ABSTRACT
Genome architecture has proven to be critical in determining gene regulation across almost all domains of life. While many of the key components and mechanisms of eukaryotic genome organization have been described, the interplay between bacterial DNA organization and gene regulation is only now being fully appreciated. An increasing pool of evidence has demonstrated that the bacterial chromosome can reasonably be thought of as chromatin, and that bacterial chromosomes contain transcriptionally silent and transcriptionally active regions analogous to heterochromatin and euchromatin, respectively. The roles played by histones in eukaryotic systems appear to be shared across a range of nucleoid-associated proteins (NAPs) in bacteria, which function to compact, structure, and regulate large portions of bacterial chromosomes. The broad range of extant NAPs, and the extent to which they differ from species to species, has raised additional challenges in identifying and characterizing their roles in all but a handful of model bacteria. Here we review the regulatory roles played by NAPs in several well-studied bacteria and use the resulting state of knowledge to provide a working definition for NAPs, based on their function, binding pattern, and expression levels. We present a screening procedure which can be applied to any species for which transcriptomic data are available. Finally, we note that NAPs tend to play two major regulatory roles – xenogeneic silencers and developmental regulators – and that many unrecognized potential NAPs exist in each bacterial species examined.
Introduction
Every cell must solve an incredible challenge: to organize negatively charged genetic material, DNA, that is orders of magnitude longer than the width of the cell. The DNA must be compacted and electrostatically neutralized, and yet it must also be accessible to enable changes in gene expression necessitated by changing environmental conditions. In eukaryotes, compaction of DNA is mediated by histones – basic proteins that form an octamer around which DNA is tightly wrapped into structural units called nucleosomes [1]. These nucleosomes then wrap to form chromatin fibers [2,3]. There are two broad categories of chromatin that largely relate to the accessibility of DNA to RNA polymerase binding and transcription: euchromatin, which is loosely packed and accessible for transcription, and heterochromatin, which is inaccessible and silent [4]. Along with histones, post-translational modifications of histones and other chromatin-associated proteins are collectively termed the “histone code”, which impacts chromatin structure and gene regulation [2,3]. The histone code is capable of both repressing and promoting transcription, flexibility provided by the varying sites and types of post-translational modifications to histones [5]. In addition to maintaining physical organization of the genome, chromatin facilitates cell-type memory as tissue-specific cells pass information within the histone code to daughter cells. This passage of information – often termed “transcriptional memory” – is critical for the viability of the organism [6,7]. In humans, changes in chromatin that lead to global genome misregulation and instability have been linked to cancer [8–11], implicating chromosome structure as an essential facet of genome maintenance and health.
In the case of bacteria, it was widely believed until relatively recently that within the membrane-free bacterial nucleoid the genome was universally accessible for transcription [12–15]. However, recent work suggests that genomic context plays a significant role in the propensity for genes at a given locus to be transcribed, irrespective of gene dosage effects [16–18]. Aside from availability for transcription, several lines of evidence have long suggested that the bacterial nucleoid has an organized topology [19–21]. With modern advances in technology, it is now clear that bacteria maintain a discrete species-specific nucleoid shape that changes through different growth phases or conditions [22–27] (Figure 1). The tools that exist to study genome architecture in eukaryotes cannot simply be applied to bacteria, so much of our understanding of genome organization in bacteria, and how it may impact gene regulation, is limited. We do understand that overall chromosome compaction is facilitated by highly abundant nucleoid associated proteins (NAPs) [28–30] (Fig. 1–2). However, NAPs exhibit broad binding specificity and distinct NAPs often overlap in terms of which genomic loci they bind. These facets of NAPs make it difficult to precisely profile a single NAP’s contribution to gene regulation (as does cross-regulation between NAPs), although the fact that they make essential contributions has long been recognized. It is also difficult to make a binary distinction between the definition of a “NAP” and a transcription factor, as there are often identifiable evolutionary connections between proteins placed in those bins [31]. Indeed, in recently considering how to distinguish between what constitutes a “transcription factor” versus a “NAP”, Dorman and colleagues rightly point out that the effort to strictly distinguish between the two categories likely constitutes a false dichotomy, citing numerous examples of overlap between characteristics of NAPs and TFs with large regulons, and the structural and evolutionary connections between members of the two categories [31]. It has even been speculated that transcriptional regulation by NAPs may be the ancestral state and that the evolution of more specific transcription factors is a more recent innovation [32]. However, even if there are not absolute distinctions between a “NAP” and a “global regulator” or “transcription factor”, we will discuss below the extent to which these categories represent useful conceptual groupings for considering the functions of different abundant DNA-binding proteins.
Figure 1.
Nucleoid associated proteins mediate a variety of DNA conformations and are abundant in different stages of growth. Overview of highly abundant NAPs and their relative abundance in the different stages of growth in E. coli. Single subunit molecular weights (kD), oligomeric states, and binding preferences were obtained from [221], binding capabilities (filaments, bendings, etc.) were obtained from [28,221]. The H-NS binding depicted is an example of a bridged DNA filament formed between Hha, StpA, and H-NS
Figure 2.
Comparison of abundance of NAPs to a local regulator. Molecules per cell (log2) of nucleoid associated proteins (NAPs; green) and the transcription factor LacI (gray) in E. coli grown in rich media (data from [221])
Here, we will explore the current state of knowledge regarding transcriptional regulation by NAPs, aiming both to provide a reader with a comprehensive survey of references covering the development of our understanding of this topic, and to provide a useful cognitive framework for considering the regulatory logic implemented by NAPs. As the body of literature contributing to this field is immense and spans more than 80 years, it is impossible to exhaustively cover every study that has contributed to the field. Rather, we will focus on summarizing what is known now and what is currently being worked out. We also extend knowledge that has been obtained mainly in a handful of model organisms to the bacterial kingdom as a whole, by providing a set of broadly applicable criteria for what could reasonably be called a NAP, and how potential NAPs may be identified in currently less-well-studied organisms.
The organized bacterial nucleoid: E. coli as a case study
The hallmark of what defines a prokaryote vs. an eukaryote is the absence of membrane bound organelles. Instead, in most prokaryotes, the single circular chromosome is contained within the nucleoid, which is an irregularly shaped, organized mixture of negatively charged DNA [24,28,33]. The precise shape of the E. coli chromosome remains somewhat controversial and may involve either the formation of compacted “macrodomains” [34,35] or a well-mixed structure with a few specific long-range contacts formed between co-regulated loci such as ribosomal RNAs [27] (Figure 1). At a mesoscale level, the overall shape undergoes characteristic changes during stress, replication, and transition through phases of growth [22,23]. Two major determining factors of the overall shape of the nucleoid are NAPs and transcription [29,36]. Many parallels have been drawn between histones and NAPs, since NAPs have DNA binding domains, broad specificity, and the ability to compact DNA. However, the in vivo interactions that NAPs have with DNA have not been completely resolved. In E. coli, there are around a dozen NAPs that interact with DNA and other nucleic acids in various ways [28–30,37–39] (Figure 1). NAPs facilitate the formation of DNA bridges, filaments, bending, supercoiling, RNA mediated interactions, puncta, phase separated droplets, and impede RNA polymerase directly [28,40], which can (depending on the context) either promote or suppress transcription [24,29]. NAPs are highly abundant in the cell, but their abundance may differ as the nucleoid responds to different conditions. For instance, the NAP Dps is especially abundant in stationary phase and becomes the main component of the nucleoid [30], sequestering iron and protecting DNA from damage [41,42] (Figure 1). Transcription itself can also impact the shape of the nucleoid, as transcription generates positive supercoils and can influence the binding of NAPs [28]. An experiment to beautifully exhibit the influence transcription has on nucleoid shape is to treat cells with rifampicin, an antibiotic that blocks and inhibits RNA polymerase. The inhibition of transcription initially results in a compaction of the nucleoid due to the reduction in expansion from transcription-translation interactions, and then an expansion occurs from ribosomes and chromosome mixing [43,44]. Together, NAPs, transcription, and DNA form the organized nucleoid, all influencing each other in the process of growth and stress.
Bacterial chromatin influences transcription
New top-down technologies have uncovered additional evidence that heterochromatin-like regions – areas that are densely bound by protein and transcriptionally silent – exist in bacteria [16,28,45,46]. Therefore, it is important to define mechanisms that maintain and regulate chromatin in bacteria.
The structure of bacterial chromatin is largely defined by supercoiling and compaction of the DNA mediated by NAPs [33,43,47–50]. However, our understanding of what occurs in vivo remains incomplete and will require adaptation of tools to capture 3D structure. The five modes by which bacterial chromatin may impact RNA polymerase binding, and thereby impact transcription, are organized and presented in [28] but will briefly be discussed here due to their relevance: (1) occlusion of RNA polymerase binding: proteins bound to a promoter or transcription start site that prohibit RNA polymerase touch down, (2) blocking RNA polymerase progression: RNA polymerase is able to bind and initiate transcription, but cannot proceed due to a protein roadblock, (3) DNA topology: positive supercoiling is generated in front of an elongation complex and negative supercoiling is generated behind; negative supercoiling supports DNA unwinding and thus facilitates transcription initiation and inhibits termination, and the opposite is true for positive supercoiling [51,52], (4) RNA-mediated silencing: transcription factors that bind to nascent RNA transcripts can interfere with RNA polymerase termination, translocation, and pausing [53,54], (5) phase-separation: the formation of DNA condensates has been primarily shown in eukaryotes to control transcription [55], where DNA is compacted in droplets. However, whether phase separation mediates transcription in bacteria is unclear and is the focus of much ongoing research (e.g., [56]). Each one of these modes has been linked to NAPs, and specific NAP roles are described in detail below. We focus our initial discussion on E. coli, as a majority of work on bacterial chromatin has focused on this species or other related enterobacteria; our view will expand to a wide range of other bacterial species in the later portions of the present review.
Nucleoid-associated proteins mediate the formation of bacterial chromatin
NAPs mediate chromosomal structure and DNA compaction across bacterial species [29,57,58]. NAPs are loosely defined as having broad DNA binding specificity (usually preferring curved DNA and/or AT-rich DNA) and being highly abundant in the cell [29] (Figure 1). The majority form multi-protomer complexes (either homo – or hetero-oligomers), with some, such as H-NS and StpA, further extending together to form DNA filaments [28,29] (Figure 1). Crystal structures and in vitro experiments have led to insights into some of the interactions between nucleic acids (RNA and DNA) and NAPs [59–61]. Increasing evidence has implicated NAPs serving important functions as regulators of expression of horizontally acquired genes and pathogenesis [62–67]. H-NS is one of the most studied NAPs and has been shown to silence horizontally acquired DNA [68–70] by virtue of its relative AT-richness. While the enumeration of NAPs in E. coli varies from study to study, several of the most frequently noted NAPs are H-NS, Fis, HU, IHF, Dps, and Hfq (Figure 1). Here, we will briefly summarize what is known for the generally accepted NAPs that make up the main components of the nucleoid (Fis, HU, IHF, Dps, H-NS) and the highly conserved RNA chaperone (and DNA binding protein) Hfq.
Factor for inversion stimulation (Fis)
Factor for inversion stimulation (Fis) was named and first identified for its role in G-loop inversion of the bacteriophage Mu [71,72], but has a broader role in organization and maintenance of nucleoid structure [73,74], acting through binding as a homodimer to DNA to directly alter DNA structure and acting as a transcription factor to modulate expression of gyrase and topoisomerase I [75]. Its role in modulating gyrase and topoisomerase I is linked with its induction by high supercoiling levels in the cell [76] Fis is one of the most abundant NAPs in the cell (>60,000 copies per cell) during rapid growth. However, Fis abundance falls drastically entering stationary phase (<100 copies per cell) [30,77] (Figure 1). Fis expression is autoregulated, and together with H-NS and (p)ppGpp, which also impact the expression of Fis itself, Fis plays a major role in environmental responses and growth-phase-dependent changes in gene expression [78–80]. Fis can have both inducing [81–83] and suppressing effects [84–86] on gene expression, which largely depend on Fis abundance [75]. It is one of the major gene regulators of the cell, with 894 Fis-associated regions across the E. coli genome [73]. In the case of CspA, which is induced by cold shock and immediately upon dilution of cells into fresh media, Fis induces expression of cspA while H-NS represses [78]. (p)ppGpp negatively regulates both H-NS and Fis, and abolishes induction of cspA by inhibiting cspA promoter activity [79]. In total, this regulatory loop is an example of the complicated regulatory influence in which multiple NAPs and regulators can simultaneously engage. The global binding of Fis positively correlates with transcriptional propensity, measured by a reporter construct randomly inserted across the genome [16], suggesting that it either forms or is associated with transcriptionally activating contexts. At the same time, Fis also serves an overlapping role with H-NS in silencing xenogenic regions of the genome, and fis mutants exhibit a loss in silencing at similar prophages to H-NS mutants [86]. Fis comprises an α-helical core with four helices and an N-terminal domain that has a β-hairpin arm that facilitates DNA inversion [87–90]. As a homodimer, Fis has been shown to bend DNA to as large as a 90° bend, which stabilizes DNA looping, thus leading to compaction of DNA and effects on transcription [66,91,92]. In total, it is clear that Fis is a major gene regulator in E. coli; however, a fis mutant is still viable. It impacts transcription through its DNA binding capacity and thus can change nucleoid shape and global gene expression. Mutations within fis and topA that led to increased DNA supercoiling resulted in increased fitness in long-term evolution experiments involving daily transfers in minimal media, again tying cell viability with DNA topology and regulation [93].
Transcriptional dual regulator HU
(sometimes referred to as heat stable or heat unstable nucleoid protein across the literature) is the most abundant and highly conserved NAP [94–96], and in E. coli exists primarily as a heterodimer composed of subunits HupA and HupB, encoded by hupA and hupB, respectively. In rare cases, homodimers of each protein can form as well [97]. Like many other NAPs, HU is known as a regulator and organizer of the E. coli nucleoid. HU can bind RNA and linear dsDNA with low affinity but prefers DNA forks, sharp bends, bulges, and kinks [28,98]. Some amount of RNA binding activity is a common feature of many NAPs, including HU. HU interacts with key RNA molecules such as the rpoS transcript, which encodes the sigma S (σS) submit of RNA polymerase, RpoS. The subunit is an alternative sigma factor that is induced upon stress responses, starvation, and stationary phase to activate genes important for these environmental stresses and growth changes [99,100]. HU aids in efficient translation of RpoS by binding to the translation initiation region, thus playing a critical role in the induction of stationary phase and stress response genes [101,102]. It has also been suggested that RNA molecules bind to HU (and other NAPs) to stabilize the nucleoid and chromosome structure by mediating local DNA structure, supercoiling domains, and overall nucleoid morphology [103,104]. In vitro, HU modulates the shape of polyamine DNA condensates, facilitating the formation of rod structures [105]. However, it is unclear if the same pattern would be observed in vivo. HU plays a largely repressive role as an accessory factor that regulates key pathways involved in replication initiation [106], stress response [107], the Gal respressome [108], and outer membrane maintenance [109]. HU’s ability to bend DNA and form higher-order nucleoprotein complexes at promoters stabilizes dense structures that prohibit the ability of transcription initiation at that site [108,110,111]. There is no known inducer for HU, but similarly to Fis, HU is one of the major components of the nucleoid during exponential growth and becomes less abundant during later stages of growth [30]. Using soft x-ray tomography and comparing WT vs. HUαE34K, a HupA variant that inhibits the formation of higher-order HU-nucleoprotein complexes, it was found that oligomeric HU is critical for remodeling of the nucleoid through various phases of growth. It was also shown that HU facilitates transitions of a dynamic nucleoid core that changes throughout growth phases and is hypothesized to coordinate gene regulation by condensing the nucleoid in a specific manner through these transitions [112]. One of HU’s main modes of impacting gene expression comes from the introduction of negative supercoiling in the presence of topoisomerase I [113–115], or in some cases, stimulating topoisomerase I to remove negative supercoiling [116] (Figure 1). Its variety of regulatory roles, homo- and heterodimers, and binding modes leaves an incomplete picture of how HU mediates nucleoid structure and regulation.
Integration host factor (IHF)
Integration host factor (IHF) forms a heterodimer with subunits IhfA and IhfB, contributes to DNA supercoiling, is more abundant in exponential phase of growth compared to stationary phase [30], has a preference for curved DNA [117], plays a role in polyamine DNA condensation [118], and is largely an accessory factor to stabilize nucleoprotein complexes [119]. It has been found to play a role in major processes such as DNA replication, recombination, and gene expression [119–121]. IHF, as the name suggests, was initially discovered to be an essential factor for site-specific recombination of phage λ [122]. The binding and bending (which can be up to a 160° bend) of DNA by IHF positions and stabilizes the DNA, directing integrase molecules to bind to the DNA [123] (Figure 1). This interaction is not likely to be mediated by direct protein–protein interactions between IHF and λ integrase, but more likely relies on IHF bending of substrate DNA, as other sources of bending independent of IHF also promote integrase function [123–125]. The crystal structure of IHF bound to DNA shows that IHF binding specificity seems to be determined by the inherent structure of DNA imposed by AT-rich regions [126–130]. In terms of motif specificity, Fis and IHF can bind the same sites across the genome, and can lead to both repressive and activating effects [131]. IHF is a dual regulator that can either transcriptionally repress or activate targets, depending on the context; it plays a particularly prominent role in facilitating activation of σ54-dependent promoters [121]. In other contexts, what causes differences between the regulatory effects of different NAPs binding similar sites, whether it be from the mode by which NAPs bind, interaction with other DNA-binding proteins, or the amount of protein bound, remains unclear.
It should be noted that HU and IHF are structurally similar. However, IHF does not compensate for a lack of HU. Thus, while there are some overlapping targets and structural similarities, they have distinct roles [132–134]. DNA binding specificity is one of the major differences between HU and IHF, where IHF has defined binding motifs compared to HU’s relatively sequence-nonspecific binding to DNA. An example of this behavior is exhibited at the origin of replication (oriC), where HU binds randomly and forms various higher-order complexes, while IHF binds the same DNA sequence reproducibly and forms a distinct protein-DNA complex [135]. Furthermore, HU impacts IHF binding to oriC in a concentration-dependent manner, where the presence of HU leads to an increase in IHF-oriC complexes [135]. However, with higher molar ratios of HU to IHF, HU inhibits IHF’s ability to bind to oriC [135].
DNA protection during starvation (Dps)
DNA protection during starvation (Dps) forms a ferritin-like dodecamer with a hollow core [136], has low sequence specificity for DNA, is required for the proteomic response to prolonged starvation [42], and makes up more than half of the protein component of the nucleoid in stationary phase [137]. DNA and Dps form a DNA-protein crystal [138] that aids in protection of the DNA [42,139,140]. Dps is a main facilitator of DNA compaction during later stages of growth, largely combating the actions of Fis [74,141–143]. Similarly to ferritin, Dps serves an important role in iron acquisition and contributes to oxidative damage protection [144–147]. Dps is induced post-transcriptionally in times of carbon or nitrogen starvation and oxidative stress [141,148,149], and in pathogenic E. coli, Dps promotes acid tolerance [150]. Regulation of Dps has been extensively studied and summarized [151–157]. During exponential growth, Dps is degraded by proteases ClpXP. However, in times of carbon starvation, proteolysis is halted to maintain proper levels of Dps for DNA protection and compaction [148]. The compaction mediated by Dps does not repress transcription in vitro [56] (Figure 1). Furthermore, Dps excludes the restriction endonuclease KpnI, but not RNA polymerase, from accessing the DNA [56]; it was proposed that this may be indicative of exclusion of most or all sequence-specific DNA binding proteins from phase separated Dps-DNA condensates [56]. The potential for Dps to distinguish between DNA binding modes of the proteins it excludes may have profound regulatory implications or may indicate a broader protection of DNA from horizontally acquired restriction endonucleases. More exploration into the impact Dps has on genome regulation is required to fully appreciate its impact on the cell. It is also notable that at least under laboratory conditions, the close E. coli relative Salmonella can tolerate an exchange of the normally diametrically opposed growth phase-dependent expression profiles of Fis and Dps, although substantial changes in gene expression were observed [158] and the regulatory exchange caused both a drop in growth in laboratory media and altered infectivity in HeLa cells. These findings demonstrate that bacterial regulatory networks are able to largely compensate for even a substantial change in the timing of expression of these two NAPs, at least under laboratory conditions, although substantial regulatory effects were observed and it is not clear how the Fis/Dps swapped cells would fare under other stress conditions.
Histone-like nucleoid structuring protein (H-NS)
Histone-like nucleoid structuring protein (H-NS) is a small basic protein known to be a major repressor of gene expression [159–161], and particularly well characterized for its role in transcriptional silencing of horizontally acquired DNA (xenogeneic silencing [68]). H-NS is present at a roughly constant amount throughout various stages of growth, aside from a spike in early lag phase [30]. It is one of the first proteins to be synthesized when starved cells are brought into rich medium [162] and is also induced by insults such as cold shock [163] and iron starvation [164]. H-NS affects transcription of factors involved in a variety of processes, such as acid resistance [165], flagellar biogenesis [166], rRNA synthesis [167], protease expression [168], and metabolism [169]. The wide range of systems H-NS regulates and sequences it binds suggest that the regulatory role of H-NS relies on its role in impacting chromosomal structure. H-NS has a strong preference for AT-rich regions including AT-rich horizontally acquired DNA, and it regulates newly acquired DNA [165,170]. H-NS contributes to the compaction [171,172] and organization [173] of the nucleoid, is capable of supercoiling DNA [50,174–176] and forms different types of DNA filaments [28,61]. H-NS can form homodimers and multi-protein complexes. There are many paralogs of H-NS which may act as co-regulators alongside it [177–181]; the most studied H-NS paralog is StpA, which together form heterodimers [182]. While sharing similar sequence and structural features [61,183], H-NS is more abundant in the cell and has a lower affinity for DNA [184]. Loss of hns leads to an increase in expression of stpA [184,185]. StpA has been shown to partially compensate [186] for the loss of hns and to repress similar genes. However, loss of stpA shows minimal phenotypic effects, likely due to its lower abundance in the cell and overlapping regulatory roles by comparison to H-NS. In vitro, H-NS forms both linear (binding one DNA fragment) and bridge (“bridging” two fragments of DNA) filaments across dsDNA (Figure 1) that have the capacity to impede transcription by interfering with RNA polymerase [28,61]. Both linear and bridged filaments can impede transcription initiation by binding throughout the promoter and transcription start site [61]. Only bridged filaments promote RNA polymerase backtracking and subsequent ρ-dependent termination [61]. StpA can form these types of filaments with H-NS [61]. Additionally, in some species Hha, which can associate with DNA by interacting with H-NS and related proteins, but does not have a DNA binding domain, supports the formation of bridged H-NS filaments [61]. StpA and H-NS have been linked to RNA chaperone activity, further deepening the mechanistic options, these proteins have on impacting bacterial chromatin [187,188]. Despite considerable research into the physiological effects of H-NS, the mechanisms by which H-NS is recruited to target DNA, the consequences of and cues for its different modes of DNA binding, and the regulation of H-NS as whole is poorly understood.
Host factor for phage Q beta (Hfq): RNA chaperone and DNA binding protein?
Hfq is a well documented, conserved RNA-binding protein whose homohexameric ring has the propensity to bind RNA in a number of different conformations [189–192]. Early studies demonstrated that Hfq can bind both RNA and DNA, and that 10–20% of the Hfq in the cell is typically associated with the nucleoid [193]. Sequence analysis of Hfq revealed that it was related to Sm proteins found in eukaryotes and archaea, which similarly form ring structures that are the main unit of spliceosomal small nuclear ribonucleoproteins (snRNPs) [194,195]. These snRNPs are key building blocks of the spliceosome, which splice and process RNA [196]. Hfq facilitates and stabilizes small RNA interactions that repress mRNA translation and promote degradation for a number of RNA transcripts [189,197]. Through these interactions, it has been shown that Hfq mediates the translation of RpoS – a stress-induced sigma factor in both Salmonella typhimurium and E. coli [198–200]. The similarities between Sm proteins and Hfq, while incredibly fruitful to understanding evolutionary links across domains of life, has led to a bias toward studies focusing solely on Hfq’s RNA interactions, leaving its DNA binding capabilities largely uncharacterized. Hfq was originally identified as a gene required for phage Q beta propagation and RNA-directed synthesis in infected E. coli [201,202] but has been connected to a number of different processes. For instance, Hfq has been shown to associate with nascent transcripts and RNA polymerase, but the connection has not been made in vivo [203,204]. Cells lacking hfq exhibit pleiotropic phenotypes, such as impacts on cell division, increased negative supercoiling, and osmosensitivity, largely thought to be linked heavily to its RNA chaperone activity [205–209] and role in ribosome assembly [210]. Hfq has been shown to form foci in response to starvation [40] and plays a role in stress-induced mutagenesis [205]. Despite the historical emphasis in the literature on its RNA binding activity, Hfq has long been known to also bind double stranded DNA (especially curved DNA [211]), and to be a highly abundant component of the nucleoid [211], although the majority of Hfq is cytoplasmic. Additionally, Hfq has been shown to compact dsDNA, and recently a structure was resolved showing its interaction with DNA [59] (Figure 1). Like H-NS, Hfq is able to bridge dsDNA [59,212]. The combination of high protein abundance, sequence-independent DNA binding activity, and structural effects on DNA exhibited by Hfq are very similar to those of the other NAPs listed above, and thus while Hfq has historically been included in some enumerations of E. coli NAPs (e.g., [211]) and excluded from others (e.g., [28]), here we include it within the category of NAPs due to the increasingly recognized importance of its DNA binding activity. One obvious complicating factor is the fact that the cellular Hfq pool is partitioned between ribosome binding, RNA chaperone activities, and DNA binding, with the direct effects arising from each of those functions difficult to tease apart. Understanding Hfq’s role in regulating genes at the level of occupancy across the genome will give insight into the mechanism behind the wide variety of effects Hfq has on the cell.
Toward a generalizable working definition for NAPs
While the E. coli proteins discussed above are generally accepted as “nucleoid-associated proteins” in the literature (with the possible exception of Hfq), a unifying definition of NAPs has remained elusive. Certainly, the term as it currently exists does not generally refer to every protein that is associated with the nucleoid, but rather, to highly abundant proteins that bind DNA with limited sequence specificity and play large-scale architectural and regulatory roles. Noting the ambiguity of the terminology, Shen and Landick [28] recently recommended instead the use of the terms “chromatin or nucleoid-organizing protein” rather than nucleoid-associated proteins. Either of those terms is more informative, however, we will continue to use the term “nucleoid-associated protein” for the remainder of this review to maintain consistency with prior literature.
Toward the goal of developing useful categories of bacterial DNA-binding proteins, NAPs can be fairly easily separated from the majority of transcription factors simply on account of regulon size, as most transcription factors are primarily local regulators acting at only a few promoters by virtue of the scale-free architecture of the transcriptional regulatory network [213]. Another key distinction to be made is how lines of separation may be drawn between NAPs and “global regulators”, with the latter category referring to highly abundant transcriptional regulators with large regulons such as E. coli Lrp, CRP, and Fur. While there is likely some overlap between the categories [31], we propose that the key distinctions between a “NAP” (or “NOP”) and a “global regulator” include: (i) Upstream signal, with global regulators generally responding to a defined signal or effector and NAPs responding more generally to DNA biophysics and changes in their own abundance; (ii) binding specificity, with NAPs tending to recognize structural characteristics and AT richness, whereas global regulators have more defined sequence specificities; (iii) abundance, with NAPs generally being more highly expressed in terms of RNA and protein copy numbers than global regulators (see below); and (iv) coherence of regulon, with global regulators tending to act mainly on pathways that respond to a single or closely related set of environmental conditions, whereas NAPs tending to act at a broader variety of sites. Although these definitions are of course imperfect and certainly some exceptions and ambiguities exist for each rule proposed above (as reviewed, e.g., in [31]), the criteria above provide a useful starting point for identifying the coherent regulatory roles played by NAPs in E. coli and extending that view to other bacterial species. We also note that it is less clear what label ought to be applied to less abundant structuring proteins (such as CbpA [214] and MatP [215]) and accessory factors that modulate NAP behavior (such as Hha [61,216]), but at least at present we consider them not to fall into the NAP umbrella primarily due to their lower abundance and locality of action.
Expression-based screening identifies potential new NAPs across a range of bacterial species
As described above, several NAPs have been characterized in E. coli, but few have been investigated in other bacteria. Most studied NAPs in bacteria other than E. coli are present in B. subtilis, as we will enumerate in the following section. Few NAPs from other bacteria have been deeply characterized. Notable examples of NAPs outside E. coli and B. subtilis that have been characterized are GapR and IHF from Caulobacter crescentus and Lsr2 from Mycobacterium tuberculosis; we also note that recognizable homologs of proteins such as HU and IHF, and xenogeneic silencers, are readily identified in many sequenced bacteria.
We suspect there is a wealth of information on DNA-binding proteins in published literature suggesting many proteins are NAPs, but that they may have been largely overlooked as NAPs due to the fact that genome-wide methods to detect protein–DNA interactions have not been widely used until fairly recently. Rather, much published literature has made use of assays that test a protein’s binding to, or protection of, specific sequences in vitro, e.g., DNAse I footprinting. Therefore, we devised a simple informatic screen to identify potential NAPs based on GO terms and transcript abundance. The list of potential NAPs that arose from our screen was then cross-referenced with literature to evaluate the applicability of the label to the identified highly expressed DNA binding proteins.
Screening proteomes for potential NAPs
To screen proteomes for potential NAPs, we searched UniProt [217] reference proteomes from several bacteria for proteins with the GO term GO:0003677, which indicates “DNA binding”, or selected child terms GO:0043565, GO:0003681, or GO:0003690 (“sequence-specific DNA binding”, “bent DNA binding”, and “double-stranded DNA binding”, respectively). We also filtered against proteins with GO terms indicating a protein was directly involved in DNA replication, transcription, or translation. The remaining proteins were then cross-referenced to transcriptomic data to determine, relative to other genes in the bacterium, the expression level of their transcripts. RNA-seq data were obtained from the studies noted below and aligned to a RefSeq reference transcriptome for the organism of interest using kallisto [218], and then simple means across replicates of the kallisto-derived TPM were used to measure transcript abundance. For the proteins that passed our GO term filters and were in the 80th percentile for transcript abundance, we considered them as possible NAPs, as we will describe in more detail below. A reference implementation of our approach, along with the specific case studies given below, is available for download from github.com/jwschroeder3/NAPper.
The results of our screen may not provide an absolutely complete list of NAPs in these bacteria (for reasons noted below), and in some cases also include many proteins which should not be considered NAPs, but given our screen’s generally high sensitivity in detecting known NAPs in well-characterized organisms (see below), we view our list as a useful provisional list of NAPs that can guide future investigations. As will be seen in the case studies below, for some organisms, the 80th percentile threshold may be overly permissive (e.g., for E. coli), but for others (such as B. subtilis) it is actually necessary and appropriate for identifying all known NAPs. In the discussion below, we provide information on the conclusions that would be reached with both the more permissive 80th percentile and more strict 90th percentile thresholds; our practical advice would be that investigators looking for potential NAPs should first prioritize candidates falling above the 90th percentile in transcript levels under at least one available condition and turn to those between the 80th and 90th percentile as a lower priority. We also note that the completeness of the list of potential NAPs returned is strongly dependent on the accuracy of the underlying GO terms themselves. While curated UniProt annotations are often a good starting point, researchers will assuredly benefit from application of additional tools to a proteome of interest if possible.
Known E. coli NAPs are recovered by our screen
Application of the procedure described above to E. coli recovers all of the classic NAPs (Figure 3, left; see also Table 1); furthermore, the classic NAPs are all above the 90th percentile for expression (at the transcript abundance level) in at least one of the two conditions that we inspected. Consistent with what is known about expression of the NAP Fis, Fis meets our criteria for potential NAPs in exponentially growing cells, but not in stationary phase (Figure 3, left). Other proteins which have sometimes been referred to as NAPs in the literature (e.g., the ter-binding protein MatP or the accessory H-NS/StpA bridging factor Hha) are lower in abundance. The transcript abundances also appear to be a good proxy for protein production levels for cells in exponential phase (Figure 3, right). The prominent position of the generally accepted NAPs in these datasets demonstrates the efficacy of our expression-based screening procedure in at least identifying NAP candidates. A small handful of other proteins also stand out as having similar expression levels to those of the known NAPs in at least one condition; these proteins can be decomposed into three groups: (i) Global regulators (e.g., Lrp, Fur, CRP), which are somewhat less abundant and more site-specific than NAPs, and respond directly to defined effectors; (ii) enzymes (SodA, DinJ) which associate with DNA as part of their function; and (iii) potential additional NAPs (notably BolA, YiaG, HspQ, and CspA). We consider the four members of the last category below.
Figure 3.
Identification of known and potential E. coli NAPs through expression analysis. Left: Plots of the expression levels of all genes flagged as NAP candidates by our screening approach (see Text for details); all data are taken in glucose minimal media (raw sequencing data are provided in the github link from the main text). Genes are categorized by manual curation (based on information from Ecocyc [221]) into global regulators, local regulators, classical nucleoid-associated proteins (NAP), possible new nucleoid-associated proteins (Potential NAP), or DNA binding proteins with non-regulatory primary functions (Other). Red dashed lines indicate the 80th and 90th percentile thresholds used to filter for potential NAPs. Right: Correlation of RNA levels and protein production per cell cycle (data from [344]) for all E. coli proteins identified in our screen; all data are taken in glucose minimal media
Table 1.
Proteins in the E. coli MG1655 reference proteome that met our GO term and transcript abundance criteria for potential NAPs. aGR = global regulator, LR = local regulator, PNAP = potential NAP. For exponential phase, the 80th and 90th percentile log2(TPM) values are 7.03 and 8.27, respectively. In stationary phase, 80th and 90th percentile log2(TPM) values are 6.30 and 7.84, respectively
| UNIPROT ID | Protein | Gene | log2 TPM (exponential phase) |
log2 TPM (stationary phase) |
Categorya |
|---|---|---|---|---|---|
| P0ACF8 | DNA-binding protein H-NS | hns | 12.0 | 11.3 | NAP |
| P0ACF4 | DNA-binding protein HU-beta | hupB | 12.5 | 8.9 | NAP |
| P0ACF0 | DNA-binding protein HU-alpha | hupA | 12.1 | 9.0 | NAP |
| P0A6X7 | Integration host factor subunit alpha (IHF-alpha) | ihfA | 11.0 | 12.2 | NAP |
| P0A6Y1 | Integration host factor subunit beta (IHF-beta) | ihfB | 10.9 | 10.8 | NAP |
| P0A9X9 | Cold shock protein CspA | cspA | 10.8 | 6.6 | PNAP |
| P00448 | Superoxide dismutase | sodA | 10.6 | 9.2 | Other |
| P0ABT2 | DNA protection during starvation protein | dps | 10.2 | 12.5 | NAP |
| P0ABE2 | DNA-binding transcriptional regulator BolA | bolA | 10.1 | 12.3 | PNAP |
| P0A6R3 | DNA-binding protein Fis | fis | 9.8 | 5.9 | NAP |
| P0ACG1 | DNA-binding protein StpA | stpA | 9.6 | 4.9 | NAP |
| P0A9V5 | Uncharacterized HTH-type transcriptional regulator YiaG | yiaG | 9.6 | 11.9 | PNAP |
| P0ACE3 | Hemolysin expression-modulating protein Hha | hha | 9.2 | 8.7 | NAP |
| P0A8U6 | Met repressor | metJ | 9.0 | 6.9 | LR |
| P0A9Q5 | Acetyl-coenzyme A carboxylase carboxyl transferase subunit beta | accD | 8.9 | 5.7 | Other |
| P0A8B5 | Nucleoid-associated protein YbaB | ybaB | 8.9 | 7.5 | PNAP |
| P0A9E5 | Fumarate and nitrate reduction regulatory protein | fnr | 8.7 | 8.3 | GR |
| P0AB20 | Heat shock protein HspQ | hspQ | 8.4 | 11.3 | PNAP |
| P0DMC7 | Transcriptional regulatory protein RcsB | rcsB | 8.4 | 7.6 | LR |
| P57998 | Insertion element IS1 4 protein InsB | insB4 | 8.1 | 5.7 | Other |
| P0ACR9 | Transcriptional repressor MprA (Protein EmrR) | mprA | 8.0 | 4.7 | LR |
| P0A9T6 | Uncharacterized HTH-type transcriptional regulator YbaQ | ybaQ | 7.9 | 8.2 | PNAP |
| P0AAR0 | Hha toxicity modulator TomB | tomB | 7.9 | 9.2 | PNAP |
| Q47150 | Antitoxin DinJ | dinJ | 7.8 | 9.5 | Other |
| P0A7H6 | Recombination protein RecR | recR | 7.7 | 5.4 | Other |
| P68767 | Cytosol aminopeptidase | pepA | 7.5 | 6.4 | Other |
| P76116 | Uncharacterized protein YncE | yncE | 7.5 | 4.9 | PNAP |
| P0A9M0 | Lon protease | lon | 7.5 | 8.2 | Other |
| P0ACN4 | HTH-type transcriptional repressor AllR | allR | 7.4 | 9.8 | LR |
| P0A7C2 | LexA repressor | lexA | 7.4 | 7.3 | GR |
| P0A8A0 | Probable transcriptional regulatory protein YebC | yebC | 7.3 | 5.4 | PNAP |
| P0A881 | Trp operon repressor | trpR | 7.3 | 3.9 | LR |
| P76062 | Prophage repressor RacR | racR | 7.3 | 4.3 | LR |
| P06966 | HTH-type transcriptional regulator DicA | dicA | 7.3 | 6.7 | LR |
| P0AED5 | Response regulator UvrY | uvrY | 7.2 | 6.3 | LR |
| P0A8A2 | Probable transcriptional regulatory protein YeeN | yeeN | 7.2 | 2.2 | LR |
| P67699 | Uncharacterized HTH-type transcriptional regulator YddM | yddM | 7.0 | 8.5 | PNAP |
| Q2EES9 | Response regulator inhibitor for tor operon (Tor inhibitor) | torI | 7.0 | 6.5 | LR |
| P06612 | DNA topoisomerase 1 | topA | 6.9 | 6.9 | Other |
| P27434 | Cytoskeleton protein RodZ | rodZ | 6.9 | 6.8 | Other |
| P63204 | Transcriptional regulator GadE | gadE | 6.7 | 8.9 | LR |
| P76268 | Transcriptional regulator KdgR | kdgR | 6.6 | 6.6 | LR |
| P0ACL2 | Exu regulon transcriptional regulator | exuR | 6.5 | 7.1 | LR |
| P15373 | Antitoxin PrlF | prlF | 6.4 | 8.1 | Other |
| P64530 | Transcriptional repressor RcnR | rcnR | 6.1 | 7.3 | LR |
| P0ACG8 | Heat shock protein 15 (HSP15) | hslR | 6.1 | 8.6 | PNAP |
| Q47149 | mRNA interferase toxin YafQ | yafQ | 6.0 | 6.5 | Other |
| P0A823 | Sugar fermentation stimulation protein A | sfsA | 5.9 | 8.2 | PNAP |
| P42641 | GTPase ObgE/CgtA | obgE | 5.8 | 7.2 | Other |
| P0A8F8 | UvrABC system protein B | uvrB | 5.3 | 6.9 | Other |
| P75993 | Probable two-component-system connector protein AriR | ariR | 5.2 | 9.2 | Other |
| P0A698 | UvrABC system protein A | uvrA | 5.0 | 6.4 | Other |
| P77569 | DNA-binding transcriptional activator MhpR | mhpR | 5.0 | 6.3 | LR |
| P33224 | Putative acyl-CoA dehydrogenase AidB | aidB | 4.4 | 7.2 | Other |
| P0ACS2 | Redox-sensitive transcriptional activator SoxR | soxR | 4.1 | 6.4 | LR |
| P06992 | Ribosomal RNA small subunit methyltransferase A | rsmA | 6.6 | 6.8 | Other |
| P22186 | Transcriptional regulator MraZ | mraZ | 8.3 | 7.9 | LR |
| P36771 | Probable HTH-type transcriptional regulator LrhA | lrhA | 6.7 | 7.0 | LR |
| P0A8D0 | Transcriptional repressor NrdR | nrdR | 7.1 | 4.7 | LR |
| Q46864 | Antitoxin MqsA | mqsA | 6.7 | 9.7 | Other |
| P0AF63 | HTH-type transcriptional repressor NsrR | nsrR | 6.2 | 6.6 | LR |
| P0ACJ8 | cAMP-activated global transcriptional regulator CRP | crp | 9.5 | 7.3 | GR |
| P0A9A9 | Ferric uptake regulation protein | fur | 9.2 | 8.7 | GR |
| P0AGK8 | HTH-type transcriptional regulator IscR | iscR | 9.0 | 8.7 | GR |
| P0A8N0 | Macrodomain Ter protein | matP | 6.4 | 6.4 | NAP |
| P0ACJ0 | Leucine-responsive regulatory protein | lrp | 9.9 | 7.5 | GR |
| P0AE72 | Antitoxin MazE | mazE | 7.1 | 6.2 | Other |
| P0ACI0 | Right origin-binding protein | rob | 7.5 | 8.4 | LR |
| P0A6X3 | RNA-binding protein Hfq | hfq | 10.6 | 10.5 | NAP |
| P36659 | Curved DNA-binding protein | cbpA | 6.2 | 7.5 | NAP |
DNA-binding transcriptional regulator BolA
BolA has long been recognized as a regulator of cellular morphology that is expressed highly in stationary phase under control of a σS dependent promoter [219–221]. More recently, combined RNA-seq and ChIP-seq experiments identified ~1,300 binding sites for BolA across the chromosome, as well as a far broader range of regulatory targets including flagellar assembly, central carbon metabolism, and biofilm development [222]. Given the conditional specificity of BolA activity, its coherent set of targets, and the presence of a reasonably informative binding sequence motif (YYGCCAGH, per [222]), BolA appears to match our definition of a global regulator rather than a NAP.
Uncharacterized HTH-type transcriptional regulator YiaG
YiaG is predicted to be a Cro-like helix-turn-helix transcriptional regulator, but to the best of our knowledge has not been substantially characterized. Gao et al. included it in a survey of poorly characterized transcription factors, including performing ChIP-exo experiments on epitope-tagged YiaG [223], but did not observe any peaks under the growth condition used (exponential growth in glucose minimal media). It is not clear whether the absence of YiaG peaks arises because it is not actually a DNA binding protein, some technical issues prevented observation of peaks in the epitope-tagged version used, or whether it would in fact show strong DNA binding under another physiological condition. Based on the gene expression data shown in Figure 3, any effort to study the role of YiaG likely ought to begin with work in stationary phase, given that during that growth condition yiaG transcript levels rival those of hns, dps, and ihfA. At present, however, while YiaG appears likely to fall into the category of either a global regulator or a NAP, it is impossible to assign without more information.
Heat shock protein HspQ
Another minimally studied small protein, HspQ, had until recently been characterized only in the context of enhancing the degradation of specific mutant DnaA proteins [224]. Recent evidence, however, has demonstrated that HspQ in fact acts as a modulator of the activity of Lon and Clp proteases in several enterobacteria [225,226]. The annotated DNA-binding activity of HspQ arises due to the presence of a hemimethylated DNA-binding domain and demonstrated hemimethylated DNA-binding activity [227]; this activity likely spatially modulates the protease-regulating activity of HspQ and may in particular explain its effects on DnaA expression. However, classification as a NAP (at least by our criteria) does not appear appropriate.
Cold shock protein CspA
CspA is a somewhat deceptively named protein in that while it was originally identified as being expressed during cold stress [228], it is in fact also highly expressed under a wide range of conditions, and is present throughout exponential growth at 37oC [229]. E. coli CspA appears to have both DNA-binding [230] and RNA chaperone [231] activities, and in its DNA binding capacity has been shown to act as a transcriptional regulator of hns [163] and gyrA [232] and to act as an antiterminator for several other cold-induced genes [233]. At the same time, high (but physiologically relevant) concentrations of CspA inhibit both transcription and translation in cell-free systems, likely through nonspecific nucleic acid-binding activity [234]. Integrating many of the above findings, Brandi and Pon speculated (and we concur) that CspA likely plays a large-scale silencing role under conditions where it is highly expressed, reducing the expression of all but a small set of genes (potentially those specifically needed for survival under some stress condition [229]). E. coli CspA thus appears likely to fall under the general umbrella of NAPs with large-scale effects on transcription as we have defined them. As cold shock proteins similar to CspA were identified in our screen as potential NAPs in a wide range of other organisms, they are discussed in more detail below.
Regulatory NAPs in Bacillus subtilis
The Firmicute Bacillus subtilis provides a well-studied counterpoint for E. coli among Gram-positive bacteria. B. subtilis contains three fairly well-characterized proteins that appear aligned with our understanding of nucleoid-associated proteins: Rok, HBsu, and Noc.
Rok broadly affects transcription in Bacillus subtilis
Repressor of ComK (Rok)
Repressor of ComK (Rok) was identified in a transposon mutagenesis screen as a repressor of genetic competence in Bacillus subtilis [235]. Since then, our understanding of Rok has expanded to view it as analogous to the H-NS present in many Gram-negative bacteria, acting as a xenogeneic silencer [236]. Rok preferentially binds AT-rich DNA and is enriched along prophages and mobile DNA in B. subtilis [237]. Loss of rok causes many changes to B. subtilis physiology, including upregulation of genes involved in genetic competence, expression of genes involved in production of secreted antibiotics, changes in colony morphology, and increased mobilization of the integrative and conjugative element, ICEBs1 [235,237–240]. DnaA directly regulates initiation of DNA replication (see [241] for review) and the expression, either directly or indirectly, of several genes involved in DNA replication and repair, cell cycle progression, stress response, and sporulation [242]. Recent evidence suggests that Rok is required not only for occupancy of DnaA at several regions of the B. subtilis genome but also for the regulation of gene expression in those regions by DnaA [242,243]. Most Rok-dependent gene regulation by DnaA is repressive [242]. Although current evidence is insufficient to assign a concise physiological role to the DnaA/Rok interaction, the set of genes regulated by DnaA in a Rok-dependent manner suggests that the interaction may be important for regulating stress-associated processes such as antibiotic production and secretion, and nucleotide, oligopeptide, and amino acid import [242]. Therefore, in general Rok is a repressor of competence and extracellular functions, and is a mediator of gene regulation by DnaA, but the mechanisms by which Rok represses expression remain poorly understood.
Roles for other B. subtilis NAPs in regulating gene expression have not been directly identified
B. subtilis HBsu
B. subtilis HBsu is homologous to E. coli HU and is a highly abundant NAP in B. subtilis. HBsu is essential, is involved in DNA compaction, and associates broadly with the B. subtilis nucleoid, with a slight preference for the SPβ prophage region [237,244,245]. Although it is currently unclear what effects HBsu has on gene expression in B. subtilis, two potential mechanisms by which HBsu could be regulated are worth discussion here. Expression of the gene encoding HBsu, hupA (also called hbs), is decreased during glucose exhaustion [246]. In addition, HBsu is acetylated in vivo, and its acetylation likely decreases its affinity for DNA in vitro and nucleoid compaction in vivo [247]. We suggest that differential expression and acetylation of HBsu may provide mechanisms by which its effect on DNA structure, and potentially gene expression, can be regulated, although obtaining direct evidence of effects of HBsu on gene expression has been difficult due to the essentiality of the gene.
B. subtilis nucleoid occlusion protein (Noc)
B. subtilis nucleoid occlusion protein (Noc) was originally characterized in its role limiting cell division over nucleoids [248]. Since its discovery, it has been found to bind many sites in the B. subtilis genome, and also to bind the cell membrane, potentially bridging bacterial DNA to the membrane [249,250]. While it has been thought that Noc prevents filamentation of the cell division protein FtsZ at improper sites, new evidence instead suggests that it provides a physical barrier to prevent already-formed FtsZ filaments from migrating away from the proper location of the future cell division plane [251], thus increasing the precision of FtsZ filament accumulation for midcell. To the best of our knowledge, no role of Noc in regulation of transcription has been proposed.
Identification of known and potential NAPs in B. subtilis
Similarly to the success observed above in E. coli, screening the B. subtilis proteome for potential NAPs following the same procedure, we verified that the known NAPs Noc, Rok, and HBsu were hits in our screen, although Noc and Rok are not as highly expressed as many E. coli NAPs, whereas HBsu (hupA) is a clear standout in terms of transcript abundance (Figure 4 and Table 2). In the process, we also identified several other DNA-binding proteins with abundance on par with HBsu, which should be considered as potential additional NAPs.
Figure 4.
Identification of known and potential B. subtilis NAPs through expression analysis. Left: Plot of the expression levels of all genes flagged as NAP candidates by our screening approach (see Text for details); vegetative growth refers to the 8-hour time point in [345], all data were drawn from [345]. Red dashed lines indicate the 80th and 90th percentile thresholds used to filter for potential NAPs. Genes were categorized by manual curation (based on information from Subtiwiki [346]) into the categories described for Figure 3. Right: Expansion of inset from plot on the left
Table 2.
Proteins in the B. subtilis 168 proteome that met our GO term and transcript abundance criteria for potential NAPs. aGR = global regulator, LR = local regulator, PNAP = potential NAP. In vegetative growth, the 80th and 90th percentile log2(TPM) values are 7.82 and 8.86, respectively. After 24 hours in biofilm conditions, the 80th and 90th percentile log2(TPM) values are 7.71 and 8.84, respectively
| UNIPROT ID | Protein | Gene name | log2(TPM) in vegetative growth | log2(TPM) 24 hours in biofilm condition | Categorya |
|---|---|---|---|---|---|
| P51777 | Cold shock protein CspB | cspB | 14.3 | 13.6 | PNAP |
| P32081 | Cold shock protein CspD | cspD | 14.4 | 14.6 | PNAP |
| P08821 | DNA-binding protein HU 1 | hupA | 13.5 | 13.1 | NAP |
| P39158 | Cold shock protein CspC | cspC | 12.5 | 13.3 | PNAP |
| P08874 | Transition state regulatory protein AbrB | abrB | 12.1 | 9.4 | PNAP |
| O32067 | Uncharacterized HTH-type transcriptional regulator YtzE | ytzE | 11.2 | 11.9 | PNAP |
| P96622 | Endoribonuclease EndoA | ndoA | 10.6 | 10.4 | Other |
| O34766 | SPbeta prophage-derived uncharacterized HTH-type transcriptional regulator YopS | yopS | 10.35 | 10.6 | Other |
| Q7WY72 | Extracellular matrix regulatory protein A | remA | 10.1 | 9.6 | GR |
| P37582 | HTH-type transcriptional regulator GlnR | glnR | 9.8 | 7.9 | LR |
| P54512 | HTH-type transcriptional regulator MntR | mntR | 9.5 | 8.8 | LR |
| O32253 | Central glycolytic genes regulator | cggR | 9.4 | 7.6 | LR |
| O07586 | HTH-type transcriptional regulator CueR | cueR | 9.01 | 8.98 | LR |
| O31771 | Uncharacterized membrane protein YmfM | ymfM | 9.0 | 8.23 | Other |
| P06533 | HTH-type transcriptional regulator SinR | sinR | 8.9 | 9.2 | GR |
| O31761 | Uncharacterized HTH-type transcriptional regulator YmfC | ymfC | 8.87 | 8.38.2 | Other |
| P24281 | Nucleoid-associated protein YaaK | yaaK | 8.78 | 8.58.4 | PNAP |
| O31417 | Uncharacterized HTH-type transcriptional regulator YazB | yazB | 8.7 | 8.63 | Other |
| P25499 | Heat-inducible transcription repressor HrcA | hrcA | 8.6 | 7.54 | LR |
| P39758 | Putative transition state regulator Abh | abh | 8.6 | 8.76 | GR |
| O05236 | Uncharacterized HTH-type transcriptional regulator YugG | alaR | 8.5 | 7.7 | Other |
| P39667 | Transcription repressor NadR | nadR | 8.5 | 8.76 | LR |
| O34857 | Repressor Rok | rok | 8.45 | 8.18.2 | NAP |
| P37524 | Nucleoid occlusion protein (Noc) | noc | 8.3 | 8.56 | NAP |
| O34835 | Transcription factor FapR | fapR | 8.31 | 6.86.6 | LR |
| P06534 | Stage 0 sporulation protein A | spo0A | 8.32 | 8.9 | GR |
| O07573 | HTH-type transcriptional regulator NsrR | nsrR | 8.32 | 8.99.0 | GR |
| P39776 | Tyrosine recombinase XerC | xerC | 8.3 | 7.75 | Other |
| P39814 | DNA topoisomerase 1 | topA | 8.3 | 7.3 | Other |
| P31080 | LexA repressor | lexA | 8.23 | 8.8 | GR |
| Q7BVT7 | Uncharacterized protein YerC | yerC | 8.23 | 8.8 | Other |
| P04831 | Small, acid-soluble spore protein A (SASP) | sspA | 8.13 | 9.58.4 | PNAP |
| P13800 | Transcriptional regulatory protein DegU | degU | 8.1 | 8.42 | GR |
| P37568 | Transcriptional regulator CtsR | ctsR | 8.07.9 | 6.36.0 | LR |
| P54476 | Probable endonuclease 4 | nfo | 8.01 | 6.76.8 | Other |
| P37551 | Pur operon repressor | purR | 7.9 | 7.4 | LR |
| Q45549 | Transcriptional repressor NrdR | nrdR | 7.89 | 7.67.7 | LR |
| P11470 | Spore germination protein GerE | gerE | 7.66.9 | 9.18.9 | GR |
| O31509 | Probable transcriptional regulatory protein YeeI | yeeI | 7.4 | 7.87 | PNAP |
| P26497 | Stage 0 sporulation protein J | spo0J | 7.43 | 7.7 | Other |
| O34692 | Uncharacterized HTH-type transcriptional regulator YvnA | yvnA | 6.1 | 8.27.9 | PNAP |
| P94433 | HTH-type transcriptional repressor YcnK | ycnK | 5.6 | 9.7 | LR |
| P40762 | HTH-type transcriptional regulator PchR | pchR | 3.57 | 9.9 | LR |
AbrB
AbrB is a transcriptional regulator governing many developmental processes as B. subtilis transitions from exponential to stationary phase, including competence, sporulation, and biofilm development. AbrB has been termed a global regulator, but we find it has NAP-like properties. Specifically, it is highly abundant, with transcript levels in the 99th percentile (Figure 4). AbrB can protect a large footprint on bound regions of DNA (≈ 25 to 80 or more bps) in DNase I footprinting assays [252,253], and binding of AbrB to regulated regions of DNA may be more dependent on shape than on sequence [253–255]. With these criteria in mind, it may be useful to think of AbrB as a regulatory NAP which, during rapid growth, represses developmental processes which occur after transition to stationary phase in B. subtilis.
LrpC does not meet our definition of a NAP
B. subtilis LrpC is homologous to E. coli Lrp [256]. LrpC has been considered a NAP due to its ability to bind DNA nonspecifically, and to bend, wrap, and positively supercoil DNA [29,257,258]. However, the lrpC transcript is not particularly abundant (≈50th percentile in the data we analyzed) and thus does not meet our criteria for a potential NAP.
SspA
SspA is a member of a group termed small, acid-soluble spore proteins (SASP). SASP are DNA binding proteins which are highly expressed during sporulation in the genera Bacillus and Clostridium [259]. Intriguingly, we find that the sspA transcript is quite abundant during vegetative growth and biofilm development (Figure 4, right). Spores are not metabolically active and thus cannot repair their DNA. This, coupled with the fact that spores may exist for extremely long periods of time and be exposed to DNA damaging agents throughout that time, necessitates that spores have mechanisms to limit damage to their DNA without expending energy. SASP fill that need, promoting spore viability against heat, hydrogen peroxide, and UV light [260,261]. SASP bind DNA nonspecifically, contorting it to the A-form [262], promoting the formation of a ring-shaped nucleoid in spores [263], protecting DNA from enzymatic and radical-induced cleavage [264], and modulating the reactivity of DNA to UV light (reviewed in [265]). Although SspC, which shares 81% amino acid identity with SspA, can inhibit in vitro transcription [264], it is not yet clear what effect SASP have on transcription in vivo.
Cold shock proteins
CspB, CspC, and CspD were among the top four hits in our B. subtilis screen. Similar to Hfq and E. coli CspA, B. subtilis CspB can bind either RNA or DNA [231,266]; thus, many of the arguments regarding E. coli CspA apply here as well, and in general, the highly abundant DNA-binding cold shock proteins present in B. subtilis may act as additional NAPs.
Case examples in other moderately studied bacteria
With the advancement of DNA sequencing technologies, there has been an expansion of exploration into a variety of different species. Here, we will explore both Gram-negative and Gram-positive bacterial species’ putative NAPs, making use of the operative definitions enumerated above wherever possible.
Pseudomonas aeruginosa
Pseudomonas aeruginosa (P. aeruginosa) is a common infectious bacterium [267], typically found in hospital patients [268], that can cause pneumonia and be multidrug resistant [269]. Two important aspects of its pathogenic response are quorum sensing (QS), a communication system related to population density, and the formation of biofilms, hardy multicellular structures that can enable colonization and growth in a variety of hosts. Expression-based screening (Figure 5 and Table 3) indicates the presence of roughly 10 proteins which meet our criteria as potential NAPs. These candidates include homologs of NAPs that have been extensively studied in other model organisms, as well as examples that have been identified in less studied organisms such as P. putida [57], including HU, IHF, and Dps (PA0962), and the potential NAP CspA. In P. aeruginosa, the CspA family of proteins has been shown to be important for the control of type III secretion systems in response to temperature shifts and host environments [270], and the high abundance of CspA itself supports the possibility of a NAP-like role similar to that hypothesized above in E. coli. PA0962, or Dps, is another conserved NAP in P. aeruginosa, and is critical for survival of reactive oxygen species, both aiding in detoxifying peroxides and contributing to antibiotic tolerance [271,272]. In addition to CspA and Dps, we identified AmrZ as one of the most highly expressed putative NAPs. AmrZ mediates the formation of biofilm development in response to the secondary messenger cyclic diguanylate (c-di-GMP), where AmrZ limits the accumulation, and represses the expression, of AmrZ dependent cyclase A (adcA) a main component leading to the production of c-di-GMP, thus delaying development of biofilms [273]. AmrZ appears to bind DNA through at least two different modes, and depending upon broader biological context can act as a transcriptional repressor or activator [274]. While generally studied alone, AmrZ has been identified as a key protein in the virulence regulatory network of P. aeruginosa [275]. Also in this network is RsaL [275], whose binding at the bidirectional promoter of rsaL-lasR negatively regulates one of the lasR/lasI QS systems [276] and is known to repress virulence gene expression [277]. However, surprisingly, a rsaL mutant exhibits impaired biofilm formation, leading to the hypothesis that RsaL may contribute to the transition of P. aeruginosa to a pathogenic state [278].
Figure 5.
Identification of putative NAPs from example Gram-negative species. Shown are all proteins in P. aeruginosa (left) and N. meningitidis that pass the NAP-screening procedures described in the text. The hits were then manually assigned to one of the functional categories shown in Figure 3 on the basis of their annotations and expression levels, with the unknown category reserved primarily for lower-expression cases with unclear functions. Red dashed lines indicate the 80th and 90th percentile thresholds used to filter for potential NAPs. All transcriptional data were processed from [347], using the “control” and “stationary phase” conditions for each organism
Table 3.
All listed proteins in the P. aeruginosa PA01 proteome that met our GO term and transcript abundance criteria for potential NAPs. aGR = global regulator, LR = local regulator, PNAP = potential NAP. Log2(TPM) values at the 80th and 90th percentile in the control condition are 6.39 and 7.48, respectively. In stationary phase, the log2(TPM) values at the 80th and 90th percentiles are 5.70 and 6.84, respectively
| UNIPROT ID | Protein | Gene | log2 TPM (control) |
log2 TPM (stationary phase) |
Category |
|---|---|---|---|---|---|
| Q06553 | HTH-type transcriptional regulator PrtR (Pyocin repressor protein) | prtR | 6.57 | 4.68 | LR |
| P55222 | cAMP-activated global transcriptional regulator Vfr | vfr | 8.98 | 7.62 | GR |
| Q51425 | Holliday junction ATP-dependent DNA helicase RuvA (EC 3.6.4.12) | ruvA | 6.76 | 5.00 | Other |
| P25084 | Transcriptional activator protein LasR | lasR | 9.29 | 9.56 | GR |
| Q51472 | Integration host factor subunit alpha (IHF-alpha) | ihfA | 11.10 | 11.00 | NAP |
| P95412 | Siroheme decarboxylase NirD subunit (EC 4.1.1.111) | nirD | 7.82 | 2.06 | Other |
| Q51423 | Transcriptional regulatory protein PmpR | pmpR | 7.43 | 5.84 | Unknown |
| P95459 | Major cold shock protein CspA | cspA | 12.81 | 10.29 | PNAP |
| G3XCY4 | Transcription factor AmrZ (Alginate and motility regulator Z) | amrZ | 12.91 | 13.68 | PNAP |
| Q9I1M3 | Bkd operon transcriptional regulator | bkdR | 6.18 | 6.00 | Unknown |
| P15276 | Transcriptional regulatory protein AlgP (Alginate regulatory protein AlgR3) | algP | 8.03 | 6.96 | Unknown |
| Q9HTL0 | DNA-binding protein HU-alpha | hupA | 11.45 | 10.97 | NAP |
| P05384 | DNA-binding protein HU-beta | hupB | 15.19 | 13.02 | NAP |
| P26275 | Positive alginate biosynthesis regulatory protein | algR | 7.63 | 7.61 | Unknown |
| P26993 | HTH-type transcriptional regulator ExsA | exsA | 6.52 | 4.40 | Unknown |
| Q51473 | Integration host factor subunit beta (IHF-beta) | ihfB | 10.68 | 10.87 | NAP |
| Q51373 | Response regulator GacA (Global activator) | gacA | 8.21 | 8.35 | GR |
| Q9I0M3 | DNA translocase FtsK | ftsK | 6.71 | 5.39 | Other |
| Q9HU78 | Histidine utilization repressor | hutC | 6.11 | 5.78 | LR |
| P37452 | LexA repressor (EC 3.4.21.88) | lexA | 6.44 | 5.14 | GR |
| Q9HWX1 | Transcriptional repressor NrdR | nrdR | 7.04 | 5.85 | LR |
| Q51481 | Denitrification regulatory protein NirQ | nirQ | 9.20 | 4.80 | LR |
| Q06552 | Transcription regulatory protein PrtN (Pyocin activator protein) | prtN | 8.57 | 7.24 | LR |
| Q9HYL3 | Regulatory protein NosR | nosR | 8.33 | 1.64 | LR |
| Q9I3I0 | Nucleoid-associated protein PA1533 | PA1533 | 10.26 | 9.21 | PNAP |
| Q9I3I7 | Probable transcriptional regulator | PA1526 | 7.75 | 7.36 | Unknown |
| Q9I233 | Probable transcriptional regulator | PA2082 | 6.61 | 4.75 | Unknown |
| Q9HZW2 | Putative repressor of atu genes | atuR | 6.63 | 6.01 | Unknown |
| Q9I0C4 | Probable transcriptional regulator | PA2718 | 6.62 | 6.59 | Unknown |
| Q9HXW2 | Probable transcriptional regulator | PA3678 | 6.41 | 5.33 | Unknown |
| Q9HVS2 | Probable transcriptional regulator | PA4499 | 7.70 | 6.96 | Unknown |
| Q9I4Y6 | Uncharacterized protein | PA0983 | 8.09 | 8.56 | Unknown |
| Q9HX68 | Two-component response regulator RocA1 | rocA1 | 7.44 | 5.57 | Unknown |
| Q9HXV1 | Probable transcriptional regulator | PA3689 | 6.65 | 5.95 | Unknown |
| Q9HTK2 | Transcriptional regulator GlcC | glcC | 6.82 | 6.50 | LR |
| Q9HTP6 | Leucine-responsive regulatory protein | lrp | 7.91 | 5.73 | GR |
| Q9HZF2 | HTH cro/C1-type domain-containing protein | PA3056 | 7.17 | 5.82 | Unknown |
| Q9I4Z7 | Probable dna-binding stress protein | PA0962 | 11.49 | 9.07 | PNAP |
| Q9I2T9 | Lon protease (EC 3.4.21.53) (ATP-dependent protease La) | lon | 7.72 | 7.10 | Other |
| Q9HVC1 | HTH cro/C1-type domain-containing protein | PA4674 | 8.41 | 7.42 | Unknown |
| Q9HXI7 | IscR | iscR | 9.97 | 9.11 | GR |
| Q9HTL4 | OxyR | oxyR | 6.81 | 5.35 | LR |
| Q9HT12 | Probable chromosome-partitioning protein ParB | spoOJ | 6.65 | 4.96 | NAP |
| Q9HTG1 | Probable transcriptional regulator | PA5403 | 6.39 | 4.38 | Unknown |
| Q9HT74 | Transcriptional regulator np20 | np20 | 7.41 | 6.48 | Unknown |
| Q9HTQ3 | Probable transcriptional regulator | PA5301 | 8.53 | 8.32 | Unknown |
| Q9HYY0 | Probable transcriptional regulator | PA3260 | 8.02 | 8.23 | Unknown |
| Q9HZF3 | Uncharacterized protein | PA3055 | 6.92 | 5.24 | Unknown |
| Q9I295 | Regulator of liu genes | liuR | 8.12 | 5.94 | Unknown |
| Q9I0Z8 | HTH tetR-type domain-containing protein | PA2484 | 7.27 | 7.54 | Unknown |
| Q9I3J3 | Probable transcriptional regulator | PA1520 | 7.20 | 6.03 | Unknown |
| Q9HY46 | NalD | nalD | 6.47 | 4.34 | Unknown |
| G3XD78 | Regulatory protein RsaL | rsaL | 12.41 | 12.35 | PNAP |
| Q9HX76 | Probable DNA binding protein | PA3940 | 11.97 | 8.83 | PNAP |
| Q9I5F9 | Lon protease (EC 3.4.21.53) (ATP-dependent protease La) | lon | 6.71 | 5.35 | Other |
PA1533 was identified as a potential NAP in our screen and is abundant in biofilms [279]. While very little is known about PA1533, its upregulation in biofilms may indicate a global role in gene regulation. P. aeruginosa can survive in water for long periods of time (>145 d), which is important considering it spreads in environments such as hospitals [268,280]. During this process, cells enter a dormant state, which aids in the survival of suboptimal conditions. Along with genes utilized for DNA replication and excision repair, the DNA binding protein PA3940 (hupN), another potential NAP identified in our screen, was induced in water conditions [281], perhaps aiding in the process of switching to a dormant state.
While P. aeruginosa does not have an annotated H-NS protein, MvaT and MvaU are generally considered xenogeneic silencers, playing similar roles in the silencing of prophages and regulating the type III secretion system, QS, and virulence gene expression, and have global impacts on gene expression [282–286]. We note that MvaT is missed in our screen and does not appear in Figure 5, solely due to an apparent under-annotation in UniProt, in that it lacks the key GO:0003677 (DNA binding) GO term, although it is annotated with GO:0032993 (protein-DNA complex, a Cellular Component term). Its expression levels would certainly put it in the realm of NAPs, with expression estimates at 214.0 TPM in exponential phase and 211.5 in stationary phase. The MvaT case provides an important reminder that our screen, as constructed, is reliant on the accuracy and completeness of existing GO term annotations, which fortunately are undergoing continuous curation and improvement [287].
Surprisingly, ParB has been identified as a potential NAP through our RNA-seq investigation as well as others [288]. Deletion of parB causes genome-wide changes to transcription. While ParB is highly conserved in other species to facilitate chromosome segregation [289], it has been shown to bind over 400 regions across the genome [288].
Neisseria meningitidis
Neisseria meningitidis (N. meningitidis), a causative bacterium of meningitis and septicemia, is a Gram-negative β-Proteobacterium which contains genes that code for putative NAPs that share high similarity in sequence and binding site preference to E. coli NAPs, such as ihfA/B, hupA/B [290,291]. N. meningitidis can be present in the normal microbial flora; however, some strains can be infectious via the bloodstream, where they adhere to and invade cells to inflict disease. IHF binding sites have been detected at the promoter of nadA, the N. meningitidis adhesin [292], an important factor for this infectious process. IHF has been shown to facilitate phase variable expression via binding of operators, enabling distant interactions of key proteins involved in the regulation of adherence and invasion [293]. Interestingly, Tn-seq implicated hupA to be an important factor in meningococcal colonization on human skin grafts and survival in the bloodstream of mice [294]. Applying our screening procedure for potential NAPs to N. meningitidis (Figure 5 and Table 4), in addition to the expected hupB and ihfA/B, we have identified the transcriptional regulator MerR (NMB-1303) and uncharacterized protein NMB-0897 as within the top 10% of expression levels, with expression of NMB-0897 on par with ihf. MerR (mer referring to mercury resistance) is a widely studied, highly conserved family of transcriptional activators linked by their N-terminal similarity [31,295]. The MerR regulators are usually found on transposable elements and activate gene expression by impacting RNA polymerase binding and initiation. Typically, these proteins will distort DNA so that RNA polymerase is forced to bind and initiate transcription at a suboptimal promoter, greatly impacting the regulation of a variety of operons [295]. MerR has not been deeply studied in N. meningitidis but given its abundance may act as a global regulator. While still uncharacterized, NMB-0897 has both been discovered to be essential and linked to the prophage IHT-E, perhaps again linking NAPs to having critical roles in prophage regulation across species [296,297], as the exceedingly high transcript level of this protein suggests a potential role as a NAP. It is interesting to note that H-NS is present in some species of Neisseria, including N. gonorrhoeae, and in that organism has been shown to regulate pilus formation (and thus likely pathogenesis) [298]. However, hns is not found in our screen because the N. meningitidis strain MC58, which is used in our work since it is the UniProt reference strain for that species, does not contain a detectable hns homolog (assessed both via annotation searches and searches for amino acid sequences resembling N. gonorrhoeae hns using promer [299]).
Table 4.
All listed proteins in the N. meningitidis MC58 proteome that met our GO term and transcript abundance criteria for potential NAPs. aGR = global regulator, LR = local regulator, PNAP = potential NAP. The log2(TPM) values at the 80th and 90th percentiles in the control condition are 8.00 and 9.15, respectively. In stationary phase, the log2(TPM) values at the 80th and 90th percentiles are 8.02 and 9.40, respectively
| UNIPROT ID | Protein | Gene | log2 TPM (control) |
log2 TPM (stationary phase) |
Category |
|---|---|---|---|---|---|
| P64389 | DNA-binding protein HU-beta | hupB | 14.97 | 14.12 | NAP |
| P0A0U2 | Integration host factor subunit beta (IHF-beta) | ihfB | 10.87 | 11.16 | NAP |
| P64393 | Integration host factor subunit alpha (IHF-alpha) | ihfA | 10.11 | 10.30 | NAP |
| P0A0Z0 | Putative HTH-type transcriptional regulator NMB1378 | NMB1378 | 7.71 | 9.94 | PNAP |
| Q9JYC7 | Probable transcriptional regulatory protein NMB1648 | NMB1648 | 8.83 | 8.27 | Unknown |
| Q9JRT9 | Pilin gene inverting protein PivNM-1A | pivNM-1B | 7.32 | 8.73 | Other |
| Q7DDC9 | Transcriptional regulator, MerR family | NMB1303 | 8.53 | 9.86 | GR |
| Q7DDQ7 | Transcriptional regulator, Crp/Fnr family | NMB0380 | 8.16 | 8.37 | GR |
| Q9JYC5 | Leucine-responsive regulatory protein | lrp | 8.68 | 9.06 | GR |
| Q9JZU4 | Uncharacterized protein | NMB0897 | 10.41 | 10.36 | PNAP |
| Q7DD45 | Uncharacterized protein | NMB2094 | 9.48 | 8.46 | PNAP |
| Q7DDA2 | Transcriptional regulator, GntR family | NMB1563 | 8.12 | 7.82 | LR |
| Q7DD98 | Transcriptional regulator MtrA | mtrA | 8.55 | 8.46 | LR |
| Q9JXT5 | Helix-turn-helix family protein | NMB1891 | 8.12 | 6.97 | Unknown |
Caulobacter crescentus
C. crescentus encodes well-described NAPs such as HU and IHF (aspects of IHF specific to C. crescentus are described below). Additionally, GapR was recently investigated in C. crescentus, and our screen suggests that several proteins acting as regulators of cell cycle progression have NAP-like properties.
The chromosome structuring protein GapR
The chromosome structuring protein GapR binds positive supercoils in DNA to promote DNA replication [300]. GapR was discovered as a NAP required for cell cycle progression in C. crescentus [301] and is present in α-Proteobacteria and several bacteriophage genomes [302]. Several studies have noted its preference for AT-rich DNA binding [300–302], but its propensity to bind at the 3' end of highly transcribed genes is of particular interest in its function as a chromosome structuring protein binding supercoiled DNA [300]. Structural evidence suggests that GapR can bind overtwisted DNA or B-form DNA [300,303,304], and its preference for binding AT-rich sequences may be an indirect effect of AT-rich DNA being more likely to assume an overtwisted conformation [300]. Loss of GapR results in a global change in gene expression, even after correcting for gene dosage effects, with genes near the origin of replication trending toward increased expression relative to genes near the replication terminus [302].
Integration host factor (IHF)
Integration host factor (IHF) has been discussed above for E. coli. Both IHF subunits were hits in our screen, and IHF has been characterized for its involvement in swarmer cell development in C. crescentus. IHF was initially identified in C. crescentus due to the presence of consensus E. coli IHF binding sites in or near promoters of genes encoding flagellar subunits in the C. crescentus genome, determination that E. coli IHF protected these sequences in DNase I footprinting assays in vitro, finding that making substitutions in these promoters yielded decreased expression from them in C. crescentus, and finally by the finding of bands in a western blot of C. crescentus lysates that cross-reacted with antibodies raised against E. coli IHF [305]. As in E. coli, C. crescentus IHF promotes site-specific λ recombination [306]. IHF plays a role in development of motility in the swarmer cell during C. crescentus differentiation; IHF expression is cell-cycle dependent, and loss of IHF leads to decreased expression of many flagellar biosynthesis genes, decreased motility, and cell filamentation [307].
Potential NAPs
We found that among the results of our screen in C. crescentus were several proteins involved in cell cycle control (Table 5). CtrA, SciP, MucR1, and MucR2 are primarily thought of as global regulators of cell cycle progression [308–310]. We feel that, while CtrA and SciP seem to fit the definition of global regulators well, they have NAP-like properties, and MucR1 and MucR2 (herein MucR1/2) should be categorized as probable NAPs. CtrA has broad effects on cell cycle-dependent gene expression, its effect on cell cycle regulation is largely dependent on its abundance throughout the cell cycle, and its binding site is degenerate and AT-rich [310–312], which are all helpful in characterizing proteins as NAPs. SciP, on the other hand, may act more as an accessory factor antagonistic to CtrA. SciP may bind some promoters directly, but overall its DNA binding seems largely dependent on CtrA also being present at promoters [309,313]. RNAP binding to promoters in vitro is abrogated when CtrA and SciP both occupy the promoter [309]. In the cases of MucR1/2, ChIP-seq suggests that MucR1/2 associate broadly with the genome, including association with a 26-kb mobile genetic element in C. crescentus [308]. It is currently unclear what effect, if any, MucR1/2 binding to the C. crescentus mobile genetic element has on its stability within the genome. Aside from their association with the C. crescentus mobile genetic element, MucR1/2 associate with promoters active during G1 phase, and loss of both mucR1 and mucR2 results in motility and buoyancy defects [308].
Table 5.
Proteins in the C. crescentus NA1000 proteome met our GO term and transcript abundance criteria for potential NAPs. Wild type C. crescentus RNA-seq data from [300] were used. aGR = global regulator, LR = local regulator, PNAP = potential NAP. bN = not identified as a nucleoid associated protein in [300], Y = identified as nucleoid associated in [300]. The 80th and 90th percentile log2(TPM) values are 7.67 and 8.61, respectively
| UNIPROT ID | Protein | Gene name | In [300]b | log2(TPM) | Categorya |
|---|---|---|---|---|---|
| A0A0H3C507 | CtrA inhibitory protein SciP | sciP | N | 13.0 | PNAP |
| A0A0H3CAE3 | DNA-binding protein HU | NA | N | 12.0 | PNAP |
| A0A0H3CFF1 | HTH XRE-family transcriptional regulator | NA | N | 10.3 | PNAP |
| B8GYD9 | Nucleoid-associated protein CCNA_00269 | NA | N | 10.2 | PNAP |
| B8GX11 | DNA-binding protein HU | hup | Y | 10.1 | NAP |
| A0A0H3C684 | ROS/MUCR transcriptional regulator MucR2 | mucR2 | N | 9.9 | PNAP |
| B8H1V7 | Transcription elongation factor GreA | greA | N | 9.9 | Other |
| B8H358 | Cell cycle transcriptional regulator CtrA | ctrA | N | 9.9 | GR |
| B8H546 | Integration host factor subunit alpha | ihfA | Y | 9.8 | NAP |
| A0A0H3CCW8 | Lrp-family transcriptional regulator | NA | N | 9.1 | GR |
| B8H6A5 | Integration host factor subunit beta | ihfB | Y | 9.0 | NAP |
| B8GX12 | Lon protease | lon | N | 9.0 | Other |
| A0A0H3C7B0 | ROS/MUCR transcriptional regulator | NA | N | 8.9 | PNAP |
| A0A0H3CF14 | SpoVT-AbrB family transcription factor, phd antitoxin | phd | N | 8.6 | GR |
| B8GYE0 | Recombination protein RecR | recR | N | 8.5 | Other |
| A0A0H3C636 | Cro/CI-family transcriptional regulator | NA | N | 8.5 | GR |
| A0A0H3C9Q5 | Xre-family transcriptional regulator | NA | N | 8.5 | GR |
| B8GZ33 | Modification methylase CcrMI | ccrMIM | N | 8.4 | Other |
| A0A0H3C9V7 | LacI-family transcriptional regulator | NA | N | 8.4 | Unknown |
| A0A0H3C569 | ROS/MUCR transcriptional regulator MucR1 | mucR1 | N | 8.4 | PNAP |
| A0A0H3CEJ4 | Two-component response regulator cenR | cenR | N | 8.3 | |
| A0A0H3CD88 | Two-component response regulator | NA | N | 8.3 | Unknown |
| B8H533 | Transcriptional repressor NrdR | nrdR | N | 8.3 | LR |
| B8H462 | Probable transcriptional regulatory protein CCNA_03352 | NA | N | 8.1 | |
| A0A0H3C803 | Hemimethylated DNA-binding protein yccV | NA | N | 8.1 | |
| A0A0H3C7Q7 | LacI-family transcriptional regulator | NA | N | 8.0 | |
| B8GW30 | Chromosome-partitioning protein ParB | parB | N | 7.9 | Other |
| A0A0H3C9J9 | Transcriptional regulator of stalk biogenesis staR | staR | N | 7.9 | |
| A0A0H3C4K4 | TetR family transcriptional regulator | NA | N | 7.8 | |
| A0A0H3C9K1 | LexA repressor | lexA | N | 7.8 | GR |
| A0A0H3C8F5 | DNA topoisomerase 4 subunit A | parC | N | 7.7 | Other |
Mycobacteria
Kriel and colleagues recently reviewed six known or probable mycobacterial NAPs [314]. Of the six NAPs discussed by Kriel and colleagues, two (HU and the xenogeneic silencer Lsr2) were retrieved by our screen in M. tuberculosis (Table 6). A third proposed NAP discussed in [314], EspR, barely missed our expression-level threshold, as its TPM value was at the 78th percentile in the data that we analyzed from [315]. The final three NAPs discussed by Kriel and colleagues had very sparse GO term annotations and fell out of our screen at the GO term filtering step; the percentile rankings of these proteins in the expression profile were 94.6 for Rv1388 (annotated as mIHF, for mycobacterial integration host factor), 79.9 for Rv0047c (annotated as NapM), and 93.5 for Rv3852 (annotated as H-NS in M. tuberculosis). Despite the lack of current GO term annotation suggesting mIHF and NapM bind DNA, evidence exists to support their classification as NAPs. In the case of mIHF, its expression in E. coli complemented loss of ihfA or ihfB in E. coli [316] and it promotes integration of mycobacteriophage L5 [317]. NapM exhibits broad genome binding in M. tuberculosis [318] and its loss in M. smegmatis leads to differential expression of 156 genes [319]. Furthermore, use of the sequence-based pipeline from COFACTOR [320] would add the “DNA binding” GO term for NapM. Rv3852 encodes a histone-like protein that binds many different DNA structures, is membrane-associated, and may be involved in tethering DNA to the membrane, biofilm formation, and regulation of motility [321,322]. Therefore, for the sparsely annotated M. tuberculosis proteome, our screen may result in more false negatives than for the species discussed above. Similar difficulties may be encountered with other sparsely annotated proteomes, but on the other hand, we expect that as computational functional annotations continue to advance in efficiency and accuracy, larger fractions of potential NAPs will be recoverable using our approach.
Table 6.
All listed M. tuberculosis proteins met our GO term and transcript abundance criteria for calling potential NAPs. aN = not discussed in [314], Y = discussed in [314]. The 80th and 90th percentile log2(TPM) values are 8.15 and 8.93, respectively
| UNIPROT ID | Protein | Gene name | In [314]a | log2(TPM) |
|---|---|---|---|---|
| P9WP75 | Probable cold shock protein A | cspA | N | 12.6 |
| P9WF43 | Transcriptional regulator WhiB1 | whiB1 | N | 11.8 |
| P9WMK7 | DNA-binding protein HU homolog | hup | Y | 11.2 |
| P9WII1 | Probable endoribonuclease MazF2 | mazF2 | N | 10.6 |
| P9WNB3 | ESX-1 secretion system protein EccCa1 | eccCa1 | N | 10.1 |
| O53238 | Probable transcriptional regulatory protein | NA | N | 10.1 |
| P9WHR7 | LexA repressor | lexA | N | 9.8 |
| P9WNB1 | Transcriptional regulator WhiB2 | whiB2 | N | 9.5 |
| O53353 | Transcriptional regulator BlaI | blaI | N | 9.5 |
| P9WMJ5 | Nucleoid-associated protein Lsr2 | lsr2 | Y | 9.4 |
| P9WIP7 | Putative DNA-binding protein Rv0500A | NA | N | 9.3 |
| P9WKT7 | ESX-1 secretion system protein EccCb1 | eccCb1 | N | 9.5 |
| P9WMH1 | Iron-dependent repressor IdeR | ideR | N | 9.2 |
| P9WF35 | Tyrosine recombinase XerC | xerC | N | 9.1 |
| P9WMH3 | CRP-like cAMP-activated global transcriptional regulator | crp | N | 9.0 |
| P96354 | Mutator family transposase | NA | N | 9.0 |
| P9WJB9 | ESX-1 secretion-associated protein EspL | espL | N | 8.9 |
| P9WGA5 | Probable transcriptional regulatory protein Rv2603c | NA | N | 8.8 |
| P9WME7 | Uncharacterized HTH-type transcriptional regulator Rv1828 | NA | N | 8.8 |
| P9WME5 | Uncharacterized HTH-type transcriptional regulator Rv1830 | NA | N | 8.7 |
| P9WKV1 | Transcriptional regulator Rv0485 | NA | N | 8.7 |
| P9WNA3 | DNA translocase FtsK | ftsK | N | 8.6 |
| P9WQ13 | Probable endonuclease 4 | end | N | 8.5 |
| Q79G00 | Probable transcriptional regulatory protein Mce1R (Probably GntR-family) | mce1R | N | 8.5 |
| P9WGM5 | Probable transcriptional regulatory protein NarL | narL | N | 8.4 |
| P9WMH7 | Transcriptional regulator ClgR | clgR | N | 8.4 |
| P9WNR9 | Nucleoid-associated protein Rv3716c | NA | N | 8.4 |
| O50462 | Antitoxin RelB | relB | N | 8.4 |
| P9WMC9 | Uncharacterized HTH-type transcriptional regulator Rv1816 | NA | N | 8.3 |
| O33333 | Probable transposase | NA | N | 8.3 |
| I6XW38 | DNA topoisomerase | NA | N | 8.3 |
| P9WHI3 | Recombination protein RecR | recR | N | 8.3 |
| P9WNA5 | ESX-5 secretion system protein EccC5 | eccC5 | N | 8.2 |
With this caveat in mind, potential mycobacterial NAPs arising from our screen and not discussed in [314] include CspA, WhiB1, and WhiB2. CspA falls into the same family of cold-shock proteins that has been discussed above and may well play a similar role in Mycobacteria; WhiB1/2 belong to an actinobacterial family of poorly characterized transcription factors that are speculated to bind DNA with low sequence specificity and can act either as repressors or transcriptional activators depending on the nature of other proteins binding at the same promoter [323].
NAPs act to suppress potentially harmful genetic elements and regulate developmental transitions
Integrating over the variety of biological processes carried out and regulated by NAPs across the examples enumerated above, coherent structural and regulatory roles for NAPs emerge. First, NAPs act as chromatin-structuring elements, with major roles in controlling both the local and large-scale conformations of the nucleoid, and making essential contributions to DNA compaction. In these structural roles, NAPs act as bacterial analogs of histone proteins (we are by no means the first to draw such analogies – others have pointed out similarity in both the structural [28] and regulatory roles [68] of NAPs). When considering the areas in which NAPs bind, several NAPs have been widely recognized as xenogeneic silencers. The enterobacterial protein H-NS is a heavily studied example [324] and represents a widely distributed family of xenogeneic silencers; flagship examples of other widespread families of xenogeneic silencers include, Mycobacteria Lsr2, Bacillus Rok, and Pseudomonas MvaT [325,326]. All four such protein families fall within the parameters of nucleoid-associated proteins as defined here. At the same time, there is some evidence that xenogeneic silencing is a coordinated effort by multiple NAPs, rather than only those that are within the classically recognized xenogeneic silencer families. For example, recent protein-occupancy profiling in E. coli demonstrated the presence of large transcriptionally silent, heterochromatin-like domains of protein occupancy [46]. But whereas only a subset of those domains match H-NS bound regions, similar levels of silencing were observed in non-H-NS regions, and must thus arise from the activity of some other silencing protein. Other E. coli NAPs have also been shown to act as xenogeneic silencers in specific contexts, such as Fis [86].
Alongside their roles in xenogeneic silencing, NAPs also play prominent roles in transitions between different cellular lifestyles. For example, E. coli H-NS regulates cell envelope composition [161] and the expression of operons important for various stages of host invasion in urovirulence [327] and enterovirulence [328]. Likewise, HU, as well as H-NS, mediates nucleoid reorganization in response to changes in environmental osmolarity, temperature, and pH to globally alter gene expression [112]. A similar role has been proposed for potential NAP CspA, which may act to globally repress gene expression during times of response to specific environmental stresses [229]. Abundant examples of similar behavior exist in other bacteria: Lsr2 regulates a range of virulence factors in Mycobacteria and secondary metabolite production in Streptomycetes [329]; IHF regulates both stationary phase entry and virulence genes in Salmonella [330]; HU acts as a regulator of virulence in a wide variety of species (recently reviewed in [331]), although in many cases insufficient data are available to determine which regulatory events reflect direct NAP binding vs. indirect network-mediated effects. Control of the stalk/swarmer transition in C. crescentus by CtrA arguably reflects yet another example, depending upon whether CtrA is classified as a global regulator or NAP. Many of the regulatory events noted here may be considered within the category of ‘developmental transitions’, as they reflect major changes in bacterial lifestyle such as activation of virulence or entry into a transiently non-growing state.
Several of the regulatory activities noted above for known xenogeneic silencers might be argued to reflect xenogeneic silencing activity. However, the fact remains that repression by xenogeneic silencers is not constitutive but permits de-repression in response to environmental cues such as changes in temperature and osmolarity, and additional binding by sequence-specific local regulators [68], as comprehensively reviewed in [332]. This provides a striking overlap between xenogenic silencers and NAPs like AbrB, which represses developmental pathways that should not be entered during rapid growth. In addition, AbrB exhibits dramatically lower expression upon transition to stationary phase [246], thus derepressing stationary phase developmental processes in B. subtilis. Likewise, it is increasingly apparent that other NAPs that are not typically considered xenogeneic silencers may also act to silence potentially harmful genetic elements and to regulate developmental transitions (as in the aforementioned case of E. coli Fis, for example).
Both of the general activities of NAPs, silencing potentially harmful DNA and regulating developmental transitions, have direct analogies to functions of histone-mediated gene regulation in eukaryotes. Constitutive heterochromatin in eukaryotes acts to silence potentially harmful mobile genetic elements [3], and facultative heterochromatin often regulates genes involved in developmental processes [333,334]. In addition, another essential aspect of histone-mediated gene regulation in eukaryotes is the presence of epigenetic marks on the histones themselves modulating gene regulatory behavior [335]; similarly, a range of post-translational modifications (PTMs) such as phosphorylation, lysine acetylation, and lysine methylation have recently been identified on many bacterial NAPs [247,336–339], and in one case post-translational modification of Mycobacterial HupB was shown to control the formation of rare epigenetically drug-resistant subpopulations [340]. Thus, despite differences in molecular mechanisms, the activity of NAPs in regulating bacterial transcription has profound parallels to histone-driven regulatory patterns in eukaryotes. The analogy must not be pushed too far, however. A particularly prominent area of difference is that the simple presence of regulation by NAPs does not necessarily imply repressive, heterochromatin-like function, as many NAPs also serve as transcriptional activators as well.
The broad and fairly nonspecific regulatory capabilities offered by NAPs (particularly the classic xenogeneic silencers) likely play a profound role in the evolution of bacterial genomes, by allowing the domestication and preliminary regulation of horizontally acquired genetic elements that imbue cells with new capabilities [159,324,332]. It appears likely that in many cases, silencing by NAPs provides an initial regulatory input into newly acquired DNA that can subsequently be modified by additional regulators that act more locally, often through interaction with the NAPs themselves (such cases have been especially clearly seen in the multitude of factors that interact with H-NS, as reviewed in [68,341]). At the same time, the regulatory capabilities of NAPs can equally well be used in the large-scale regulation of native genes to aid in stress responses (as, for example, in the case of CspA noted above [229]).
Outlook
As we have seen above, NAPs fulfill both structural and regulatory roles in bacteria that are in many ways analogous to the roles played by histones in eukaryotes, despite the completely independent molecular implementation. Many open questions remain regarding both how broadly and how deeply this analogy holds. While the molecular substrates necessary for a “NAP code” such as post-translational modifications exist, it is unclear if PTMs of NAPs enable reading/writing of epigenetic information, trans-generational memory, etc. It is also possible that the modifications are purely stochastic or that they simply arise as directed responses to the environment. A key set of related questions arise regarding the spatial distribution of different modifications – do copies of a particular protein with the same PTM co-localize in the cell, either in linear or three-dimensional space? Are PTMs involved in defining features such as transcriptionally silent regions [16], chromosomal interaction domain boundaries [25,26], or spatial clusters of distant genetic loci [342,343]? Are NAPs acting as dual regulators switched between repressive and activating roles by a PTM? It is also notable that the number of NAPs apparent in E. coli is substantially larger than the known list of NAPs in many other species. The full extent to which similar proteins play equivalent functional roles in other bacteria remains to be seen, although we have shown here that several other proteins in a variety of species appear to share the characteristics of E. coli NAPs in terms of both regulatory and structural roles. The full extent and balance of the various mechanisms by which NAPs might regulate transcription (i.e., by blockage of RNA polymerase recruitment, polymerase elongation, effects on DNA topology, RNA binding, and phase separation) remains to be determined, as is the full extent to which the regulatory logic of NAPs is determined by the cell’s transcriptional regulatory network. For all of the questions posed above, ongoing multimodal investigations will be needed to merge information on transcriptional regulatory states, protein modification states, and protein-nucleic acid interactions into a coherent model of the regulation of bacterial chromatin by NAPs.
Acknowledgments
We are grateful to members of the Freddolino laboratory for many helpful discussions, and also to the generations of scientists who have contributed to our tremendous body of knowledge on bacterial NAPs. Work in the authors’ laboratory is supported by NIH R35GM128637 and R01AI13467801. The E. coli RNA-seq datasets used in this work were generated by Dr Judith Kribelbauer and PLF in the lab of Saeed Tavazoie (Columbia U.).
Funding Statement
This work was supported by the National Institute of Allergy and Infectious Diseases [R01AI13467801]; National Institute of General Medical Sciences [R35GM128637].
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