To identify possible instances of regulatory integration between iron and Trp starvation in C. trachomatis, we optimized a stress response condition that preceded the development of a characteristically persistent phenotype. We reasoned that in order to effectively identify regulatory integration, we would need to investigate the bacterium under stressed, but not aberrant, growth conditions such that we could distinguish primary stress responses from abnormal growth. To specifically investigate the possible contribution of iron limitation to a broader immunological (e.g. IFN-γ-mediated) stress, we utilized the membrane-permeable iron chelator 2,2-bipyridyl (Bpdl), which has the advantage of rapidly and homogeneously starving C. trachomatis of iron (Thompson and Carabeo, 2011). We chose to starve C. trachomatis serovar L2 of iron starting at 12 hr post-infection (hpi), or roughly at the beginning of mid-cycle growth. At this point the chlamydial organisms represent a uniform population of replicative RBs that are fully competent, both transcriptionally and translationally, to respond to stress. We treated infected HeLa cell cultures with 100 μM Bpdl or mock for either 6 or 12 hr (hrs) to determine a condition sufficient to limit iron to C. trachomatis without inducing hallmark persistent phenotypes. We stained infected cells seeded on glass coverslips with convalescent human sera and analyzed chlamydial inclusion morphology under both Bpdl- and mock-treated conditions by laser point-scanning confocal microscopy (Figure 1A). Following 6 hr of Bpdl treatment, chlamydial inclusions were largely indistinguishable from mock-treated inclusions, containing a homogeneous population of larger organisms, consistent with RBs in mid-cycle growth. However, by 12 hr of Bpdl treatment, the inclusions began to display signs of aberrant growth: they were perceptibly smaller, more comparable in size to 18 hpi, and contained noticeably fewer organisms, perhaps indicating a defect in bacterial replication or RB-to-EB differentiation. These observations were consistent with our subsequent analysis of genome replication by quantitative PCR (qPCR; Figure 1B). At 6 hr of Bpdl treatment, there was no statistically distinguishable difference in genome copy number when compared to the equivalent mock-treated time-point. However, by 12 hr of treatment, genome copy number was significantly reduced 4.47-fold in the Bpdl-treated group relative to mock-treatment (p=0.00151). We then assayed the transcript expression of two markers for persistence by reverse transcription quantitative PCR (RT-qPCR): the early gene euo, encoding a transcriptional repressor of late-cycle genes (Figure 1C), and the adhesin omcB, which is expressed late in the developmental cycle (Figure 1D). Characteristic persistence would display elevated euo expression late into infection and suppressed omcB expression throughout development. We observed that at 6 hr of Bpdl treatment, there was no statistically distinguishable difference in either euo or omcB expression when compared to the mock-treatment. Still at 12 hr of Bpdl treatment, euo expression was unchanged. However, omcB expression was significantly up-regulated following 12 hr of Bpdl-treatment (p=0.0024). This was unexpected, but we note that omcB expression has been shown to vary between chlamydial serovars and species when starved for iron (Pokorzynski et al., 2017). The decision to begin our brief iron starvation at 12 hpi may produce notable transcriptional differences from previous studies in which iron starvation was induced at the beginning of the chlamydial developmental cycle, and thereby prevented the establishment of a normal transcriptional program by Chlamydia. Collectively, these data indicated that 6 hr of Bpdl treatment was a more suitable time-point at which to monitor iron-limited stress responses.

Figure 1 with 1 supplement see all

Download asset Open asset

Figure 1—source data 1

Source data for Figure 1A–D, Figure 1—figure supplement 1.

https://doi.org/10.7554/eLife.42295.005

Download elife-42295-fig1-data1-v2.xlsx

We additionally assayed these same metrics following 6 or 12 hr of Trp starvation by culturing cells in either Trp-replete or Trp-depleted DMEM-F12 media supplemented with fetal bovine serum (FBS) pre-dialyzed to remove amino acids. We observed no discernable change in inclusion morphology out to 12 hr of Trp starvation (Figure 1—figure supplement 1A), but genome copy numbers were significantly reduced 2.7-fold at this time-point (p=0.00612; Figure 1—figure supplement 1B). The transcript expression of euo (Figure 1—figure supplement 1C) and omcB (Figure 1—figure supplement 1D) did not significantly change at either treatment duration, but Trp-depletion did result in a 1.88-fold reduction in omcB expression (p=0.0544), consistent with a more characteristic persistent phenotype. These data therefore also indicated that 6 hr of treatment would be ideal to monitor non-persistent responses to Trp limitation.

We next sought to determine whether our brief 6 hr Bpdl treatment was sufficient to elicit a transcriptional iron starvation phenotype. We chose to analyze the expression of three previously identified iron-regulated transcripts, ytgA (Figure 2A), ahpC (Figure 2B) and devB (Figure 2C), by RT-qPCR under Bpdl- and mock-treated conditions (Dill et al., 2009; Thompson and Carabeo, 2011). In addition, we analyzed the expression of one non-iron-regulated transcript, dnaB (Figure 2D), as a negative control (Brinkworth et al., 2018). Following 6 hr of Bpdl treatment, we observed that the transcript expression of the periplasmic iron-binding protein ytgA was significantly elevated 1.75-fold relative to the equivalent mock-treated time-point (p=0.0052). However, we did not observe induction of ytgA transcript expression relative to the 12 hpi time-point. Here, we considered that apparent increases in transcription could be due to two factors: developmental regulation and transcriptional response to stress. Therefore, expression of genes of interest were monitored over time, for example 18 versus 12 hpi, in addition to single-timepoint comparisons, for example 18 hpi only. While we did not observe induction of ytgA over time, which would be more consistent with an iron-starved phenotype (i.e. ‘turning on’ gene expression), we reason that this is a consequence of the brief treatment period. This is in agreement with the need to prolong iron chelation to observe the transcriptional induction of ytgA (Miller et al., 2009; Raulston et al., 2007; Thompson and Carabeo, 2011). Similarly, we observed that the transcript expression of the thioredoxin ahpC was significantly elevated 2.15-fold relative to the equivalent mock-treated time-point (p=0.038) but was not induced relative to the 12 hpi time-point. The modestly elevated expression of these genes likely represents bona fide transcriptional responses to iron starvation given that the treatment condition was optimized to avoid gross changes in chlamydial development. The transcript expression of devB, encoding a 6-phosphogluconolactonase involved in the pentose phosphate pathway, was not observed to significantly respond to our brief iron limitation condition, suggesting that it is not a component of the primary iron starvation stress response in C. trachomatis. As expected, the transcript expression of dnaB, a replicative DNA helicase, was not altered by our iron starvation condition, consistent with its presumably iron-independent regulation (Brinkworth et al., 2018). Overall, these data confirmed that our 6 hr Bpdl treatment condition was suitable to produce a mild iron starvation phenotype at the transcriptional level, facilitating our investigation of iron-dependent regulatory integration.

Figure 2

Download asset Open asset

Figure 2—source data 1

Source data for Figure 2A–D.

https://doi.org/10.7554/eLife.42295.007

Download elife-42295-fig2-data1-v2.xlsx

Upon identifying an iron limitation condition that produced a relevant transcriptional phenotype while avoiding the onset of persistent development, we aimed to investigate whether the immediate response to iron starvation in C. trachomatis would result in the consistent induction of pathways unrelated to iron utilization/acquisition, but nevertheless important for surviving immunological stress. The truncated Trp biosynthetic operon, trpRBA (Figure 3A), has been repeatedly linked to the ability of genital and LGV serovars (D-K and L1-3, respectively) of C. trachomatis to counter IFN-γ-mediated stress. This is due to the capacity of the chlamydial Trp synthase in these serovars to catalyze the β synthase reaction, that is the condensation of indole to the amino acid serine to form Trp (Fehlner-Gardiner et al., 2002). In the presence of exogenous indole, C. trachomatis is therefore able to biosynthesize Trp such that it can prevent the development of IFN-γ-mediated persistence. Correspondingly, the expression of trpRBA is highly induced following IFN-γ stimulation of infected cells (Belland et al., 2003; Østergaard et al., 2016). These data have historically implicated Trp starvation as the primary mechanism by which persistence develops in C. trachomatis following exposure to IFN-γ. However, these studies have routinely depended on prolonged treatment conditions that monitor the terminal effect of persistent development, as opposed to the immediate molecular events which may have important roles in the developmental fate of Chlamydia. As such, these studies may have missed the contribution of other IFN-γ-stimulated insults such as iron limitation.

Figure 3

Download asset Open asset

Figure 3—source data 1

Source data for Figure 3B–C.

https://doi.org/10.7554/eLife.42295.009

Download elife-42295-fig3-data1-v2.xlsx

To decouple Trp limitation from iron limitation and assess their relative contribution to regulating a critical pathway for responding to IFN-γ-mediated stress, we monitored the transcript expression of the trpRBA operon under brief Trp or iron starvation by RT-qPCR. Here again, we analyzed changes in transcript levels at the 18 hpi time-point and between the 12 hpi and 12 hpi + 6 hr time-points. This allowed us to determine if differences in expression could be accounted for by reduced, maintained or induced expression relative to 12 hpi. When starved for Trp for 6 hr, we observed that the expression of trpR, trpB and trpA were all significantly induced greater than 5.18-fold relative to 12 hpi (p=0.0040, 0.020 and 0.0036, respectively; Figure 3B). All three ORFs were also significantly elevated relative to the equivalent mock-treated time-point (p=0.0039, 0.019 and 0.0035, respectively). This result demonstrated that a relatively brief duration of Trp starvation was sufficient to induce trpRBA transcription and highlights the highly attuned sensitivity of C. trachomatis to even moderate changes in Trp levels.

We then performed the same RT-qPCR analysis on the expression of the trpRBA operon in response to 6 hr of iron limitation via Bpdl treatment (Figure 3C). While we observed that the transcript expression of all three ORFs was significantly elevated at least 2.1-fold relative to the equivalent mock-treated time-point (p=0.015, 0.00098 and 0.0062, respectively), we made the intriguing observation that only the expression of trpB and trpA was significantly induced relative to 12 hpi (p=0.00383 and 0.0195, respectively). The marginal elevation in trpR expression at the 18 hpi time-point was surprising given that this gene was not identified as iron-responsive in a recent genome-wide RNA-sequencing study (Brinkworth et al., 2018). Our results suggested that while the trpRBA operon is responsive to iron limitation, trpBA may have a more complex mode of regulation given the additional induction observed relative to trpR, which only maintained expression between 12 hpi and 12 hpi +6 hr Bpdl time-points. Taken together, these findings demonstrate that an important stress response pathway, the trpRBA operon, is regulated by the availability of both Trp and iron, consistent with the notion that the pathway may be cooperatively regulated to respond to various stress conditions. Notably, iron-dependent regulation of Trp biosynthesis has not been previously documented in other prokaryotes.

We hypothesized that the specific iron-related induction of trpBA expression relative to trpR expression may be attributable to an iron-regulated alternative transcriptional start site (alt. TSS) downstream of the trpR ORF. Indeed, a previous study reported the presence of an alt. TSS in the trpRBA IGR, located 214 nucleotides upstream of the trpB translation start position (Carlson et al., 2006). However, a parallel study could not identify a TrpR binding site in the trpRBA IGR (Akers and Tan, 2006). We reasoned that a similar alt. TSS may exist in the IGR that controlled the iron-dependent expression of trpBA. We therefore performed Rapid Amplification of 5’-cDNA Ends (5’-RACE) on RNA isolated from C. trachomatis L2-infected HeLa cells using the SMARTer 5’/3’ RACE Kit workflow (Takara Bio). Given the low expression of the trpRBA operon during normal development, we utilized two sequential gene-specific amplification steps (nested 5’-RACE) to identify 5’ cDNA ends in the trpRBA operon. These nested RACE conditions resulted in amplification that was specific to infected-cells (Figure 4—figure supplement 1A). Using this approach, we analyzed four conditions: 12 hpi, 18 hpi, 12 hpi + 6 hr of Bpdl treatment, and 12 hpi + 6 hr of Trp-depletion (Figure 4A). We observed three RACE products that migrated with an apparent size of 1.5, 1.1 and 1.0 kilobases (kb). At 12 and 18 hpi, all three RACE products exhibited low abundance, even following the nested PCR amplification. This observation was consistent with the expectation that the expression of the trpRBA operon is very low under normal, iron and Trp-replete conditions. However, we note that the 6 hr difference in development did appear to alter the representation of the 5’ cDNA ends, which may suggest a stage-specific promoter utilization within the trpRBA operon. In our Trp starvation condition, we observed an apparent increase in the abundance of the 1.5 kb RACE product, which was therefore presumed to represent the primary TSS upstream of trpR, at nucleotide position 511,389 (C. trachomatis L2 434/Bu). Interestingly, the 1.0 kb product displayed a very similar apparent enrichment following Bpdl treatment, suggesting that this RACE product represented a specifically iron-regulated TSS. Both the 1.5 and 1.0 kb RACE products were detectable in the Trp-depleted and iron-depleted conditions, respectively, during the primary RACE amplification, consistent with their induction under these conditions (Figure 4—figure supplement 1B).

Figure 4 with 2 supplements see all

Download asset Open asset

Figure 4—source data 1

Source data for Figure 4B.

https://doi.org/10.7554/eLife.42295.013

Download elife-42295-fig4-data1-v2.xlsx

If iron depletion was inducing trpBA expression independent of trpR, we reasoned that we would observe specific enrichment of trpB transcripts in our 5’-RACE cDNA samples relative to trpR transcripts. We again utilized RT-qPCR to quantify the abundance of trpB transcripts relative to trpR transcripts in the 5’-RACE total RNA samples (Figure 4B). In agreement with our model, only under iron starved conditions did we observe a significant enrichment of trpB relative to trpR (p<0.01). Additionally, we observed that at 12 and 18 hpi in iron-replete conditions, the ratio of trpB to trpR was approximately 1.0, suggesting non-preferential basal expression across the three putative TSSs. Another factor contributing to this ratio is the synthesis of the full-length trpRBA polycistron. In support of this, the trpB to trpR ratio remained near 1.0 under the Trp-starved condition, which would be expected during transcription read-through of the whole operon. The apparent lack of preferential promoter utilization as described above could be attributed to the relatively low basal expression of the operon at 12 and 18 hpi under Trp- and iron-replete conditions, thus precluding quantitative detection of differential promoter utilization in this assay.

To determine the specific location of the 5’ cDNA ends within the trpRBA operon, we isolated the 5’-RACE products across all conditions by gel extraction and cloned the products into the pRACE vector supplied by the manufacturer. We then sequenced the ligated inserts and BLASTed the sequences against the C. trachomatis L2 434/Bu genome to identify the location of the 5’-most nucleotides (Figure 4C). These data are displayed as a statistical approximation of the genomic regions most likely to be represented by the respective 5’-RACE products in both histogram (semi-continuous) and density plot (continuous) format (See Supplementary file 1 for a description of all mapped 5’-RACE products). As expected, the 1.5 kb product mapped in a distinct and tightly grouped peak near the previously annotated trpR TSS, with the mean and modal nucleotide being 511,388 and 511,389, respectively (Figure 4—figure supplement 2A). Surprisingly, we found that neither the 1.1 or 1.0 kb RACE product mapped to the previously reported alt. TSS in the trpRBA IGR, at position 511,826. Instead, we observed that the 1.1 kb product mapped on average to nucleotide position 511,878, with the modal nucleotide being found at 511,898 (Figure 4—figure supplement 2B). The 1.0 kb product mapped with a mean nucleotide position of 512,013, with the modal nucleotide being 512,005 (Figure 4—figure supplement 2C), only 35 bases upstream of the trpB coding sequence. Interestingly, the 1.0 kb product mapped to a region of the trpRBA IGR flanked by consensus σ⁶⁶ -10 and −35 promoter elements, found at positions 512,020–5 and 511,992–7, respectively (Ricci et al., 1995). In C. trachomatis, σ⁶⁶ is the major housekeeping sigma factor, homologous to E. coli σ⁷⁰. In silico analyses did not reveal the presence of any promoter elements near the 1.1 kb product, however the mean nucleotide position is 50 bp downstream of the previously identified palindrome suspected to have a TrpR operator function (Carlson et al., 2006). These data collectively pointed toward the 1.0 kb 5’-RACE product representing a novel, iron-regulated alt. TSS and bona fide σ⁶⁶-dependent promoter element that allows for the specific iron-dependent expression of trpBA.

As the only known iron-dependent transcriptional regulator in Chlamydia, we hypothesized that YtgR may regulate the iron-dependent expression of trpBA from the putative promoter element we characterized by 5’-RACE. Using bioinformatic sequence analysis, we investigated whether the trpRBA IGR contained a candidate YtgR operator sequence. By local sequence alignment of the putative YtgR operator sequence (Akers et al., 2011) and the trpRBA IGR, we identified a high-identity alignment (76.9% identity) covering 67% of the putative operator sequence (Figure 5A). Interestingly, this alignment mapped to the previously identified palindrome suspected to have operator functionality (Carlson et al., 2006). By global sequence alignment of the YtgR operator to the palindromic sequence, an alignment identical to the local alignment was observed, which still displayed relatively high sequence identity (43.5% identity). We hypothesized that this sequence functioned as a YtgR operator, despite being located 184 bp upstream of the trpBA alt. TSS.

Figure 5 with 2 supplements see all

Download asset Open asset

Figure 5—source data 1

Source data for Figure 5B–E.

https://doi.org/10.7554/eLife.42295.017

Download elife-42295-fig5-data1-v2.xlsx

To investigate the ability of YtgR to bind and repress transcription from the putative trpBA promoter, we implemented a heterologous in vivo two-plasmid assay that reports on YtgR repressor activity as a function of β-galactosidase expression (Thompson et al., 2012). In brief, a candidate DNA promoter element was cloned into the pCCT expression vector between an arabinose-inducible pBAD promoter and the reporter gene lacZ. This plasmid was co-transformed into BL21 (DE3) E. coli along with an IPTG-inducible pET151 expression vector with (pET151-YtgR) or without (pET151-EV) the C-terminal 139 amino acid residues of CTL0325 (YtgC). Note that we have previously demonstrated that this region is a functional iron-dependent repressor domain (Thompson et al., 2012). To verify the functionality of this assay, we determined whether ectopic YtgR expression could repress pCCT reporter gene expression in the presence of three candidate DNA elements: a no-insert empty vector (pCCT-EV), the putative promoter element for C. trachomatis dnaB (pCCT-dnaB), and the promoter region of the ytgABCD operon (pCCT-ytgABCD; Figure 5B). As expected, from the pCCT-EV reporter construct, ectopic YtgR expression did not significantly reduce the activity of β-galactosidase. Additionally, reporter gene expression from pCCT-dnaB, containing the promoter of non-iron-regulated dnaB, was not affected by ectopic expression of YtgR. In contrast, in the presence of pCCT-ytgABCD, induction of YtgR expression produced a significant decrease in β-galactosidase activity (p=0.03868) consistent with its previously reported auto-regulation of this promoter (Thompson et al., 2012).

Using this same assay, we then inserted into the pCCT reporter plasmid 1) the trpR promoter element (pCCT-trpR), 2) the putative trpBA promoter element represented by the IGR (pCCT-trpBA), and 3) the same putative trpBA promoter element with a mutated YtgR operator sequence that was diminished for both palindromicity and A-T richness, two typical features of prokaryotic promoter elements (pCCT-trpBAΔOp; Figure 5C) (Schmitt, 2002; Tao et al., 1992). Note that the trpRBA IGR is >99% conserved at the nucleotide level across urogenital, ocular and LGV serovars of C. trachomatis, and the putative YtgR operator sequence is 100% identical (Figure 5—figure supplement 1). When YtgR was ectopically expressed in the pCCT-trpR background, we observed no statistically distinguishable change in β-galactosidase activity, indicating YtgR could not regulate transcription from the trpR promoter. However, in the pCCT-trpBA background, ectopic YtgR expression significantly reduced β-galactosidase activity at levels similar to those observed in the pCCT-ytgABCD background (p=0.01219). This suggested that YtgR was capable of repressing transcription from the trpBA promoter element specifically. Interestingly, this repression phenotype was abrogated in the pCCT-trpBAΔOp background, where we observed no statistically meaningful difference in β-galactosidase activity, demonstrating that YtgR transcriptional repression of trpBA is operator-dependent. We subsequently addressed whether the region of the trpRBA IGR containing the YtgR operator site was sufficient to confer YtgR repression in this assay (Figure 5—figure supplement 2). We cloned three fragments of the trpRBA IGR into the pCCT reporter plasmid: the first fragment represented the 5’-end of the IGR containing the operator site at the 3’-end (pCCT-IGR1), the second fragment represented a central region of the IGR containing the operator site at the 5’-end (pCCT-IGR2), and the third fragment represented the 3’-end of the IGR and did not contain the operator site (pCCT-IGR3). Surprisingly, we observed that none of these fragments alone were capable of producing a significant repression phenotype in our reporter system.

To verify that YtgR repression of the trpBA promoter element was dependent upon iron availability, we assessed repressor activity in the presence or absence of 500 µM Bpdl, mimicking previously utilized approaches for assessing iron-dependent repressor activity of the homologous DtxR (Ding et al., 1996). YtgR is known to bind cognate DNA elements in an iron-dependent fashion (Thompson et al., 2012), and as such any repressor activity of YtgR should be a direct consequence of its ability to bind DNA in the presence of iron. When we treated co-transformed E. coli expressing YtgR and harboring the pCCT-trpBA reporter plasmid, we observed a modest but statistically significant increase in β-galactosidase activity (Figure 5D; p=0.01409), consistent with the alleviation of YtgR repression at the trpBA promoter element.

To demonstrate that the repression phenotype observed in this reporter system was attributable to DNA-binding of YtgR, we optimized a targeted chromatin immunoprecipitation (ChIP) qPCR method to detect the abundance of co-immunoprecipitated promoter fragments with the recombinant YtgR domain. As before, we co-transformed the pET151-YtgR expression vector with pCCT-trpBA, pCCT-trpBA∆Op, pCCT-trpR or pCCT-dnaB and then fixed the co-transformed cells with formaldehyde prior to immunoprecipitation of the cross-linked 6xHis-YtgR-DNA complexes. Using this system, we observed a significant enrichment of trpBA promoter relative to the dnaB negative control promoter (p=0.0018; Figure 5E). However, enrichment of the trpR promoter was marginal and not statistically distinguishable from that of dnaB. Consistent with its requirement for repression, enrichment of trpBA∆Op was also marginal and statistically indistinguishable from trpR or dnaB, suggesting that mutation of the putative operator sequence alone is sufficient to abrogate YtgR DNA-binding to the trpBA promoter. In conjunction with the IGR fragment analysis, these findings indicated that while the operator site was necessary for YtgR DNA-binding and transcriptional repression, further unknown structural elements in the trpRBA IGR may be required for repression. Nonetheless, this demonstrated the existence of a functional YtgR binding site that conferred transcriptional regulation to trpBA, independent of the major trpR promoter. Collectively, the most parsimonious model derived from these data is that specific and direct iron-dependent DNA-binding of YtgR at the identified operator site acts to repress expression of genes downstream of the trpBA promoter element.

We hypothesized that YtgR binding at the trpRBA YtgR operator site may disadvantage the processivity of RNAP reading-through the IGR from the upstream trpR promoter, possibly leading to transcript termination. Similar systems of RNAP read-through blockage have been reported; the transcription factor Reb1p ‘roadblocks’ RNAPII transcription read-through in yeast by promoting RNAP pausing and subsequent degradation (Colin et al., 2014). To investigate this question, we first returned to RNA-Sequencing data we generated to define the immediate iron-dependent transcriptional regulon in C. trachomatis (Brinkworth et al., 2018). Using data obtained from C. trachomatis-infected HeLa cells at 12 hpi +6 hr mock or Bpdl treatment, we mapped the sequenced reads in batch across three biological replicates to the C. trachomatis L2 434/Bu genome (NC_010287) which we modified to include annotations for non-operonic IGRs. Using this coverage map, we sought to semi-quantitatively assess the mapping of reads across the trpRBA IGR to gain insight regarding possible transcription readthrough. Our analysis revealed that under Bpdl-treated conditions, there was a 2.21-fold increase in reads mapping to the trpRBA IGR (IGR_trpB) relative to mock treatment (Figure 6A). This observation is consistent with the alleviation of YtgR repression under iron-starved conditions permitting readthrough of transcription from the upstream trpR promoter. However, due to high variation and the low number of reads mapping to this region, we were unable to detect a significant difference in coverage (p=0.11) using the genomewide RNA-seq analysis toolkit in the CLC Genomics Workbench. Regardless, a comparable increase in reads mapping to the upstream trpR CDS was not observed (1.54-fold increase, p=0.32), suggesting that under iron replete conditions, transcripts originating from the primary trpR promoter may be terminated before reading through the IGR, thereby accounting for the increase in reads mapping to IGR_trpB following Bpdl treatment.

Figure 6 with 4 supplements see all

Download asset Open asset

We additionally assessed the read coverage of the IGRs upstream of euo (IGR_euo; not iron-regulated) and lpdA (IGR_lpdA; iron-regulated, Brinkworth et al., 2018) which are similarly configured compared to the trpRBA IGR (i.e. the IGRs are between two ORFs in the same coding orientation). For both IGR_euo (1.17-fold, p=0.58) and IGR_lpdA (1.40-fold, p=0.37), we did not observe a similar increase in read coverage following Bpdl treatment indicating that the increased coverage at IGR_trpB is likely specific (Figure 6—figure supplement 1A–B). Qualitatively, we note that only IGR_trpB displays relatively uniform read coverage across the defined region, while both IGR_euo and IGR_lpdA have non-uniform coverage, reinforcing the idea that the reads mapping to IGR_trpB originate upstream in the trpR ORF and readthrough the entire region, thus presenting the opportunity for premature termination. Furthermore, the upstream ORFs for IGR_euo and IGR_lpdA did not display robust fold-increases; recJ, upstream of IGR_euo, was 1.29-fold increased (p=0.17) while CTL0819, upstream of IGR_lpdA, was 1.13-fold increased (p=0.56). We extracted individual RPKM values for each of these regions from this dataset and observed the same trend in mean fold-change differences reported from the genomewide analysis: only IGR_trpB was substantially increased relative to its upstream ORF (Figure 6—figure supplement 2). Despite being informative, genomewide RNA-Seq is ultimately insufficient to elucidate particular mechanistic details of transcriptional regulation in the trpRBA IGR and therefore we turned to more sensitive and quantitative methods to investigate possible transcript termination within the trpRBA IGR.

To identify transcription termination sites (TTSs) in the trpRBA operon in C. trachomatis, we utilized 3’-RACE to map the 3’-ends of transcripts using gene-specific primers within the trpR CDS (Figure 6B; lower panel). We again utilized two RACE amplification cycles to generate distinct, specific bands suitable for isolation and sequencing (Figure 6—figure supplement 3B–C). By gel electrophoresis of the 3’-RACE products, we observed the appearance of four distinct bands that migrated with an apparent size of 0.55, 0.45, 0.40 and 0.20 kb. In our Trp-depleted condition, we observed only a very weak amplification of the 2.5–3 kb full-length trpRBA message by 3’-RACE (Figure 6—figure supplement 3A). However, we did observe it across all replicates. To confirm that the full-length product was specific to the Trp-depleted treatment, we amplified the trpRBA operon by RT-PCR from the 3’-RACE total RNA (Figure 6B; upper panel). As expected, only in the Trp-depleted sample did we observe robust amplification of the full-length trpRBA message. We note however that image contrast adjustment reveals a very weak band present in all experimental samples. Therefore, the specific 3’-RACE analysis identified novel transcription termination events within the trpRBA operon.

To identify the specific TTS locations, we gel extracted the four distinct 3’-RACE bands across all conditions and cloned them into the pRACE sequencing vector as was done for the 5’-RACE experiments. We then sequenced the inserted RACE products and mapped them to the C. trachomatis L2 434/Bu genome (Figure 6C). This revealed a highly dynamic TTS landscape within the trpRBA IGR, which has not previously been investigated (For a full description of mapped 3’-RACE products, see Supplementary file 2). The 0.20 kb RACE product mapped to the 3’-end of the trpR CDS, with a mean nucleotide position of 511,665 and a modal nucleotide position of 511,667 (Figure 6—figure supplement 4A). Contrastingly, the other three 3’-RACE products did not map in such a way so as to produce specific, unambiguous modal peaks. Instead, their distribution was broader and more even, with only a few nucleotide positions mapping more than once. Accordingly, the 0.45 kb product mapped with an average nucleotide position of 511,889, just downstream of the 1.1 kb 5’-RACE product (Figure 6—figure supplement 4C), while the 0.55 kb product mapped with an average nucleotide position of 511,986, upstream of the 1.0 kb 5’-RACE product (Figure 6—figure supplement 4D). Interestingly, the 0.40 kb product mapped to a region directly overlapping the putative YtgR operator site, with a mean nucleotide position of 511,810 (Figure 6—figure supplement 4B). We therefore reasoned that this putative TTS may have an iron-dependent function controlled by YtgR.

We hypothesized that transcript termination at the YtgR operator site was regulated in an iron-dependent manner by YtgR binding to the operator DNA and blocking upstream transcription readthrough. Under iron-replete conditions, YtgR would be bound to the operator DNA and transcript termination would occur at a greater frequency, preventing readthrough of the transcription machinery initiated at the upstream trpR promoter. Under iron-depleted conditions, the inactivation of YtgR DNA-binding activity would allow the transcription machinery to readthrough the YtgR operator site to the downstream sequence, including trpBA. To test this model, we utilized RT-qPCR to quantify the amount of readthrough at the YtgR operator site in iron-depleted C. trachomatis-infected HeLa cells and in our two-plasmid E. coli system where the expression of YtgR could be controlled.

We tested whether nutrient availability altered the extent of readthrough in the IGR at the YtgR operator site using a RT-qPCR-based quantification of various mRNA species (Figure 7A). Levels of each intermediate transcript species indicated by unique non-overlapping amplicons were reported as ratios against a common upstream amplicon. Thus, for amplicons downstream of termination sites, a higher downstream amplicon-to-common amplicon ratio would indicate increased readthrough. Note that this value has a theoretical limit of 1.0, where all transcripts would be at least as long as the downstream amplicon being measured. We designed primers to amplify regions immediately 5’ and 3’ to the YtgR operator and its corresponding TTS. Additionally, we analyzed the very 3’ end of trpA, which would be expected to only monitor complete full-length transcripts as well as alternative transcription initiation from the trpBA promoter.

Figure 7 with 1 supplement see all

Download asset Open asset

Figure 7—source data 1

Source data for Figure 7B and D–E, Figure 7—figure supplement 1.

https://doi.org/10.7554/eLife.42295.025

Download elife-42295-fig7-data1-v2.xlsx

First, we utilized our infected cell-culture model to assess the iron-dependency of transcription readthrough at the YtgR operator site. As before, we assayed four conditions: 12 hpi, 18 hpi, 12 hpi +6 hr Bpdl and 12 hpi +6 hr Trp-depletion. Under normal developmental conditions, we expected that there would not be dramatic changes in readthrough, though we note that changes in development or nutrient availability could have this affect under normal conditions. Under Trp-depleted conditions, we predicted that the inactivation of the TrpR repressor would produce a robust readthrough phenotype that, in accordance with the literature and evidence provided herein, would transcribe the full-length trpRBA message. Thus, Trp-depletion served as a positive control for our readthrough analysis. We hypothesized that under iron-depleted conditions, we would not observe a significant increase in readthrough 5’ of the YtgR operator site, but when YtgR is inactivated, we should be able to detect an increase in readthrough 3’ of the operator site as the transcription machinery is no longer blocked by the bound repressor and is permitted to continue transcription into the downstream sequence.

Consistent with our model, readthrough 5’ of the YtgR operator site was not affected by Bpdl treatment, whereas Trp-depletion resulted in significantly more readthrough than all other conditions (Figure 7B; p<0.05 for all comparisons). However, 3’ of the YtgR operator site, Bpdl treatment significantly increased readthrough compared to 12 and 18 hpi (p<0.05 for all comparisons), suggesting that inactivation of YtgR by iron depletion alleviates transcription termination at this site. We performed this same analysis with an alternative amplicon 3’ of the YtgR operator site and observed a similar increase in readthrough, confirming that the effect was not unique to the amplicon we used (Figure 7—figure supplement 1; p<0.005 for all comparisons). When this readthrough analysis is applied 3’ of the alternative trpB TSS, we observe that following Bpdl-treatment, the readthrough value substantially exceeds 1.0, consistent with transcription of trpBA independent of trpR under this condition. Moreover, only the Bpdl-treated group is significantly different from the other treatment conditions (p<0.05 for all comparisons). Notably, Trp-depletion did not increase the readthrough value, suggesting that Trp-depletion does not relieve YtgR repression at the alt. TSS and the levels of trpA remain relatively constant to those of trpR. It is therefore possible that under Trp-depleted, but iron-replete conditions, YtgR repression of the alternative trpBA promoter acts as a rheostat for full-length trpRBA transcription, helping to maintain a constant ratio of trpR to trpBA. Together, these analyses indicated that iron limitation resulted in transcription readthrough specifically at the YtgR operator site.

To investigate whether or not this readthrough phenotype was dependent upon YtgR, we again turned to our heterologous two-plasmid reporter system. We designed a reporter vector (pCCT-RT) that harbored the entire trpR-IGR DNA sequence such that any transcription initiated at the upstream arabinose-inducible pBAD promoter would have to readthrough the entire trpR ORF and the IGR before reaching the reporter gene lacZ (Figure 7C). Note that this is functionally similar to expression of the trpRBA operon under Trp-starved conditions: the major Trp-dependent promoter upstream of trpR would be activated and initiate readthrough independent of the presence of YtgR bound downstream in the IGR. As such, we performed the same RT-qPCR readthrough analysis on RNA harvested from BL21 (DE3) E. coli co-transformed with pCCT-RT and either pET151-EV or pET151-YtgR. We observed that ectopic expression of YtgR significantly reduced readthrough into the lacZ gene by RT-qPCR (Figure 7D; p=0.003561), consistent with YtgR DNA-binding specifically inhibiting readthrough via a mechanism of transcript termination at the operator site. In support of this result, we additionally observed a significant decrease in β-galactosidase activity as measured by the Miller assay from the pCCT-RT vector when YtgR was ectopically expressed (Figure 7E; p=0.01723), indicating that YtgR-dependent inhibition of readthrough limits the expression of downstream genes. In sum, YtgR binding to the trpRBA IGR at the predicted operator site concomitantly represses transcription from the alternative trpBA promoter while blockading transcription readthrough from the upstream trpR promoter, ultimately rendering the expression of trpBA susceptible to changes in iron availability. Importantly, alleviated transcript termination at the YtgR operator site may offer a mechanistic explanation for the moderate elevation in trpR expression observed following iron starvation, as enhanced readthrough at the YtgR operator site under this condition may produce more stable mRNA species relative to normally developing C. trachomatis. Thus, YtgR may function as an iron-dependent attenuator of trpRBA expression.

Human cervical epithelial adenocarcinoma HeLa (ATCC CCL-2; Purchased 08/2016; Last tested for Mycoplasma 07/2018) cells were cultured at 37° C with 5% atmospheric CO₂ in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10 μg/mL gentamicin, 2 mM L-glutamine, and 10% (v/v) filter sterilized fetal bovine serum (FBS). For all experiments, HeLa cells were cultured between passage numbers 4 and 16. Chlamydia trachomatis serovar L2 (434/Bu) was originally obtained from Dr. Ted Hackstadt (Rocky Mountain National Laboratory, NIAID). Chlamydial EBs were isolated from infected HeLa cells at 36–40 hr post-infection (hpi) and purified by density gradient centrifugation essentially as described (Caldwell et al., 1981).

For the infection of 6-well tissue culture plates, HeLa cells cultured to 80–90% confluency were first washed with pre-warmed Hanks Buffered Saline Solution (HBSS) prior to the monolayer being overlaid with inoculum (un-supplemented DMEM) at the indicated multiplicity of infection (MOI). Tissue culture plates were then centrifuged at 4° C with a speed of 1000 RPM (Eppendorf 5810 R table top centrifuge, A-4–81 rotor) for 5 min to synchronize the infection. Inoculum was aspirated and cells were washed again with pre-warmed HBSS prior to the media being replaced with pre-warmed complete DMEM. Infected cultures were then returned to the tissue culture incubator until the indicated times post-infection. This procedure was replicated exactly for the infection of 24-well tissue culture plates.

Chlamydia trachomatis L2-infected HeLa cell cultures were starved for iron by supplementation of the media with the iron chelator 2,2-bipyridyl (Bpdl; Sigma Aldrich, St. Louis, MO, USA; CAS: 366-18-7) essentially as described (Thompson and Carabeo, 2011). Briefly, at the indicated times post-infection, infected cell cultures were washed with pre-warmed HBSS prior to the addition of complete DMEM (mock) or complete DMEM supplemented with 100 μM Bpdl. Infected cell cultures were returned to the incubator for the indicated treatment periods. Bpdl was prepared as a 100 mM stock solution in 100% ethanol and stored at −20° C for no longer than 6 months.

Chlamydia trachomatis L2-infected HeLa cell cultures were starved for tryptophan by replacement of complete DMEM with tryptophan-depleted medium. In brief, Tryptophan-replete or –deplete DMEM-F12 (U.S. Biological Life Sciences, Salem, MA, USA) powder media was prepared following manufacture instructions and supplemented with 10% (v/v) filter-sterilized FBS which had been previously dialyzed 16–20 hr at 4° C in PBS in a 10 kDa MWCO dialysis cassette. Media was then further supplemented with 10 μg/mL gentamicin. At the indicated times post-infection, complete DMEM was aspirated and wells were washed with pre-warmed HBSS prior to the addition of tryptophan-replete or –deplete medium. Infected cell cultures were returned to the incubator for the indicated treatment periods.

All constructs were cloned using standard molecular cloning techniques, e.g. restriction enzyme, homology-directed, etc. All primers and plasmids used in this study can be found in Supplementary file 5 and 6, respectively. All pCCT constructs were cloned by amplifying the promoter region of interest with 5’ and 3’ flanking KpnI sites, which were then KpnI digested (New England Biolabs, Ipswich, MA, USA) along with the pCCT-ytgA vector (to excise the ytgA promoter fragment). The vector was then treated with antarctic phosphatase (New England Biolabs) prior to having the promoter of interest ligated into the pCCT backbone. Inserted promoters were verified to be in the correct orientation and free of sequence errors by PCR and sequencing. All pET vectors were cloned following manufacturer instructions (Invitrogen, ThermoFisher Scientific, Waltham, MA, USA).

At the indicated times post-infection, C. trachomatis L2-infected HeLa cell cultures seeded on glass coverslips in 24-well tissue cultures plates were first washed with pre-warmed HBSS prior to fixation with 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS) for 20 min at RT ° C. Fixation solution was aspirated and wells were washed with PBS prior to permeabilization with 0.2% Triton X-100 in PBS for 5 min at RT° C. Permeabilization solution was then decanted and cells were washed with PBS. The coverslips were blocked for 30 min with 1% bovine serum albumin (BSA) in PBS at RT° C. To stain for Chlamydia, coverslips were washed with PBS and PBS supplemented with 1% BSA and 1:500 convalescent human sera was added to wells and incubated at RT° C for 1 hr with rocking. Primary antibody solution was decanted and coverslips were again washed with PBS. Goat anti-human Alexa-647 (Invitrogen, ThermoFisher Scientific) diluted 1:1000 in PBS with 1% BSA was then added to the wells and incubated in the dark for another hour at RT° C with rocking. Secondary antibody solution was then decanted, coverslips were washed again with PBS and coverslips were either immediately mounted on microscopy slides using ImmuMount (ThermoFisher Scientific) or VectaShield H-1000 (Vector Laboratories, Burlingame, CA, USA) or stored in the dark at 4° C until mounting. All images were acquired on a Leica TCS SP8 laser scanning confocal microscope, using identical settings, in the Integrative Physiology and Neuroscience Advanced Imaging Center at Washington State University. All images are Z-projections and were processed in Fiji (Schindelin et al., 2012) and Adobe Creative Suite identically for each comparative time-point.

RNA was harvested from C. trachomatis-infected HeLa cell monolayers by scraping 3 wells of a 6-well plate in ice-cold Trizol Reagent (ThermoFisher Scientific). Samples were then pooled and split into two technical replicates (RT-qPCR) or kept as one biological replicate (RACE). Trizol-extracted samples were then thoroughly vortexed with a 100 μL volume of Zirconia beads prior to chloroform extraction. 100% ethanol was added to the aqueous phase and RNA was isolated using the Ambion RiboPure RNA Purification kit for bacteria following manufacturer instructions (ThermoFisher Scientific). DNA was removed from RNA samples using the Invitrogen DNA-free DNA Removal Kit following manufacturer instructions (ThermoFisher Scientific). RNA was stored at −20° C until further use. For E. coli, RNA was harvested using the Ambion RiboPure RNA Purification kit for bacteria following manufacturer instructions (ThermoFisher Scientific) from 9 mL of bacterial culture prepared as described below for the Two-Plasmid Reporter Assay. RNA was subsequently DNased as described above. cDNA was generated using either SuperScript IV Reverse Transcriptase (RT-qPCR; ThermoFisher Scientific) or SMARTScribe Reverse Transcriptase (RACE and RACE-specific qRT-PCR); Takara Bio, Kusatsu, Shiga Prefecture, Japan) essentially as described by the respective manufacturers. For cDNA generated for RT-qPCR, 650 ng of total RNA was used as a template in a 20 μL total reaction volume. For every RT reaction, a ‘no-RT’ control, generated from 350 ng of total RNA template in a 10 μL total volume, was included. For 5’-RACE, cDNA was generated from 250 ng of total RNA using random primers in a 10 μL total volume and further processed in the RACE workflow. cDNA was stored at −20° C.

gDNA was harvested from C. trachomatis-infected HeLa cell monolayers by scraping 3 wells of a 6-well plate in ice-cold PBS + 10% Proteinase K (ThermoFisher Scientific). Samples were then pooled and split into two technical replicates for analysis of genome copy number by qPCR. gDNA was isolated using the DNeasy Blood and Tissue Kit following manufacturer protocols (QIAGEN, Hilden, Germany). gDNA was stored at −20° C until further use.

Reverse Transcription Quantitative Polymerase Chain Reaction (RT-qPCR) cDNA (or gDNA in qPCR), prepared as described above, was diluted 1:10 or 1:100 in nuclease-free H₂O depending on the experimental condition being assayed (e.g. treatment, point in development cycle, etc.). On ice, 3.3 μL of diluted sample was added to 79 μL of PowerUp SYBR Green Master Mix (ThermoFisher Scientific) with specific qPCR primers diluted to 500 nM. From this master mix, each experimental sample was assayed in triplicate 25 μL reactions. Assays were run on an Applied Biosystems 7300 Real Time PCR System with cycling conditions as follows: Stage 1: 50.0° C for 2 min, one rep. Stage 2: 95.0° C for 10 min, one rep. Stage 3: 95.0° C for 15 s, 40 reps. Stage 4: 60.0° C for 1 min, one rep. Primers were subjected to dissociation curve analysis to ensure that a single product was generated. For each primer set, a standard curve was generated using purified C. trachomatis L2 gDNA from EB preparations diluted from 2 × 10⁻³ to 2 × 10⁰ ng per reaction. C_t values generated from each experimental reaction were then fit to standard curves (satisfying an efficiency of 95 ± 5%) for the respective primer pair and from the calculated ng quantities, transcript or genome copy number was calculated as follows:

Where df = dilution factor and the number of copies/ng DNA is calculated based on the size of the C. trachomatis L2 genome assuming that the molar mass per base pair is 650 (g/mol)/bp (note that this value should be the same for any single-copy ORF on the genome). All quantifications of genome copy number were determined using the ahpC qPCR primer set. Values from replicate assays were averaged, and values from replicate RNA/gDNA isolations were averaged to obtain the mean and standard deviation for one biological replicate. For some experiments, to account for batch effects across biological replicates, data was transformed such that the mean of all samples in each replicate was identical. In some instances, batch correcting generated negative values, and in this case data sets were scaled such that the lowest value equaled 1.0. HeLa cells were infected at an MOI of 2 for all RT-qPCR studies.For analysis of transcriptional readthrough, RT-qPCR was performed as described above and the readthrough value was computed as:

Readthrough values were then batch corrected such that the mean of each replicate was identical.

All RACE studies were performed using the SMARTer RACE 5’/3’ Kit (Takara Bio). To observe 5’-RACE products from the trpRBA operon, a ‘nested’ RACE protocol was used as outlined in the SMARTer RACE 5’/3’ Kit user manual. Briefly, 1.25–2.5 μL of cDNA generated for RACE was added to a 25 μL reaction volume and run in a thermal cycler for 40 cycles using the touch-down PCR conditions described by the manufacturer. In brief, five cycles were run at an annealing temperature of both 72° C and 70° C prior to 30 cycles run with an annealing temperature of 68° C. Following this primary amplification, the RACE products were diluted 1:50 in Tricine-EDTA Buffer supplied by the manufacturer, and 2.5 μL of diluted primary RACE product was added to a 25 μL reaction volume and subjected to another 20 cycles of nested PCR, as described by the manufacturer, using primers designed within the amplicon of the primary RACE products. Samples were electrophoresed on a 2% agarose gel for visualization and analysis. HeLa cells were infected at a MOI of 5 for all RACE studies.

3’-RACE studies were performed essentially identical to 5’-RACE with the exception that total RNA was subjected to poly(A) tailing with a Poly(A) Polymerase following manufacturer instructions (New England Biolabs). In brief, at least 3.5 μg of total RNA was incubated at 37° C with Poly(A) Polymerase in reaction buffer supplemented with ATP and murine RNase Inhibitor (New England Biolabs) for 30 min prior to heat-inactivation at 65° C for 20 min. RNA was re-isolated through an RNA clean-up filter cartridge (Ambion, ThermoFisher Scientific). A total of 125 ng of poly(A)-tailed total RNA was then used to generate 3’-RACE ready cDNA in a 10 μL reaction volume following manufacturer instructions. Primary and nested RACE was performed using 3’-RACE gene-specific primers following the same protocol for amplification described for 5’-RACE, with the exception that the extension time was adjusted to accommodate amplification of the full ~3 kb trpRBA polycistronic message.

5’-RACE products generated from either primary or nested RACE reactions were excised from the agarose gel and DNA was isolated using the NucleoSpin Gel and PCR Clean-up kit (Macherey-Nagel, Takara Bio). The isolated RACE products were then cloned into the pRACE vector supplied in the SMARTer RACE 5’/3’ Kit using the In-Fusion HD cloning kit (Takara Bio). Ligated vectors were transformed into chemically competent Stellar E. coli cells by heat shock. Transformed bacteria were plated on LB agar containing 50 μg/mL carbenicillin and incubated overnight at 37° C. Colonies were selected and screened for relevant inserts by PCR. Positive colonies were cultured overnight at 37° C in LB liquid broth containing 50 μg/mL carbenicillin and plasmids were isolated using the QIAprep Spin Miniprep kit (QIAGEN). Inserts were then sequenced by Eurofins Genomics using the default M13 Reverse sequencing primer. Returned sequencing data was aligned to the C. trachomatis L2 (434/Bu) genome (NCBI Accession: NC_010287) by BLAST and the most 5’ aligned nucleotide was considered the 5’ end of the insert. In the case of 3’-RACE data, the reverse complement sequence was first generated prior to alignment. Grouping of individual products was determined 1.) by clusters being greater than 30 nucleotides apart and 2.) by the specific RACE band that the alignment was derived from. These two criteria were not both satisfied in all cases and in those cases criteria 1.) was favored.

All C. trachomatis L2 434/Bu genome sequences were obtained from NCBI Accession NC_010287. Global pairwise sequence alignments were made using the EMBOSS Needle algorithm. Alignment parameters were set as follows: Matrix: DNAfull, Gap Open: 20, Gap Extend: 0.8, Output Format: pair, End Gap Penalty: True, End Gap Open: 10, End Gap Extend: 0.5. These conditions were sufficient to replicate the previously published alignment between the putative YtgR operator sequence and the TroR operator (Akers et al., 2011). Local pairwise sequence alignments were made using the EMBOSS Water algorithm. The putative YtgR operator was aligned to the entire 348 bp intergenic region of the trpRBA operon (C. trachomatis L2 [434/Bu] genome position 511,692–512,039). The alignment parameters were set as follows: Matrix: DNAfull, Gap Open:10, Gap Extend: 0.5, Output Format: pair. These are the default conditions and were chosen to remove bias from the alignment results.

The YtgR-binding reporter assay was performed essentially as described, with minor modifications (Thompson et al., 2012). Promoter regions of interest were amplified from the C. trachomatis L2 (434/Bu) genome by PCR using the indicated primer sets, which included KpnI restriction endonuclease sites at the 5’ and 3’ ends of the promoter amplicon. The amplified fragments and the pCCT-EV plasmid were then KpnI-digested and the promoters ligated into the vector using T4 or Quick Ligase (New England BioLabs). Insert directionality was confirmed by directional colony PCR and positive clones were sequence verified. pCCT-trpBAΔOperator was cloned by amplifying two fragments of the pCCT-trpBA vector with one ~ 60 mer primer containing the bases to be substituted for each fragment. Thus, the whole vector was split into two half-fragments containing the substituted bases. The two fragments were then cloned back together using In-Fusion Homology-Directed cloning (Takara Bio) to yield the final vector. Electrocompetent BL21(DE3) E. coli (Sigma Aldrich) were co-transformed by electroporation with the pCCT reporter plasmid and the pET151 expression vector (-EV or –YtgR) and plated on double selective LB agar containing 50 μg/mL carbenicillin and 15 μg/mL tetracycline. Prior to plating of transformed cells, 50 μL of 40 mg/mL X-Gal in DMSO (EMD Millipore, Burlington, MA, USA) was applied to the plate for colorimetric determination of β-galactosidase expression. Transformants were incubated overnight at 37° C. The following evening, blue colonies from each experimental condition were selected and cultured overnight in LB liquid broth containing 0.2% (w/v) D-glucose (for catabolite repression of expression vectors), 50 μg/mL carbenicillin and 15 μg/mL tetracycline. Cultures were incubated overnight at 37° C. The following morning, overnight cultures were spun down to remove glucose-containing media and sub-cultured in LB liquid broth medium containing 50 μM FeSO₄, 50 μg/mL carbenicillin and 15 μg/mL tetracycline to an OD₆₀₀ of 0.45. Cultures were incubated for 1 hr at 37° C and sub-cultured a second time in the same media to an OD₆₀₀ of 0.1. Cultures were returned to the incubator for another hour prior to the addition of 500 μM isopropyl β-D-1-thiogalactopyranoside (IPTG) to induce pET151 expression from the lac promoter. Cultures were incubated another hour prior to the addition of 0.2% L-arabinose to induce lacZ expression from the araBAD promoter. Cultures were incubated a final 2 hr prior to the collection of a 0.1 mL volume of cells for assaying β-galactosidase activity by the Miller Assay (Miller, 1972). Cell pellets were stored at −80° C prior to being assayed. To assay β-galactosidase activity, cell pellets were first re-suspended in Z-buffer (pH 7.0, 60 mM Na₂HPO₄, 40 mM NaH₂PO₄, 10 mM KCl, 1 mM MgSO₄ and 2.7 μL/mL β-mercaptoethanol). 50 μL of 0.1% SDS and 100 μL of chloroform were then added to each sample prior to thorough vortexing. Samples were equilibrated for 5 min at 30° C and 200 μL of 4 mg/mL ortho-nitrophenyl-β-galactoside (ONPG) prepared in Phosphate Buffer (pH 7.0, 60 mM Na₂HPO₄, 40 mM NaH₂PO₄) were added to the samples to initiate the reaction. Reactions were stopped by the addition of 500 μL 1 M Na₂CO₃. Absorbance was measured on a FLUOStar Optima plate reader (BMG Labtech, Offenburg, Germany) at 420 nm and Miller Units were calculated as:

Where t = reaction time, v = volume of cells and OD_final = OD₆₀₀ at the time of sample collection. It was empirically determined that the subtraction of absorbance at 550 nm had a negligible effect on the calculated value. A blank sample lacking cells was included in each experimental batch and used as a reference for absorbance. For each experimental condition, three independent co-transformed colonies were assayed in technical triplicate. In some instances, significantly high Miller Unit outliers were excluded by Grubb’s Test (p<0.05) under the assumption that extreme lacZ expression may reflect plasmid copy number or reporter gene expression issues.

RNA-Sequencing experiments were performed as described in their original publication (Brinkworth et al., 2018). Raw and processed sequencing files were submitted to the NCBI Gene Expression Omnibus (GEO) as a Superseries and can be found at accession number GSE106763. Coverage maps were generated by mapping all reads across three biological replicates to a single reference file in CLC Genomics Workbench v11. To facilitate easy analysis of IGR boundaries, the C. trachomatis L2 434/Bu genome (Accession: NC_010287) was modified to contain annotations for intergenic regions that fell between two genes in the same coding orientation, and this genome was used as the reference for read mapping. Read mapping and differential expression analysis was performed using default settings in CLC Genomics Workbench. Data aggregation in the Reads track was set to aggregate above 1 bp.

All graphs were generated using the ggplot2 package (Wickham, 2009) in R Studio, and/or in the Adobe Creative Suite. All line plots and bar graphs represent the mean ± one standard deviation unless otherwise noted. All box and whisker plots represent the distribution of data between the 1^st and 3^rd quartile range within the box, while the whiskers represent data within 1.5 interquartile ranges of the 1^st or 3^rd quartile. Extreme values outside this range are plotted as open circles. The 2^nd quartile (median) is plotted as a black line within the box. Histogram plots were generated with a bin width of 20 and are plotted on a density scale. The overlaid density plots represent a statistical approximation of the data over a continuous scale. All statistical analyses were carried out in R Studio. All statistical computations were performed on the mean values of independent biological replicates calculated from the indicated number of respective technical replicates. For single pairwise comparisons, a two-sided unpaired Student’s t-test with Welch’s correction for unequal variance was used to determine statistical significance. For multiple pairwise comparisons, a One-Way Analysis of Variance (ANOVA) was conducted to identify significant differences within groups. If a significant difference was detected, then the indicated post-hoc pairwise test was used to identify the location of specific statistical differences. A p-value less than 0.05 was considered statistically significant. For all figures, *=p < 0.05, **=p < 0.01, and ***=p < 0.005.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Liam Caven, Korinn Murphy and Matthew Romero for critical review of this manuscript; Dr. Christopher C Thompson for the establishment of the E. coli YtgR reporter system and generation of the pCCT construct; Dr. Scot P Ouellette for support, critical feedback and advice; and Dr. David Dewitt for expert advice, training and maintenance of equipment in the IPN Advanced Imaging Center. This work was supported by NIH grants R01-AI065545 and R01-AI132406 to RAC; F31-AI136295 and T32-GM008336 to NDP; NDP was also supported by an Achievement Reward for College Scientists (ARCS; Seattle Chapter) Fellowship.

© 2019, Pokorzynski et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.