The btm gene cluster in S. scabies encodes a single putative regulator, BtmL, a 20.5 kDa protein that contains a C-terminal domain of unknown function (DUF2087 or PF09860), which has been associated with putative transcriptional regulators and is proposed to bind nucleic acids (Rigden, 2011). To date (July 2019), over 6,500 DUF2087-containing proteins have been sequenced and deposited in Genbank, of which more than 2,000 have additional DNA-binding domains. In silico analysis of BtmL using both Phyre2 (Crone et al., 2012; Kelley et al., 2015) and I-TASSER (Yang et al., 2015) predicted that the N-terminus of the protein has structural homology with SmtB-ArsR-like repressors, and therefore would feature a winged helix-turn-helix (wHTH) domain characteristic of this family of transcriptional regulators (Busenlehner et al., 2003; Osman and Cavet, 2010; Chakravorty and Merz, 2014). However, no canonical wHTH domain was detected by sequence analysis and the protein appears to lack the conserved residues characteristically involved in ArsR-family metal binding (Osman and Cavet, 2010; Crone et al., 2012). Despite this, previous empirical evidence showed that addition of cobalt(II) to the production medium increased bottromycin yields (Crone et al., 2012), inferring that the ArsR-like structure of BtmL may control cluster expression via a metal-binding mechanism. This hypothesis was especially compelling given that several genes in the btm cluster encode class B radical SAM methyltransferases, which are cobalamin-dependent enzymes (Bauerle et al., 2015).

To investigate the role of BtmL in bottromycin biosynthesis, a mutant S. scabies strain carrying an in-frame deletion of btmL was generated (Crone et al., 2012) and its ability to produce bottromycin was assessed using liquid chromatography - mass spectrometry (LC-MS). Inactivation of btmL did not abolish bottromycin biosynthesis, but we could observe a moderate and consistent decrease in production levels to approximately 40% of wild type (WT) levels (Figure 2A). This result suggests that btmL acts as a positive modulator of bottromycin biosynthesis, but is not the master activator of the pathway. WT levels of bottromycin production were restored in ΔbtmL upon in trans complementation with a copy of btmL under the control of the constitutive promoter ermE^∗p (Bibb et al., 1985), confirming that the phenotype was due to the deletion of btmL. Surprisingly, when that same btmL expression construct was introduced in the WT strain (generating strain WT + L), there was no increase in bottromycin levels (Figure 2A).

To evaluate quantitatively whether btmL could be acting as a transcriptional regulator, qRT-PCR experiments were carried out with RNA samples from both the WT and ΔbtmL strains (Figure 2B). The btm gene cluster is organized in two divergent groups of genes: btmA and btmB on one strand and btmC-btmM on the other strand (Figure 1). We therefore measured the expression of btmB from the smaller block, and btmC, btmD (the gene encoding the precursor peptide), btmE and btmL from the rest of the cluster. Expression values of these genes were normalized using the expression of hrdB, which encodes the principal sigma factor of RNA polymerase. A time course experiment determined that the optimal time to measure btm gene expression was at 72 h post-inoculation of bottromycin production medium (Supplementary Figure S1). At this time point, it could be observed that deletion of btmL has no significant effect on most of the genes tested, with one notable exception: btmD, the gene encoding the bottromycin precursor peptide. Transcription of btmD is reduced to approximately 40% of WT levels, which correlates with the reduction in bottromycin production (Figure 2B). Furthermore, qRT-PCR analysis of the WT + L strain revealed that transcription of btmD is significantly increased in this strain, confirming that BtmL is a specific and positive regulator of btmD. Interestingly, upstream and downstream genes (btmB, btmC, and btmE) are only slightly overexpressed or not overexpressed at all, suggesting the presence of an alternative operon encompassing only btmD.

After confirming that btmL is involved in bottromycin regulation, we assessed whether cobalt(II) levels influence pathway productivity, and whether this effect was mediated by btmL. Bottromycin production was measured in S. scabies WT and ΔbtmL cultured in bottromycin production medium (BPM) supplemented with different concentrations of CoCl₂ (Figure 3A). In absolute terms, bottromycin production increased upon addition of cobalt(II), reaching a maximum at 15–25 μg/mL CoCl₂ after which production steadily decreased until almost disappearing at 200 μg/mL CoCl₂. However, the increase in production simply corresponded to a proportional increase in mycelial growth upon addition of up to 25 μg/mL of cobalt(II) in the medium (Figure 3A). At higher cobalt(II) concentrations mycelial growth was sustained, so the drop in production was probably due to a general inhibitory effect of excess metal in the medium, a phenomenon reported for the biosynthesis of other antibiotics (Rogers and Birnbaum, 1974; Abbas and Edwards, 1990). The growth-dependent production increase was also noticeable in ΔbtmL, which followed a similar pattern, albeit at a reduced level of production. Moreover, a comparison of qRT-PCR data for the expression levels of several pathway genes in the presence or absence of added cobalt(II) (Figure 3B) showed that these are nearly identical in either condition for both the WT and ΔbtmL strains, discarding any specific regulatory effect of this metal. This result is in accordance with the observation that btmL lacks the conserved metal-binding residues characteristic of ArsR-SmtB regulators (Osman and Cavet, 2010; Crone et al., 2012).

These results ruled out cobalt(II)-dependent pathway regulation by BtmL, which is instead a cobalt-independent transcriptional modulator that selectively increases btmD transcript levels in order to generate high levels of the precursor peptide in a feed-forward loop. Other positive loop systems have been reported in the biosynthesis of several RiPPs like microbisporicin and planosporicin, where the autoinduction mechanism relies of the detection of small quantities of precursor peptide by a sigma-antisigma complex (Sherwood and Bibb, 2013; Fernández-Martínez et al., 2015), or cinnamycin, where production is launched after onset of immunity to the compound in the producer strain (O’Rourke et al., 2017).

The discordance between the increase in btmD expression and the lack of change in bottromycin production in the WT + L strain (Figure 2A) led us to postulate that the potential increase in available BtmD might not be effectively channeled by the pathway and exported out of the cell, leading to increased degradation of pathway intermediates and generation of shunt metabolites. Supporting this hypothesis, previous work showed that overexpression of botT, the major facilitator superfamily (MFS) transporter gene (Kumar et al., 2016; Quistgaard et al., 2016) in the bottromycin cluster from Streptomyces sp. BC16019 increased bottromycin production in a heterologous host, although it still did not reach wild type production levels (Huo et al., 2012). We therefore assessed whether overexpressing the homolog of this gene (btmA, 88% identity to botT) in the S. scabies btm gene cluster might increase the efficiency of the pathway and self-resistance to potentially toxic levels of the antibiotic.

Expression of a second copy of btmA under the control of the constitutive promoter ermE^∗p provided a 25% increase in bottromycin production with respect to the WT strain (Figure 4A), an effect that was maintained, but not improved, when it was expressed in WT + L (generating WT + L + A). When the quantification was extended to other bottromycin related metabolites described previously (Crone et al., 2016; Eyles et al., 2018) this pattern was still observed for the total set of mature bottromycins (those including the main posttranslational modifications) (Figure 4A). However, in the case of the pathway

intermediates and shunt metabolites the trend was not present, with WT + L and WT + A producing more of them than the WT and the double overexpression strain WT + L + A (Figure 4B). This result indicates that efficiency can be increased to some extent by improving export of the molecule, but this is not the major bottleneck for bottromycin production. The main limiting factor for bottromycin production is likely to be the post-translational maturation of the precursor peptide instead, as we previously showed that the bottromycin pathway inefficiently stalls at numerous biosynthetic steps, even in the WT strain (Crone et al., 2016).

Given that btmD appears to be situated within a larger operon, the specific increase in btmD transcription following btmL overexpression was unexpected and prompted us to investigate the transcriptional organization of the btm gene cluster. This cluster contains 13 genes, where 11 (btmC-M) are arranged in the same orientation and appear tightly clustered, with btmE to btmJ physically overlapping and btmK starting 8 bp after the btmJ stop codon. To assess the possibility of co-transcription of the remaining genes, RT-PCR experiments were performed on 72 h cultures of both WT and ΔbtmL strains with specific primer pairs for the intergenic regions btmA-B, btmC-D, btmD-E, and btmK-L (Figure 5A). These showed co-transcription of all of the tested regions except btmA-B, indicating that genes btmC-M co-transcribe and can behave as a single 14.5 kb operon, whereas btmA and btmB are transcribed separately. The gene cluster was analyzed for putative terminators using WebGeSTer (Mitra et al., 2011). Consistent with the RT-PCR results, terminators were predicted spanning up to 90 bp away from the stop codon of btmA (ΔG = −19.5), between btmA and btmB (ΔG = −19.43) and 149 bp away from the stop codon of btmM (ΔG = −19.27) (Figure 5A).

This evidence for btmC-M co-transcription did not exclude the possibility of alternative operons, and the specific increase in btmD transcription following btmL overexpression hinted toward an alternative transcription start site (TSS) for btmD that would explain how it is differentially regulated from its surrounding genes. In order to map the TSSs for the whole cluster we employed 5′-tag-RNA-seq (also called tagRNA-seq) (Innocenti et al., 2015) that enables the identification of TSSs in an untargeted, genome-wide fashion. This powerful technique has the advantage of differentially distinguishing primary transcripts (those generated by an RNA polymerase and therefore coming from true TSSs) from processed transcripts, which arise upon degradation or RNase mediated cleavage of the original transcripts at specific processing sites (PSs). In this particular case, with an operon spanning over 10 kb, degradation or post-transcriptional modification of the resulting transcript could also account for the difference between btmD expression and the remainder of the btmC-M operon.

5′-tag-RNA-seq reads mapped to the S. scabies 87-22 genome clearly showed the presence of three TSSs in the btm cluster: one preceding btmB, one before btmC, and an extra one preceding btmD and within the coding region of the upstream gene, btmC (Figure 5B). Unexpectedly, no TSS was found before btmA, in contrast with what was expected from the intergenic RT-PCR result (Figure 5A). To independently confirm the existence of these TSSs, we performed 5′-RACE (Rapid amplification of cDNA ends) experiments for btmB_TSS, btmC_TSS, and btmD_TSS, which yielded identical results to the 5′tag-RNA-seq (Figures 5C,D). btmC_TSS is in position -32 with respect to the btmC start codon, and a prediction of putative σ70 promoters in the btmB-btmC intergenic region using BPROM (Solovyev and Salamov, 2011) identified a nearly canonical −35 sequence (TTGACT) and a poorly conserved −10 region (TCCTACGGT) that are consistent with the location of the btmC_TSS (Figure 5C). It was not possible to confidently identify −10 and −35 boxes for the btmB_TSS using BPROM or other prediction tools, so potential sequences were identified manually (Figure 5C). The btmD_TSS is located 348 bp away from the start of btmD and 215 bp before the end of btmC coding sequence (Figure 5D), but we could not identify any suitable −10 or −35 regions for this TSS, either manually or with predictive tools. Unexpectedly, btmD_TSS appears both in the WT and in ΔbtmL, albeit with lower intensity in ΔbtmL (Figure 5B), which indicates that although btmL does affect the transcription of btmD, it is not essential for the expression of this alternative transcript.

Interestingly, the 5′tagRNA-seq results showed that at 72 h of growth there is an extremely high level of transcription of some btm genes, in particular btmD. Transcription initiation preceding this gene seems rather promiscuous with some background noise (Figure 5B). Strikingly, an analysis of the full transcriptome to assess levels of gene expression as reads per kilobase per million reads (RPKM) and transcripts per million (TPM) (Wagner et al., 2012), showed that btmD and btmC are the 4th and 9th most highly transcribed genes in the genome at this time point (Supplementary Datasheet S1). Transcription after btmD drops considerably, showing that even upon overexpression of btmL, expression of btmE did not increase in a comparable way to btmD, in accordance with the qRT-PCR results (Figure 2B). The transcriptomic data also shows that all downstream genes in the operon are transcribed at lower levels than their preceding genes. This may be explained by a transcription attenuator downstream of btmD that likely serves to dampen transcription of downstream genes in the cluster. Accordingly, a putative terminator sequence between btmD and btmE (ΔG = −21.22) is predicted by WebGeSTer (Mitra et al., 2011).

To further characterize the main promoter regions in the pathway, promoter activity was measured using the β-glucuronidase reporter gene gusA (Myronovskyi et al., 2011). A 414 bp fragment containing the whole intergenic region btmB-btmC, including btmC_TSS but not its RBS, was cloned in vector pIJ10742 (Feeney et al., 2017) to create a transcriptional fusion with gusA. This construct was introduced in S. scabies WT, ΔbtmL, and WT + L to test whether the different genetic backgrounds would affect promoter activity. As a positive control, vector pIJ10741 carrying gusA under the control of the constitutive promoter ermE^∗p (Feeney et al., 2017) was used, while empty pIJ10742 with no promoter was employed as a negative control. btmCp proved to be an exceptionally strong promoter in both in solid and in liquid culture (Figure 6), and at 72 h of growth its activity was comparable to ermE^∗p. This result correlates with the high levels of expression of btmC observed in the transcriptomic analysis. The same gusA reporter experiment was performed with a 545 bp region that was predicted to contain the promoter preceding btmD_TSS, but no reporter activity could be detected in any of the genetic backgrounds tested (Figure 6).

Therefore, further reporter constructs were made that progressively extended the region tested until it included the whole coding sequence of btmC (but not btmCp), but these provided identical negative results (Supplementary Figures S4, S5). This result pointed to the possibility that transcription from btmD_TSS is not driven from its own promoter, but rather from btmCp, where some sort of secondary structure might help enhance transcription of this particular gene. To test this, we assessed the promoter activity of the whole region (btmCp, btmC, and the intergenic region before btmD) and compared its promoter activity to that of the original btmCp reporter. Two promoter versions were generated, one containing the WT sequence, and another one where several synonymous point mutations were introduced in the region of btmDTSS via yeast refactoring (Supplementary Figure S4) in order to disrupt possible DNA secondary structures in this region. Although these new transcriptional fusions were more active than all the other sequences lacking btmCp, they showed drastically reduced activity when compared to the original btmCp reporter, indicating that the additional DNA sequence dampened the promoter activity of btmCp. The mutated btmDTSS region did not significantly affect this phenotype (Supplementary Figures S4, S5).

Our transcriptional analysis combined with the data from S. scabies ΔbtmL and WT + L show that the master regulator(s) of the btm cluster are not encoded by the btm cluster and are therefore unlikely to be pathway-specific. Identification of such regulators could aid in improving the yield of bottromycin. Given the difficulty in predicting binding sequences for transcriptional activators in streptomycetes without extensive transcriptomic and metabolic datasets (Iqbal et al., 2012; Hwang et al., 2019), the intergenic region btmB-C was used as a probe in a DNA-binding protein capture experiment. This region was amplified using biotinylated primers and the resulting fragment was immobilized on streptavidin coated magnetic beads. The beads were then incubated with lysates of 72 h cultures of S. scabies WT grown in BPM. As a negative control, the same promoter region was incubated with lysates of cultures grown in GYM medium, where the cluster is not expressed and therefore there is no bottromycin production (Supplementary Figure S6). After incubation and several washes with competitor DNA to eliminate proteins binding non-specifically to the probe, the remaining proteins were eluted, washed again and analyzed using quantitative proteomics. A total of 321 proteins were identified in the 2 sets of samples, 120 of which appeared in both conditions, whereas 177 where present exclusively in the BPM samples and only 24 were specific to GYM samples. These results were filtered to identify annotated regulatory proteins or uncharacterized proteins binding to btmCp specifically in bottromycin production conditions. This reduced the candidates to 21 proteins (Supplementary Table S3), of which three were annotated as regulators, while the remainder were uncharacterized proteins.

Based on the intensity of the signal obtained or the biological relevance of the hit, we selected four promising candidates for further genetic analysis to assess their involvement in bottromycin biosynthesis: MtrA (SCAB_55281), GlnK (SCAB_61751), a putative regulatory protein (SCAB_ 85931) and an uncharacterized protein that binds specifically to btmCp under production conditions (SCAB_51451). MtrA forms a three-component system with the histidine kinase MtrB and LpqB, a lipoprotein that is involved in signal transduction (Hoskisson and Hutchings, 2006; Nguyen et al., 2010). This system is widely conserved in Actinobacteria, where it has been shown to regulate both development and antibiotic production in Streptomyces venezuelae and in Streptomyces coelicolor (Som et al., 2017; Zhang et al., 2017). SCAB_61751 encodes a homolog of GlnK, a signal transduction protein with a central role in nitrogen metabolism (Thomas et al., 2000). SCAB_ 85931 is annotated as a putative regulatory protein and contains domains characteristic of RsbR-like anti-anti-sigma factors. Rsb proteins form part of the “stressosome,” which responds to bacterial stress and ultimately leads to the activation of the alternative sigma factor σ^B (Lee et al., 2004). SCAB_51451 contains no conserved domains. In order to test the role of these genes in bottromycin biosynthesis, each of them was constitutively overexpressed under the control of ermE^∗p in S. scabies WT using the integrative plasmid pIB139-RBS, a derivative of pIB139 (Wilkinson et al., 2002). The potential involvement of the MtrAB two-component system prompted us to also express mtrB, which encodes the cognate histidine kinase. Despite the evidence provided by DNA binding, the bottromycin titres were comparable to the WT under the conditions tested (Supplementary Figure S7).

Unless otherwise specified, antibiotics, media components and chemical reagents used in this work were purchased from Sigma-Aldrich (United Kingdom) with the exception of soy flour, which was purchased from Holland & Barret (United Kingdom). Enzymes and molecular biology kits were purchased from New England Biolabs and Promega Healthcare, respectively.

The following strains were used in this work: S. scabies DSM 41658, both the wild-type (WT) and the mutant strain ΔbtmL (ΔL), whose construction was described previously (Crone et al., 2012). Additionally, Escherichia coli K-12 strain DH5α (Invitrogen) was used for plasmid propagation and DNA manipulation, and the methylation deficient strain E. coli ET12567 containing pUZ8002 (Paget et al., 1999) was used for intergeneric conjugal transfer of genetic material to S. scabies, which was performed following standard procedures (Kieser et al., 2000). E. coli culture media used in this work are described in Sambrook et al. (1989). Several Streptomyces culture media were used: mannitol soya flour medium (SFM) (Kieser et al., 2000) was used for Streptomyces propagation and conjugations, and instant potato mash agar [2% Smash (Premier Foods), 2% agar] was used to grow S. scabies for spore harvesting. GYM medium (0.4% glucose, 0.4% yeast extract, and 1.0% malt extract, in Milli-Q water) and bottromycin production medium (BPM: 1% glucose, 1.5% soluble starch, 0.5% yeast extract, 1.0% soy flour, 0.5% NaCl, and 0.3% CaCO₃, in Milli-Q water) were used for bottromycin production. Production experiments were performed as follows: 30 μL of concentrated spores were used to inoculate 10 mL of GYM medium in 50 mL flasks and were incubated for 48 h at 30°C and 250 rpm. 250 μL of seed culture were used to inoculate 10 mL BPM in 50 mL Falcon tubes covered with foam bungs instead of caps. Alternatively, 1 mL of seed culture was used to inoculate 50 mL of BPM in 250 mL flasks containing a spring. When necessary, cultures were supplemented with appropriate concentrations of CoCl₂ (25 μg/mL unless otherwise stated). Triplicate production cultures were incubated for 5–6 days at 28°C and 230 rpm, at which point samples were collected and processed immediately, or frozen at −20°C until further processing and analysis. When antibiotic selection was necessary, culture media were supplemented with the appropriate antibiotics at the following final concentrations: kanamycin at 50 μg/mL, apramycin at 50 μg/mL, hygromycin at 50 μg/mL, chloramphenicol at 25 μg/mL, and nalidixic acid at 25 μg/mL.

All the primers used for gene amplification and generation of the following constructs are listed in Supplementary Table S1. The gene expression vectors used in this work were pIB139-RBS-btmD (Crone et al., 2016) and pIJ10257 (Hong et al., 2005), which are both integrative plasmids containing the constitutive ermE^∗ promoter. pIB139-RBS-btmD is a derivative of pIB139 (Wilkinson et al., 2002) that carries the btmD gene preceded by a ribosome binding site (RBS) and an NdeI restriction site installed to facilitate cloning and improve gene expression (Crone et al., 2016). This plasmid was linearized using restriction enzymes NdeI and EcoRI, releasing btmD and allowing the introduction of the gene of interest, either via ligation of an EcoRI/NdeI treated PCR fragment or by Gibson assembly, following published protocols (Crone et al., 2012). The ligation method was used to introduce btmL and generate pIB139-RBS-btmL, which was used to complement the mutant ΔbtmL and to overexpress this gene in the WT strain (WT + L). The Gibson assembly method was used to introduce mtrA, mtrB, glnB, SCAB_85931, and SCAB_51451, into pIB139-RBS for gene overexpression the WT strain. An empty version of pIB139-RBS was generated to use as a control in production experiments, via Gibson assembly, using the NdeI/EcoRI linearized vector and primers pIB-RBS_fw and pIB-RBS_rv, carrying the RBS sequence. pIJ10257 was used for the expression of btmA. This gene was amplified using primers btmA-start and btmA-end, digested with NdeI and HindIII and ligated into NdeI/HindIII digested pIJ10257. Correct construction of all expression plasmids was confirmed using colony PCR and sequencing, after which they were transferred to E. coli ET12567/pUZ8002 cells for conjugation into S. scabies (WT or ΔbtmL as appropriate). pIB139 integrates in the ϕC31 phage integration site, and pIJ10257 integrates in the ϕBT1 site, which allowed for the simultaneous overexpression of two genes when necessary. Correct integration of the plasmids was verified by colony PCR using ermEp_chk1 in combination with the reverse amplification primer for each of the genes. As a control for the production experiments, a strain carrying the corresponding empty vector was generated in each case.

Bottromycin production culture samples (1 mL) were extracted with an equal volume of methanol, mixed with shaking for a minimum of 10 min. The mixtures were then centrifuged for 4 min at 13,000 rpm to pellet cellular material and other particulate contaminants. 2 μL of the resulting supernatants were injected onto a Phenomenex Kinetex 2.6 μm XB-C18 column (50 mm × 2.1 mm, 100 Å) attached to a Shimadzu Nexera X2 UHPLC and eluted with a linear gradient of 5 to 95% acetonitrile (ACN) in water + 0.1% formic acid (FA) over 6 min, with a flow-rate of 0.6 mL/min. MS data were obtained in positive mode using a Shimadzu IT-TOF mass spectrometer coupled to the UHPLC and analyzed using LabSolutions software (Shimadzu). Bottromycin production was plotted in peak area units. To normalize production values across samples, culture growth was quantified by measuring DNA concentration with an adaptation of the Burton diphenylamine colorimetric assay (Zhao et al., 2013).

Two milliliter samples of S. scabies liquid cultures were harvested after 72 h incubation in conditions as described above. Samples were washed with an equal volume of RNAlater (Thermo Fisher Scientific) and stored at −80°C until further processing. RNA was then extracted following previous protocols (Crone et al., 2016), resuspending the mycelium in 1 mL of RLT buffer from the RNeasy Kit (Qiagen) and homogenizing the sample in lysing matrix B tubes (MP Biomedicals) using a FastPrep instrument and a program of 3 × 30 s pulses at 6 m/s with 1 min cooling intervals on ice. The lysates were then centrifuged at 13,000 rpm and 700 μL of supernatant from each sample were then transferred to spin tubes from the RNeasy Kit to undergo purification following manufacturer instructions. Chromosomal DNA contamination was eliminated with on-column DNase I treatment (Qiagen) and a further cleanup step using TURBO DNA-free Kit (Ambion, Invitrogen). RNA concentration in the samples was quantified measuring A₂₆₀ using a Nanodrop. In the case of the samples for 5′tag-RNAseq, further quantity and integrity measurements were performed with RNA ScreenTape (Agilent).

Both RT-PCR and qRT-PCR analyses were carried out using 250 ng of total RNA as template in a two-step protocol. The first step consisted of cDNA synthesis with the QuantiTect Reverse Transcription Kit (Qiagen), as detailed in the manufacturer’s instructions. For RT-PCR analyses the cDNA was then used as a template for PCR reactions using Taq polymerase and specific primers for the regions to test (Supplementary Table S1). The amplification conditions were as follows: initial denaturation at 95°C for 3 min followed by 33 cycles of 95°C for 30 s, 58–62°C for 30 s and 72°C for 40–60 s, with a final extension step at 72°C for 5 min. The resulting RT-PCR products were separated in 2% agarose gels and stained with ethidium bromide for visualization. For qRT-PCR analyses the aforementioned cDNA was used as template in quantitative reactions with the SensiFAST SYBR No-ROX Kit (Bioline) following the manufacturer’s instructions. The reactions were run in a Bio-Rad CFX96 thermocycler and the amplification protocol was a 2-step cycling PCR program: 1 cycle at 95°C for 2 min followed by 40 cycles of 5 s at 95°C and 30 s at 60°C. An additional melting curve step was used at the end of the reaction to assess the specificity of the amplified products. The qRT-PCR results were analyzed with CFX Manager software (Bio-Rad). In both cases, negative control samples of the cDNA synthesis step with no retrotranscriptase were included, in order to control for the presence of contaminating chromosomal DNA in the RNA samples. Primers used in these analyses (Supplementary Table S1) were designed with a preference for 17–23 mers and T_m ∼ 65°C with the help of Vector NTI Advance 11.5 (Invitrogen) and Primer3 software (Untergasser et al., 2012) and validated used the online tool NetPrimer (Premier Biosoft). In the specific case of qRT-PCR primers, these were designed to amplify fragments of ∼100 bp and their efficiency was tested using serial dilutions of chromosomic DNA as template. For both RT-PCR and qRT-PCR, primers for hrdB, encoding a housekeeping sigma factor, were used as an internal control to assess the quality of RNA and in the latter case to normalize gene expression levels.

The TSSs identified in the 5′tag-RNA-seq experiment were validated using a 5′-RACE system for rapid amplification of cDNA ends (Invitrogen), using the manufacturer’s instructions (version 2.0). 1 μg of total RNA from S. scabies WT and ΔbtmL harvested after 72 h of growth at 28°C and 230 rpm was used to carry out cDNA synthesis with specific primers for each of the promoters tested (Supplementary Table S1). The cDNA was purified and treated with terminal deoxynucleotidyl transferase (TdT) to add poly(dC) tails to its 3′ends. After an initial PCR amplification of the tailed fragments with the 5′-RACE abridged anchor primer and subsequent amplifications with the universal amplification primer (both provided by the kit) and specific nested primers (Supplementary Table S1), defined amplification products were observed. These products were gel purified and submitted for Sanger sequencing to confirm the position of the TSSs.

Putative promoter regions of btmC (btmCp, 414 bp) and btmD (btmDp, 545 bp) and extended promoter regions spanning different lengths of btmC were amplified from S. scabies genomic DNA or from a refactored bottromycin cluster containing 33 synonymous point mutations in the btmD_TTS region (Supplementary Figure S4) with primers containing NdeI and XhoI restriction sites (Supplementary Table S1). These were verified by sequencing, and then ligated or assembled into NdeI/XhoI digested pIJ10742, which contains a promoterless copy of the reporter gene gusA (Feeney et al., 2017). These plasmids were introduced via intergeneric conjugation into S. scabies (WT, ΔbtmL, and WT + L) using E. coli ET12567/pUZ8002, where they integrated in the ϕBT1 phage integration site. Hygromycin resistant exconjugants were analyzed by colony PCR with primers pGUS_chk_fw and pGUS_chk_fw. β-glucuronidase assays in solid and liquid medium were carried out as described previously (Sherwood and Bibb, 2013). In the case of the liquid assays, reporter activity was represented as Miller units/mg protein. In both cases, pIJ10742 carrying no promoter was used as a negative control, and pIJ10741, carrying gusA under the control of ermE^∗p (Feeney et al., 2017), was included as a positive control of promoter activity.