RNA Management During T7 Infection

Abstract

Post-transcriptional regulation (PTR) determines the fate of RNA in the cell and represents an important control point in the flow of genetic information and thus underpins many, if not all, aspects of cell function. Host takeover by phages through misappropriation of the bacterial transcription machinery is a relatively advanced area of research. However, several phages encode small regulatory RNAs, which are major mediators of PTR, and produce specific proteins to manipulate bacterial enzymes involved in RNA degradation.^1–4 However, PTR during phage development still represents an understudied area of phage–bacteria interaction biology. In this study, we discuss the potential role PTR could play in determining the fate of RNA during the lifecycle of the prototypic phage T7 in Escherichia coli.

Regulation of RNA Synthesis by T7—the Knowns

Phages have evolved complex and elegant strategies to manipulate bacterial metabolic processes to divert host resources to benefit phage development and progeny production.^5,6 Transcription represents the first step of gene expression when RNA is synthesized. Unsurprisingly, a common theme by which phages modulate bacterial physiology is through the bacterial RNA polymerase (RNAP), the multisubunit enzyme that catalyzes DNA transcription.⁷ Gene specificity upon the bacterial RNAP is conferred by one of the several sigma (σ) factors present in bacterial genomes, which direct their cognate RNAPs to express different sets of genes to meet cellular requirements. In fact, many phages encode σ-factor like proteins to reprogram the specificity of the bacterial RNAP to benefit phage development.^8,9 Phages also encode small proteins that specifically modulate or interfere with the activity of the bacterial RNAP to facilitate and coordinate phage transcriptional programs.

A paradigmatic example of this is provided by T7 and we refer interested readers to a recent detailed overview of this topic by Tabib-Salazar et al.⁷ In brief, the transcription of T7 genes occurs in a temporally coordinated manner and proceeds uniformly from left to right on the T7 genome. Hence, the T7 genes are designated early, middle, and late based on the timing of their expression during the phage development process (Fig. 1). The transcription of early genes is catalyzed by the Escherichia coli RNAP containing the housekeeping σ factor, σ⁷⁰, whereas the transcription of middle and late genes require the T7 RNAP, the product of the T7 early gene 1, gp1. The coordination between the transcription of early T7 genes and middle and late T7 genes, that is, the switching of the dependency from the E. coli RNAP to T7 RNAP, is orchestrated by three T7 encoded E. coli RNAP interacting proteins, encoded by early T7 gene gp0.7 and middle genes gp2 and gp5.7.

FIG. 1.

Schematic diagram showing how the Escherichia coli RNAP is regulated by Gp0.7, Gp2 and Gp5.7 to coordinate transcription of T7 genes. We refer interested readers to Tabib-Salazar et al⁷ where more details are provided. RNAP, RNA polymerase.

As the σ⁷⁰-RNAP is not required for the transcription of T7 middle and late genes, it becomes initially inhibited by the action of Gp0.7 and Gp2. This inhibition is important for T7 development as it prevents any aberrant σ⁷⁰-RNAP activity into the regions of the T7 genome that contain the middle and late genes, which could thwart phage replication and progeny development.¹⁰ Gp2 binds tightly to the RNAP and inhibits several obligatory conformational changes in σ⁷⁰ that are required for transcription.^11,12 In contrast, Gp0.7, a serine/threonine kinase, phosphorylates the RNAP (among other host proteins involved in RNA metabolism—see hereunder), and thereby enhances the ability of the RNAP to terminate at specific termination sites located between the T7 early and middle genes.¹³

Hence, Gp2 is an essential T7 protein under any growth condition, whereas Gp0.7 only becomes essential when the activity of Gp2 is compromised or under compromised growth conditions (nutrient depletion, elevated temperature, etc.).^10,13,14 It is known that T7 infection of E. coli results in the accumulation of (p)ppGpp, the bacterial stress-signaling nucleotide guanosine pentaphosphate.^15,16 One of the major consequences of (p)ppGpp accumulation is that it directly contributes to the accumulation of σ³⁸, which competes with σ⁷⁰ for the bacterial RNAP. The σ³⁸-RNAP can transcribe T7 early genes as effectively as the σ⁷⁰-RNAP. However, Gp2 is a poor inhibitor of the σ³⁸-RNAP. As such, this poses a further challenge for T7 to ensure that aberrant transcription by bacterial RNAP into regions of the genomes transcribed by the T7 RNAP does not occur, and this is when Gp5.7 is required. Unlike Gp2, Gp5.7 is a specific inhibitor of the σ³⁸-RNAP.¹⁵

Thus, Gp0.7, Gp2, and Gp5.7 collaborate to initially takeover but subsequently fully inhibit the host RNAP to benefit optimal execution of the T7 transcription program. A succinct diagrammatic summary of how the transcription of T7 early, middle, and late genes are coordinated by Gp0.7, Gp2, and Gp5.7 is provided in Figure 1.

Post-Transcriptional Regulation of RNA in T7-Infected E. coli—the Unknowns

Although the regulatory basis by which T7 RNA synthesis in E. coli is managed is clearly well understood, the role of post-transcriptional regulation (PTR) in T7 development in E. coli has not been studied in any detail. Non-coding regulatory RNA molecules, called small RNAs (sRNAs), play a pivotal role in PTR and identify which and to what extent mRNA molecules targeted for PTR are translated, translationally repressed or degraded.¹⁷ For all these functions, sRNAs rely on RNA binding “chaperone” proteins to facilitate the base pairing between them and their target mRNAs. In E. coli and related bacteria, most sRNA-mediated PTR is facilitated by the ubiquitous RNA binding protein Hfq.¹⁸

Therefore, to investigate whether PTR plays any role in T7 development in E. coli, we used hfq as a surrogate and measured the ability of T7 to develop in bacteria devoid of hfq in a T7 plaque growth assay. This assay allows assessment of the efficacy of T7 plaque formation on a lawn of E. coli as a function of time.¹⁵ In brief, E. coli MG1655 wild-type and Δhfq cultures were grown to an OD₆₀₀ of 0.45 in lysogeny broth (LB) at 37°C and aliquots of the culture was taken out and T7 lysate (sufficient to produce ∼10 plaques) was added. This phage–bacteria mixture was incubated 37°C for 10 min to allow the phage to adsorb to the bacteria. The mixture was then plated onto LB agar plates. The inoculated plates were then put on a standard office scanner placed in a 30°C incubator and images of the plate was taken every 2 h during a 72-h period of analysis. Results did not reveal any obvious differences (density, appearance, color, etc.) in the lawn formed by wild-type and Δhfq bacteria and the rate of plaque formation by T7 on both lawns was initially indistinguishable (Fig. 2). However, over time, the plaques produced by T7 on wild-type bacteria continued to enlarge, but the plaques formed by T7 on Δhfq bacteria slowed down and eventually ceased to grow. When plasmid-borne hfq was provided to Δhfq bacteria, the rate of plaque formation resembled that seen in wild-type bacteria. It seems that hfq is a novel host factor for T7 and that PTR might indeed play a role in efficient T7 development in E. coli.

FIG. 2.

Schematic showing the T7 plaque growth assay.¹⁵ Representative scanned images of T7 plaques formed on a lawn of wild-type and Δhfq Escherichia coli over a 72-h incubation period. The graph shows average (n = 5) plaque size as percentage of final plaque size formed by T7 on a lawn of wild-type bacteria after 72 h of incubation.

An open question remains whether PTR during T7 development in E. coli involves any sRNA of T7 origin and/or whether bacterial sRNAs are misappropriated to do so. The former is a possibility as sRNAs of phage origin have been documented before.^2–4 A phage plaque is a clearing in a bacterial lawn and plaques form via an outward diffusion of progeny virions that prey on surrounding bacteria. One of the limiting factors of plaque growth is the physiological status of the bacteria on the lawn, which concomitantly become nutrient starved with the aging of the lawn. We note that the differences in the rate of T7 plaque formation on wild-type and Δhfq bacteria become more pronounced as the cells age and thus become growth attenuated. Hence, PTR could have a more prominent role in T7 development in the natural physiological context where bacteria are often in a growth-attenuated state.

However, given the functional pleiotropy of hfq, any interpretation based on the ability of T7 to develop in Δhfq bacteria must be made cautiously. For instance, it is well established that the composition of the bacterial cell wall is highly heterogeneous and dynamically regulated by mechanisms involving Hfq-sRNA-mediated PTR upon stress.¹⁹ Hence, the differences we see in T7 plaque formation on wild-type and Δhfq bacteria could be owing to differences in the cell wall composition that prevent T7 binding (absorption) to aged Δhfq bacterial cells—an aspect that yet awaits investigation.

As the E. coli RNAP becomes inhibited in the T7-infected bacteria and the E. coli chromosome is hydrolyzed by T7-encoded exonucleases, it is likely that Hfq-sRNA-mediated PTR solely serves to fulfill the gene expression requirements of T7 and to provide and divert cellular resources to do so. Hfq-sRNA-mediated PTR during T7 development in E. coli could thus involve preferential translation of T7 mRNAs and, conversely, translational repression or degradation of bacterial mRNAs. In E. coli, the Hfq-sRNA-guided degradation of target mRNA by the RNA degradosome represents a major aspect of Hfq-sRNA-mediated PTR. The canonical RNA degradosome complex consists of RNase E, the RNA helicase RhlB and the 3′-5′ exoribonuclease polynucleotide phosphorylase (PNPase). The interaction between a Hfq-sRNA–mRNA complex and the RNA degradosome is facilitated by the Hfq bound sRNA, which interacts with the carboxyl-terminal domain (CTD) of RNase E (where RhlB and the PNPase also bind).

Intriguingly, Gp0.7 also phosphorylates, among ∼90 other host proteins, RNase E (in the CTD) and RhlB as well as several components of the translation apparatus.²⁰ Neither the precise phosphorylation sites on and percentage phosphorylation of target proteins nor the full biological function of Gp0.7 during T7 development is currently known. However, the timing of expression of Gp0.7 and the biological functions of many of its phosphorylation targets suggest an important role for Gp0.7 in Hfq-sRNA-mediated PTR during T7 development. In addition to the established role of Gp0.7 in transcription regulation (see previously), two mutually inclusive roles for Gp0.7 in PTR during T7 development can be considered. First, as mentioned earlier, Gp0.7 is not essential for T7 development in actively growing E. coli, but it becomes essential under stress or nutrient-starved growth conditions that lead to growth attenuation.^21,22 A possible explanation for this change in requirement could be because global RNA degradation generally becomes greatly enhanced under such conditions. In addition to Hfq-sRNA-guided target RNA cleavage, RNase E can also directly sense and cleave target RNAs with a monophosphorylated 5′-end (generated from primary transcripts by pyrophosphohyrolase or other RNases). Therefore, the phosphorylation of RNase E and RhlB by Gp0.7 could attenuate (but not fully inhibit) the hydrolytic activity and/or compromise the composition of the RNA degradosome so that any collateral chances that T7 mRNAs destined for translation/translational repression are degraded remain low. Indeed, a role for Gp0.7 in stabilizing T7 mRNAs over bacterial mRNAs has been previously suggested.²³ However, a recent study contested this view and reported that most if not all T7 early and middle gene mRNAs become degraded shortly after synthesis to potentially favor translation of T7 late mRNAs that encode structural proteins for the assembly of progeny T7 virions.²⁴

Second, during T7 development in E. coli, mRNA targeted for both translation and degradation are simultaneously exposed to the bacterial RNA degradosome. Conversely, the translation apparatus can access any mRNA destined for degradation. Therefore, ensuring that the mRNAs destined for degradation (that would be predominantly bacterial mRNA) are distinguished from mRNA that are destined for translation or translational repression (that would predominantly be T7 mRNAs) could represent a conundrum for T7. It is thus possible that phosphorylation of the RNA degradosome and the translation apparatus by Gp0.7 serves to change the subcellular distribution and localization of either one or both of these macromolecular machineries so that they are physically separated from one another during T7 development. Indeed, the subcellular compartmentalization of the RNA degradosome has been observed in bacteria under diverse growth conditions.^25–28

Conclusion

Clearly, much research is warranted to establish how mRNA is managed in T7-infected E. coli. Recent high-resolution next-generation sequencing methods to capture the interactomes of RNA binding proteins and identities of RNA targeted for degradation and translation provide tractable experimental methods to investigate the many unknowns of PTR during T7 development discussed here. These could be combined with single molecule imaging techniques in live bacteria to study the subcellular distribution of enzymes involved in PTR during T7 infection. Nonetheless, the management of the fate of RNA in the phage-infected bacterial cell is perhaps an aspect of host takeover by phages that is still in its infancy and one that could offer new opportunities to uncover novel paradigms of gene regulation in bacteria.

Authors' Contributions

Conceptualization, methodology investigation, formal analysis, and writing (A.T.-S. and S.W.); funding acquisition (S.W.).

Author Disclosure Statement

No competing financial interests exist.

Funding Information

The author received funding from the Wellcome Trust (100958), Leverhulme Trust (RPG-2020-050) and BBSRC (BB/V000284/1).