ABSTRACT
Though bacteria in nature are often nutritionally limited and growing slowly, most of our understanding of core cellular processes such as transcription comes from studies in a handful of model organisms doubling rapidly under nutrient-replete conditions. We previously identified a small protein of unknown function, called SutA, in a global screen of proteins synthesized in Pseudomonas aeruginosa under growth arrest (Babin BM, et al. (2016) SutA is a bacterial transcription factor expressed during slow growth in Pseudomonas aeruginosa. PNAS 113(5):E597-605). SutA binds RNA polymerase (RNAP), causing widespread changes in gene expression, including upregulation of the ribosomal RNA (rRNA) genes. Here, using biochemical and structural methods, we examine how SutA interacts with RNAP and the functional consequences of these interactions. We show that SutA consists of a central α-helix with unstructured N- and C-terminal tails, and binds to the β1 domain of RNAP. It activates transcription from the P. aeruginosa rrn promoter by both the housekeeping sigma factor holoenzyme (Eσ70) and the general stress response sigma factor holoenzyme (EσS) in vitro, and its N-terminal tail is required for activation in both holoenzyme contexts. However, we find that the interaction between SutA and each holoenzyme is distinct, with the SutA C-terminal tail and an acidic loop unique to σ70 playing the determining roles in these differences. Our results add SutA to a growing list of transcription regulators that use their intrinsically disordered regions to remodel transcription complexes.
SIGNIFICANCE Little is known about how bacteria regulate their activities during periods of dormancy, yet growth arrest dominates bacterial existence in most environments and is directly relevant to the problem of physiological antibiotic tolerance. Though much is known about transcription in the model organism, Escherichia coli, even there, our understanding of gene expression during dormancy is incomplete. Here we explore how transcription under growth arrest is modulated in Pseudomonas aeruginosa by the small acidic protein, SutA. We show that SutA binds to RNA polymerase and controls transcription by a mechanism that is distinct from other known regulators. Our work underscores the potential for fundamental, mechanistic discovery in this important and understudied realm of bacterial physiology.
INTRODUCTION
Despite the fact that most natural environments do not allow bacteria to double every 20-30 minutes, our understanding of essential cellular processes—such as DNA replication, transcription, and translation—has been shaped by studies of a few model organisms growing exponentially at these rates, or responding to a rapid shift from exponential to slow growth. We do not know how the molecular machines responsible for transcription and translation (processes that are necessary to maintain homeostasis even when cell division is not occurring) adapt to long periods of reduced activity and low or uneven substrate availability (2). P. aeruginosa and many other members of the Pseudomonadales order are notable opportunists, capable of using diverse substrates for rapid growth but also able to persist in dormancy for long periods (3), making them attractive model systems for addressing such questions. A better understanding of slow-growing or dormant states in P. aeruginosa also has clinical importance, as these states are thought to contribute to this organism’s antibiotic tolerance in chronic infections (4–6).
Accordingly, in previous work, we used a proteomics-based screen to identify P. aeruginosa regulators that are preferentially expressed during hypoxia-induced growth arrest. We identified an RNAP-binding protein, SutA, that had broad impacts on gene expression and affected that affected the ability of P. aeruginosa to form biofilms and produce virulence factors. Notably, SutA expression led to upregulation of the rRNA genes under slow-growth conditions, and ChIP data showed both that SutA localized to rRNA promoters and that higher levels of RNAP localized to rRNA promoters when SutA was present (1). In this study, we investigate whether, as these results suggest, SutA directly impacts transcription initiation at the rrn promoters, and how these effects are carried out.
The regulation the rrn promoters in E. coli is one of the best-studied examples of growth-rate-responsive control of bacterial gene expression. While they can drive extremely high levels of expression (up to about 70% of all transcription) during exponential growth, they are rapidly and strongly repressed entry into stationary phase (7). This behavior depends on an extremely unstable open complex (OC) formed at rrn P1, which facilitates high levels of expression by enabling rapid promoter clearance by RNAP but also sensitizes initiation to conditions encountered during nutrient downshifts, such as decreased concentrations of the initiating nucleotides ([iNTPs]) (8). A second rrn promoter that drives low levels of expression and is relatively insensitive to regulatory inputs, P2, has been proposed as the mechanism by which some rRNA transcription can be maintained during stationary phase (9, 10), but this would imply that expression levels in E. coli are not actively modulated during protracted dormancy.
Expression of rrn is further modulated by diverse regulators acting at different stages of transcription initiation in different organisms. In many cases, the unstable OC is the target of additional regulation. For example, in E. coli, the signaling molecule (p)ppGpp and its co-regulator DksA bind to RNAP during early stationary phase and further destabilize the final rrn OC (11); the identities of the iNTPs (adenosine or guanosine) allow for direct coordination with the diminished energy stores available to drive translation (7). Also, in many clades outside the Gammaproteobacteria, homologs of the global regulator CarD can enhance rRNA expression by directly stabilizing the OC (12). By contrast, some factors that activate rrn P1 during rapid exponential growth in E. coli (e.g., Fis and DNA supercoiling) exert their effects before the final OC has formed, by helping to recruit RNAP or facilitating the initial opening of the double-stranded DNA (7, 13).
SutA lacks sequence or structural homology to any known transcription factor, raising the possibility that its mode of action is unique. Here, we report that SutA binds to a site on RNAP that is distinct from the binding sites of other regulators, that its activation of rrn transcription depends on its intrinsically disordered N- and C-terminal tails, and that its activity is modulated by the identity of the σ factor. Though our work focuses on a specific transcription factor and promoter in P. aeruginosa, the topic it tackles and the questions it raises are broadly relevant to understanding how bacteria survive periods of slow growth or dormancy in diverse environments.
RESULTS
SutA consists of a conserved alpha helix flanked by flexible N- and C-terminal tails
Because SutA is a small protein (105 amino acids) with no similarity to any known domains, we first explored its structural characteristics. We began by looking at structure predictions (using the Jpred4 algorithm for secondary structure and DISOPRED3 for intrinsic disorder) and sequence conservation (14–16). SutA homologs are found in most organisms in the “Pseudomonadales-Oceanospirallales” clade of Gammaproteobacteria (17). Residues 56-76 are predicted to form an α-helix, followed by a β-strand comprising residues 81-84, but the rest of the protein has no predicted secondary structural elements, and residues 1-50 and 101-105 are predicted to be intrinsically disordered (Figure 1A). While the central, potentially structured region is reasonably well conserved, some homologs completely lack the last 15-18 residues, and others lack most or all of the first 40 residues (Figure S1). This suggests that the N-and C-terminal tails (N-tail and C-tail) might function independently and that their removal might not affect folding/function of other regions.
For structural characterization by NMR, we purified 15N- and 13C-labeled full-length SutA, as well as a 15N- and 13C-labeled construct that lacked most of the predicted disordered residues: SutA 46-101. We also constructed two deletion mutants (Figure 1B): SutA ∆N, retaining residues 41-105, and SutA ∆C, retaining residues 1-87.
We were able to assign resonances and determine backbone chemical shifts for about 85% of the residues of the full-length protein (Table 1). Low sequence complexity and large regions of disorder caused a high degree of overlap in the spectra and made assignment difficult; spectra from the 46-101 variant were easier to assign, and served as a starting point for assignment of the full-length SutA. We focused on measuring secondary-structure chemical shift index values, R2 relaxation rates, and 1H-15N NOE magnitude and sign to determine secondary-structure elements and degree of disorder for each residue that we could assign. We also embedded the protein in a stretched polyacrylamide gel to achieve weak alignment, and calculated residual dipolar couplings (RDCs) by measuring differences in in-phase–antiphase spectra between the isotropic solution sample and the anisotropic stretched-gel sample (Figure 1C). The results of these analyses lend credence to the bioinformatics predictions. Residues 56-76 show the positive Cα and CO and negative Hα secondary chemical shifts associated with an α-helix structure (18), and also show fast R2 relaxation rates and positive (1H-15N)NOE, suggesting that they are not disordered (19). RDCs for the helix region are also positive, as has been observed for α-helical regions of a partially denatured protein (20). The short β-strand is less strongly supported, but secondary shifts for those residues are mostly of the appropriate sign for a β-strand (albeit of small magnitudes). In the N-tail, a small number of residues have a positive NOE signal or secondary shifts that are not near zero, but in general, the residues of this region have the low R2, secondary shift, and RDC values that are characteristic of disorder. The C-tail has several residues that show somewhat higher R2 values and non-zero RDCs, suggestive of some degree of structure, but classic secondary structure elements are not apparent. We also compared 15N HSQC spectra for 15N-labeled ∆N and ∆C mutants to the full-length SutA (Figures S2 and S3). Deletion of either tail had minimal impact, affecting only the 2-4 residues adjacent to the newly created terminus.
The difficulty of making unambiguous assignments for all residues and the high likelihood that much of the protein is intrinsically disordered precluded building a full NMR-based structural model of SutA. To model some of the conformations that might be adopted by SutA, we used the Robetta Server and PyRosetta to perform low-resolution Monte Carlo–based modeling, using the chemical shifts and RDC values from our NMR analysis to guide fragment library construction (21–23). Figure 1D shows a resulting model. The most highly conserved residues are found in the α-helix, and the C-tail is also highly conserved among homologs that have it. The N-tail is less conserved and varies in length, but is generally quite acidic. Figure S4 shows additional models (see Extended Materials and Methods for modeling details).
SutA binds to the β1 domain of RNAP
To investigate the binding interaction between SutA and RNAP, we used cross-linking and protein footprinting to map the region of RNAP with which SutA interacts. First we used the homobifunctional reagent bis(sulfosuccinimidyl)suberate (BS3), which cross-links primary amines that are within about 25 Å of each other (24). We added BS3 to complexes formed with purified core RNAP and SutA (Figure S2A), used the peptidase Glu-C to digest cross-linked complexes, and subjected the resulting fragments to LC-MS/MS. Analysis with the software package Protein Prospector (25) identified species that comprised one peptide from SutA and one peptide from RNAP (see Extended Materials and Methods and Figures S5 and S7), which allowed mapping of cross-link sites. We also used the photoreactive non-canonical amino acid p-benzoyl-L-phenylalanine (BPA), which, when irradiated with UV light, can form covalent bonds with a variety of moieties within 10 Å (26, 27). We introduced BPA at 9 different positions of SutA (residue 6, 11, 22, 54, 61, 74, 84, 89, or 100); we then formed complexes with purified core RNAP and each of the BPA-modified SutA proteins, irradiated them with UV light, and visualized cross-linked species following SDS-PAGE (Figure S6). For the most efficient cross-linkers (BPA at positions 54 and 84), we determined the sites of the cross-links on RNAP by identifying cross-linked peptides via StavroX (28) analysis of LC-MS/MS data after tryptic digest of the complexes (Figure S8).
Both cross-linking approaches identified interactions between the central region of SutA and the β1 domain or nearby regions of the β subunit of RNAP (Figure 2A and B, green and orange). All SutA residues participating in the cross-links were within the α-helix region (BS3) or just outside it (BPA). BPA cross-linking is sensitive to the orientations of the residues, so BPA residues within the helix that did not cross-link efficiently may not have been oriented optimally.
To identify the positions of the N- and C-tails, we designed variants of SutA for affinity cleavage experiments. We introduced cysteine residues at SutA position 2, 32, or 98 and conjugated the chelated iron reagent, iron-(S)-1-[p-(bromoacetamido)benzyl]EDTA (FeBABE), to these cysteines. FeBABE catalyzes localized (estimated to occur within 12 Å of the FeBABE moiety) hydroxyl radical cleavage (29). We assembled complexes with the FeBABE-modified SutA variants and core RNAP, initiated the cleavage reactions, and analyzed the cleavage products by SDS-PAGE followed by Western blotting with a monoclonal antibody against a peptide near the C-terminus of β. To map the cleavage sites, products were compared to C-terminal β fragments with known N-terminal endpoints (Figure S9). While the primary cleavage product of the N-terminal FeBABE (at residue 2; N-Fe) was in the cleft between the β1 and β2 (a.k.a. β lobe) domains, the strongest cleavage product of the C-terminal FeBABE (at residue 98; C-Fe) was in the long α-helix on the inside surface of β1 (designated α6 (30)), amongst BS3 and BPA cross-linking sites (Figure 2A, blue). The FeBABE at residue 32 induced cleavage at both β positions, suggesting that the N-tail remains mobile to some degree even when bound to RNAP.
We also detected possible interactions with βi9, which is an insertion in the β flap domain (31): BPA at residue 84 crosslinked to β967, and weak cleavage products were detected at β721 for the N-Fe variant and β1058 for the C-Fe variant (Figure 2B). β967 and β484/493 residues that were strongly cross-linked to BPA84 are too far apart to be reached from a single, stably bound position of SutA 84. However, we did not detect more than one shifted band after cross-linking with the 54 or 84 BPA variants (Figure S2B), suggesting that two separate sites on β are not likely to be occupied by two SutA molecules at the same time. Instead, it may be that SutA’s inherent flexibility, combined with a binding interaction with a surface on the outside of the β1 domain that allows some rotation or translation of SutA, could allow for all of the observed cross-links and cleavages.
To corroborate SutA-β interaction without cross-linking or cleavage and assess which residues of SutA might directly participate, we conducted an NMR experiment. We were able to purify only a small amount of soluble β1 domain (colored darker blue in Figure 2C), which we mixed with an equimolar amount of 15N-labeled full-length SutA. As a control to rule out non-specific interactions, we mixed SutA with an equimolar amount of σS, which does not appear to bind SutA. Several SutA residues showed chemical shift perturbations in the β1 mixture, compared with the σS mixture (Figure 2D). Three of these residues, K95, D97 and K99, would be on the same side of an extended peptide chain in the C-tail, suggesting that this tail could directly interact with β1 in an extended conformation. However, the C-Fe SutA variant induced weaker cleavage than the N-Fe variant, suggesting that this interaction is probably not the only binding determinant. The other perturbed residues flank the α-helix, suggesting that the regions at the junctions with the flexible tails may change conformation upon binding to β.
SutA activates the rrn promoter in vitro
Next we investigated SutA effects on transcription. We focused on the rRNA promoter because our ChIP data suggested that SutA directly affects rrn initiation (1). We first asked whether SutA affects transcription by the closely related E. coli RNAP, for which extensive in vitro tools are available. Overexpressing SutA in E. coli did not lead to rrn upregulation in vivo as it did in P. aeruginosa (Figure S10), necessitating the use of a cognate P. aeruginosa in vitro transcription system. We purified the core RNAP natively from a ∆sutA strain using a protocol originally designed for E. coli RNAP and previously used to purify RNAP from P. aeruginosa (32–34). The P. aeruginosa homologs of σS, σ70, and DksA (as well as SutA) were heterologously expressed in E. coli with cleavable N-terminal 6xHis tags and purified by metal affinity and size-exclusion chromatography.
Unlike its well-studied E. coli counterpart, the P. aeruginosa rrn initiation region has not been characterized. We mapped the dominant rrn transcription start site using 5’-RACE to a cytidine 8 bp downstream of a -10 consensus sequence (Figure S11). We produced linear templates of 120-170 bp containing the rrn promoters and 42 bp of transcribed sequence for use in single-turnover initiation experiments (see supplement for details).
Transcription initiation proceeds via a multi-step pathway consisting of: 1)formation of a closed complex between the double-stranded DNA and the RNAP holoenzyme; 2)initial DNA strand separation, followed by isomerization through several open intermediates into a final open complex (OC) in which the +1 position of the template DNA strand is loaded into the active site and the downstream DNA duplex is stably held by RNAP; 3)initial abortive rounds of nucleotide addition; and 4)promoter clearance and transition into the elongation phase. Any of these steps could theoretically be affected by a regulator, and the details of this pathway differ at different promoters (35). Because much of the control of the E. coli P1 depends on the inherent instability of its OC (7), we sought to determine 1) whether the P. aeruginosa rrn has similar structure and response to regulatory inputs, and 2) whether SutA affects the rrn OC stability. The P. aeruginosa rrn promoter shares some of the features known to contribute to E. coli rrn P1 OC instability (Figure 3A): suboptimal spacing (16 nt vs the optimal 17-18 nt) between near-consensus -35 and -10 hexamers, a GC-rich discriminator region, and a C residue 2 nt downstream of the -10 hexamer that cannot make productive contacts to σ70 (36). However, the P. aeruginosa discriminator is 7 nt, which is longer than the optimal 6 nt, but still one base shorter than that of E. coli. Also, the initiating nucleotide is a cytidine rather than the adenosine or guanine iNTPs found in E. coli, indicating potentially different regulatory connections to cellular energy levels.
First, we directly measured the half-life of the heparin-resistant Eσ70 OC in a transcription-based assay. In contrast to what has been seen in E. coli (37), we detected some OC at standard salt concentrations and on a linear template, but its half-life was quite short, at about 45 seconds. Addition of SutA at 125 or 500 nM had no significant effect (Figures 3B and S12). Next, we measured effects of ppGpp/DksA and increasing [iNTPs], which repress and activate transcription from the E. coli rrn P1, respectively, in the presence or absence of SutA (Figures 3C, S13, and S14). As observed in E. coli, rrn transcription was strongly repressed at low [iNTPs] and by DksA/ppGpp, but SutA did not significantly counter these effects. Taken together, these results suggest that while the P. aeruginosa rrn promoter forms an inherently unstable OC, which is sensitive to regulatory inputs that utilize its instability, SutA does not alter the OC stability.
To directly measure the effects of SutA on transcription initiation, we performed single turnover initiation assays using the wild type (WT) SutA and the ∆N- and ∆C-tail variants described in Figure 1. Because EσS binds the rrn locus in vivo during stationary phase in E. coli (38), we wanted to investigate whether SutA effects on rrn transcription in vivo could be mediated through EσS or Eσ70, or both. We found that WT SutA increased transcription by both holoenzymes in vitro, but the magnitude of the effect was much larger for EσS (up to 400% increase) than for Eσ70 (up to 70% increase) (Figures 3D, S15). In both cases, the effect saturated at concentrations of SutA between 125 and 500 nM. The acidic N-tail is strictly required for activation, as the ∆N mutant inhibited transcription in a dose-dependent manner. The ∆C mutant was still able to enhance transcription, albeit with a small shift in the concentration dependence evident with EσS. This shift may reflect C-tail interactions with EσS: we observed that the chemical shifts of three residues in the C-tail were perturbed upon mixing with the β1 domain.
A disordered acidic loop in σ70 modulates SutA binding
We wondered what difference between σ70 and σS could explain the difference in SutA’s impact on rrn initiation by Eσ70 compared to EσS. Domains 2, 3, and 4 are highly similar, and both σ70 and σS have unstructured acidic regions, referred to as 1.1, near their N-termini (39). However, σ70 contains a large (~245 amino acids) insertion, termed the “non-conserved region” or NCR, which is not present in σS (Figure 4A). Crystal and cryoEM structures show that most of the NCR is situated relatively far from the β1 binding site of SutA, contacting the β’ subunit on the opposite side of the main channel of RNAP, but an acidic stretch of ~40 residues within the NCR is too flexible to be resolved in these structures (herein AL for Acidic Loop) (40, 41).
To investigate possible interactions between the AL of σ70 and SutA, we threaded the P. aeruginosa sequence onto the β subunit of an E. coli RNAP crystal structure (42), docked that model into the recent cryoEM structure of the E. coli Eσ70 OC (41), and modeled the missing AL (using the E. coli sequence for both the structured and flexible regions of σ70) using the MODELLER software suite (43). The highly flexible AL could occupy a wide range of positions (e.g., Figure 4A, top), some of which would stay well above the DNA in the main channel (position 1) and some of which would clash with the DNA (position 2), and it could reach the β1 residues that participate in SutA cross-links, especially in the absence of DNA. In contrast, σS has no corresponding flexible region and remains far from the SutA cross-links (Figure 4A, bottom).
To determine whether the AL might contribute to the observed differences between Eσ70 and EσS activation by SutA, we constructed and purified a P. aeruginosa σ70 mutant lacking residues 171-214 (∆AL), which correspond to the region missing in the E. coli structure, and repeated our cross-linking and cleavage assays using Eσ70, EσS, or Eσ70∆AL holoenzymes instead of just the core enzyme. In the absence of DNA, SutA L54BPA cross-linked to Eσ70 less efficiently than to E or EσS. Interestingly, Eσ70∆AL largely restored the cross-linking to the levels seen with E or EσS (Figure 4C, lanes 1-4), suggesting that the σ70 AL modulates the SutA interaction with Eσ70. The difference in cross-linking efficiency between Eσ70 and Eσ70∆AL decreased at higher SutA concentrations, as might be expected if SutA and AL are competing to occupy a similar space.
SutA competes with DNA in the final open complex
Our cross-linking and cleavage results suggested that SutA’s position on RNAP might allow it to compete with the promoter DNA. To explore this possibility, we added to our crosslinking assay either a double-stranded (ds) rrn promoter DNA or a bubble template in which the region of the DNA that forms the transcription bubble in the OC was non-complementary (Figure 4B). The dsDNA requires σ to melt the DNA strands, and will support a native population of promoter complex intermediates. By contrast, the bubble template obviates the need for σ and would be expected to stabilize an OC formed with the holoenzyme, but this complex may not represent the dominant native complex, as the E. coli rrn P1 does not form a stable final OC (37). The addition of the bubble DNA had a large negative effect on SutA binding that was synergistic with the presence of σ (Figure 4C, lanes 5-8). Cross-linking could still be readily detected in the absence of σ, and to a lesser extent when σS was present, but not with either σ70 or σ70∆AL; longer exposures revealed that cross-linking did occur at low efficiency (Figure S16). Addition of dsDNA allowed more SutA binding, but still less than in the absence of DNA (Figure 4C, lanes 9-11).
The flexible SutA tails approach the transcription bubble
The flexible, acidic σ1.1 region competes with promoter DNA for binding to the main channel of RNAP, but still enhances initiation from some promoters (44). We wondered whether SutA might likewise use its acidic N-tail to compete with promoter DNA, but enhance initiation at the rrn promoter. The BPA crosslinking reports on the interactions established by the central region of SutA, but gives no information on the position of its flexible tails, and a decrease in crosslinking could be due to a loss of binding or a change of SutA conformation. We used FeBABE cleavage assays with the N-Fe and C-Fe SutA variants (Figures 4D, S17) to address these questions. We found that the addition of σ70 had a much larger negative effect on cleavage induced by C-Fe than on cleavage induced by N-Fe. The σ70∆AL mutant partially restored C-Fe cleavage levels to those observed with the core enzyme or EσS, but caused a decrease compared to σ70 in the N-Fe cleavage at residue 721. This suggests that the σ70AL does not fully displace SutA, but instead interferes with a binding interaction of the C-tail (Figure 4F; compare top right to top left panel), decreasing the crosslinking efficiency of the 54BPA variant, consistent with our observation that the ∆C mutant required higher concentrations for maximal activity on EσS but not on Eσ70 (Figure 3D).
By contrast, the effects of template DNA were similar for both C-Fe and N-Fe cleavage reactions, as well as for BPA crosslinking. This suggested that DNA might induce SutA dissociation, rather than its subtle repositioning (Figure 4F; compare top to bottom panels), prompting us to investigate whether SutA and DNA could form a ternary complex with RNAP holoenzyme. We measured FeBABE SutA-dependent cleavage of the template and non-template DNA strands using primer extension. We saw stronger cleavage with EσS than with Eσ70, but in both cases the signal was relatively weak, as might be expected for a factor that does not directly bind DNA (Figures 4E, S19). In the EσS complex, C-Fe induced cleavage of both strands between residues -8 and -12, suggesting that it remains near the upstream fork junction of the transcription bubble. N-Fe cleaved the template strand near the upstream junction but also cleaved both strands further downstream. For Eσ70, the cleavage was weaker overall and showed a different pattern; for C-Fe in particular, more cleavage took place on the downstream region of the non-template strand. This difference could reflect the AL-mediated repositioning of the C-tail.
Our results argue that SutA may not stably bind the final OC. However, SutA-induced DNA cleavage demonstrates that SutA does bind to some promoter complexes, in turn suggesting that an earlier intermediate in the initiation pathway may be the main target of SutA activity (Figure 4F).
DISCUSSION
As part of their response to fluctuating environmental conditions, bacterial cells produce regulators that directly bind RNAP and modify its behavior, eliciting global changes in gene expression patterns, in addition to producing different DNA-binding transcription factors that help recruit RNAP to specific genes (reviewed in (45)). We previously identified SutA as a global regulator that binds RNAP and contributes to a broad response to protracted growth arrest, enhancing ongoing, low-level expression of housekeeping genes (1). SutA directly affects initiation at the rrn promoter, prompting comparison to other regulators that affect rRNA expression by directly binding RNAP, such as DksA, ppGpp, and CarD. DksA and ppGpp, which appear to operate similarly in P. aeruginosa and E. coli, broadly destabilize OCs, leading to repression of rrn P1 and activation of amino acid biosynthesis genes in response to nutrient downshifts (46). In contrast, CarD, constitutively expressed in many non-Gammaproteobacteria, broadly stabilizes OCs and modestly enhances rRNA transcription (approximately 3-fold in vitro) (12). SutA is distinct from both of these examples: though it acts on a P. aeruginosa rrn promoter that also forms an unstable OC with Eσ70, its activity does not affect the stability of this complex. This difference is perhaps unsurprising, as both its structure and interactions with RNAP are also distinct, as discussed below.
A model for SutA interactions with promoter complexes
Unlike DksA and CarD, which have well-defined structures (12, 47), SutA is largely intrinsically disordered, with its flexible tails playing key functional roles. SutA binds to the β1 domain of RNAP; its crosslinking and cleavage interactions suggest a binding site that is close to but distinct from that of CarD (12), and far from the sites occupied by DksA and ppGpp (11, 48). Although the extreme flexibility of SutA and the relatively large distances over which our cross-linking and cleavage reagents could act (10-25 Å) preclude precise docking of SutA onto RNAP, a binding site on the outside of the β1 domain is consistent with our data. SutA failed to activate rrn transcription in E. coli in vivo (Figure S10) and failed to bind the E. coli Eσ70 in vitro (Figures S16 and S17), suggesting that its binding site is in a region that is different between the two polymerases. Most of the β residues are identical (72%) or similar (87%) between E. coli and P. aeruginosa, but two β1 loops that contain residues involved in BS3 cross-linking (K45 and K116) are among a small number of amino acid sequences with reduced similarity (Figure S20). From such a binding site for the SutA helix, its flexible tails could reach into the main channel of RNAP through the cleft between β1 and β2. The fact that FeBABE-modified SutA variants could catalyze cleavage of the rrn DNA suggests that the tails approach the DNA in some promoter complexes.
The β1/β2 cleft has been shown to accommodate the non-template strand during the early rounds of nucleotide addition, when it must scrunch to allow additional bases of the downstream DNA to enter the enzyme before the upstream contacts are released (49); on E. coli rrnB P1, scrunching occurs even before initiation (50). Interestingly, the β1/β2 cleft is also the site of several point mutations that suppress the auxotrophy phenotype of a ∆dksA mutant in E. coli, consistent with a model where modulating its interaction with the DNA could be functionally important in growth-phase-dependent gene expression regulation (51). Our results show that the fully melted promoter DNA inhibits SutA binding, suggesting that DNA and SutA may compete for similar contacts with RNAP on β1 or near the β1/β2 cleft. SutA (and especially its N-tail) could positively influence the formation of the rrn OC through interaction with early promoter complex intermediates, and then be displaced as the final OC forms, potentially leading to its dissociation (Figure 4F). This is analogous to the regulatory mechanism of σ70 1.1, an acidic flexible region that binds in the main RNAP channel in early promoter complexes and must be ejected to accommodate the promoter DNA that binds to the same site in the final OC (44, 52–55). Like SutA, σ70 1.1 stimulates initiation at some promoters but does not affect the stability of their OCs (44).
The roles of acidic disordered regions in SutA regulation and beyond
In addition to the critical role of SutA’s unstructured acidic N-tail in mediating its enhancement of rrn transcription, we also found that an unstructured, acidic region of σ70, the AL, directly modulates SutA’s activity, possibly by interfering with the ability of the C-tail to bind β1. The AL is part of the NCR region that is unique to σ70 (56), so this interaction may contribute to the difference in the SutA’s effects on initiation by Eσ70 versus EσS. The NCR makes contacts with the upstream DNA duplex (41), and our modeling suggests that the AL could be positioned near the upstream junction of the transcription bubble. Like region 1.1 and the SutA N-tail, the σ70 AL is a highly dynamic element that could modulate the DNA trajectory in early intermediates in open complex formation, before the bubble is locked in place in the final OC, and we found that in addition to affecting the RNAP-SutA interaction, σ70 ∆AL also has mild defects in initiation at the rrn promoter on its own (Figure S18). Since the OC formation pathways and the relative occupancies of the intermediates vary among promoters (37), the effects of these unstructured elements are expected to be distinct for different promoters.
Dynamic interactions of intrinsically disordered (and often highly acidic) modules play key roles in eukaryotic transcriptional regulators, bacteriophage proteins, and σ factors. Unstructured regions can gain access to and remodel dynamic regions of transcription complexes, as in the case of the phage proteins Gp2 and Nun, leading to inhibition of RNA synthesis (57, 58). They also can bind or mimic flexible nucleic acid sequences, as in the case of the λN protein (59) or σ1.1 (55), and activate transcription. In the case of eukaryotic transcriptional activators such as Gcn4 or Ino2, they can serve as flexible protein-protein interaction domains, capable of mediating interactions whose structural constraints vary depending on nuances of nucleic acid sequences, chromatin states, and other aspects of the intracellular environment (60–62). While their disordered nature has made many of these domains difficult to study using traditional structural and biochemical approaches, it is becoming increasingly clear that they play critical roles in many aspects of transcriptional regulation in all domains of life, and SutA adds to this growing body of evidence.
Implications for the in vivo role of SutA during slow growth
Our finding that SutA can differentially impact transcription in a σ-dependent manner could have important implications in vivo during growth arrest. σ70 and σS are closely related σ factors with partially overlapping promoter specificities (40, 56, 63). Our previous RNA-Seq results imply that SutA interactions with both holoenzymes are likely to be functionally relevant, as some affected genes are bona fide σS regulon members (63), but overall the affected genes are biased toward classic targets of σ70 (1). In E. coli, the activities and relative abundances of EσS and Eσ70 change throughout different growth phases, with σS upregulated at the transition to stationary phase (64, 65), and much of Eσ70 sequestered by the 6S RNA late in stationary phase (66). In addition, EσS and Eσ70 appear to be differentially sensitive to changes in cellular conditions that occur during stationary phase, such as an overall decrease in the negative supercoiling of the chromosome, shifts in patterns of nucleoid associated proteins, and different concentrations of solutes. Moreover, in specific cases that have been examined in vitro, EσS initiation efficiency increases under the stationary phase-associated condition, while Eσ70 initiation efficiency decreases (67, 68). These characteristics of EσS may in part explain why σS ChIP signal at the rrn promoters increases in stationary phase in E. coli and σ70 ChIP signal decreases (38). The ability of SutA to enhance initiation by EσS and Eσ70 differentially could allow greater flexibility during different stages of growth arrest. For example, SutA may enable baseline levels of housekeeping gene expression regardless of which holoenzyme ismost available and active, or could allow for combinatorial control whereby promoter activity during dormancy would be synergistically affected by σ preference and SutA interaction. More work will be required to fully understand how SutA contributes to the regulatory architecture that allows P. aeruginosa to thrive during dormancy, but this study represents important mechanistic insight into the function of this global regulator.
MATERIALS AND METHODS
See Extended Materials and Methods in SI for additional details about all experiments, for strain construction details, and for tables of strains and primers used.
Protein purifications
P. aeruginosa core RNAP was purified as previously described(32–34). N-terminal 6xHis-tagged SutA, SutA variants, DksA, σS, σ70, σ70∆AL, and β1 were heterologously expressed in E. coli and purified by standard metal affinitiy chromatography followed by cleavage of the 6xHis tag with TEV protease and size exclusion chromatography. For NMR experiments, cells were grown in minimal media prepared with 15NH4Cl or 13C glucose or both. For BPA crosslinking, amber stop codons were introduced at positions of interest and BPA was incorporated via amber suppression following co-transformation of the SutA plasmid with pEVOL-pBpF as previously described (26). For preparation of FeBABE variants, cysteine residues were introduced at positions of interest (SutA has no natural cysteines) and following purification of the protein, the FeBABE moeity was conjugated to the cysteine as previously described (29). Conjugation efficiencies were estimated to be 57%, 38%, and 76% respectively for the residue 2, 32, and 98 variants.
NMR experiments
Data were collected from SutA proteins at concentrations of 300 µM in a buffer containing 20 mM sodium phosphate, pH 7.0, 100 mM sodium chloride, and 10% D2O. For the 46-101 variant, the following spectra were acquired on a Varian Inova 600 MHz NMR with a triple resonance inverse probe running VnmrJ 4.2A: 15N HSQC, 13C HSQC, HNCO, HNCA, HNCACB, CBCACONH, HNCOCA, HNCACO, CCONH, and 15N HSQC experiments modified for measurement of T2 and of 15N-1H NOE. For the full-length protein, 15N HSQC, 13C HSQC, HNCACB, and CBCACONH spectra were acquired at 7 °C on a Bruker AV III 700 MHz spectrometer with a TCI cryoprobe running Topspin 3.2, but 15N HSQC experiments modified for measurement of T2 and of 15N-1H NOE were collected on the Varian Inova 600 MHz NMR, as were 15N HSQC spectra for the SutA ∆N and SutA ∆C SutA proteins. 15N13C-labeled full-lengthSutA was embedded in a stretched polyacrylamide gel for measurement of residual dipolar couplings as previously described (20), using the Varian Inova 600 MHz NMR. To assess SutA binding to β1 by NMR, 15N-labeled SutA and β1 fragment were buffer exchanged into 20 mM sodium phosphate, pH 7.0, 100 mM sodium chloride and the resulting complex was isolated by size exclusion chromatography, resulting in a final concentration of complex of approximately 25 µM. In addition, 15N-labeled SutA was mixed with σS at 50 µM each. 15N HSQC spectra were acquired on a Bruker 800 MHZ AV III HD spectrometer with a TCI cryoprobe at 25 °C. Peak assignments and analysis were done using the PINE Server, CcpNmr Analysis Suite, and MestreNova software.
Crosslinking experiments and data analysis
BS3 crosslinking of core RNAP and SutA was carried out as previously described (24) with modifications. Crosslinked complexes were subjected to in-solution digestion by the Glu-C peptidase, and the resulting fragments were analyzed by LC-MS/MS on an Orbitrap Elite Hybrid Ion Trap MS. Crosslinked pepties were identified as previously described, with modifications (25). BPA crosslinking was achieved by irradiating RNAP core or holoenzyme complexes with UV light from an Omnicure S2000 lamp, complexes were digested in solution with trypsin, and analyzed by LC-MS/MS on a Q Exactive HF Orbitrap MS. Crosslinked peptides were identified using the StavroX software package (28).
FeBABE experiments and analysis
Cleavage reactions of holoenzyme complexes assembled in TGA buffer were initiated by the addition of ascorbate and hydrogen peroxide to final concentrations of 5 mM each, as previously described (29). For measuring protein cleavage, reactions were quenched by the addition of SDS loading buffer and were evaluated by SDS-PAGE followed by Western blotting, using a monoclonal antibody raised against a peptide from the extreme C-terminus of E. coli β (EPR18704 from Abcam). To generate standards for size comparison, several different C-terminal fragments of RpoB with endpoints ranging from aa 355 to aa 1062 were overexpressed in E. coli and crude lysates from these strains were subjected to SDS-PAGE and Western blotting alongside the FeBABE cleavage products. For measuring DNA cleavage, reactions were quenched with thiourea and treated with proteinase K. The DNA was precipitated, and subjected to primer extension using Cy3- or Cy5-labeled primers against the non-template or template strand respectively.. Products were separated on denaturing 12% polyacrylamide gels and imaged by laser scanner.
ACKNOWLEDGMENTS
We thank Ben Ramirez (University of Illinois at Chicago) for helping us with preliminary NMR studies of SutA, Jacqueline Barton (Caltech) for giving us access to her lab to perform experiments involving radioactivity, Nate Glasser for help with HPLC measurements to quantify SutA, Hsiau-Wei (Jack) Lee and Aimee Marceau (University of California, Santa Cruz) for help with the NMR binding experiment using the Bruker AVIII HD 800 MHz NMR, Weidong Hu (City of Hope) for help with NMR experiments using the Bruker AV III 700 MHz spectrometer, and Julia Kardon and Niels Bradshaw (Brandeis University) and members of the Newman lab for feedback on the project at different stages. MB was supported by a post-doctoral fellowship from the Cystic Fibrosis Foundation. Grants from the NIH (GM067153) to IA and grants from the HHMI and NIH (5R01HL117328-03 and 1R01AI127850-01A1) to DKN supported this work. The Proteome Exploration Laboratory is supported by the Beckman Institute and NIH 1S10OD02001301. This work was also supported by the Institute for Collaborative Biotechnologies through grant W911NF-09-0001 from the U.S. Army Research Office. The content of the information does not necessarily reflect the position or the policy of the Government, and no official endorsement should be inferred.
Footnotes
Some data has been removed and the remaining data more thoroughly described; Supplemental data moved to a separate file.