ORIGINAL RESEARCH article
Defining the Transcriptional and Post-transcriptional Landscapes of Mycobacterium smegmatis in Aerobic Growth and Hypoxia
- 1Department of Biology and Biotechnology, Worcester Polytechnic Institute, Worcester, MA, United States
- 2Program in Bioinformatics and Computational Biology, Worcester Polytechnic Institute, Worcester, MA, United States
The ability of Mycobacterium tuberculosis to infect, proliferate, and survive during long periods in the human lungs largely depends on the rigorous control of gene expression. Transcriptome-wide analyses are key to understanding gene regulation on a global scale. Here, we combine 5′-end-directed libraries with RNAseq expression libraries to gain insight into the transcriptome organization and post-transcriptional mRNA cleavage landscape in mycobacteria during log phase growth and under hypoxia, a physiologically relevant stress condition. Using the model organism Mycobacterium smegmatis, we identified 6,090 transcription start sites (TSSs) with high confidence during log phase growth, of which 67% were categorized as primary TSSs for annotated genes, and the remaining were classified as internal, antisense, or orphan, according to their genomic context. Interestingly, over 25% of the RNA transcripts lack a leader sequence, and of the coding sequences that do have leaders, 53% lack a strong consensus Shine-Dalgarno site. This indicates that like M. tuberculosis, M. smegmatis can initiate translation through multiple mechanisms. Our approach also allowed us to identify over 3,000 RNA cleavage sites, which occur at a novel sequence motif. To our knowledge, this represents the first report of a transcriptome-wide RNA cleavage site map in mycobacteria. The cleavage sites show a positional bias toward mRNA regulatory regions, highlighting the importance of post-transcriptional regulation in gene expression. We show that in low oxygen, a condition associated with the host environment during infection, mycobacteria change their transcriptomic profiles and endonucleolytic RNA cleavage is markedly reduced, suggesting a mechanistic explanation for previous reports of increased mRNA half-lives in response to stress. In addition, a number of TSSs were triggered in hypoxia, 56 of which contain the binding motif for the sigma factor SigF in their promoter regions. This suggests that SigF makes direct contributions to transcriptomic remodeling in hypoxia-challenged mycobacteria. Taken together, our data provide a foundation for further study of both transcriptional and posttranscriptional regulation in mycobacteria.
Tuberculosis is a disease of global concern caused by Mycobacterium tuberculosis (Mtb). This pathogen has the ability to infect the human lungs and survive there for long periods, often by entering into non-growing states. During infection, Mtb must overcome a variety of stressful conditions, including nutrient starvation, low pH, oxygen deprivation and the presence of reactive oxygen species. Consequently, the association of Mtb with its host and the adaptation to the surrounding environment requires rigorous control of gene expression.
As the slow growth rate and pathogenicity of Mtb present logistical challenges in the laboratory, many aspects of its biology have been studied in other mycobacterial species. One of the most widely used models is Mycobacterium smegmatis, a non-pathogenic fast-growing bacterium. While there are marked differences between the genomes of Mtb and M. smegmatis, such as the highly represented PE/PPE-like gene category and other virulence factors present in Mtb and poorly represented or absent in M. smegmatis, these organisms have at least 2,117 orthologous genes (Prasanna and Mehra, 2013) making M. smegmatis a viable model to address certain questions about the fundamental biology of mycobacteria. Indeed, studies using M. smegmatis have revealed key insights into relevant aspects of Mtb biology including the Sec and ESX secretion systems involved in transport of virulence factors (Coros et al., 2008; Rigel et al., 2009), bacterial survival during anaerobic dormancy (Dick et al., 1998; Bagchi et al., 2002; Trauner et al., 2012; Pecsi et al., 2014) and the changes induced during nutrient starvation (Elharar et al., 2014; Wu et al., 2016; Hayashi et al., 2018). However, the M. smegmatis transcriptome has been less extensively studied than that of Mtb.
Identification of transcription start sites (TSSs) is an essential step toward understanding how bacteria organize their transcriptomes and respond to changing environments. Genome-wide TSS mapping studies have been used to elucidate the general transcriptomic features in many bacterial species, leading to the identification of promoters, characterization of 5′ untranslated regions (5′ UTRs), identification of RNA regulatory elements and transcriptional changes in different environmental conditions (examples include (Albrecht et al., 2009; Mitschke et al., 2011; Cortes et al., 2013; Schlüter et al., 2013; Dinan et al., 2014; Ramachandran et al., 2014; Sass et al., 2015; Shell et al., 2015b; Thomason et al., 2015; Berger et al., 2016; Čuklina et al., 2016; D’arrigo et al., 2016; Heidrich et al., 2017; Li et al., 2017). To date, two main studies have reported the transcriptomic landscape in Mtb during exponential growth and carbon starvation (Cortes et al., 2013; Shell et al., 2015b). These complementary studies revealed that, unlike most bacteria, a substantial percentage (∼25%) of the transcripts are leaderless, lacking a 5′ UTR and consequently a Shine-Dalgarno ribosome-binding site. In addition, a number of previously unannotated ORFs encoding putative small proteins were found (Shell et al., 2015b), showing that the transcriptional landscape can be more complex than predicted by automated genome annotation pipelines. Thus, TSS mapping is a powerful tool to gain insight into transcriptomic organization and identify novel genes. Less is known about the characteristics of the M. smegmatis transcriptome. A recent study reported a number of M. smegmatis TSSs in normal growth conditions (Li et al., 2017). However, this work was limited to identification of primary gene-associated TSSs and lacked an analysis of internal and antisense TSSs, as well as characterization of promoter regions and other relevant transcriptomic features. In addition, Potgieter et al. (2016) validated a large number of annotated ORFs using proteomics and were able to identify 63 previously unannotated leaderless ORFs.
To achieve a deeper characterization of the M. smegmatis transcriptional landscape, we combined 5′-end-mapping and RNAseq expression profiling under two different growth conditions. Here we present an exhaustive analysis of the M. smegmatis transcriptome during exponential growth and hypoxia. Unlike most transcriptome-wide TSS analyses, our approach allowed us to study not only the transcriptome organization in different conditions, but also the frequency and distribution of RNA cleavage sites on a genome wide scale. Whereas regulation at the transcriptional level is assumed to be the main mechanism that modulates gene expression in bacteria, post-transcriptional regulation is a key step in the control of gene expression and has been implicated in the response to host conditions and virulence in various bacterial pathogens (Kulesekara et al., 2006; Mraheil et al., 2011; Heroven et al., 2012; Schifano et al., 2013; Holmqvist et al., 2016). Here we show that the predominant RNA cleavage sequence motif in M. smegmatis is distinct from what has been reported for other bacteria. We also show that RNA cleavage decreases during adaptation to hypoxia, suggesting that RNA cleavage may be a refinement mechanism contributing to the regulation of gene expression in harsh conditions.
Materials and Methods
Strains and Growth Conditions Used in This Study
Mycobacterium smegmatis strain mc2155 was grown in Middlebrook 7H9 supplemented with ADC (Albumin Dextrose Catalase, final concentrations 5 g/L bovine serum albumin fraction V, 2 g/L dextrose, 0.85 g/L sodium chloride, and 3 mg/L catalase), 0.2% glycerol and 0.05% Tween 80. For the exponential phase experiment (Dataset 1), 50 ml conical tubes containing 5 ml of 7H9 were inoculated with M. smegmatis to have an initial OD = 0.01. Cultures were grown at 37°C and 200 rpm. Once cultures reached an OD of 0.7–0.8, they were frozen in liquid nitrogen and stored at -80°C until RNA purification. For hypoxia experiments (Dataset 2), a protocol similar to the Wayne model (Wayne and Hayes, 1996) was implemented. Briefly, 60 ml serum bottles (Wheaton, product number 223746, actual volume to top of rim 73 ml) were inoculated with 36.5 ml of M. smegmatis culture with an initial OD = 0.01. The bottles were sealed with rubber caps (Wheaton, W224100-181 Stopper, 20 mm) and aluminum caps (Wheaton, 20 mm aluminum seal) and cultures were grown at 37°C and 125 rpm to generate hypoxic conditions. Samples were taken at an early stage of oxygen depletion when growth had slowed but not completely stopped (15 h) and at a later stage when a methylene blue indicator dye was fully decolorized and growth had ceased (24 h). These time points were experimentally determined according to growth curve experiments (see Supplementary Figure S1). 15 ml of each culture were sampled and frozen immediately in liquid nitrogen until RNA extraction.
RNA was extracted as follows: frozen cultures stored at -80°C were thawed on ice and centrifuged at 4,000 rpm for 5 min at 4°C. The pellets were resuspended in 1 ml Trizol (Life Technologies) and placed in tubes containing Lysing Matrix B (MP Bio). Cells were lysed by bead-beating twice for 40 s at 9 m/sec in a FastPrep 5G instrument (MP Bio). 300 μl chloroform was added and samples were centrifuged for 15 min at 4,000 rpm at 4°C. The aqueous phase was collected and RNA was purified using Direct-Zol RNA miniprep kit (Zymo) according to the manufacturer’s instructions. Samples were then treated with DNase Turbo (Ambion) for 1 h and purified with an RNA Clean & Concentrator-25 kit (Zymo) according to the manufacturer’s instructions. RNA integrity was checked on 1% agarose gels and concentrations were determined using a Nanodrop instrument. Prior to library construction, 5 μg RNA was used for rRNA depletion using Ribo-Zero rRNA Removal Kit (Illumina) according to the manufacturer’s instructions.
Construction of 5′-End-Mapping Libraries
After rRNA depletion, RNA samples from each biological replicate were split in three, in order to generate two 5′-end differentially treated libraries and one RNAseq expression library (next section). RNA for library 1 (“converted” library) was treated either with RNA 5′ pyrophosphohydrolase RPPH (NEB) (exponential phase experiment, Dataset 1), or with 5′ polyphosphatase (Epicenter) (hypoxia experiment, Dataset 2), in order to remove the native 5′ triphosphates of primary transcripts, whereas RNA for Library 2 (“non-converted” library) was subject to mock treatment. Thus, the converted libraries capture both 5′ triphosphates (converted to monophosphates) and native 5′ monophosphate transcripts, while non-converted libraries capture only native 5′ monophosphates (see scheme in Supplementary Figure S2A). Library construction was performed as described by Shell et al. (2015a). A detailed scheme showing the workflow of 5′-end libraries construction, the primers and adapters used in each step, and modifications to the protocol are shown in Supplementary Figure S2B.
Construction of RNAseq Expression Libraries
One third of each rRNA-depleted RNA sample was used to construct RNAseq expression libraries. KAPA stranded RNA-Seq library preparation kit and NEBNext Ultra RNA library prep kit for Illumina (NEB) were used for Dataset 1 and Dataset 2, respectively, according to manufacturer’s instructions. The following major modifications were introduced into the protocols: (i) For RNA fragmentation, in order to obtain fragments around 300 nt, RNA was mixed with the corresponding buffer and placed at 85°C for 6 min (Dataset 1), or at 94°C for 12 min (Dataset 2). (ii) For library amplification, 10 or 19–23 PCR cycles were used for Dataset 1 and Dataset 2, respectively. The number of cycles was chosen according to the amount of cDNA obtained for each sample. After purification, DNA concentration was measured in a Qubit instrument before sequencing.
Libraries Sequencing and Quality Assessment
For 5′-end-mapping libraries from Dataset 1, Illumina MiSeq paired-end sequencing producing 100 nt reads was used. For 5′ end directed libraries from Dataset 2 as well as for all expression libraries, Illumina HiSeq 2000 paired-end sequencing producing 50 nt reads was used. Sequencing was performed at the UMass Medical School Deep Sequencing Core Facility. Quality of the generated fastq files was checked using FastQC.
Identification of 5′ Ends and Discrimination Between Transcription Start Sites (TSSs) and Cleavage Sites (CSs)
Paired-end reads generated from 5′-end-directed libraries were mapped to the M. smegmatis mc2155 NC_008596 reference genome. In order to reduce noise from the imprecision of transcriptional initiation, only the coordinate with the highest coverage in each 5 nt window was used for downstream analyses. For read filtering, different criteria were used for the 2 datasets according to the library depth and quality (see Supplementary Figure S3). In order to discriminate between TSSs and CSs, the ratio of the coverage in converted/non-converted libraries for each detected 5′ end was calculated. To focus our analyses on the 5′ends that are relatively abundant in their local genomic context, we employed a filter based on the ratio of 5′ end coverage to expression library coverage in the preceding 100 nt. 5′ ends for which this ratio was ≤0.05 were removed. After this filter, 15,720 5′ ends remained and were further analyzed using a Gaussian mixture modeling to differentiate TSSs from CSs with a high confidence in Dataset 1 (Figure 1A). For this analysis, we used the iterative expectation maximization (EM) algorithm in the mixtools package (Benaglia et al., 2009) for R (version 1.1.0) to fit the mixture distributions.
Figure 1. Mapping and categorization of transcription start sites in M. smegmatis. (A) Diagram showing the ratios of coverage in the converted/non-converted libraries for each coordinate. Gaussian mixture modeling was used to discriminate between TSSs and CSs. For this analysis, the 15,720 coordinates from Dataset 1 were used. (B) Abundance of the ANNNT promoter motif located between bases –13 to –6 upstream of the 15,720 coordinates. The light blue dashed line indicates the percentage of coordinates in the genome of M. smegmatis that have at least one ANNNT motif located between bases –13 to –6 upstream (9.7%). (C) Base frequency at the +1 position among the 15,720 5′ ends from Dataset 1. (D) Categories for TSS annotation based on the genomic context. TSSs were classified according to their relative position to genes as primary (pTSSs, red), internal (iTSSs, green), antisense (aTSSs, light blue) and orphan (oTSSs, violet). (E) Distribution of TSSs among the different categories.
Analysis of Expression Libraries
Reads were aligned to the Mycobacterium smegmatis str. mc2155 reference genome (Accession number NC_008596) using Burrows-Wheeler Aligner (Li and Durbin, 2009). For comparison of gene expression levels according to presence or absence of Shine-Dalgarno sequences, RPKMs were calculated for all genes. The DEseq2 pipeline was used to evaluate the changes in gene expression in hypoxia (Love et al., 2014).
Transcription Start Sites Categorization
For analysis in Figure 1D, TSSs were classified as follows: those coordinates located ≤500 bp upstream from an annotated gene were considered to be primary TSSs (pTSS). Coordinates located within an annotated gene were classified as internal (iTSS) or N-associated internal TSSs (N-iTSSs) if they were located within the first 25% of the annotated coding sequence. N-iTSSs were considered for reannotation as a pTSSs only if their associated gene lacked a pTSS. TSSs located on the antisense strand of a coding sequence, 5′ UTR, or 3′ UTR were considered as antisense (aTSS). 5′ UTRs boundaries were assigned after assignment of pTSSs to genes annotated in the mc2155 reference genome (accession number NC_008596). When a gene had more than one pTSS, the longest of the possible 5′ UTRs was used for assignment of aTSSs. In the case of genes for which we did not identify a pTSS, we considered a hypothetical leader sequence of 50 bp for assignment of aTSSs. For assignment of aTSSs in 3′ UTRs, we arbitrarily considered a sequence of 50 bp downstream the stop codon of the gene to be the 3′ UTR. Finally, TSSs not belonging to any of the above-mentioned categories were classified as orphan (oTSSs).
Adjacent genes with the same orientation were considered to be co-transcribed if there were at least 5 spanning reads between the upstream and the downstream gene in at least one of the replicates in the expression libraries from Dataset 1. After this filtering, a downstream gene was excluded from the operon if: (1) it had a TSS ≤ 500 bp upstream the annotated start codon on the same strand, and/or (2) had a TSS within the first 25% of the gene on the same strand, and/or (3) the upstream gene had a TSS within the last 50–100% of the coding sequence. Finally, the operon was assigned only if the first gene had a primary TSS with a confidence ≥95% according to the Gaussian mixture modeling.
Cleavage Sites Categorization
For CS categorization in Figure 4D, we established stringent criteria in order to determine the frequency of CSs in each location category relative to the amount of the genome comprising that location category. For 3′ UTR regions, we considered only CSs that were located between 2 convergent genes. To assess frequency relative to the whole genome, we considered the sum of all regions located between two convergent genes. For 5′ UTRs we considered all CSs located between 2 divergent genes, and the sum of all leader lengths for genes having a pTSS whose upstream gene is in the opposite strand (divergent) determined in this study was used for assessing relative frequency. For 5′ ends corresponding to cleavages between co-transcribed genes we used the operon structures determined in this study, and the sum of all their intergenic regions was used for assessing relative frequency. Finally, for CSs located within coding sequences all genes were considered, as all of them produced reads in the expression libraries. The sum of all coding sequences in NC_008596 genome was used for assessing relative frequency, after subtracting overlapping regions to avoid redundancy.
5′ RACE (Rapid Amplification of cDNA Ends)
For validation of TSSs and CSs, RNA samples from M. smegmatis were split in two and treated with or without RPPH (NEB) in order to remove the native 5′ triphosphates of primary transcripts or not, respectively. Then, an adapter oligo SSS1016 (CTGGAGCACGAGGACACTGACATGGACTGAAGGAGTrArGrArArA, where nts preceded by “r” are ribonucleotides and the rest of the oligo is composed of deoxyribonucleotides) was ligated to the RNA 5′ ends using T4 RNA ligase (NEB). Prior to ligation, 8 μl of RNA sample were combined with 1 μl of 1 μg/μl adapter oligo and incubated at 65°C for 10 min. For ligation, the 9 μl of RNA-oligo mix were combined with: 10 μl 50% PEG8000, 3 μl 10X ligase buffer, 3 μl 10 mM ATP, 3 μl DMSO, 1 μl Murine RNase inhibitor (NEB), and 1 μl T4 ligase (NEB). Ligation reactions were incubated at 20°C overnight and then cleaned using RNA Clean and Concentrator 25 kit (Zymo). Both RPPH-treated and mock-treated samples were used for cDNA synthesis. Reactions in absence of reverse transcriptase were performed to control for genomic DNA contamination. For amplification of specific 5′ ends, PCR was done using a forward primer SSS1017 binding to the adapter oligo (CTGGAGCACGAGGACACTGA) and a reverse (specific) primer binding near the predicted 5′ end (see Supplementary Table S1). For PCRs, a touchdown protocol in which the annealing temperature was reduced 1°C every cycle was performed as follows: (i) initial step of DNA denaturation at 95°C for 5 min, (ii) 17 cycles of 95°C for 30 s, 72–55°C (touchdown) for 20 s and 68°C for 25 s, (iii) 20 cycles of 95°C for 30 s, 55°C for 20 s and 68°C for 25 s and (iv) a final elongation step at 68°C for 5 min. Each amplified fragment was sequenced using the specific primer. A TSS or CS was validated if (i) the 5′ end position coincided with that mapped the 5′ end libraries and (ii) the PCR product was more abundant in the RPPH than in the no RPPH treatment (TSS) or the PCR product was equally abundant in the RPPH and in the no RPPH treatment (CS).
For validation of the MSMEG_0063 promoter, an M. smegmatis mutant strain lacking the region comprising the genes MSMEG_0062-MSMEG_0066 was transformed with either of the 3 following constructs: (i) Wt promoter, which has the gene MSMEG_0063 with the native predicted promoter region and the downstream genes MSMEG_0064-MSMEG_0066, (ii) Δpromoter, in which the predicted promoter region for MSMEG_0063 was deleted, and (iii) Mutated promoter, in which two point mutations were introduced in the predicted -10 region of the MSMEG_0063 promoter. These constructs were inserted in the L5 site of the M. smegmatis genome.
Mapping, Annotation, and Categorization of Transcription Start Sites
In order to study the transcriptome structure of M. smegmatis, RNAs from triplicate cultures in exponential phase were used to construct 5′ end mapping libraries (Dataset 1) according to our previously published methodologies (Shell et al., 2015a,b) with minor modifications. Briefly, our approach relies on comparison of adapter ligation frequency in a dephosphorylated (converted) library and an untreated (non-converted) library for each sample. The converted libraries capture both 5′ triphosphate and native 5′ monophosphate-bearing transcripts, while the non-converted libraries capture only native 5′ monophosphate-bearing transcripts (Supplementary Figure S2). Thus, assessing the ratios of read counts in the converted/non-converted libraries permits discrimination between 5′ triphosphate ends (primary transcripts from transcription start sites) and 5′ monophosphate ends (cleavage sites). By employing a Gaussian mixture modeling analysis (Figure 1A) we were able to identify 5,552 TSSs in M. smegmatis with an observed probability of being a TSS ≥ 0.95 (high confidence TSSs, Supplementary Table S2). A second filtering method allowed us to obtain 222 additional TSSs from Dataset 1 (Supplementary Figure S3). A total of 5,774 TSSs were therefore obtained from Dataset 1. In addition, data from separate libraries constructed as controls for the hypoxia experiment (Dataset 2) in “The Transcriptional Landscape Changes in Response to Oxygen Limitation’ were also included in this analysis to obtain TSSs. After noise filtering (Supplementary Figure S3), 4,736 TSSs from Dataset 2 were identified. The union of the two datasets yielded a total of 6,090 non-redundant high confidence TSSs, of which 4,420 were detected in both datasets (Supplementary Figure S4 and Supplementary Table S2).
Although not all 5′ ends could be classified with the Gaussian mixture modeling, we were able to assign 57% of the 5′ ends in Dataset 1 to one of the two 5′ end populations with high confidence (5,552 TSSs and 3,344 CSs). To validate the reliability of the Gaussian mixture modeling used to classify 5′ ends, we performed two additional analyses. First, according to previous findings in Mtb (Cortes et al., 2013) and other well studied bacteria (Sass et al., 2015; Berger et al., 2016; Čuklina et al., 2016; D’arrigo et al., 2016), we anticipated that TSSs should be enriched for the presence of the ANNNT -10 promoter consensus motif in the region upstream. Evaluation of the presence of appropriately-spaced ANNNT sequences revealed that 5′ ends with higher probabilities of being TSSs are enriched for this motif, whereas for those 5′ ends with low probabilities of being TSSs (and thus high probabilities of being CSs) have ANNNT frequencies similar to that of the M. smegmatis genome as a whole (Figure 1B). Secondly, we predicted that TSSs should show enrichment for A and G nts at the +1 position, given the reported preference for bacterial RNA polymerases to initiate transcription with these nts (Lewis and Adhya, 2004; Mendoza-Vargas et al., 2009; Mitschke et al., 2011; Cortes et al., 2013; Shell et al., 2015b; Thomason et al., 2015; Berger et al., 2016). Thus, we analyzed the base enrichment in the +1 position for the 5′ ends according to the p-value in the Gaussian mixture modeling (Figure 1C). These results show a clear increase in the percentage of G and A bases in the position +1 as the probability of being a TSS increases, while the percentage of sequences having a C at +1 increases as the probability of being a TSS decreases. These two analyses show marked differences in the sequence contexts of TSSs and CSs and further validate the method used for categorization of 5′ ends.
To study the genome architecture of M. smegmatis, the 6,090 TSSs were categorized according to their genomic context (Figures 1D,E and Supplementary Table S2). TSSs located ≤500 nt upstream of an annotated gene start codon in the M. smegmatis str. mc2155 (accession NC_008596) reference genome were classified as primary TSSs (pTSS). TSSs within annotated genes on the sense strand were denoted as internal (iTSS). When an iTSS was located in the first quarter of an annotated gene, it was sub-classified as N-terminal associated TSS (N-iTSS), and was further examined to determine if it should be considered a primary TSS (see below). TSSs located on the antisense strand either within a gene or within a 5′ UTR or 3′ UTR were grouped as antisense TSSs (aTSSs). Finally, TSSs located in non-coding regions that did not meet the criteria for any of the above categories were classified as orphan (oTSSs). When a pTSS also met the criteria for classification in another category, it was considered to be pTSS for the purposes of downstream analyses. A total of 4,054 distinct TSSs met the criteria to be classified as pTSSs for genes transcribed in exponential phase. These pTSSs were assigned to 3,043 downstream genes, representing 44% of the total annotated genes (Supplementary Table S3). This number is lower than the total number of genes expressed in exponential phase, in large part due to the existence of polycistronic transcripts (see operon prediction below). Interestingly, 706 (23%) of these genes have at least two pTSSs and 209 (7%) have three or more, indicating that transcription initiation from multiple promoters is common in M. smegmatis. We used 5′ RACE to confirm seven selected pTSSs (Supplementary Table S1), all of which mapped to the same position by both methods. Four of these were novel TSSs not reported by Li et al. (2017).
A total of 995 iTSSs (excluding the iTSSs that were also classified as a pTSS of a downstream gene, see Supplementary Figure S5 for classification workflow) were identified in 804 (12%) of the annotated genes, indicating that transcription initiation within coding sequences is common in M. smegmatis. iTSSs are often considered to be pTSSs of downstream genes, to be spurious events yielding truncated transcripts, or to be consequences of incorrect gene start annotations. However, there is evidence supporting the hypothesis that iTSSs are functional and highly conserved among closely related bacteria (Shao et al., 2014), highlighting their potential importance in gene expression.
We were also able to detect antisense transcription in 12.5% of the M. smegmatis genes. Antisense transcription plays a role in modulation of gene expression by controlling transcription, RNA stability, and translation (Morita et al., 2005; Kawano et al., 2007; Andre et al., 2008; Fozo et al., 2008; Giangrossi et al., 2010) and has been found to occur at different rates across bacterial genera, ranging from 1.3% of genes in Staphylococcus aureus to up to 46% of genes in Helicobacter pylori (Beaume et al., 2010; Sharma et al., 2010). Of the 1,006 aTSSs identified here (excluding those that were primarily classified as pTSSs), 881 are within coding sequences, 120 are within 5′ UTRs and 72 are located within 3′ UTRs (note that some aTSS are simultaneously classified in more than one of these three subcategories, Supplementary Figure S6). While we expect that many of the detected antisense transcripts have biological functions, it is difficult to differentiate antisense RNAs with regulatory functions from transcriptional noise. In this regard, Lloréns-Rico et al. (2016) reported that most of the antisense transcripts detected using transcriptomic approaches are a consequence of transcriptional noise, arising at spurious promoters throughout the genome. To investigate the potential significance of the M. smegmatis aTSSs, we assessed the relative impact of each aTSS on local antisense expression levels by comparing the read depth upstream and downstream of each aTSS in our RNAseq expression libraries. We found 318 aTSSs for which expression coverage was ≥10-fold higher in the 100 nt window downstream of the TSS compared to the 100 nt window upstream (Supplementary Table S4). Based on the magnitude of the expression occurring at these aTSS, we postulate that they could represent the 5′ ends of candidate functional antisense transcripts rather than simply products of spurious transcription. However, further work is needed to test this hypothesis. Finally, 78 oTSSs were detected across the M. smegmatis genome. These TSSs may be the 5′ ends of non-coding RNAs or mRNAs encoding previously unannotated ORFs.
Out of the 995 iTSSs identified, 457 were located within the first quarter of an annotated gene (N-iTSSs). In cases where we could not predict a pTSS with high confidence, we considered the possibility that the start codon of the gene was misannotated and the N-iTSS was in fact the primary TSS. Although we do not discount the possibility that functional proteins can be produced when internal transcription initiation occurs far downstream of the annotated start codon, we only considered N-iTSSs candidates for gene start reannotation when there was a start codon (ATG, GTG, or TTG) in-frame with the annotated gene in the first 30% of the annotated sequence. In this way, we suggest re-annotations of the start codons of 213 coding sequences (see Supplementary Figure S5 and Supplementary Table S5). These N-iTSSs were considered to be pTSSs (N-iTSSs → pTSSs) for all further analyses described in this work.
To predict operon structure, we combined 5′ end libraries and RNAseq expression data. We considered two or more genes to be co-transcribed if (1) they had spanning reads that overlapped both the upstream and downstream gene in the expression libraries, (2) at least one TSS was detected in the 5′ end-directed libraries for the first gene of the operon, and (3) the downstream gene(s) lacked pTSSs and iTSSs (for more detail, see section “Materials and Methods”). Thus, we were able to identify and annotate 294 operons with high confidence across the M. smegmatis genome (Supplementary Table S6). These operons are between 2 and 4 genes in length and comprise a total of 638 genes. Our operon prediction methodology has some limitations. For example, operons not expressed in exponential growth phase could not be detected in our study. Furthermore, internal promoters within operons can exist, leading to either monocistronic transcripts or suboperons (Guell et al., 2009; Paletta and Ohman, 2012; Skliarova et al., 2012). We limited our operon predictions to genes that appear to be exclusively co-transcribed, excluding those cases in which an internal gene in an operon can be alternatively transcribed from an assigned pTSS. Finally, our analysis did not capture operons in which the first gene lacked a high-confidence pTSS. Despite these limitations, our approach allowed us to successfully identify new operons as well as previously described operons. Previously reported operons that were captured by our predictions included the furA-katG (MSMEG_6383-MSMEG_6384) operon involved in oxidative stress response (Milano et al., 2001), the vapB-vapC (MSMEG_1283-MSMEG_1284) Toxin–Antitoxin module (Robson et al., 2009) operon, and the ClpP1-ClpP2 (MSMEG_4672-MSMEG_4673) operon involved in protein degradation (Raju et al., 2012).
Characterization of M. smegmatis Promoters Reveals Features Conserved in M. tuberculosis
Most bacterial promoters have two highly conserved regions, the -10 and the -35, that interact with RNA polymerase via sigma factors. However, it was reported that the -10 region is necessary and sufficient for transcription initiation by the housekeeping sigma factor SigA in mycobacteria, and no SigA -35 consensus motifs were identified in previous studies (Cortes et al., 2013; Newton-Foot and Gey van Pittius, 2013; Li et al., 2017; Zhu et al., 2017). To characterize the core promoter motifs in M. smegmatis on a global scale we analyzed the 50 bp upstream of the TSSs. We found that 4,833 of 6,090 promoters analyzed (79%) have an ANNNT motif located between positions -6 to -13 upstream the TSSs (Figure 2A). In addition, 63% of the promoters with ANNNT motifs have a thymidine preceding this sequence (TANNNT). This motif is similar to that previously described in a transcriptome–wide analysis for Mtb (Cortes et al., 2013) and for most bacterial promoters that are recognized by the σ70 housekeeping sigma factor (Ramachandran et al., 2014; Sass et al., 2015; Berger et al., 2016; Čuklina et al., 2016; D’arrigo et al., 2016). However, no apparent bias toward specific bases in the NNN region was detected in our study or in Mtb, while in other bacteria such as Escherichia coli, Salmonella enterica, Burkholderia cenocepacia, Pseudomonas putida, and Bacillus subtilis an A/T preference was observed in this region (Jarmer et al., 2001; Ramachandran et al., 2014; Sass et al., 2015; Berger et al., 2016; D’arrigo et al., 2016). We were unable to detect a consensus motif in the -35 region either using MEME server (Bailey et al., 2015) or manually assessing the possible base-enrichment in the -35 region. Analysis of the sequences in the immediate vicinity of TSSs revealed that G and A are the most frequent bases at the +1 position, and C is considerably more abundant at -1 (Figure 2B).
Figure 2. M. smegmatis promoter -10 regions are dominated by the ANNNT motif. (A) Identification of promoter motifs. Consensus motifs were identified by using MEME. The 20 nt upstream the 6,090 TSSs were used for the initial analysis. Those sequences lacking an ANNNT –10 motif between positions –13 and –6 (1,257) were used to identify other conserved promoter sequences. Motif 2 (20 nt length) and Motif 4 (18 nt length) are located immediately upstream of the TSS (at the –1 position), while the spacing of Motif 5 varies from –4 to –1 relative to the TSS, with –3 being the dominant position (75% of the motifs). (B) The sequences flanking 3,500 randomly chosen TSSs were used to create a sequence logo by WebLogo 3 (Crooks et al., 2004), revealing the two dominant spacings for the ANNNT motif and base preferences in the immediate vicinity of the TSS. (C) Comparison of apparent promoter activity for different motifs. Mean normalized read depth in the converted libraries from Dataset 1 was compared for TSSs having or lacking the ANNNT motif in the –10 region, and ANNNT-associated TSSs were further subdivided into those containing the extended TANNNT motif or conversely the VANNNT sequence (where V = A, G or C). Motifs 2, 4, and 5 in Figure 2A are also included. ∗∗∗∗p < 0.0001, ∗∗∗p < 0.001, ∗∗p < 0.01, ∗p < 0.05 (Kruskal–Wallis test with post-test for multiple comparisons).
Interestingly, we identified several alternative motifs in the -10 promoter regions of transcripts lacking the ANNNT motif (Figure 2A). One of these, (G/C)NN(G/C)NN(G/C), is likely the signature of M. smegmatis’ codon bias in the regions upstream of iTSSs. The other three sequences are candidate binding sites for alternative sigma factors, which are known to be important in regulation of transcription under diverse environmental conditions. However, the identified consensus sequences differ substantially from those previously described in mycobacteria (Raman et al., 2001, 2004; Sun et al., 2004; Lee et al., 2008a,b; Song et al., 2008; Veyrier et al., 2008; Humpel et al., 2010; Gaudion et al., 2013). The TSSs having these sigma factor motifs and the associated genes are listed in Supplementary Table 7. We next examined the relationship between promoter sequence and promoter strength, as estimated by the read depths in the 5′ end converted libraries. As shown in Figure 2C, the expression levels of transcripts with ANNNT -10 motifs are on average substantially higher than those lacking this sequence. In addition, promoters with the full TANNNT motif are associated with more highly abundant transcripts compared to those having a VANNNT sequence, where V is G, A or C. These results implicate TANNNT as the preferred -10 sequence for the housekeeping sigma factor, SigA, in M. smegmatis. As shown in Figure 2C, expression levels of transcripts having the motif 2 in Figure 2A were significantly increased when compared to the total pool of transcripts lacking the ANNNT motif.
Leaderless Transcription Is a Prominent Feature of the M. smegmatis Transcriptome
5′ UTRs play important roles in post-transcriptional regulation and translation, as they may contain regulatory sequences that can affect mRNA stability and/or translation efficiency. Whereas in most bacteria 5′ UTR-bearing (“leadered”) transcripts predominate, this is not the case for Mtb, in which near one quarter of the transcripts have been reported to be leaderless (Cortes et al., 2013; Shell et al., 2015b). To investigate this feature in M. smegmatis, we analyzed the 5′ UTR lengths of all genes that had at least one pTSS. We found that for 24% of the transcripts the TSS coincides with the translation start site or produces a leader length ≤5 nt, resulting in leaderless transcripts (Figure 3A). This is less than the 40% reported for M. smegmatis in a smaller TSS-mapping study (Li et al., 2017), and suggests that the proportions of leaderless transcripts are in fact similar for M. smegmatis and Mtb. A total of 1,099 genes (including those re-annotated in section “Mapping, Annotation, and Categorization of Transcription Start Sites”) have leaderless transcripts, and 155 of those (14%) are also transcribed as leadered mRNAs from separate promoters. Two of the pTSSs we validated by 5′ RACE (Supplementary Table S1) belong to leaderless transcripts. For leadered transcripts, the median 5′ UTR length was 69 nt. Interestingly, 15% of the leaders are >200 nt, suggesting that these sequences may contain potential regulatory elements. We then sought to compare the leader lengths of M. smegmatis genes with the leader lengths of their homologs in Mtb. For this analysis we used two independent pTSS mapping Mtb datasets obtained from Cortes et al. (2013) and Shell et al. (2015b) (Figure 3B). To avoid ambiguities, we used only genes that had a single pTSS in both species. Our results show a statistically significant correlation of leader lengths between species, suggesting that similar genes conserve their transcript features and consequently may have related regulatory mechanisms. Additionally, comparison of leaderless transcription in M. smegmatis and Mtb revealed that 62% or 73% of the genes that are only transcribed as leaderless in M. smegmatis also lack a 5′ UTR in MTB, according to Cortes et al. (2013) or Shell et al. (2015b), respectively (Supplementary Table S8). We next assessed if leaderless transcripts are associated with particular gene categories, and found the distribution across categories was uneven (Figure 3C). The three categories “DNA metabolism,” “Amino acid biosynthesis,” and “Biosynthesis of cofactors, prosthetic groups and carriers” were significantly enriched in leaderless transcripts (p-value < 0.05, hypergeometric test), while “Signal transduction,” “Transcription,” and “Transport and binding proteins” appear to have fewer leaderless transcripts.
Figure 3. Leader features are conserved in mycobacteria. (A) Leader length distribution. The 4,054 pTSSs and the pTSSs of the 213 reannotated genes (N-iTSSs → pTSSs) were used. (B) Leader length correlation between M. smegmatis and Mtb genes. The leader sequences of genes having a single unique pTSS in both species (leader length ≥ 0 and ≤500 nt) were used. 508 homologous genes in Cortes et al. (2013) (left figure) and 251 homologous genes in Shell et al. (2015b) (right figure) were used. When a gene in M. smegmatis had more than one homolog in Mtb, that with the highest identity was considered. Spearman r p-value < 0.00001 in both cases. (C) Distribution of leaderless transcripts among different functional TIGRfam functional categories (Haft et al., 2001). 557 genes having TIGRfam categories were used for this analysis. Genes having both leadered and leaderless transcripts were excluded. The black dashed line indicates the expected proportion of leaderless genes (25%) according to the global analysis performed in this study. The numbers above each bar indicate the total number of genes used for this analysis in each category (leaderless + leadered). ∗∗∗∗p < 0.0001, ∗∗∗p < 0.001 (Chi-Square test with Bonferroni correction for multiple comparisons). (D) RNA levels vary according to leader status. Mean expression levels were compared for genes expressed with leaders containing a canonical SD sequence (SD) or not (No SD) or lacking leaders (leaderless). Gene expression was quantified by RNAseq. Genes were classified as containing an SD sequence if at least one of the three tetramers AGGA, GGAG, or GAGG (core sequence AGGAGG) were present in the region –6 to –17 nt relative to the start codon. rRNAs, tRNAs, sRNAs, and genes expressed as both leadered and leaderless transcripts were excluded. ∗∗∗∗p < 0.0001, ∗∗p < 0.005; ns: not significant. (Kruskal–Wallis test with post-test for multiple comparisons).
We next evaluated the presence of the Shine-Dalgarno ribosome-binding site (SD) upstream of leadered coding sequences. For this analysis, we considered those leaders containing at least one of the three tetramers AGGA, GGAG or GAGG (core sequence AGGAGG) in the region -6 to -17 relative to the start codon to possess canonical SD motifs. We found that only 47% of leadered coding sequences had these canonical SD sequences. Thus, considering also the leaderless RNAs, a large number of transcripts lack canonical SD sequences, suggesting that translation initiation can occur through multiple mechanisms in M. smegmatis. We further compared the relative expression levels of leaderless and leadered coding sequences subdivided by SD status. Genes expressed as both leadered and leaderless transcripts were excluded from this analysis. We found that on average, expression levels were significantly higher for those genes with canonical SD sequences than for those with leaders but lacking this motif and for those that were leaderless (Figure 3D). Together, these data suggest that genes that are more efficiently translated have also higher transcript levels. Similar findings were made in Mtb, where proteomic analyses showed increased protein levels for genes with SD sequences compared to those lacking this motif (Cortes et al., 2013).
Identification of Novel Leaderless ORFs in the M. smegmatis Genome
As GTG or ATG codons are sufficient to initiate leaderless translation in mycobacteria (Shell et al., 2015b; Potgieter et al., 2016), we used this feature to look for unannotated ORFs in the M. smegmatis NC_008596 reference genome. Using 1,579 TSSs that remained after pTSS assignment and gene reannotation using N-iTSSs (see Supplementary Figure S5) we identified a total of 66 leaderless ORFs encoding putative proteins longer than 30 amino acids, 5 of which were previously identified (Shell et al., 2015b). 83% of these ORFs were predicted in other annotations of the M. smegmatis mc2155 or MKD8 genome [NC_018289.1, (Gray et al., 2013)], while 10 of the remaining ORFs showed homology to genes annotated in other mycobacterial species and Helobdella robusta and two ORFs did not show homology to any known protein. The TSS of ORF15 was validated by 5′RACE. These results show that automatic annotation of genomes can be incomplete and highlight the utility of transcriptomic analysis for genome (re)annotation. Detailed information on these novel putative ORFs is provided in Supplementary Table S9.
Endonucleolytic RNA Cleavage Occurs at a Distinct Sequence Motif and Is Common in mRNA Regulatory Regions
As our methodology allows us to precisely map RNA cleavage sites in addition to TSSs, we sought to analyze the presence and distribution of cleavage sites in the M. smegmatis transcriptome. mRNA processing plays a crucial role in regulation of gene expression, as it is involved in mRNA maturation, stability and degradation (Arraiano et al., 2010). Mixture modeling identified 3,344 CSs with a posterior probability ≥0.9 (high confidence CSs) (Figure 1A and Supplementary Table S10). To determine the sequence context of the CSs, we used the regions flanking the 5′ ends to generate a sequence logo (Figure 4A). There was a strong preference for a cytosine in the +1 position (present in more than the 90% of the CSs) (Figure 4B), suggesting that it may be structurally important for RNase recognition and/or catalysis.
Figure 4. Cleavage site positions are biased with respect to sequence context and genetic location. (A) Sequence context of cleavage sites. The sequences flanking the 3,344 high-confidence CSs were used to create the sequence logo with WebLogo 3 (Crooks et al., 2004). (B) Base preference for RNA cleavage. The base frequencies for the –2 to +2 positions were determined. (C) Cleavage site categories based on the genetic context. CSs are denoted with arrows. 5′ UTR: the CS is within the leader of a gene, and the genes upstream and downstream of the CS are divergent (Gene 1 and Gene 2, red arrow). CDS: The CS is within a coding sequence (green arrow). 3′ UTR: the genes upstream and downstream of the CS are convergent (Gene 2 and Gene 3, light blue arrow). Operon: The CS is between two genes with the same orientation and the first gene in the operon has a pTSS according to Supplementary Table S6 (violet arrow). (D) Distribution of cleavage sites. The frequency of CSs in each location was normalized to the proportion of the genome that the location category comprised. The proportions were then normalized to the CDS category, which was set as 1. ∗∗∗∗p < 0001, ∗p < 0.01 (Chi-square test).
Cleaved 5′ ends can represent either degradation intermediates or transcripts that undergo functional processing/maturation. In an attempt to investigate CS function, we classified them according to their locations within mRNA transcripts (Figure 4C and Supplementary Table S10). We found that, after normalizing to the proportion of the expressed transcriptome that is comprised by each location category, cleaved 5′ ends are more abundant within 5′ UTRs and intergenic regions of operons than within coding sequences and 3′ UTRs (Figure 4D). Stringent criteria were used in these analyses to avoid undesired bias (Figure 4C and see section “Materials and Methods”). While one would expect the CSs associated with mRNA turnover to be evenly distributed throughout the transcript, enrichment of CSs within the 5′ UTRs as well as between two co-transcribed genes may be indicative of cleavages associated with processing and maturation. Alternatively, these regions may be more susceptible to RNases due to lack of associated ribosomes. Here we predicted with high confidence that at least 101 genes have one or more CSs in their 5′ UTRs (Supplementary Table S11).
We detected cleaved 5′ ends within the coding sequences of 18% of M. smegmatis genes, ranging from 1 to over 40 sites per gene. We analyzed the distribution of CSs within coding sequences (Supplementary Figure S7), taking into consideration the genomic context of the genes. When analyzing the distribution of CSs within the coding sequences of genes whose downstream gene has the same orientation, we observed an increase in CS frequency in the region near the stop codon (Supplementary Figure S7A). However, when only coding sequences having a downstream gene on the opposite strand (convergent) were considered, the distribution of CSs through the coding sequences was significantly different (p-value < 0.0001, Kolmogorov-Smirnov D test) with the CSs more evenly distributed throughout the coding sequence (Supplementary Figure S7B). This suggests that the cleavage bias toward the end of the genes observed in Supplementary Figure S7A may be due to the fact that many of these CSs are actually occurring in the 5′ UTRs of the downstream genes. In cases where the TSS of a given gene occurs within the coding sequence of the preceding gene, a CS may map to both the coding sequence of the upstream gene and the 5′ UTR of the downstream gene. In these cases, we cannot determine in which of the two transcripts the cleavage occurred. However, cleavages may also occur in polycistronic transcripts. We therefore assessed the distributions of CSs in the operons predicted above. The distribution of CSs in genes co-transcribed with a downstream gene showed a slight increase toward the last part of the gene (Supplementary Figure S7C). This may reflect cases in which polycistronic transcripts are cleaved near the 3′ end of an upstream gene, as has been reported for the furA-katG operon, in which a cleavage near the stop codon of furA was described (Milano et al., 2001; Sala et al., 2008; Taverniti et al., 2011). The furA-katG cleavage was identified in our dataset, located 1 nt downstream of the previously reported position. A similar enrichment of CSs toward stop codons was also observed in a recent genome-wide RNA cleavage analysis in S. enterica (Chao et al., 2017), although in this case the high frequency of cleavage may be also attributed to the U preference of RNase E in this organism, which is highly abundant in these regions.
Prediction of Additional TSSs and CSs Based on Sequence Context
The sequence contexts of TSSs (Figure 2B) and CSs (Figure 4A) were markedly different, as G and A were highly preferred in the TSS +1 position whereas C was highly preferred in the CS +1 position, and TSSs were associated with a strong overrepresentation of ANNNT -10 sites while CSs were not. These sequence-context differences not only provide validation of our methodology for distinguishing TSSs from CSs, as discussed above, but also provide a means for making improved predictions of the nature of 5′ ends that could not be categorized with high confidence based on their converted/non-converted library coverage alone. Taking advantage of these differences, we sought to obtain a list of additional putative TSSs and CSs. Thus, of the 5′ ends that were not classified with high confidence by mixture modeling, we selected those that had an appropriately positioned ANNNT motif upstream and a G or an A in the +1 position and classified them as TSSs with medium confidence (Supplementary Table S12). In the same way, 5′ ends with a C in the +1 position and lacking the ANNNT motif in the region upstream were designated as medium confidence CSs (Supplementary Table S13). In this way, we were able to obtain 576 and 4,838 medium confidence TSSs and CSs, respectively. Additional validation of a medium confidence TSS was performed for gene MSMEG_0063 using 5′RACE. We were able to corroborate that, as predicted, transcription of this gene is initiated 139 bp upstream the coding sequence and that either deletion or mutation of the predicted -10 promoter region dramatically decreased transcription initiation (Supplementary Figure S8). These results support the value of TSS prediction based on -10 promoter region motif and base composition at +1 position, and highlight the importance of the -10 ANNNT promoter motif for mycobacterial transcription. Three medium confidence CSs (86927+, 87293+, and 5038902-) were also validated using 5′ RACE. Although we are aware of the limitations of these predictions, these lists of medium confidence 5′ ends provide a resource that may be useful for guiding further studies. 5′ ends that did not meet the criteria for high or medium confidence TSSs or CSs are reported in Supplementary Table S14.
The Transcriptional Landscape Changes in Response to Oxygen Limitation
We sought to study the global changes occurring at the transcriptomic level in oxygen limitation employing a system similar to the Wayne model (Wayne and Hayes, 1996) (see section “Materials and Methods”). Two timepoints were experimentally determined in order to evaluate transcriptomic changes during the transition into hypoxia (Supplementary Figure S1). A different enzyme was used for conversion of 5′ triphosphates to 5′ monophosphates in these 5′-end libraries, and it appeared to be less effective than the enzyme used for the 5′ end libraries in Dataset 1. As a consequence, our ability to distinguish TSSs from CSs de novo in these datasets was limited. However, we were able to assess changes in abundance of the 5′ ends classified as high-confidence TSSs or CSs in Dataset 1, as well as identify a limited number of additional TSSs and CSs with high confidence (Supplementary Figure S4 and Supplementary Table S3). Corresponding RNAseq expression libraries revealed that, as expected, a large number of genes were up and downregulated in response to oxygen limitation (Supplementary Figure S9 and Supplementary Table S15). We next investigated the transcriptional changes in hypoxia by assessing the relative abundance of TSSs in these conditions. We found 318 high-confidence TSSs whose abundance varied substantially between exponential phase and hypoxia (Supplementary Table S16). A robust correlation was observed between the pTSS peak height in the 5′-end-directed libraries and RNA levels in the expression libraries for hypoxia (Supplementary Figure S10). In an attempt to identify promoter motifs induced in hypoxia, we analyzed the upstream regions of those TSSs whose abundance increased (fold change ≥2, adjusted p-value ≤ 0.05). Interestingly, we detected a conserved GGGTA motif in the -10 region of 56 promoters induced in hypoxia using MEME (Figure 5A and Supplementary Table S16). This motif was reported as the binding site for alternative sigma factor SigF (Rodrigue et al., 2007; Hartkoorn et al., 2010; Humpel et al., 2010). Additionally, the extended -35 and -10 SigF motif was found in 44 of the 56 promoter sequences (Figure 5A and Supplementary Table S16). SigF was shown to be induced in hypoxia at the transcript level in Mtb (Iona et al., 2016) and highly induced at the protein level under anaerobic conditions using the Wayne model in Mycobacterium bovis BCG strain and Mtb (Michele et al., 1999; Galagan et al., 2013). In M. smegmatis, SigF was shown to play a role under oxidative stress, heat shock, low pH and stationary phase (Gebhard et al., 2008; Humpel et al., 2010; Singh et al., 2015) and sigF RNA levels were detected in exponential phase at a nearly comparable level to sigA (Singh and Singh, 2008). Here, we did not detect significant changes in expression of the sigF gene in hypoxia at the transcript level. However, this is consistent with reported data showing that sigF transcript levels remain unchanged under stress conditions in M. smegmatis (Gebhard et al., 2008), as it was postulated that SigF is post-transcriptionally modulated via an anti-sigma factor rather than through sigF transcription activation (Beaucher et al., 2002). We noted that, in the case of TSSs whose abundance was reduced in hypoxia, almost the totality of the promoters contains the -10 ANNNT σ70 binding motif. We then examined the presence of SigF motif in the regions upstream of 5′ ends that were not classified as high confidence TSSs. We speculate that 5′ ends associated with this motif may be potential TSSs triggered by hypoxia. We found 96 additional putative TSSs that were (1) overrepresented in hypoxia and (2) associated with appropriately-spaced SigF motifs (Supplementary Table S17). Three of the hypoxia-induced genes with SigF motifs (MSMEG_3460, MSMEG_4195 and MSMEG_5329) have homologous genes induced in hypoxia in Mtb (Park et al., 2003; Rustad et al., 2008).
Figure 5. The transcriptional landscape substantially changes upon oxygen limitation. (A) TSSs significantly increased or decreased in hypoxia. 132 TSSs were overrepresented (upper panel) and 186 were underrepresented (lower panel) in different hypoxia stages. The upstream regions of these TSSs were used to search for promoter motifs using MEME. (B) The mean normalized read depths for each 5′ end in the non-converted libraries were compared between hypoxia and normoxia. Graphics show the Log2 of the ratios of read depth for each CSs at 15 h (upper left) and 24 h (upper right), and the Log2 of the ratios of the read depth for each TSSs at 15 h (lower left) and 24 h (lower right) compared to normoxia. (C) Normalized read depth at high-confidence cleavage sites under normoxia and the transition into hypoxia. ∗∗∗∗p < 0.0001, ∗∗∗p < 0.001; ns, not significant (non-parametric Wilcoxon matched-pairs signed rank test).
It is well known that under anaerobic conditions mycobacteria induce the DosR regulon, a set of genes implicated in stress tolerance (Rosenkrands et al., 2002; O’Toole et al., 2003; Park et al., 2003; Roberts et al., 2004; Rustad et al., 2008; Honaker et al., 2009; Leistikow et al., 2010). The DosR transcriptional regulator was highly upregulated at both hypoxic timepoints in the expression libraries (13 and 18-fold at 15 and 24 h, respectively, Supplementary Figure S9) and 30 out of the 49 DosR-activated genes (Berney et al., 2014) were upregulated in our dataset. Thus, we hypothesized that the DosR binding motif should be present in a number of regions upstream the TSSs that were upregulated in hypoxia. Analysis of the 200 bp upstream the TSSs using the CentriMo tool for local motif enrichment analysis (Bailey and Machanick, 2012) allowed us to detect putative DosR motifs in 13 or 53 promoters, depending on whether a stringent (GGGACTTNNGNCCCT) or a weak (RRGNCYWNNGNMM) consensus sequence was used as input (Lun et al., 2009; Berney et al., 2014; Gomes et al., 2014) (Supplementary Table S16). At least two of the 13 genes downstream of these TSSs were previously reported to have DosR motifs by Berney et al. (2014) and RegPrecise Database (Novichkov et al., 2013) and two others are homologs of genes in the Mtb DosR regulon that were not previously described in M. smegmatis as regulated by DosR (Supplementary Table S16).
We then used CentriMo to search for DosR motifs in the regions upstream of 5′ ends that were not classified as high confidence TSSs, given that TSSs derived from hypoxia-specific promoters may have been absent from Dataset 1. We found 36 putative TSSs associated with 20 different genes (Supplementary Table S18), of which 11 have been shown to have DosR binding motifs (Berney et al., 2014). Five of these are homologs of genes in the Mtb DosR regulon.
M. smegmatis Decreases RNA Cleavage Under Oxygen Limitation
There is evidence that mycobacterial mRNA is broadly stabilized under hypoxia and other stress conditions (Rustad et al., 2013; Ignatov et al., 2015). Thus, we anticipated that RNA cleavage should be reduced under hypoxia as a strategy to stabilize transcripts. We compared the relative abundance of each high confidence CS in stress and in exponential phase (Figure 5B) and found that RNA cleavage is significantly reduced in both hypoxia 15 and 24h on a global scale (Figure 5C). In contrast, relative abundance of TSSs did not decrease in these conditions, indicating that the reduction in CSs is not an artifact of improper normalization (Figure 5B). When the ratios of CSs abundance in hypoxia/normal growth of individual genes were analyzed, we observed the same behavior (Supplementary Figure S11). These results indicate that the number of cleavage events per gene decreases during adaptation to hypoxia, which could contribute to the reported increases in half-life (Rustad et al., 2013).
In recent years, genome-wide transcriptome studies have been widely used to elucidate the genome architecture and modulation of transcription in different bacterial species (Albrecht et al., 2009; Mendoza-Vargas et al., 2009; Mitschke et al., 2011; Cortes et al., 2013; Schlüter et al., 2013; Dinan et al., 2014; Ramachandran et al., 2014; Innocenti et al., 2015; Sass et al., 2015; Thomason et al., 2015; Berger et al., 2016; Čuklina et al., 2016; D’arrigo et al., 2016; Heidrich et al., 2017; Li et al., 2017; Zhukova et al., 2017). Here we combined 5′-end-directed libraries and RNAseq expression libraries to shed light on the transcriptional and post-transcriptional landscape of M. smegmatis in different physiological conditions.
The implementation of two differentially treated 5′-end libraries followed by Gaussian mixture modeling analysis allowed us to simultaneously map and classify 5′ ends resulting from nucleolytic cleavage and those resulting from primary transcription with high confidence. We were able to classify 57% of the 5′ ends in Dataset 1 with high confidence. In addition, we elaborated a list of medium confidence TSSs and CSs (Supplementary Tables S12, S13). These lists constitute a valuable resource for the research community.
Analysis of TSS mapping data allowed us to identify over 4,000 primary TSSs and to study the transcript features in M. smegmatis. The high proportion of leaderless transcripts, the lack of a consensus SD sequence in half of the leadered transcripts, and the absence of a conserved -35 consensus sequence indicate that the transcription-translation machineries are relatively robust in M. smegmatis. These findings are consistent with a recent study that mapped a 2,139 TSSs in M. smegmatis (Li et al., 2017). The apparent robustness of translation is shared with Mtb, where 25% of the transcripts lack a leader sequence (Cortes et al., 2013; Shell et al., 2015b). In addition, high abundances of transcripts lacking 5′ UTRs have been reported in other bacteria including Corynebacterium diphtheria, Leptospira interrogans, Borrelia burgdorferi, and Deinococcus deserti, the latter having 60% leaderless transcripts (de Groot et al., 2014; Adams et al., 2017; Zhukova et al., 2017; Wittchen et al., 2018). Considering the high proportion of leaderless transcripts and the large number of leadered transcripts that lack a SD sequence (53%), it follows that an important number of transcripts are translated without canonical interactions between the mRNA and anti-Shine-Dalgarno sequence, suggesting that M. smegmatis has versatile mechanisms to address translation. A computational prediction showed that the presence of SD can be very variable between prokaryotes, ranging from 11% in Mycoplasma to 91% in Firmicutes (Chang et al., 2006). Cortes et al. (2013) reported that the 55% of the genes transcribed with a 5′ UTR lack the SD motif. The correlation of leader lengths for homologous genes in M. smegmatis and M. tuberculosis (Figure 3B) suggests that some genes may share additional UTR-associated regulatory features, although further work is required to investigate the possible regulatory roles of 5′ UTRs in both species.
To begin to understand the role of RNA cleavage in mycobacteria, we identified and classified over 3,000 CSs throughout the M. smegmatis transcriptome, presenting the first report of an RNA cleavage map in mycobacteria. The most striking feature of the CSs was a cytidine in the +1 position, which was true in over 90% of the cases. While the RNases involved in global RNA decay in mycobacteria have not been yet elucidated, some studies have implicated RNase E as a major player in RNA processing and decay (Kovacs et al., 2005; Zeller et al., 2007; Csanadi et al., 2009; Taverniti et al., 2011), given its central role in other bacteria such as E. coli and its essentiality for survival in both M. smegmatis and Mtb (Sassetti et al., 2003; Sassetti and Rubin, 2003; Griffin et al., 2011; Taverniti et al., 2011; DeJesus et al., 2017). It is therefore possible that mycobacterial RNase E, or other endonucleases with dominant roles, favor cytidine in the +1 position. Interestingly, the sequence context of cleavage found here is different from that described for E. coli, for which the consensus sequence is (A/G)N↓AU (Mackie, 2013) or S. enterica, in which a marked preference for uridine at the+2 position and AU-rich sequences are important for RNase E cleavage (Chao et al., 2017).
RNA cleavage is required for maturation of some mRNAs (Li and Deutscher, 1996; Condon et al., 2001; Gutgsell and Jain, 2010; Moores et al., 2017). Therefore, the observation that CSs are enriched in 5′ UTRs and intergenic regions suggests that processing may play roles in RNA maturation, stability, and translation for some transcripts in M. smegmatis. A high abundance of processing sites around the translation start site was also observed in P. aeruginosa and S. enterica in transcriptome-wide studies (Chao et al., 2017; Gill et al., 2018), suggesting that 5′ UTR cleavage may be a widespread post-transcriptional mechanism for modulating gene expression in bacteria.
Regulation of RNA decay and processing plays a crucial role in adaptation to environmental changes. We present evidence showing that RNA cleavage is markedly reduced in conditions that result in growth cessation. It was previously demonstrated that in low oxygen concentrations mycobacteria reduce their RNA levels (Ignatov et al., 2015) and mRNA half-life is strikingly increased (Rustad et al., 2013), likely as a mechanism to maintain adequate transcript levels in the cell without the energy expenditures that continuous transcription would require. While several traits are involved in the regulation of transcript abundance and stability, the observation that cleavage events are pronouncedly reduced in these conditions pinpoint this mechanism as a potential way to control RNA stability under stress. In agreement with this hypothesis, RNase E was modestly but significantly decreased at the transcript level in early and late hypoxia (fold change = 0.63 and 0.56, respectively, p-value adjusted <0.05), suggesting that reducing the RNase E abundance in the cell may be a strategy to increase transcript half-life. Further study is needed to better understand the relationship between transcript processing and RNA decay in normoxic growth as well as stress conditions.
Hypoxic stress conditions were also characterized by major changes in the TSSs. 5′-end-mapping libraries revealed that over 300 TSSs varied substantially when cultures were limited in oxygen. We found that 56 transcripts triggered in hypoxia contain the SigF promoter binding motif, indicating that this sigma factor plays a substantial role in the M. smegmatis hypoxia response. While previous work revealed increased expression of SigF itself in hypoxia in Mtb (Galagan et al., 2013; Iona et al., 2016; Yang et al., 2018), this is the first report demonstrating the direct impact of SigF on specific promoters in hypoxic conditions in mycobacteria. Further work is needed to better understand the functional consequences of SigF activation in both organisms in response to hypoxia.
The work reported here represents the most complete M. smegmatis transcriptome map to date. We have almost doubled the number of mapped TSSs, and report the presence and locations of internal and antisense TSSs as well as primary TSSs. Comparison of TSSs used in log phase and hypoxia revealed a signature of SigF activity in hypoxia, which has not been previously reported. We report the presence of locations of thousands of RNA cleavage sites, which reveals for the first time the consensus sequence recognized by the major mycobacterial RNase(s) that produces monophosphorylated 5′ ends. Cleavage sites are enriched in 5′ UTRs and intergenic regions, suggesting that these locations are more accessible to RNases and/or subject to regulation by RNA processing. Cleaved RNAs are relatively less abundant in hypoxic M. smegmatis cultures, suggesting that RNase activity is reduced as part of the phenotypic transition into hypoxia-induced growth arrest.
All next-generation sequencing data are available in raw and processed forms ion the GEO site, accession number GSE128412.
MM, YZ, and SS conceived and designed the experiments. MM and YZ performed the experiments. MM, HS, and SS analyzed the data. MM and SS wrote the manuscript.
This work was supported by NSF CAREER award 1652756 to SS and by institutional startup funds from WPI.
Conflict of Interest Statement
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
We are grateful to Zheyang Wu and Thomas Ioerger for helpful advice on the mixture modeling and Michael Chase for helpful advice on other aspects of our data analysis pipeline. We thank Todd Gray and Keith Derbyshire for the kind gift of the msmeg_0062-msmeg_0066 deletion strain used for promoter validation. We thank members of the Shell lab for technical assistance and helpful discussions. All Illumina sequencing was performed by the UMass Medical School Deep Sequencing Core.
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2019.00591/full#supplementary-material
Adams, P. P., Flores Avile, C., Popitsch, N., Bilusic, I., Schroeder, R., Lybecker, M., et al. (2017). In vivo expression technology and 5’ end mapping of the Borrelia burgdorferi transcriptome identify novel RNAs expressed during mammalian infection. Nucleic Acids Res. 45, 775–792. doi: 10.1093/nar/gkw1180
Albrecht, M., Sharma, C. M., Reinhardt, R., Vogel, J., and Rudel, T. (2009). Deep sequencing-based discovery of the chlamydia trachomatis transcriptome. Nucleic Acids Res. 38, 868–877. doi: 10.1093/nar/gkp1032
Andre, G., Even, S., Putzer, H., Burguiere, P., Croux, C., Danchin, A., et al. (2008). S-box and T-box riboswitches and antisense RNA control a sulfur metabolic operon of Clostridium acetobutylicum. Nucleic Acids Res. 36, 5955–5969. doi: 10.1093/nar/gkn601
Arraiano, C. M., Andrade, J. M., Domingues, S., Guinote, I. B., Malecki, M., Matos, R. G., et al. (2010). The critical role of RNA processing and degradation in the control of gene expression. FEMS Microbiol. Rev. 34, 883–923. doi: 10.1111/j.1574-6976.2010.00242.x
Bagchi, G., Das, T. K., and Tyagi, J. S. (2002). Molecular analysis of the dormancy response in Mycobacterium smegmatis: expression analysis of genes encoding the DevR–DevS two-component system, Rv3134c and chaperone α-crystallin homologues. FEMS Microbiol. Lett. 211, 231–237.
Beaucher, J., Rodrigue, S., Jacques, P. E., Smith, I., Brzezinski, R., and Gaudreau, L. (2002). Novel Mycobacterium tuberculosis anti-sigma factor antagonists control sigmaF activity by distinct mechanisms. Mol. Microbiol. 45, 1527–1540. doi: 10.1046/j.1365-2958.2002.03135.x
Beaume, M., Hernandez, D., Farinelli, L., Deluen, C., Linder, P., Gaspin, C., et al. (2010). Cartography of methicillin-resistant S. aureus transcripts: detection, orientation and temporal expression during growth phase and stress conditions. PLoS One 5:e10725. doi: 10.1371/journal.pone.0010725
Berger, P., Knödler, M., Förstner, K. U., Berger, M., Bertling, C., Sharma, C. M., et al. (2016). The primary transcriptome of the Escherichia coli O104: H4 pAA plasmid and novel insights into its virulence gene expression and regulation. Sci. Rep. 6:35307. doi: 10.1038/srep35307
Berney, M., Greening, C., Conrad, R., Jacobs, W. R. Jr., and Cook, G. M. (2014). An obligately aerobic soil bacterium activates fermentative hydrogen production to survive reductive stress during hypoxia. Proc. Natl. Acad. Sci. U.S.A. 111, 11479–11484. doi: 10.1073/pnas.1407034111
Chang, B., Halgamuge, S., and Tang, S. L. (2006). Analysis of SD sequences in completed microbial genomes: non-SD-led genes are as common as SD-led genes. Gene 373, 90–99. doi: 10.1016/j.gene.2006.01.033
Chao, Y., Li, L., Girodat, D., Förstner, K. U., Said, N., Corcoran, C., et al. (2017). In vivo cleavage map illuminates the central role of RNase E in coding and non-coding RNA Pathways. Mol. Cell 65, 39–51. doi: 10.1016/j.molcel.2016.11.002
Condon, C., Brechemier-Baey, D., Beltchev, B., Grunberg-Manago, M., and Putzer, H. (2001). Identification of the gene encoding the 5S ribosomal RNA maturase in Bacillus subtilis: mature 5S rRNA is dispensable for ribosome function. RNA 7, 242–253. doi: 10.1017/S1355838201002163
Coros, A., Callahan, B., Battaglioli, E., and Derbyshire, K. M. (2008). The specialized secretory apparatus ESX-1 is essential for DNA transfer in Mycobacterium smegmatis. Mol. Microbiol. 69, 794–808. doi: 10.1111/j.1365-2958.2008.06299.x
Cortes, T., Schubert, O. T., Rose, G., Arnvig, K. B., Comas, I., Aebersold, R., et al. (2013). Genome-wide mapping of transcriptional start sites defines an extensive leaderless transcriptome in Mycobacterium tuberculosis. Cell Rep. 5, 1121–1131. doi: 10.1016/j.celrep.2013.10.031
Čuklina, J., Hahn, J., Imakaev, M., Omasits, U., Förstner, K. U., Ljubimov, N., et al. (2016). Genome-wide transcription start site mapping of Bradyrhizobium japonicum grown free-living or in symbiosis–a rich resource to identify new transcripts, proteins and to study gene regulation. BMC Genomics 17:302. doi: 10.1186/s12864-016-2602-9
D’arrigo, I., Bojanovič, K., Yang, X., Holm Rau, M., and Long, K. S. (2016). Genome-wide mapping of transcription start sites yields novel insights into the primary transcriptome of Pseudomonas putida. Environ. Microbiol. 18, 3466–3481. doi: 10.1111/1462-2920.13326
de Groot, A., Roche, D., Fernandez, B., Ludanyi, M., Cruveiller, S., Pignol, D., et al. (2014). RNA sequencing and proteogenomics reveal the importance of leaderless mRNAs in the radiation-tolerant bacterium Deinococcus deserti. Genome Biol. Evol. 6, 932–948. doi: 10.1093/gbe/evu069
DeJesus, M. A., Gerrick, E. R., Xu, W., Park, S. W., Long, J. E., Boutte, C. C., et al. (2017). Comprehensive essentiality analysis of the mycobacterium tuberculosis genome via saturating transposon mutagenesis. MBio 8:e2133-16. doi: 10.1128/mBio.02133-16
Dinan, A. M., Tong, P., Lohan, A. J., Conlon, K. M., Miranda-CasoLuengo, A. A., Malone, K. M., et al. (2014). Relaxed selection drives a noisy noncoding transcriptome in members of the Mycobacterium tuberculosis complex. MBio 5:e1169-14. doi: 10.1128/mBio.01169-14
Elharar, Y., Roth, Z., Hermelin, I., Moon, A., Peretz, G., Shenkerman, Y., et al. (2014). Survival of mycobacteria depends on proteasome-mediated amino acid recycling under nutrient limitation. EMBO J. 33, 1802–1814. doi: 10.15252/embj.201387076
Fozo, E. M., Kawano, M., Fontaine, F., Kaya, Y., Mendieta, K. S., Jones, K. L., et al. (2008). Repression of small toxic protein synthesis by the Sib and OhsC small RNAs. Mol. Microbiol. 70, 1076–1093. doi: 10.1111/j.1365-2958.2008.06394.x
Galagan, J. E., Minch, K., Peterson, M., Lyubetskaya, A., Azizi, E., Sweet, L., et al. (2013). The Mycobacterium tuberculosis regulatory network and hypoxia. Nature 499, 178–183. doi: 10.1038/nature12337
Gaudion, A., Dawson, L., Davis, E., and Smollett, K. (2013). Characterisation of the Mycobacterium tuberculosis alternative sigma factor SigG: its operon and regulon. Tuberculosis 93, 482–491. doi: 10.1016/j.tube.2013.05.005
Gebhard, S., Humpel, A., McLellan, A. D., and Cook, G. M. (2008). The alternative sigma factor SigF of Mycobacterium smegmatis is required for survival of heat shock, acidic pH and oxidative stress. Microbiology 154, 2786–2795. doi: 10.1099/mic.0.2008/018044-0
Giangrossi, M., Prosseda, G., Tran, C. N., Brandi, A., Colonna, B., and Falconi, M. (2010). A novel antisense RNA regulates at transcriptional level the virulence gene icsA of Shigella flexneri. Nucleic Acids Res. 38, 3362–3375. doi: 10.1093/nar/gkq025
Gomes, A. L., Abeel, T., Peterson, M., Azizi, E., Lyubetskaya, A., Carvalho, L., et al. (2014). Decoding ChIP-seq with a double-binding signal refines binding peaks to single-nucleotides and predicts cooperative interaction. Genome Res. 24, 1686–1697. doi: 10.1101/gr.161711.113
Gray, T. A., Palumbo, M. J., and Derbyshire, K. M. (2013). Draft genome sequence of MKD8, a conjugal recipient Mycobacterium smegmatis strain. Genome Announc. 1:e0014813. doi: 10.1128/genomeA.00148-13
Griffin, J. E., Gawronski, J. D., Dejesus, M. A., Ioerger, T. R., Akerley, B. J., and Sassetti, C. M. (2011). High-resolution phenotypic profiling defines genes essential for mycobacterial growth and cholesterol catabolism. PLoS Pathog. 7:e1002251. doi: 10.1371/journal.ppat.1002251
Guell, M., van Noort, V., Yus, E., Chen, W. H., Leigh-Bell, J., Michalodimitrakis, K., et al. (2009). Transcriptome complexity in a genome-reduced bacterium. Science 326, 1268–1271. doi: 10.1126/science.1176951
Haft, D. H., Loftus, B. J., Richardson, D. L., Yang, F., Eisen, J. A., Paulsen, I. T., et al. (2001). TIGRFAMs: a protein family resource for the functional identification of proteins. Nucleic Acids Res. 29, 41–43. doi: 10.1093/nar/29.1.41
Hartkoorn, R. C., Sala, C., Magnet, S. J., Chen, J. M., Pojer, F., and Cole, S. T. (2010). Sigma factor F does not prevent rifampin inhibition of RNA polymerase or cause rifampin tolerance in Mycobacterium tuberculosis. J. Bacteriol. 192, 5472–5479. doi: 10.1128/JB.00687-10
Hayashi, J. M., Richardson, K., Melzer, E. S., Sandler, S. J., Aldridge, B. B., Siegrist, M. S., et al. (2018). Stress-induced reorganization of the mycobacterial membrane domain. MBio 9:e1823-17. doi: 10.1128/mBio.01823-17
Heidrich, N., Bauriedl, S., Barquist, L., Li, L., Schoen, C., and Vogel, J. (2017). The primary transcriptome of Neisseria meningitidis and its interaction with the RNA chaperone Hfq. Nucleic Acids Res. 45, 6147–6167. doi: 10.1093/nar/gkx168
Heroven, A., Sest, M., Pisano, F., Scheb-Wetzel, M., Böhme, K., Klein, J., et al. (2012). Crp induces switching of the CsrB and CsrC RNAs in Yersinia pseudotuberculosis and links nutritional status to virulence. Front. Cell. Infect. Microbiol. 2:158. doi: 10.3389/fcimb.2012.00158
Holmqvist, E., Wright, P. R., Li, L., Bischler, T., Barquist, L., Reinhardt, R., et al. (2016). Global RNA recognition patterns of post-transcriptional regulators Hfq and CsrA revealed by UV crosslinking in vivo. EMBO J. 35, 991–1011. doi: 10.15252/embj.201593360
Honaker, R. W., Leistikow, R. L., Bartek, I. L., and Voskuil, M. I. (2009). Unique roles of DosT and DosS in DosR regulon induction and Mycobacterium tuberculosis dormancy. Infect. Immun. 77, 3258–3263. doi: 10.1128/IAI.01449-08
Humpel, A., Gebhard, S., Cook, G. M., and Berney, M. (2010). The SigF regulon in Mycobacterium smegmatis reveals roles in adaptation to stationary phase, heat, and oxidative stress. J. Bacteriol. 192, 2491–2502. doi: 10.1128/JB.00035-10
Ignatov, D. V., Salina, E. G., Fursov, M. V., Skvortsov, T. A., Azhikina, T. L., and Kaprelyants, A. S. (2015). Dormant non-culturable Mycobacterium tuberculosis retains stable low-abundant mRNA. BMC Genomics 16:954. doi: 10.1186/s12864-015-2197-6
Innocenti, N., Golumbeanu, M., Fouquier d’Herouel, A., Lacoux, C., Bonnin, R. A., Kennedy, S. P., et al. (2015). Whole-genome mapping of 5’ RNA ends in bacteria by tagged sequencing: a comprehensive view in Enterococcus faecalis. RNA 21, 1018–1030. doi: 10.1261/rna.048470.114
Iona, E., Pardini, M., Mustazzolu, A., Piccaro, G., Nisini, R., Fattorini, L., et al. (2016). Mycobacterium tuberculosis gene expression at different stages of hypoxia-induced dormancy and upon resuscitation. J. Microbiol. 54, 565–572. doi: 10.1007/s12275-016-6150-4
Jarmer, H., Larsen, T. S., Krogh, A., Saxild, H. H., Brunak, S., and Knudsen, S. (2001). Sigma A recognition sites in the Bacillus subtilis genome. Microbiology 147, 2417–2424. doi: 10.1099/00221287-147-9-2417
Kawano, M., Aravind, L., and Storz, G. (2007). An antisense RNA controls synthesis of an SOS-induced toxin evolved from an antitoxin. Mol. Microbiol. 64, 738–754. doi: 10.1111/j.1365-2958.2007.05688.x
Kulesekara, H., Lee, V., Brencic, A., Liberati, N., Urbach, J., Miyata, S., et al. (2006). Analysis of Pseudomonas aeruginosa diguanylate cyclases and phosphodiesterases reveals a role for bis-(3′-5′)-cyclic-GMP in virulence. Proc. Natl. Acad. Sci. U.S.A. 103, 2839–2844. doi: 10.1073/pnas.0511090103
Leistikow, R. L., Morton, R. A., Bartek, I. L., Frimpong, I., Wagner, K., and Voskuil, M. I. (2010). The Mycobacterium tuberculosis DosR regulon assists in metabolic homeostasis and enables rapid recovery from nonrespiring dormancy. J. Bacteriol. 192, 1662–1670. doi: 10.1128/JB.00926-09
Lloréns-Rico, V., Cano, J., Kamminga, T., Gil, R., Latorre, A., and Chen, W. H. (2016). Bacterial antisense RNAs are mainly the product of transcriptional noise. Sci. Adv. 2:e1501363. doi: 10.1126/sciadv.1501363
Lun, D. S., Sherrid, A., Weiner, B., Sherman, D. R., and Galagan, J. E. (2009). A blind deconvolution approach to high-resolution mapping of transcription factor binding sites from ChIP-seq data. Genome Biol. 10:R142. doi: 10.1186/gb-2009-10-12-r142
Mendoza-Vargas, A., Olvera, L., Olvera, M., Grande, R., Vega-Alvarado, L., Taboada, B., et al. (2009). Genome-wide identification of transcription start sites, promoters and transcription factor binding sites in E. coli. PLoS One 4:e7526. doi: 10.1371/journal.pone.0007526
Michele, T. M., Ko, C., and Bishai, W. R. (1999). Exposure to antibiotics induces expression of the Mycobacterium tuberculosis sigF gene: implications for chemotherapy against mycobacterial persistors. Antimicrob. Agents Chemother. 43, 218–225. doi: 10.1128/AAC.43.2.218
Milano, A., Forti, F., Sala, C., Riccardi, G., and Ghisotti, D. (2001). Transcriptional regulation of furA and katG upon oxidative stress in Mycobacterium smegmatis. J. Bacteriol. 183, 6801–6806. doi: 10.1128/JB.183.23.6801-6806.2001
Mitschke, J., Georg, J., Scholz, I., Sharma, C. M., Dienst, D., Bantscheff, J., et al. (2011). An experimentally anchored map of transcriptional start sites in the model cyanobacterium Synechocystis sp. PCC6803. Proc. Natl. Acad. Sci. U.S.A. 108, 2124–2129. doi: 10.1073/pnas.1015154108
Moores, A., Riesco, A. B., Schwenk, S., and Arnvig, K. B. (2017). Expression, maturation and turnover of DrrS, an unusually stable, DosR regulated small RNA in Mycobacterium tuberculosis. PLoS One 12:e0174079. doi: 10.1371/journal.pone.0174079
Morita, T., Maki, K., and Aiba, H. (2005). RNase E-based ribonucleoprotein complexes: mechanical basis of mRNA destabilization mediated by bacterial noncoding RNAs. Genes Dev. 19, 2176–2186. doi: 10.1101/gad.1330405
Mraheil, M. A., Billion, A., Mohamed, W., Mukherjee, K., Kuenne, C., Pischimarov, J., et al. (2011). The intracellular sRNA transcriptome of Listeria monocytogenes during growth in macrophages. Nucleic Acids Res. 39, 4235–4248. doi: 10.1093/nar/gkr033
Novichkov, P. S., Kazakov, A. E., Ravcheev, D. A., Leyn, S. A., Kovaleva, G. Y., Sutormin, R. A., et al. (2013). RegPrecise 3.0–a resource for genome-scale exploration of transcriptional regulation in bacteria. BMC Genomics 14:745. doi: 10.1186/1471-2164-14-745
O’Toole, R., Smeulders, M. J., Blokpoel, M. C., Kay, E. J., Lougheed, K., and Williams, H. D. (2003). A two-component regulator of universal stress protein expression and adaptation to oxygen starvation in Mycobacterium smegmatis. J. Bacteriol. 185, 1543–1554. doi: 10.1128/JB.185.5.1543-1554.2003
Paletta, J. L., and Ohman, D. E. (2012). Evidence for two promoters internal to the alginate biosynthesis operon in Pseudomonas aeruginosa. Curr. Microbiol. 65, 770–775. doi: 10.1007/s00284-012-0228-y
Park, H. D., Guinn, K. M., Harrell, M. I., Liao, R., Voskuil, M. I., Tompa, M., et al. (2003). Rv3133c/dosR is a transcription factor that mediates the hypoxic response of Mycobacterium tuberculosis. Mol. Microbiol. 48, 833–843. doi: 10.1046/j.1365-2958.2003.03474.x
Pecsi, I., Hards, K., Ekanayaka, N., Berney, M., Hartman, T., Jacobs, W. R., et al. (2014). Essentiality of succinate dehydrogenase in Mycobacterium smegmatis and its role in the generation of the membrane potential under hypoxia. MBio 5:e1093-14. doi: 10.1128/mBio.01093-14
Potgieter, M. G., Nakedi, K. C., Ambler, J. M., Nel, A. J., Garnett, S., Soares, N. C., et al. (2016). Proteogenomic analysis of Mycobacterium smegmatis using high resolution mass spectrometry. Front. Microbiol. 7:427. doi: 10.3389/fmicb.2016.00427
Raju, R. M., Unnikrishnan, M., Rubin, D. H., Krishnamoorthy, V., Kandror, O., Akopian, T. N., et al. (2012). Mycobacterium tuberculosis ClpP1 and ClpP2 function together in protein degradation and are required for viability in vitro and during infection. PLoS Pathog. 8:e1002511. doi: 10.1371/journal.ppat.1002511
Ramachandran, V. K., Shearer, N., and Thompson, A. (2014). The primary transcriptome of Salmonella enterica serovar Typhimurium and its dependence on ppGpp during late stationary phase. PLoS One 9:e92690. doi: 10.1371/journal.pone.0092690
Raman, S., Hazra, R., Dascher, C. C., and Husson, R. N. (2004). Transcription regulation by the Mycobacterium tuberculosis alternative sigma factor SigD and its role in virulence. J. Bacteriol. 186, 6605–6616. doi: 10.1128/JB.186.19.6605-6616.2004
Raman, S., Song, T., Puyang, X., Bardarov, S., Jacobs, W. R. Jr., and Husson, R. N. (2001). The alternative sigma factor SigH regulates major components of oxidative and heat stress responses in Mycobacterium tuberculosis. J. Bacteriol. 183, 6119–6125. doi: 10.1128/JB.183.20.6119-6125.2001
Rigel, N. W., Gibbons, H. S., McCann, J. R., McDonough, J. A., Kurtz, S., and Braunstein, M. (2009). The accessory SecA2 system of mycobacteria requires ATP binding and the canonical SecA1. J. Biol. Chem. 284, 9927–9936. doi: 10.1074/jbc.M900325200
Roberts, D. M., Liao, R. P., Wisedchaisri, G., Hol, W. G., and Sherman, D. R. (2004). Two sensor kinases contribute to the hypoxic response of Mycobacterium tuberculosis. J. Biol. Chem. 279, 23082–23087. doi: 10.1074/jbc.M401230200
Robson, J., McKenzie, J. L., Cursons, R., Cook, G. M., and Arcus, V. L. (2009). The vapBC operon from Mycobacterium smegmatis is an autoregulated toxin-antitoxin module that controls growth via inhibition of translation. J. Mol. Biol. 390, 353–367. doi: 10.1016/j.jmb.2009.05.006
Rodrigue, S., Brodeur, J., Jacques, P. E., Gervais, A. L., Brzezinski, R., and Gaudreau, L. (2007). Identification of mycobacterial sigma factor binding sites by chromatin immunoprecipitation assays. J. Bacteriol. 189, 1505–1513. doi: 10.1128/JB.01371-06
Rosenkrands, I., Slayden, R. A., Crawford, J., Aagaard, C., Clifton, III E., and Andersen, P. (2002). Hypoxic response of Mycobacterium tuberculosis studied by metabolic labeling and proteome analysis of cellular and extracellular proteins. J. Bacteriol. 184, 3485–3491. doi: 10.1128/JB.184.13.3485-3491.2002
Rustad, T. R., Minch, K. J., Brabant, W., Winkler, J. K., Reiss, D. J., Baliga, N. S., et al. (2013). Global analysis of mRNA stability in Mycobacterium tuberculosis. Nucleic Acids Res. 41, 509–517. doi: 10.1093/nar/gks1019
Sala, C., Forti, F., Magnoni, F., and Ghisotti, D. (2008). The katG mRNA of Mycobacterium tuberculosis and Mycobacterium smegmatis is processed at its 5′ end and is stabilized by both a polypurine sequence and translation initiation. BMC Mol. Biol. 9:33. doi: 10.1186/1471-2199-9-33
Sass, A. M., Van Acker, H., Förstner, K. U., Van Nieuwerburgh, F., Deforce, D., Vogel, J., et al. (2015). Genome-wide transcription start site profiling in biofilm-grown Burkholderia cenocepacia J2315. BMC Genomics 16:775. doi: 10.1186/s12864-015-1993-3
Schifano, J. M., Edifor, R., Sharp, J. D., Ouyang, M., Konkimalla, A., Husson, R. N., et al. (2013). Mycobacterial toxin MazF-mt6 inhibits translation through cleavage of 23S rRNA at the ribosomal A site. Proc. Natl. Acad. Sci. U.S.A. 110, 8501–8506. doi: 10.1073/pnas.1222031110
Schlüter, J.-P., Reinkensmeier, J., Barnett, M. J., Lang, C., Krol, E., Giegerich, R., et al. (2013). Global mapping of transcription start sites and promoter motifs in the symbiotic α-proteobacterium Sinorhizobium meliloti 1021. BMC Genomics 14:156. doi: 10.1186/1471-2164-14-156
Shao, W., Price, M. N., Deutschbauer, A. M., Romine, M. F., and Arkin, A. P. (2014). Conservation of transcription start sites within genes across a bacterial genus. MBio 5:e1398-14. doi: 10.1128/mBio.01398-14
Sharma, C. M., Hoffmann, S., Darfeuille, F., Reignier, J., Findeiss, S., Sittka, A., et al. (2010). The primary transcriptome of the major human pathogen Helicobacter pylori. Nature 464, 250–255. doi: 10.1038/nature08756
Shell, S. S., Wang, J., Lapierre, P., Mir, M., Chase, M. R., Pyle, M. M., et al. (2015b). Leaderless transcripts and small proteins are common features of the mycobacterial translational landscape. PLoS Genet. 11:e1005641. doi: 10.1371/journal.pgen.1005641
Singh, A. K., Dutta, D., Singh, V., Srivastava, V., Biswas, R. K., and Singh, B. N. (2015). Characterization of Mycobacterium smegmatis sigF mutant and its regulon: overexpression of SigF antagonist (MSMEG_1803) in M. smegmatis mimics sigF mutant phenotype, loss of pigmentation, and sensitivity to oxidative stress. Microbiologyopen 4, 896–916. doi: 10.1002/mbo3.288
Skliarova, S. A., Kreneva, R. A., Perumov, D. A., and Mironov, A. S. (2012). The characterization of internal promoters in the Bacillus subtilis riboflavin biosynthesis operon. Genetika 48, 1133–1141.
Song, T., Song, S. E., Raman, S., Anaya, M., and Husson, R. N. (2008). Critical role of a single position in the -35 element for promoter recognition by Mycobacterium tuberculosis SigE and SigH. J. Bacteriol. 190, 2227–2230. doi: 10.1128/JB.01642-07
Sun, R., Converse, P. J., Ko, C., Tyagi, S., Morrison, N. E., and Bishai, W. R. (2004). Mycobacterium tuberculosis ECF sigma factor sigC is required for lethality in mice and for the conditional expression of a defined gene set. Mol. Microbiol. 52, 25–38. doi: 10.1111/j.1365-2958.2003.03958.x
Taverniti, V., Forti, F., Ghisotti, D., and Putzer, H. (2011). Mycobacterium smegmatis RNase J is a 5’-3’ exo-/endoribonuclease and both RNase J and RNase E are involved in ribosomal RNA maturation. Mol. Microbiol. 82, 1260–1276. doi: 10.1111/j.1365-2958.2011.07888.x
Thomason, M. K., Bischler, T., Eisenbart, S. K., Förstner, K. U., Zhang, A., Herbig, A., et al. (2015). Global transcriptional start site mapping using differential RNA sequencing reveals novel antisense RNAs in Escherichia coli. J. Bacteriol. 197, 18–28. doi: 10.1128/JB.02096-14
Trauner, A., Lougheed, K. E., Bennett, M. H., Hingley-Wilson, S. M., and Williams, H. D. (2012). The dormancy regulator DosR controls ribosome stability in hypoxic mycobacteria. J. Biol. Chem. 287:24053. doi: 10.1074/jbc.M112.364851
Wayne, L. G., and Hayes, L. G. (1996). An in vitro model for sequential study of shiftdown of Mycobacterium tuberculosis through two stages of nonreplicating persistence. Infect. Immun. 64, 2062–2069.
Wittchen, M., Busche, T., Gaspar, A. H., Lee, J. H., Ton-That, H., Kalinowski, J., et al. (2018). Transcriptome sequencing of the human pathogen Corynebacterium diphtheriae NCTC 13129 provides detailed insights into its transcriptional landscape and into DtxR-mediated transcriptional regulation. BMC Genomics 19:82. doi: 10.1186/s12864-018-4481-8
Wu, M.-L., Gengenbacher, M., and Dick, T. (2016). Mild nutrient starvation triggers the development of a small-cell survival morphotype in mycobacteria. Front. Microbiol. 7:947. doi: 10.3389/fmicb.2016.00947
Yang, H., Sha, W., Liu, Z., Tang, T., Liu, H., Qin, L., et al. (2018). Lysine acetylation of DosR regulates the hypoxia response of Mycobacterium tuberculosis. Emerg. Microbes Infect. 7:34. doi: 10.1038/s41426-018-0032-2
Zeller, M. E., Csanadi, A., Miczak, A., Rose, T., Bizebard, T., and Kaberdin, V. R. (2007). Quaternary structure and biochemical properties of mycobacterial RNase E/G. Biochem. J. 403, 207–215. doi: 10.1042/BJ20061530
Zhu, Y., Mao, C., Ge, X., Wang, Z., Lu, P., Zhang, Y., et al. (2017). Characterization of a minimal type of promoter containing the -10 element and a guanine at the -14 or -13 position in mycobacteria. J. Bacteriol. 199:e385-17. doi: 10.1128/JB.00385-17
Zhukova, A., Fernandes, L. G., Hugon, P., Pappas, C. J., Sismeiro, O., Coppée, J. Y., et al. (2017). Genome-wide transcriptional start site mapping and sRNA Identification in the pathogen leptospira interrogans. Front. Cell Infect. Microbiol. 7:10. doi: 10.3389/fcimb.2017.00010
Keywords: tuberculosis, Mycobacterium smegmatis, transcription start sites (TSSs), RNA cleavage, RNA processing and decay, hypoxia, transcriptome, leaderless translation
Citation: Martini MC, Zhou Y, Sun H and Shell SS (2019) Defining the Transcriptional and Post-transcriptional Landscapes of Mycobacterium smegmatis in Aerobic Growth and Hypoxia. Front. Microbiol. 10:591. doi: 10.3389/fmicb.2019.00591
Received: 25 September 2018; Accepted: 08 March 2019;
Published: 26 March 2019.
Edited by:Pere-Joan Cardona, Germans Trias i Pujol Health Sciences Research Institute (IGTP), Spain
Reviewed by:Andaleeb Sajid, National Institutes of Health (NIH), United States
María Carmen Menéndez, Autonomous University of Madrid, Spain
Copyright © 2019 Martini, Zhou, Sun and Shell. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Scarlet S. Shell, email@example.com