al., 2016). In addition to DAP, S. roseosporus also produces an undesired by‐product red pigment, which interferes with isolation and purification process and affects DAP quality. Recently, we identified the BGC responsible for red pigment synthesis in S. roseosporus by disrupting the core structural gene SSIG_RS15030 using a group II intron‐based gene editing tool and demonstrated that red pigment synthesis inhibits DAP production (Sang et al., 2024).

DAP is regarded as the best substituent to vancomycin and the last‐line of defence against multidrug‐resistant Gram‐positive pathogens due to its unique mechanism of action and low susceptibility to cross‐resistance with other antibiotics (Zuttion et al., 2020). The rapid emergence of multidrug‐resistant pathogens worldwide has greatly increased the clinical demand for DAP in recent years. Therefore, there is an urgent need to construct DAP high‐yielding strains. Although many efforts have been made to improve DAP yield by traditional random mutagenesis (Yu et al., 2011), optimizing fermentation process (Ng et al., 2014), increasing supply of non‐proteinogenic amino acid precursor kynurenine (Kyn) (Liao et al., 2013), increasing resistance to toxic precursor decanoic acid (Lee et al., 2016), engineering regulatory genes (Chen et al., 2022; Huang et al., 2017; Luo et al., 2018; Mao et al., 2015, 2017; Yan et al., 2020; Yuan et al., 2016; Zhang et al., 2015), disrupting biosynthesis of red pigment (Lyu et al., 2022; Sang et al., 2024), and utilizing metabolic engineering (Lyu et al., 2022) and synthetic biology (Ji et al., 2022) approaches, DAP yield remains low. The highest reported DAP titres were only 230 μg/mL in shake‐flask culture (Ji et al., 2022) and 812 μg/mL in fed‐batch fermentation (Ng et al., 2014).

DAP consists of 13 amino acids with a decanoic acid chain and is synthesized by non‐ribosomal peptide synthases (NRPSs) (Robbel & Marahiel, 2010). A cyclic polypeptide of 10 amino acids is formed via an ester bond between Kyn13 and threonine4 (Thr4), leaving an N‐terminal three‐amino‐acid tail with a decanoic acid moiety attached to tryptophan1 (Trp1). Of the 13 amino acids, three are aspartates (Asp3, Asp7, Asp9), suggesting that Asp is the most critical amino acid precursor for DAP biosynthesis. We thus suppose that increase of Asp supply would lead to increased DAP production. However, an effective strategy for increasing Asp precursor supply in Streptomyces has not been documented.

In this study, we enhanced Asp precursor supply in S. roseosporus by inhibiting genes involved in Asp degradation and competitive pathways using CRISPRi and overexpressing genes involved in Asp synthetic pathways, thereby increasing DAP production. We also abolished red pigment production by deleting a 21.1 kb region of its BGC and engineered the DAP BGC (termed dpt cluster) by promoter engineering. Finally, we cloned and integrated an extra copy of the engineered DAP BGC (termed dpt* cluster) into S. roseosporus chromosome. These multilevel metabolic engineering strategies, in combination, greatly enhanced DAP production (Figure 1). Our strategies described here will be useful for improving the production of other antibiotics using Asp as a precursor.

FIGURE 1.

FIGURE 1

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Overview of the multilevel metabolic engineering strategies for high DAP production in S. roseosporus. Enzymes encoded by genes: dldh (SSIG_RS25800): D‐lactate dehydrogenase; poxB (SSIG_RS04730): Pyruvate oxidase; pta (SSIG_RS08715): Phosphate acetyltransferase; acyP (SSIG_RS0234230): Acylphosphatase; ackA (SSIG_RS08720): Acetate kinase; acsA1, acsA4 (SSIG_RS14540, SSIG_RS04585): Acetyl‐CoA synthetase; nadB (SSIG_RS18555): Asp oxidase; pyrB (SSIG_RS28925): Asp carbamoyltransferase; pyrC (SSIG_RS28930): Dihydroorotase; ecaA: Carbonic anhydrase; ppc (SSIG_RS20110): Phosphoenolpyruvate carboxylase; aspC (SSIG_RS12365): Asp aminotransferase; gdhA (SSIG_RS21020): NADH‐specific glutamate dehydrogenase. dpt cluster: DAP BGC. dpt* cluster: The engineered DAP BGC, in which the native dptEp was replaced by strong constitutive promoter kasOp*.

EXPERIMENTAL PROCEDURES

Strains, plasmids, primers, and growth conditions

Strains and plasmids used in this work are listed in Table S1, and primers in Table S2. S. roseosporus strains were cultured as previously described (Yan et al., 2020; Zhang et al., 2015). DA1 agar (Zhang et al., 2015) was used for sporulation and phenotype observation of S. roseosporus strains. For DAP production, seed medium (3% Trypticase soy broth, 2.5% dextrin) and fermentation medium (1.1% yeast extract, 0.086% Fe(NH4)2(SO4)2·6H2O, 7.2% dextrin, 1.07% glucose, 1% sucrose) were used.

Escherichia coli strains JM109, EPI300, and ET12567 (Macneil & Klapko, 1987) were cultured in Luria‐Bertani (LB) medium for, respectively, routine cloning, cloning of the dpt* cluster, and propagation of non‐methylated plasmids for transformation into S. roseosporus strains.

Construction of CRISPRi strains

The CRISPRi plasmids were constructed based on pSET‐dcas9 (Zhao et al., 2018) by inserting a 20‐nt specific guide sequence (N20) of sgRNA targeting the non‐template (NT) strand of gene of interest. The N20 guide sequences of sgRNAs were designed using online software CRISPy‐web (http://crispy.secondarymetabolites.org) (Blin et al., 2016) and were synthesized by PCR annealing using primer pairs LXW1A/B~LXW10A/B, respectively, and ligated into BsaI‐digested pSET‐dcas9 (in which dcas9 and sgRNA were respectively driven by strong constitutive promoters ermEp* and j23119p) to construct CRISPRi plasmids: pdldhi, pacsA4i, pacsA1i, pptai, pacyPi, packAi, pnadBi, ppyrBi, ppyrCi, and ppoxBi. These plasmids were separately transformed into protoplasts of S. roseosporus wild‐type (WT) strain NRRL11379 to generate corresponding strains dldhi, acsA4i, acsA1i, ptai, acyPi, ackAi, nadBi, pyrBi, pyrCi, and poxBi.

For construction of CRISPRi plasmids containing multiple sgRNAs, the cassette j23119p‐sgRNAacsA1‐T0 terminator for targeting acsA1 was amplified by PCR from pacsA1i using primer pair LXW11A/LXW11B and ligated into EcoRV‐digested pacsA4i to construct pacsA1iacsA4i. The cassette j23119p‐sgRNAacyP‐T0 terminator was amplified from pacyPi using the same primers and ligated into pptai to construct pacyPiptai. The cassette j23119p‐sgRNAackA‐T0 terminator was amplified from packAi, and ligated into pacyPiptai to construct packAiacyPiptai. The cassette j23119p‐sgRNApyrC‐T0 terminator was amplified from ppyrCi and ligated into ppyrBi to construct ppyrBipyrCi. The cassette j23119p‐sgRNApta‐T0 terminator was amplified from pptai, and ligated into ppyrBipyrCi to construct pptaipyrBipyrCi. The cassette j23119p‐sgRNAacsA4‐T0 terminator was amplified from pacsA4i, and ligated into pptaipyrBipyrCi to construct pacsA4iptaipyrBipyrCi. Plasmids pacsA1iacsA4i, packAiacyPiptai, ppyrBipyrCi, and pacsA4iptaipyrBipyrCi were separately transformed into WT strain to generate corresponding strains acsA1iacsA4i, ackAiacyPiptai, pyrBipyrCi, and acsA4iptaipyrBipyrCi.

Construction of aspC, gdhA, ppc, and ecaA overexpression strains

For overexpression of aspC, ppc, or gdhA, DNA fragments aspC (1227 bp), ppc (2730 bp), and gdhA (5016 bp) were PCR amplified from genomic DNA of S. roseosporus WT strain using primer pairs LXW13A/LXW13B, LXW14A/LXW14B, and LXW15A/LXW15B, respectively. Streptomyces high‐efficiency constitutive promoter kasOp* (stronger than ermEp*) (Wang et al., 2013) was amplified from pKC‐kasOp*‐aveC8m (Hao et al., 2022) using primer pair LXW12A/LXW12B. kasOp* fragment was assembled separately with fragments aspC, ppc, and gdhA by overlap extension PCR using primer pairs LXW12A//LXW13B, LXW12A//LXW14B, and LXW12A/LXW15B. The resulting fragments kasOp*‐aspC, kasOp*‐ppc, and kasOp*‐gdhA were cloned into NdeI‐digested pIJ10500 (Pullan et al., 2011) to construct plasmids pIJ‐kasOp*‐aspC, pIJ‐kasOp*‐ppc, and pIJ‐kasOp*‐gdhA, which were separately transformed into WT to obtain strains OaspC, Oppc, and OgdhA.

For co‐overexpression of aspC, gdhA, and ppc, DNA fragments kasOp*‐aspC and kasOp*‐ppc were amplified from pIJ‐kasOp*‐aspC and pIJ‐kasOp*‐ppc using primer pairs LXW16A/LXW16B and LXW17A/LXW17B, respectively. Fragment kasOp*‐aspC was cloned into SpeI‐digested pIJ‐kasOp*‐gdhA to construct plasmid pIJ‐kasOp*‐aspC‐gdhA. Fragment kasOp*‐ppc was cloned into SpeI‐digested pIJ‐kasOp*‐aspC‐gdhA to construct plasmid pIJ‐kasOp*‐aspC‐gdhA‐ppc. Plasmids pIJ‐kasOp*‐aspC‐gdhA and pIJ‐kasOp*‐aspC‐gdhA‐ppc were separately transformed into WT to obtain strains OaspC‐gdhA and OaspC‐gdhA‐ppc.

For overexpression of ecaA, a 795‐bp DNA fragment was amplified from pIJ‐pkn5p‐ecaA (Hao et al., 2022) using primer pair LXW18A/LXW18B. Promoter kasOp* was assembled with ecaA fragment by overlap extension PCR using primer pairs LXW12A/LXW18B. kasOp*‐ecaA fragment was cloned into SpeI‐digested pIJ‐kasOp*‐aspC‐gdhA‐ppc to construct plasmid pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA, which was transformed into WT to obtain strain OaspC‐gdhA‐ppc‐ecaA.

Construction of red pigment BGC deletion strain

The main genes responsible for red pigment biosynthesis extend ~21.1 kb region within its BGC (Figure S1A). To delete this region, homologous recombination strategy was used. Two homologous arms flanking the region were prepared by PCR from WT genomic DNA. A 1074‐bp 5′‐flanking fragment (positions +1225 to +2298 relative to SSIG_RS15060 start codon) was amplified using primer pair LXW19A/LXW19B, and a 1026‐bp 3′‐flanking fragment (positions −139 to +887 relative to SSIG_RS36015 start codon) was amplified using primer pair LXW20A/LXW20B. The two fragments were ligated by fusion PCR using primer pair LXW19A/LXW20B and cloned into NcoI‐digested pCIMt005 (Li et al., 2015) to construct deletion vector pΔRED, which was then transformed into WT. The expected mutant, termed ΔRED, was confirmed by PCR using primer pairs LXW21A/LXW21B (flanking the deletion region) and LXW22A/LXW22B (located within the deletion region) (Figure S1B), followed by DNA sequencing. Use of LXW21A/LXW21B generated a 144‐bp band in ΔRED, whereas no band was detected in WT. When LXW22A/LXW22B was used, only WT produced a 407‐bp band (data not shown).

Construction of strains with replacement of dptEp by kasOp*

To replace promoter dptEp in situ with kasOp*, two fragments flanking dptEp were generated by PCR from WT genomic DNA. A 745‐bp 5′ flanking region (positions −1121 to −377 relative to dptE start codon) was amplified using primer pair LXW23A/LXW23B and a 714‐bp 3′ flanking region (positions +1 to +714 relative to dptE start codon) was amplified using primer pair LXW25A/LXW25B. kasOp* fragment was amplified from pKC‐kasOp*‐aveC8m using primer pair LXW24A/LXW24B. The three fragments were ligated into NcoI‐digested pCIMt005 using a seamless assembly cloning kit (Clone Smarter, USA) to construct dptEp replacement vector pkasOp*‐dptE, which was then transformed into WT. The expected mutant, termed kasOp*‐dptE, was confirmed by PCR using primer pairs LXW26A/LXW26B (flanking the exchange regions) (Figure S2), followed by DNA sequencing. Use of LXW26A/LXW26B generated a 2028‐bp band in WT and a 1758‐bp band in kasOp*‐dptE (data not shown).

To delete the 21.1 kb region of red pigment BGC in kasOp*‐dptE, the vector pΔRED was transformed into kasOp*‐dptE protoplasts. The expected mutant, termed kasOp*‐dptE/ΔRED, was isolated by selection of ΔRED and confirmed by PCR using the same primers. Plasmids pacsA4iptaipyrBipyrCi and pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA were separately or co‐transformed into kasOp*‐dptE/ΔRED to obtain strains XW1, XW2, and XW1‐2.

Direct cloning and integration of the engineered dpt* cluster

Cas9‐Assisted Targeting of Chromosome Segments (CATCH) technique (Jiang et al., 2015) was used to capture ~60‐kb fragment containing the engineered dpt* cluster from strain kasOp*‐dptE. Briefly, mycelia of kasOp*‐dptE were collected from 2‐day culture in fermentation medium, and genomic DNA plugs were prepared using the CHEF Bacterial Genomic DNA Plug Kit (Bio‐Rad, USA). Two sgRNA sequences (sgRNA‐dapF and sgRNA‐dapR) were designed for targeting both flanking regions of the dpt* cluster. The DNA templates of sgRNA‐dapF and sgRNA‐dapR in vitro transcription were generated by overlapping PCR using three primers: sgRNA‐dapF or sgRNA‐dapR, guide RNA‐F, and guide RNA‐R. Then, in vitro transcription of sgRNAs by T7 RNA polymerase was performed using the HiScribe™ T7 Quick High Yield RNA Synthesis Kit (New England Biolabs, USA) according to the manufacturer's protocol. The kasOp*‐dptE genomic DNA plug was digested with Cas9, together with sgRNA‐dapF and sgRNA‐dapR, as previously described (Jiang et al., 2015). The digested DNA was then precipitated with ethanol and suspended in 20 μL DNase‐free water. Plasmid pSET156 was constructed from pSET152 (Bierman et al., 1992) by replacing the high copy pUC replicon with the low copy pSC101 replicon of E. coli. The linear backbone of pSET156 was amplified from plasmid pSET156 by PCR using primers dap‐156‐F and dap‐156‐R, each of which contains a ~30 bp overlap with one end of the target gene cluster fragment. Approximately 50 ng of the backbone DNA and 1 μg of the digested genomic DNA were assembled using Gibson assembly method, and the resulting product was transformed into E. coli EPI300 competent cells by electroporation. The correct clones containing recombinant plasmid pSET156‐dpt* (Figure S3A) were verified by PCR using primer pairs L‐dap yzf /L‐dap yzr, R‐dap yzf /R‐dap yzr, dap1 yzf/dap1 yzr, and dap2 yzf/dap2 yzr (Figure S3B). After isolation, pSET156‐dpt* was transferred into kasOp*‐dptE/ΔRED by conjugation (Kieser et al., 2000) to obtain strain XW3, which contains an extra copy of the dpt* cluster.

For combination of Asp precursor supply strategies in strain XW3, ~6 kb DNA fragment containing dcas9‐sgRNA expression cassette for targeting acsA4, pta, pyrB, and pyrC was amplified from pacsA4iptaipyrBipyrCi using primer pair LXW27A/LXW27B and cloned into HindIII‐digested pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA to construct plasmid pXW1‐2, which was then transformed into XW3 to obtain strain XW1‐2‐3.

Production and analysis of DAP

Fermentation of S. roseosporus strains and measurement of DAP yield in fermentation broth by HPLC were performed as previously described (Zhang et al., 2015).

RNA extraction and RT‐qPCR analysis

For RNA extraction, S. roseosporus strains were cultured in fermentation medium for various times, and mycelia were collected and ground in liquid nitrogen. Total RNA was extracted from mycelia using Trizol reagent (Tiangen, China). Then crude RNA sample was treated with DNase I (TaKaRa, China) to eliminate genomic DNA contamination. Transcription levels of tested genes were determined by RT‐qPCR analysis as previously described (Zhang et al., 2015) using corresponding primers listed in Table S2. Relative expression value of each gene was normalized internally to the value of gene reference hrdB (SSIG_RS06665). Each experiment was conducted in triplicate.

Analysis of intracellular L‐Asp

Streptomyces roseosporus mycelia cultured in fermentation medium for various times were ground in liquid nitrogen. The powder was suspended in PBS buffer and sonicated on ice for 5 min. After centrifugation, the supernatant was taken to determine the content of L‐Asp using the Amplite™ Fluorimetric L‐Asp Assay Kit (AAT Bioquest, USA). Fluorescence intensity was measured by multifunctional microplate reader (SpectraMax i3x, Austria), with an excitation wavelength 540 nm and an emission wavelength 590 nm. L‐Asp content was calculated from a standard curve constructed using L‐Asp standard provided in the kit.

RESULTS

Exogenous addition of Asp promotes DAP production

Structural analysis of DAP (Figure 2A) suggests that Asp plays a critical role in DAP biosynthesis. To test this, we added different concentrations (30, 60, 90 mg/L) of Asp to fermentation medium of S. roseosporus WT strain based on the study of Zhu et al. (2021) and measured DAP titres from 10‐day culture by HPLC. The results showed that supplementation of Asp benefited DAP production, and addition of 60 mg/L Asp resulted in the highest titre: 134.8 μg/mL, which was 41.3% higher than the control value (95.4 μg/mL) (Figure 2B). Therefore, insufficient Asp supply might limit DAP production, and enhancement of Asp supply could promote DAP production.

FIGURE 2.

FIGURE 2

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Structure of DAP (A) and effect of Asp addition (30, 60, 90 mg/L) on DAP production (B). (A) DAP consists of 13 amino acids and a decanoic acid chain. The amino acids are in the order: Trp1, D‐Asparagine2 (D‐Asn2), Asp3, Thr4, Glycine5 (Gly5), Ornithine6 (Orn6), Asp7, D‐Alanine8 (D‐Ala8), Asp9, Glycine10 (Gly10), D‐Serine11 (D‐Ser11), Methylglutamate12 (MeGlu12), Kyn13. Blue colour: Asp. Green colour: Decanoic acid chain. (B) S. roseosporus WT strain was cultured in fermentation medium for 10 days. No addition of Asp (0 mg/L) was used as control. *p < 0.05; **p < 0.01 for comparison with control value (Student's t‐test). Error bars: SD from three replicates.

Downregulation of Asp degradation pathway genes promotes DAP production

Although exogenous addition of Asp could promote DAP production, it is uneconomical and inconvenient for scaled‐up fermentation. Therefore, we attempted to enhance Asp supply for DAP production by metabolic pathway optimization strategy. Based on the predicted pathways related to Asp synthesis in S. roseosporus (Figure 3), Asp can be degraded into dihydroorotate under the catalysis of Asp carbamoyltransferase (encoded by pyrB) and dihydroorotase (encoded by pyrC). Asp can also be converted to oxaloacetate (OAA) by Asp oxidase (encoded by nadB). To investigate the effect of pyrB, pyrC, and nadB on DAP production, we downregulated each of these three genes by CRISPRi. One sgRNA was designed for each gene to target the NT strand of the coding region close to the start codon. Three CRISPRi plasmids ppyrBi, ppyrCi, and pnadBi were accordingly constructed and separately transformed into S. roseosporus WT, resulting in CRISPRi strains pyrBi, pyrCi, and nadBi. pSET‐dcas9 without 20 nt specific guide sequence of sgRNA was also transformed into WT to obtain plasmid control strain WT/dcas9.

FIGURE 3.

FIGURE 3

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Strategies for enhanced Asp precursor supply for improvement of DAP production (schematic). Red colour: Ten genes selected as CRISPRi targets. Green colour: Four genes selected for overexpression.

RT‐qPCR analysis showed that transcription levels of pyrB, pyrC, and nadB were all lower in the corresponding CRISPRi strains than in WT grown in fermentation medium at both time points day 2 (exponential phase) and day 6 (stationary phase) (Figure S4), indicating successful inhibition of these genes in the CRISPRi strains. HPLC analysis of final DAP titres from 10‐day culture revealed that control strain WT/dcas9 showed no significant titre difference from WT, whereas inhibition of pyrB, pyrC, and nadB (strains pyrBi, pyrCi, and nadBi) all resulted in increased DAP titres (Figure 4A). Among the three CRISPRi strains, pyrCi showed the highest DAP titre: 148.4 μg/mL – 53.8% higher than WT value (96.5 μg/mL). pyrBi had the second highest DAP titre: 129.8 μg/mL – 34.5% higher than WT value. nadBi showed only 10.1% increase of DAP titre: 106.2 μg/mL. Simultaneous inhibition of pyrB and pyrC located in the same branch pathway (strain pyrBipyrCi) further increased DAP titre to 154.7 μg/mL – 60.3% higher than WT value (Figure 4A). These findings indicate that inhibition of Asp degradation pathway genes is an effective strategy for enhancement of DAP production.

FIGURE 4.

FIGURE 4

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Effects of inhibition of Asp degradation and competitive pathway genes on DAP production, cell growth, and Asp level in WT. (A–C) DAP titres for WT and its derived CRISPRi strains on day 10. WT/dcas9: WT with CRISPRi control plasmid pSET‐dcas9. (D) Growth curves of WT and acsA4iptaipyrBipyrCi grown in fermentation medium. Biomass is detected as dry cell weight. (E) Time course of DAP titre for WT and acsA4iptaipyrBipyrCi. (F) Asp levels for WT and acsA4iptaipyrBipyrCi on days 2, 4, 6, and 8. Asp level is expressed as mg per g dry cell weight (DCW). NS, not significant; **p < 0.01; ***p < 0.001 (t‐test). Error bars: SD from three replicates.

Downregulation of competitive pathway genes for Asp synthesis promotes DAP production

As a key direct precursor of Asp synthesis, intracellular supply of OAA plays a critical role in Asp accumulation (Piao et al., 2019; Zhu et al., 2021). OAA is located at the key node of central carbon metabolism pathways involving glycolysis and tricarboxylic acid (TCA) cycle. In S. roseosporus, pyruvate (the key precursor of acetyl‐CoA) can be converted to by‐products D‐lactate and acetate by D‐lactate dehydrogenase (encoded by dldh) and pyruvate oxidase (encoded by poxB), respectively; acetyl‐CoA (the key precursor of TCA) can be converted to by‐product acetate by phosphate acetyltransferase‐acetate kinase (encoded by pta‐ackA), phosphate acetyltransferase‐acylphosphatase (encoded by pta‐acyP), or acetyl‐CoA synthetase (encoded by acsA1 or acsA4) (Figure 3). To redirect more carbon flux towards OAA pool and thereby enhance Asp supply for DAP production, we downregulated each of the above seven competitive pathway genes by CRISPRi in S. roseosporus WT to obtain strains dldhi, poxBi, ptai, ackAi, acyPi, acsA1i, and acsA4i. Successful inhibition of dldh, poxB, pta, ackA, acyP, acsA1, or acsA4 in the corresponding CRISPRi strains was demonstrated by RT‐qPCR analysis (Figure S5).

Shake‐flask fermentation results showed that final DAP titre was increased in all the seven CRISPRi strains (Figure 4B). Strain ptai showed the highest DAP titre: 142.1 μg/mL – 47.3% higher than WT value (96.5 μg/mL). Strain acsA4i had the second highest DAP titre: 121.7 μg/mL – 26.1% higher than WT value. Simultaneous inhibition of ackA‐acyP‐pta or acsA1‐acsA4 located in the same branch pathway further increased DAP titre relative to single inhibition of these genes. Titre for ackAiacyPiptai was 151 μg/mL – 56.5% higher than WT value and higher than ackAi (112.1 μg/mL), acyPi (113.8 μg/mL), and ptai values. Titre for acsA1iacsA4i was 130.1 μg/mL – 34.8% higher than WT value and higher than acsA1i (111 μg/mL) and acsA4i values (Figure 4B). These findings indicate that inhibition of competitive pathway genes for Asp synthesis effectively enhances DAP production.

Combined inhibition of Asp degradation and competitive pathway genes further promotes DAP production

To further increase DAP production, we attempted to inhibit both Asp degradation and competitive pathway genes in S. roseosporus WT. However, simultaneous inhibition of the above ten genes using our CRISPRi system is difficult due to homologous recombination among sgRNA expression cassettes. Therefore, we selected the two best genes, respectively from Asp degradation (i.e., pyrB and pyrC) and competitive pathways (i.e., acsA4 and pta) for combined inhibition. Final DAP titre for the resulting strain acsA4iptaipyrBipyrCi was further increased to 167.4 μg/mL – respectively 73.5%, 28.7%, 10.9%, and 8.2% higher than values for WT, acsA1iacsA4i, ackAiacyPiptai, and pyrBipyrCi (Figure 4C). acsA4iptaipyrBipyrCi did not display notable phenotypic difference from WT grown on DA1 plate (Figure S6), and its biomass value (dry cell weight) was comparable to that of WT grown in fermentation medium (Figure 4D), whereas combined inhibition of acsA4, pta, pyrB, and pyrC significantly increased DAP titre on days 8 and 10 (Figure 4E).

To investigate whether inhibition of Asp degradation and competitive pathway genes increases Asp supply for DAP production, we measured Asp levels in WT and acsA4iptaipyrBipyrCi cultured in fermentation medium for 2, 4, 6, and 8 days. Asp levels were higher in acsA4iptaipyrBipyrCi than in WT at all four time points (Figure 4F). These findings indicate that inhibition of Asp degradation and competitive pathway genes leads to increased Asp precursor levels, and consequent promotion of DAP production.

Strengthening Asp synthetic pathways promotes DAP production

To further increase Asp supply, we attempted to strengthen Asp synthetic pathways. As shown in Figure 3, Asp can be synthesized directly from OAA under the catalysis of Asp aminotransferase (AspC) using L‐glutamate (L‐Glu) as an amino group donor. L‐Glu dehydrogenase (GDH) catalyses the formation of L‐Glu from α‐ketoglutarate (α‐KG) using NADH/NADPH as cofactor. However, TCA cycle might not produce sufficient α‐KG/L‐Glu pool to promote AspC‐catalysed transamination (Piao et al., 2019). S. roseosporus contains NADH‐specific GDH gene gdhA, which can be used to regenerate L‐Glu for AspC and NAD+ for glycolysis (Figure 5A). Therefore, we overexpressed aspC and gdhA in WT, respectively, using high‐efficiency promoter kasOp* and integrative plasmid pIJ10500 to strengthen Asp synthesis. Successful overexpression of aspC and gdhA in the resulting strains OaspC and OgdhA was demonstrated by RT‐qPCR analysis (Figure S7). Introduction of control plasmid pIJ10500 (strain WT/pIJ10500) had no effect on DAP production, whereas overexpression of aspC and gdhA (strains OaspC and OgdhA) both promoted DAP production (Figure 5B). Final DAP titre was 124.3 μg/mL for OaspC (30.0% higher than WT value 95.6 μg/mL) and 124.2 μg/mL for OgdhA (29.9% higher than WT value). Co‐overexpression of aspC and gdhA (strain OaspC‐gdhA) further increased DAP titre to 149.1 μg/mL – 56.0% higher than WT value (Figure 5B).

FIGURE 5.

FIGURE 5

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Effects of overexpression of Asp synthetic pathway genes on DAP production and Asp level in WT. (A) Schematic of L‐Asp formation reaction catalysed by AspC using a cofactor self‐sufficient system. NADH‐dependent GDH (encoded by gdhA) can regenerate L‐Glu and NAD+. PEP carboxylation reaction is catalysed by PEP carboxylase encoded by ppc, and carbonic anhydrase (encoded by ecaA) converts CO2 to bicarbonate for activation of PEP carboxylase. (B) DAP titres for WT and its derived overexpression strains on day 10. WT/pIJ10500: WT with control plasmid pIJ10500. NS, not significant; *p < 0.05; **p < 0.01; ***p < 0.001 (t‐test). (C) Asp levels for WT and OaspC‐gdhA‐ppc‐ecaA on days 2, 4, 6, and 8. Error bars (panels B, C): SD from three replicates.

In addition to TCA, OAA (the critical precursor for Asp synthesis) can also be synthesized by carboxylation of pyruvate or phosphoenolpyruvate (PEP). S. roseosporus contains PEP carboxylase gene ppc (Figures 3 and 5A), but not pyruvate carboxylase gene. We therefore overexpressed kasOp*‐driven ppc to divert carbon flux from PEP towards OAA and Asp. The resulting strain Oppc displayed DAP titre 110.7 μg/mL – 15.8% higher than WT value (Figure 5B). Co‐overexpression of ppc with aspC and gdhA in WT (strain OaspC‐gdhA‐ppc) further increased DAP titre to 154.1 μg/mL – 61.2% higher than WT value (Figure 5B).

The enzymes responsible for carboxylation reactions generally require bicarbonate and ATP for activation. However, the conversion of intracellular CO2 to bicarbonate usually has low efficiency, limiting efficient carboxylation reaction. Carbonic anhydrase (encoded by ecaA) from Anabaena sp. PCC7120 can efficiently catalyse the conversion of CO2 to bicarbonate (SoltesRak et al., 1997). We previously introduced codon optimized ecaA gene into S. avermitilis to increase the carboxylation efficiency of acetyl‐CoA and propionyl‐CoA carboxylases, thereby increasing supply of malonyl‐ and methylmalonyl‐CoA precursors for avermectin production (Hao et al., 2022). To increase the efficiency of PEP carboxylase‐catalysed carboxylation reaction, we co‐overexpressed kasOp*‐driven codon optimized ecaA with aspC, gdhA, and ppc in WT, resulting in strain OaspC‐gdhA‐ppc‐ecaA. Heterologous expression of ecaA in OaspC‐gdhA‐ppc‐ecaA was confirmed by RT‐qPCR (Figure S7). OaspC‐gdhA‐ppc‐ecaA showed no obvious phenotype and growth changes from WT (Figure S8A,B), but had a final DAP titre 168 μg/mL – respectively 75.7% and 9.0% higher than WT and OaspC‐gdhA‐ppc values (Figure 5B). Asp level was higher for OaspC‐gdhA‐ppc‐ecaA than for WT (Figure 5C) and contributed to enhanced DAP production.

Construction of a chassis strain favourable for DAP production

To achieve more promoting effect on DAP production by enhancing Asp supply, it is important to construct a suitable chassis strain. Red pigment is the main by‐product of S. roseosporus that inhibits DAP production (Sang et al., 2024), and its biosynthesis should be abolished. The entire red pigment BGC predicted by antiSMASH software (Blin et al., 2021) is ~72.5 kb, of which the main biosynthetic genes are concentrated in a 21.1 kb region (Figure S1A). To construct a chassis strain for DAP production, we deleted this 21.1 kb region by homologous recombination (Figure 6A and Figure S1B). The resulting strain ΔRED showed a complete loss of red pigment production (Figure 6B), whereas its final DAP titre reached 157.7 μg/mL – 59.8% higher than WT value (98.7 μg/mL) (Figure 6C), confirming our previous finding (Sang et al., 2024).

FIGURE 6.

FIGURE 6

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Effects of abolishing red pigment production and increasing transcription level of dptE operon on phenotype, cell growth, and DAP production in WT. (A) Schematic of construction of strains ΔRED, kasOp*‐dptE, and kasOp*‐dptE/ΔRED. Red block: Red pigment BGC. Green block: DAP BGC. ΔRED: WT with deleted 21.1 kb region of red pigment BGC. kasOp*‐dptE: WT with replacement of dptEp by kasOp*. kasOp*‐dptE/ΔRED: kasOp*‐dptE with deleted 21.1 kb region of red pigment BGC. (B) Phenotypes of WT, ΔRED, kasOp*‐dptE, and kasOp*‐dptE/ΔRED grown on DA1 plate for 7 days. (C) DAP titres for the four strains on day 10. *p < 0.05; **p < 0.01; ***p < 0.001 (t‐test). (D) Growth curves of WT and kasOp*‐dptE/ΔRED grown in fermentation medium. Error bars (panels C, D): SD from three replicates.

To further increase DAP production, we next aimed to increase the expression levels of dpt genes within DAP BGC (i.e., dpt cluster). The dpt cluster contains 12 genes, and dptEp is the most important promoter in the cluster because it drives the expression of dptE‐dptF‐dptA‐dptBC‐dptD‐dptG‐dptH operon (termed dptE operon), which contains seven structural genes required for DAP biosynthesis (Figure S2A) (Gal et al., 2006). Transcriptional regulation of dptEp is very complex; to date, six transcriptional regulators AtrA (Mao et al., 2015), DepR1 (Yuan et al., 2016), DepR2 (Mao et al., 2017), PhaR (Luo et al., 2018), BldD (Yan et al., 2020), and DasR (Chen et al., 2022) have been identified to target this promoter. To bypass the complex regulation, we replaced the native dptEp on S. roseosporus WT genome with strong constitutive promoter kasOp* by homologous recombination (Figure 6A and Figure S2B). The resulting strain kasOp*‐dptE produced fewer spores on DA1 plate (Figure 6B), whereas its final DAP titre reached 168.6 μg/mL – 70.8% higher than WT value (Figure 6C). Relative to transcription levels in WT, levels of seven dpt genes within dptE operon were all upregulated in kasOp*‐dptE (Figure S9), accounting for enhanced DAP production. Furthermore, kasOp*‐dptE maintained genetic stability after at least five successive passage generations (data not shown).

Although DAP production in kasOp*‐dptE was significantly increased, it produced more red pigment than WT with unknown reasons (Figure 6B). Therefore, we also deleted the 21.1 kb region within red pigment BGC in kasOp*‐dptE (Figure 6A). As expected, the resulting strain kasOp*‐dptE/ΔRED abolished red pigment production (Figure 6B), whereas DAP titre was further increased to 185.8 μg/mL – respectively 88.2%, 17.8%, and 10.2% higher than WT, ΔRED, and kasOp*‐dptE values (Figure 6C). Although kasOp*‐dptE/ΔRED also produced fewer spores like its parental kasOp*‐dptE (Figure 6B), its growth was similar to that of WT (Figure 6D). Thus, kasOp*‐dptE/ΔRED can serve as a high‐yielding chassis strain for further enhancing DAP production by metabolic engineering or synthetic biology strategies.

Asp precursor supply strategies promote DAP production in chassis strain

In view of the significantly increased DAP production in WT strain achieved using our Asp precursor supply strategies, we applied these strategies in the high‐yielding chassis strain kasOp*‐dptE/ΔRED. Plasmids pacsA4iptaipyrBipyrCi and pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA were separately transformed into kasOp*‐dptE/ΔRED to generate strains XW1 and XW2 (Figure 7A). Final DAP titres for these two strains were 244.5 μg/mL and 248.6 μg/mL – respectively 31.6% and 33.8% higher than the value for parental kasOp*‐dptE/ΔRED (Figure 7B). Possible integrative effect was investigated by co‐transforming plasmids pacsA4iptaipyrBipyrCi and pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA into kasOp*‐dptE/ΔRED. The resulting strain XW1‐2 had final DAP titre 302 μg/mL – respectively 62.5% and 2.1‐fold higher than kasOp*‐dptE/ΔRED and WT values, whereas co‐transformation of control plasmids pSET‐dcas9 and pIJ10500 into kasOp*‐dptE/ΔRED (strain XW1‐2‐ck) had no effect on DAP titre (Figure 7B). Thus, our Asp precursor supply strategies effectively enhance DAP production in both WT and high‐yielding strains.

FIGURE 7.

FIGURE 7

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Effects of combination of Asp precursor supply strategies and the dpt* cluster duplication on DAP production in the chassis strain kasOp*‐dptE/ΔRED. (A) Schematic of construction of strains XW1, XW2, XW1‐2, XW3, and XW1‐2‐3. XW1: kasOp*‐dptE/ΔRED with plasmid pacsA4iptaipyrBipyrCi. XW2: kasOp*‐dptE/ΔRED with plasmid pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA. XW1‐2: kasOp*‐dptE/ΔRED with plasmids pacsA4iptaipyrBipyrCi and pIJ‐kasOp*‐aspC‐gdhA‐ppc‐ecaA. XW3: kasOp*‐dptE/ΔRED with an extra copy of the dpt* cluster. XW1‐2‐3: XW3 with plasmid pXW1‐2. (B) DAP titres for WT, kasOp*‐dptE/ΔRED, XW1, XW2, XW1‐2, XW1‐2‐ck, XW3, and XW1‐2‐3 on day 10. XW1‐2‐ck: kasOp*‐dptE/ΔRED with control plasmids pSET‐dcas9 and pIJ10500. NS, not significant; ***p < 0.001 (t‐test). Error bars: SD from three replicates.

Introduction of an extra copy of the engineered dpt* cluster further promotes production

Increasing copy number of BGC is an effective strategy for improving antibiotic production (Li, Gao, et al., 2022; Li, Pan, et al., 2022). As in situ replacement of dptEp with kasOp* increased expression of seven core dpt genes and thereby increased DAP production, we directly cloned this engineered dpt* cluster from strain kasOp*‐dptE into integrative plasmid pSET156 to construct pSET156‐dpt* (Figure S3), which was then transferred into the chassis strain kasOp*‐dptE/ΔRED by conjugation (Figure 7A). DAP titre for the resulting strain XW3 was further increased to 274.6 μg/mL – 47.8% higher than the value for parental kasOp*‐dptE/ΔRED (Figure 7B).

Finally, we attempted to combine Asp precursor supply strategies in strain XW3. Because CRISPRi plasmid pacsA4iptaipyrBipyrCi and pSET156‐dpt* both contain apramycin resistance gene aac(3)IV and φC31 integrase gene, it was not possible to transform pacsA4iptaipyrBipyrCi into strain XW3. We therefore amplified dcas9‐sgRNA expression cassette for targeting acsA4, pta, pyrB, and pyrC from pacsA4iptaipyrBipyrCi and ligated it into plasmid pIJ‐kasOp*‐gdhA‐aspC‐ppc‐ecaA to construct pXW1‐2, which was then transformed into XW3. The resulting strain XW1‐2‐3 was similar to WT in terms of cell growth (Figure S10) but had a final DAP titre 350.7 μg/mL – respectively 2.6‐fold, 43.4%, 41.1%, and 27.7% higher than values for WT, XW1, XW2, and XW3 (Figure 7B).

DISCUSSION

As secondary metabolites, antibiotics are generally synthesized during stationary phase of Streptomyces fermentation, but their precursors (e.g., amino acids, acyl‐CoAs) are generated by primary metabolism (which declines in stationary phase) and are also essential building blocks for cell growth. Therefore, it is necessary to optimize precursor supply for increasing antibiotic production (Bu et al., 2021; Li et al., 2021). Asp is the most important precursor for DAP biosynthesis. In the present study, we enhanced Asp precursor supply by inhibiting Asp degradation and competitive pathway genes and overexpressing Asp synthetic pathway genes, thereby significantly increasing DAP titre in S. roseosporus WT and high‐yielding strains. Our Asp precursor supply strategies effectively enhanced Asp level and are presumably applicable in other Streptomyces species that require Asp precursor for antibiotic production.

The ten CRISPRi strains with a single targeted gene involved in Asp degradation or competitive pathway all showed increased DAP titre relative to WT value, and DAP production was more strongly promoted by simultaneous inhibition of pyrB, pyrC, acsA4, and pta. We selected only four genes for combined inhibition due to potential homologous recombination between sgRNA expression cassettes. Recently, Whitford et al. (2023) developed a scalable multiplexed CRISPR‐base editing system in Streptomyces. Using this system, multiple sgRNAs were co‐transcribed from a single promoter and processed by Csy4 (the type I‐F CRISPR‐associated endoribonuclease) to achieve simultaneous base editing of up to 17 target sites in model species Streptomyces coelicolor. Based on these findings, Csy4‐mediated processing of sgRNAs may also be used in CRISPRi system to achieve simultaneous inhibition of more than ten genes and further increase DAP production in S. roseosporus.

As Asp is an intermediate product of diverse metabolic reactions, it is difficult to maximize metabolic flux for Asp production and identify key bottleneck for Asp accumulation. Besides the ten targets we inhibited, previous studies showed that deletion of Asp catabolism pathway gene aspA (encoding Asp ammonia‐lyase responsible for the conversion of Asp to fumarate) increased Asp level in E. coli (Piao et al., 2019) and Bacillus licheniformis (Zhu et al., 2021). S. roseosporus genome doesn't contain aspA gene, but the predicted KEGG pathway suggests that Asp may be converted to fumarate by adenylosuccinate synthase‐adenylosuccinate lyase (encoded by purA‐purB) or argininosuccinate synthase‐argininosuccinate lyase (encoded by aroG‐argH). These genes will be inhibited with CRISPRi in future studies to investigate their effects on Asp supply and DAP production.

We used two approaches to enhance Asp synthesis for DAP production in S. roseosporus: (i) strengthening Asp formation from OAA; (ii) increasing OAA supply. There are two kinds of enzymes responsible for the formation of Asp from OAA: AspC and Asp dehydrogenase (AspDH). Because S. roseosporus doesn't have aspDH gene, we overexpressed native aspC to enhance Asp synthesis. We also overexpressed native GDH gene gdhA to simultaneously regenerate L‐Glu for AspC and NAD+ for glycolysis. Co‐overexpression of aspC and gdhA had stronger enhancing effect on DAP titre than separate overexpression of these two genes, indicating that AspC coupled with this cofactor self‐sufficient system could efficiently drive Asp formation. AspDH is a rare amino acid dehydrogenase that directly uses ammonia as amino donor. AspDH from Pseudomonas aeruginosa could use both NADH and NADPH as cofactors (Li et al., 2011). Heterologous overexpression of aspDH in B. licheniformis contributed to Asp accumulation and therefore benefited synthesis of target product bacitracin (Zhu et al., 2021). Heterologous expression of aspDH in S. roseosporus may further enhance Asp supply and DAP production. As for the second approach, we didn't find pyruvate carboxylase gene in S. roseosporus and therefore overexpressed native PEP carboxylase gene ppc to increase OAA supply and heterologous carbonic anhydrase gene ecaA to increase the carboxylation efficiency of PEP carboxylase. S. coelicolor contains pyruvate carboxylase gene pyc. Heterologous overexpression of S. coelicolor pyc in S. roseosporus will be conducted in the future to compare its promoting effect on DAP production with native ppc overexpression.

During S. roseosporus fermentation, production of DAP accompanies with generation of a considerable amount of visible red pigment. Biosynthesis of DAP and red pigment may compete for energy and common precursors. Therefore, abolishment of red pigment biosynthesis resulted in greatly increased DAP titre. In addition to DAP and red pigment BGCs, S. roseosporus genome contains 29 other secondary metabolite BGCs and 11 of predicted products belong to NRPS or NRPS‐like types (Sang et al., 2024), which may compete for precursors and/or energy with DAP pathway. Thus, deletion of these 11 BGCs may further increase DAP production.

Antibiotic production usually correlates with the transcription levels of its biosynthetic genes (Ji et al., 2022). To increase the expression level of dpt cluster, we replaced native dptEp with strong constitutive promoter kasOp* in WT and red pigment BGC deletion mutant ΔRED, which increased DAP titre in both the resulting strains. However, kasOp* may not be optimal for DAP production and DAP titre may be further increased by stronger constitutive promoters such as stnYp (Guo et al., 2023) or certain strong native temporal promoters. We have cloned the engineered dpt* cluster into E. coli‐Streptomyces shuttle vector pSET156, thereby enabling easy promoter replacement in vitro using established CATCH method (Jiang et al., 2015; Wei et al., 2022).

We also increased DAP titre by introducing an extra copy of the engineered dpt* cluster, suggesting that multicopy BGC can efficiently enhance DAP production. Multicopy antibiotic BGC can be achieved by ZouA‐mediated tandem amplification system and integrase‐mediated site‐specific recombination system. ZouA system could increase BGC copy number to 4–12 and thus successfully enhance production of actinorhodin (Murakami et al., 2011), validamycin A (Zhou et al., 2014), bleomycin (Li, Gao, et al., 2022), and spinosad (Li, Pan, et al., 2022). Based on integrase‐mediated site‐specific recombination system, Li et al. (2019) developed an advanced multiplex site‐specific genome engineering (aMSGE) toolkit allowing for integration of 1–4 extra copies of antibiotic BGCs into Streptomyces genomes. Application of these two systems will achieve the possible copy number of the engineered dpt cluster with optimal promoter for the maximized DAP production.

In summary, we obtained maximal DAP titre 350.7 μg/mL (2.6‐fold higher than WT value) by a combination strategy involving enhancement of Asp precursor supply, removal of by‐product red pigment synthesis, engineering of dpt cluster by promoter replacement, and duplication of the engineered dpt* cluster. Our work contributes strategies to titre improvement of other Asp‐related antibiotics.

AUTHOR CONTRIBUTIONS

Xingwang Li: Data curation; formal analysis; investigation; validation; visualization; writing – original draft. Ziwei Sang: Data curation; investigation; formal analysis; validation. Xuejin Zhao: Conceptualization; supervision; writing – review and editing. Ying Wen: Conceptualization; funding acquisition; supervision; resources; writing – original draft; writing – review and editing.

CONFLICT OF INTEREST STATEMENT

The authors declare that they have no competing interests.

Supporting information

Appendix S1

MBT2-17-e70038-s001.docx (1.4MB, docx)

ACKNOWLEDGEMENTS

This study was supported by the National Key Research and Development Program of China (Grant No. 2023YFC3402400) and the National Natural Science Foundation of China (Grant No. 32170081).

Li, X. , Sang, Z. , Zhao, X. & Wen, Y. (2024) Metabolic engineering of Streptomyces roseosporus for increased production of clinically important antibiotic daptomycin. Microbial Biotechnology, 17, e70038. Available from: 10.1111/1751-7915.70038

Contributor Information

Xuejin Zhao, Email: zhaoxj@im.ac.cn.

Ying Wen, Email: wen@cau.edu.cn.

DATA AVAILABILITY STATEMENT

The data that supports the findings of this study are available in the supplementary material of this article.

REFERENCES

  1. An, Z. , Tao, H. , Wang, Y. , Xia, B. , Zou, Y. , Fu, S. et al. (2021) Increasing the heterologous production of spinosad in Streptomyces albus J1074 by regulating biosynthesis of its polyketide skeleton. Synthetic and Systems Biotechnology, 6, 292–301. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Baltz, R.H. (2009) Daptomycin: mechanisms of action and resistance, and biosynthetic engineering. Current Opinion in Chemical Biology, 13, 144–151. [DOI] [PubMed] [Google Scholar]
  3. Bierman, M. , Logan, R. , O'Brien, K. , Seno, E.T. , Rao, R.N. & Schoner, B.E. (1992) Plasmid cloning vectors for the conjugal transfer of DNA from Escherichia coli to Streptomyces spp. Gene, 116, 43–49. [DOI] [PubMed] [Google Scholar]
  4. Blin, K. , Pedersen, L.E. , Weber, T. & Lee, S.Y. (2016) CRISPy‐web: an online resource to design sgRNAs for CRISPR applications. Synthetic and Systems Biotechnology, 1, 118–121. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Blin, K. , Shaw, S. , Kloosterman, A.M. , Charlop‐Powers, Z. , van Wezel, G.P. , Medema, M.H. et al. (2021) antiSMASH 6.0: improving cluster detection and comparison capabilities. Nucleic Acids Research, 49, W29–W35. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bu, Q.T. , Li, Y.P. , Xie, H. , Li, J.F. , Lv, Z.Y. , Su, Y.T. et al. (2021) Rational engineering strategies for achieving high‐yield, high‐quality and high‐stability of natural product production in actinomycetes. Metabolic Engineering, 67, 198–215. [DOI] [PubMed] [Google Scholar]
  7. Cao, Z. , Yu, J. , Wang, W. , Lu, H. , Xia, X. , Xu, H. et al. (2020) Multi‐scale data‐driven engineering for biosynthetic titer improvement. Current Opinion in Biotechnology, 65, 205–212. [DOI] [PubMed] [Google Scholar]
  8. Chen, Q. , Zhu, J.Y. , Li, X.W. & Wen, Y. (2022) Transcriptional regulator DasR represses daptomycin production through both direct and cascade mechanisms in Streptomyces roseosporus . Antibiotics (Basel), 11, 1065. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Gal, M.C. , Thurston, L. , Rich, P. , Miao, V. & Baltz, R.H. (2006) Complementation of daptomycin dptA and dptD deletion mutations in trans and production of hybrid lipopeptide antibiotics. Microbiology (Reading), 152, 2993–3001. [DOI] [PubMed] [Google Scholar]
  10. Gavriilidou, A. , Kautsar, S.A. , Zaburannyi, N. , Krug, D. , Muller, R. , Medema, M.H. et al. (2022) Compendium of specialized metabolite biosynthetic diversity encoded in bacterial genomes. Nature Microbiology, 7, 726–735. [DOI] [PubMed] [Google Scholar]
  11. Gonzalez‐Ruiz, A. , Seaton, R.A. & Hamed, K. (2016) Daptomycin: an evidence‐based review of its role in the treatment of gram‐positive infections. Infection and Drug Resistance, 9, 47–58. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Guo, W. , Xiao, Z. , Huang, T. , Zhang, K. , Pan, H.X. , Tang, G.L. et al. (2023) Identification and characterization of a strong constitutive promoter stnYp for activating biosynthetic genes and producing natural products in Streptomyces . Microbial Cell Factories, 22, 127. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Hao, Y. , You, Y. , Chen, Z. , Li, J. , Liu, G. & Wen, Y. (2022) Avermectin B1a production in Streptomyces avermitilis is enhanced by engineering aveC and precursor supply genes. Applied Microbiology and Biotechnology, 106, 2191–2205. [DOI] [PubMed] [Google Scholar]
  14. Huang, X. , Ma, T. , Tian, J. , Shen, L. , Zuo, H. , Hu, C. et al. (2017) wblA, a pleiotropic regulatory gene modulating morphogenesis and daptomycin production in Streptomyces roseosporus . Journal of Applied Microbiology, 123, 669–677. [DOI] [PubMed] [Google Scholar]
  15. Ji, C.H. , Kim, H. , Je, H.W. , Kwon, H. , Lee, D. & Kang, H.S. (2022) Top‐down synthetic biology approach for titer improvement of clinically important antibiotic daptomycin in Streptomyces roseosporus . Metabolic Engineering, 69, 40–49. [DOI] [PubMed] [Google Scholar]
  16. Ji, C.H. , Kim, J.P. & Kang, H.S. (2018) Library of synthetic Streptomyces regulatory sequences for use in promoter engineering of natural product biosynthetic gene clusters. ACS Synthetic Biology, 7, 1946–1955. [DOI] [PubMed] [Google Scholar]
  17. Jiang, W. , Zhao, X. , Gabrieli, T. , Lou, C. , Ebenstein, Y. & Zhu, T.F. (2015) Cas9‐assisted targeting of chromosome segments CATCH enables one‐step targeted cloning of large gene clusters. Nature Communications, 6, 8101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Kieser, T. , Bibb, M.J. , Chater, K.F. , Butter, M.J. , Hopwood, D.A. & Bittner, M. (2000) Practical Streptomyces genetics: a laboratory manual. Norwich, UK: John Innes Foundation. [Google Scholar]
  19. Lee, S.K. , Kim, H.R. , Jin, Y.Y. , Yang, S.H. & Suh, J.W. (2016) Improvement of daptomycin production via increased resistance to decanoic acid in Streptomyces roseosporus . Journal of Bioscience and Bioengineering, 122, 427–433. [DOI] [PubMed] [Google Scholar]
  20. Li, H. , Gao, W.Y. , Cui, Y.F. , Pan, Y.Y. & Liu, G. (2022) Remarkable enhancement of bleomycin production through precise amplification of its biosynthetic gene cluster in Streptomyces verticillus . Science China‐ Life Sciences, 65, 1248–1256. [DOI] [PubMed] [Google Scholar]
  21. Li, H. , Pan, Y.Y. & Liu, G. (2022) Multiplying the heterologous production of spinosad through tandem amplification of its biosynthetic gene cluster in Streptomyces coelicolor . Microbial Biotechnology, 15, 1550–1560. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Li, L. , Wei, K. , Liu, X. , Wu, Y. , Zheng, G. , Chen, S. et al. (2019) aMSGE: advanced multiplex site‐specific genome engineering with orthogonal modular recombinases in actinomycetes. Metabolic Engineering, 52, 153–167. [DOI] [PubMed] [Google Scholar]
  23. Li, P.W. , Li, J.N. , Guo, Z.Y. , Tang, W. , Han, J.S. , Meng, X.X. et al. (2015) An efficient blue‐white screening based gene inactivation system for Streptomyces . Applied Microbiology and Biotechnology, 99, 1923–1933. [DOI] [PubMed] [Google Scholar]
  24. Li, S. , Li, Z. , Pang, S. , Xiang, W. & Wang, W. (2021) Coordinating precursor supply for pharmaceutical polyketide production in Streptomyces . Current Opinion in Biotechnology, 69, 26–34. [DOI] [PubMed] [Google Scholar]
  25. Li, Y.X. , Kawakami, N. , Ogola, H.J.O. , Ashida, H. , Ishikawa, T. , Shibata, H. et al. (2011) A novel L‐aspartate dehydrogenase from the mesophilic bacterium PAO1: molecular characterization and application for L‐aspartate production. Applied Microbiology and Biotechnology, 90, 1953–1962. [DOI] [PubMed] [Google Scholar]
  26. Liao, G. , Wang, L. , Liu, Q. , Guan, F. , Huang, Y. & Hu, C. (2013) Manipulation of kynurenine pathway for enhanced daptomycin production in Streptomyces roseosporus . Biotechnology Progress, 29, 847–852. [DOI] [PubMed] [Google Scholar]
  27. Liu, G. , Chater, K.F. , Chandra, G. , Niu, G.Q. & Tan, H.R. (2013) Molecular regulation of antibiotic biosynthesis in Streptomyces . Microbiology and Molecular Biology Reviews, 77, 112–143. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Lucas, X. , Senger, C. , Erxleben, A. , Gruning, B.A. , Doring, K. , Mosch, J. et al. (2013) StreptomeDB: a resource for natural compounds isolated from Streptomyces species. Nucleic Acids Research, 41, D1130–D1136. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Luo, S. , Chen, X.A. , Mao, X.M. & Li, Y.Q. (2018) Transposon‐based identification of a negative regulator for the antibiotic hyper‐production in Streptomyces . Applied Microbiology and Biotechnology, 102, 6581–6592. [DOI] [PubMed] [Google Scholar]
  30. Lyu, Z.Y. , Bu, Q.T. , Fang, J.L. , Zhu, C.Y. , Xu, W.F. , Ma, L. et al. (2022) Improving the yield and quality of daptomycin in Streptomyces roseosporus by multilevel metabolic engineering. Frontiers in Microbiology, 13, 872397. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Macneil, D.J. & Klapko, L.M. (1987) Transformation of Streptomyce avermitilis by plasmid DNA. Journal of Industrial Microbiology & Biotechnology, 2, 209–218. [Google Scholar]
  32. Mao, X.M. , Luo, S. & Li, Y.Q. (2017) Negative regulation of daptomycin production by DepR2, an ArsR‐family transcriptional factor. Journal of Industrial Microbiology & Biotechnology, 44, 1653–1658. [DOI] [PubMed] [Google Scholar]
  33. Mao, X.M. , Luo, S. , Zhou, R.C. , Wang, F. , Yu, P. , Sun, N. et al. (2015) Transcriptional regulation of the daptomycin gene cluster in Streptomyces roseosporus by an autoregulator, AtrA. Journal of Biological Chemistry, 290, 7992–8001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Murakami, T. , Burian, J. , Yanai, K. , Bibb, M.J. & Thompson, C.J. (2011) A system for the targeted amplification of bacterial gene clusters multiplies antibiotic yield in Streptomyces coelicolor . Proceedings of the National Academy of Sciences of the United States of America, 108, 16020–16025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Ng, I.S. , Ye, C. , Zhang, Z. , Lu, Y. & Jing, K. (2014) Daptomycin antibiotic production processes in fed‐batch fermentation by Streptomyces roseosporus NRRL11379 with precursor effect and medium optimization. Bioprocess and Biosystems Engineering, 37, 415–423. [DOI] [PubMed] [Google Scholar]
  36. Piao, X. , Wang, L. , Lin, B. , Chen, H. , Liu, W. & Tao, Y. (2019) Metabolic engineering of Escherichia coli for production of L‐aspartate and its derivative beta‐alanine with high stoichiometric yield. Metabolic Engineering, 54, 244–254. [DOI] [PubMed] [Google Scholar]
  37. Pullan, S.T. , Chandra, G. , Bibb, M.J. & Merrick, M. (2011) Genome‐wide analysis of the role of GlnR in Streptomyces venezuelae provides new insights into global nitrogen regulation in actinomycetes. BMC Genomics, 12, 175. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Robbel, L. & Marahiel, M.A. (2010) Daptomycin, a bacterial lipopeptide synthesized by a nonribosomal machinery. Journal of Biological Chemistry, 285, 27501–27508. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Sang, Z. , Li, X. , Yan, H. , Wang, W. & Wen, Y. (2024) Development of a group II intron‐based genetic manipulation tool for Streptomyces . Microbial Biotechnology, 17, e14472. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. SoltesRak, E. , Mulligan, M.E. & Coleman, J.R. (1997) Identification and characterization of a gene encoding a vertebrate‐type carbonic anhydrase in cyanobacteria. Journal of Bacteriology, 179, 769–774. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Urem, M. , Swiatek‐Polatynska, M.A. , Rigali, S. & van Wezel, G.P. (2016) Intertwining nutrient‐sensory networks and the control of antibiotic production in Streptomyces . Molecular Microbiology, 102, 183–195. [DOI] [PubMed] [Google Scholar]
  42. Wang, W. , Li, S. , Li, Z. , Zhang, J. , Fan, K. , Tan, G. et al. (2020) Harnessing the intracellular triacylglycerols for titer improvement of polyketides in Streptomyces . Nature Biotechnology, 38, 76–83. [DOI] [PubMed] [Google Scholar]
  43. Wang, W. , Li, X. , Wang, J. , Xiang, S. , Feng, X. & Yang, K. (2013) An engineered strong promoter for streptomycetes . Applied and Environmental Microbiology, 79, 4484–4492. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Wei, W. , Wang, W. , Li, C. , Tang, Y. , Guo, Z. & Chen, Y. (2022) Construction and heterologous expression of the di‐AFN a(1) biosynthetic gene cluster in Streptomyces model strains. Chinese Journal of Natural Medicines, 20, 873–880. [DOI] [PubMed] [Google Scholar]
  45. Whitford, C.M. , Gren, T. , Palazzotto, E. , Lee, S.Y. , Tong, Y.J. & Weber, T. (2023) Systems analysis of highly multiplexed CRISPR‐base editing in Streptomycetes . ACS Synthetic Biology, 12, 2353–2366. [DOI] [PubMed] [Google Scholar]
  46. Yan, H. , Lu, X.R. , Sun, D. , Zhuang, S. , Chen, Q. , Chen, Z. et al. (2020) BldD, a master developmental repressor, activates antibiotic production in two Streptomyces species. Molecular Microbiology, 113, 123–142. [DOI] [PubMed] [Google Scholar]
  47. Yu, G.H. , Jia, X.Q. , Wen, J.P. , Lu, W.Y. , Wang, G.Y. , Caiyin, Q. et al. (2011) Strain improvement of Streptomyces roseosporus for daptomycin production by rational screening of He‐Ne laser and NTG induced mutants and kinetic modeling. Applied Biochemistry and Biotechnology, 163, 729–743. [DOI] [PubMed] [Google Scholar]
  48. Yuan, P.H. , Zhou, R.C. , Chen, X. , Luo, S. , Wang, F. , Mao, X.M. et al. (2016) DepR1, a TetR family transcriptional regulator, positively regulates daptomycin production in an industrial producer, Streptomyces roseosporus SW0702. Applied and Environmental Microbiology, 82, 1898–1905. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Zhang, Q.L. , Chen, Q. , Zhuang, S. , Chen, Z. , Wen, Y. & Li, J.L. (2015) A MarR family transcriptional regulator, DptR3, activates daptomycin biosynthesis and morphological differentiation in Streptomyces roseosporus . Applied and Environmental Microbiology, 81, 3753–3765. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Zhao, X.J. , Zong, Y.Q. , Lou, Q.L. , Qin, C.R. & Lou, C.B. (2024) A flexible, modular and versatile functional part assembly toolkit for gene cluster engineering in Streptomyces . Synthetic and Systems Biotechnology, 9, 69–77. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Zhao, Y.W. , Li, L. , Zheng, G.S. , Jiang, W.H. , Deng, Z.X. , Wang, Z.J. et al. (2018) CRISPR/dCas9‐mediated multiplex gene repression in Streptomyces . Biotechnology Journal, 13, e1800121. [DOI] [PubMed] [Google Scholar]
  52. Zhou, T.C. , Kim, B.G. & Zhong, J.J. (2014) Enhanced production of validamycin a in Streptomyces hygroscopicus 5008 by engineering validamycin biosynthetic gene cluster. Applied Microbiology and Biotechnology, 98, 7911–7922. [DOI] [PubMed] [Google Scholar]
  53. Zhu, J. , Li, L. , Wu, F. , Wu, Y. , Wang, Z. , Chen, X. et al. (2021) Metabolic engineering of aspartic acid supply modules for enhanced production of bacitracin in Bacillus licheniformis . ACS Synthetic Biology, 10, 2243–2251. [DOI] [PubMed] [Google Scholar]
  54. Zuttion, F. , Colom, A. , Matile, S. , Farago, D. , Pompeo, F. , Kokavecz, J. et al. (2020) High‐speed atomic force microscopy highlights new molecular mechanism of daptomycin action. Nature Communications, 11, 6312. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Appendix S1

MBT2-17-e70038-s001.docx (1.4MB, docx)

Data Availability Statement

The data that supports the findings of this study are available in the supplementary material of this article.

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