# Rho-dependent transcription termination: mechanisms and roles in bacterial fitness and adaptation to environmental changes

Thuy Duong Do, Nara Figueroa-Bossi, Johnathan Black, Eric Eveno, Mildred

Delaleau, Lionello Bossi, Marc Boudvillain

► To cite this version:

Thuy Duong Do, Nara Figueroa-Bossi, Johnathan Black, Eric Eveno, Mildred Delaleau, et al.. Rho-dependent transcription termination: mechanisms and roles in bacterial fitness and adaptation to environmental changes. RNA, 2025, 31, pp.rna.080486.125. 10.1261/rna.080486.125. hal-05097114

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1 Rho-dependent transcription termination: mechanisms and roles in bacterial
2 fitness and adaptation to environmental changes

3

4 Thuy Duong Do¹,², Nara Figueroa-Bossi³, Johnathan C. Black¹,², Eric Eveno¹, Mildred

5 Delaleau¹,², Lionello Bossi³, and Marc Boudvillain¹,²,⁴

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8 ¹Centre de Biophysique Moléculaire, CNRS UPR4301, affiliated with Université d'Orléans, rue

9 Charles Sadron, 45071 Orléans cedex 2, France.

10 ²ED 549, Sciences Biologiques & Chimie du Vivant, Université d'Orléans, France.

11 ³Université Paris-Saclay, CEA, CNRS, Institut de Biologie Intégrative de la Cellule (I2BC), 91190

12 Gif-sur-Yvette, France.

13 ⁴Corresponding author (marc.boudvillain@cnrs.fr)
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ABSTRACT

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The bacterial transcription termination factor Rho is a rare example of an RNA helicase that functions as a ring-shaped ATP-powered six-subunit motor. Recent studies have linked Rho's distinctive architecture to a variety of regulatory mechanisms that shape the bacterial transcriptome at the global scale and control the transcription of individual genes in a context-dependent manner. In this review, we provide a comprehensive overview of the molecular mechanisms by which Rho triggers transcription termination. We examine the two prevailing modes of Rho's action: the "catch-up" mode, where Rho actively translocates along RNA and collides with the RNA polymerase to terminate transcription, and the "stand-by" mode where Rho, recruited by transcription elongation factor NusG, remains poised to engage RNA polymerase at specific sites or under particular constraints. Additionally, we highlight Rho's interplay with nucleoid-structuring protein H-NS in the regulation of bacterial chromatin transcription, as well as the crucial role played by Rho in the conditional regulation of specific genomic loci. We discuss how these mechanisms contribute to the fine-tuning of gene activity and integrate into broader regulatory networks, supporting bacterial adaptation to environmental changes and resilience to external challenges.
## 30 INTRODUCTION

Bacteria are highly adaptable microorganisms capable of surviving and thriving in diverse, often hostile environments. To cope with ecosystem fluctuations, such as changes in temperature, hydration, pH, redox potential, light exposure, nutrient availability, competition, or host immune response, they have evolved a variety of general and taxon-specific adaptive mechanisms. These mechanisms enable rapid gene expression adjustments in response to external stimuli, allowing bacteria to fine-tune metabolism, activate stress responses, and produce the virulence factors that optimize the balance between growth and survival (Zhu and Dai 2024).

A key player in this dynamic regulatory landscape is the transcriptional machinery, which governs RNA synthesis from the bacterial chromosome(s) and, when present, adaptive plasmid(s). Transcription termination is a crucial step in this process, ensuring proper gene and operon delineation while regulating their timely expression. For instance, premature termination prevents transcription from extending into the structural portions of genes or operons through a regulatory mechanism known as transcription attenuation (Turnbough 2019). Triggered by specific stimuli, transcription attenuation shuts down gene expression at an early stage, making it particularly relevant in bacteria, where transcription and translation are often tightly coupled (Woodgate and Zenkin 2023) and where the synthesis of polycistronic RNAs imposes a high metabolic cost.

Two primary mechanisms govern transcription termination in bacteria (Ray-Soni et al. 2016). Intrinsic (Rho-independent) termination relies on specific transcript motifs –a stable hairpin structure followed by a run of uracil residues- sufficient to disrupt the transcription elongation complex (TEC). In contrast, Rho-dependent termination (RDTT) requires the ring-shaped, ATP-dependent translocase/helicase Rho. Rho binds to nascent transcripts at poorly defined Rho utilization (Rut) sites and uses its ATP-dependent motor activity to disrupt the TEC (see below for details). Rho is broadly conserved across bacteria but is noticeably absent in *Cyanobacteriota*, *Mycoplasmatota*, or *Streptococcaceae* (D'Heygere et al. 2013; Moreira et al. 2024). Rho is also dispensable in *Bacillota* under laboratory growth conditions, even though its inactivation significantly alters the transcriptome in species such as *Bacillus subtilis* (Nicolas et al. 2012; Bidnenko et al. 2023) and *Staphylococcus aureus* (Mader et al. 2016; Nagel et al.
60 2018). While the relative contributions of intrinsic termination and RDTT vary across taxa
61 (Ahmad et al. 2023), recent findings reveal a composite regulatory landscape where these
62 mechanisms are not strictly independent (Mandell et al. 2022b). Notably, transcription
63 factors, including Rho, can enhance termination at weak intrinsic terminators in *B. subtilis*
64 (Mandell et al. 2022a) and *Escherichia coli* (Ahmad et al. 2022), blurring the distinction
65 between factor-dependent and independent pathways. In this review, we summarize the
66 current understanding of the mechanisms of RDTT and highlight recent examples of its
67 integration into complex regulatory networks, underscoring its critical role in bacterial
68 adaptation and fitness.
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70 THE ARCHITECTURE OF THE RHO MOTOR AND ITS MECHANISM(S) OF ACTION

71 The Rho factor is a ring-shaped assembly of six identical subunits. Each Rho subunit is divided
72 structurally and functionally into two major domains (Figure 1A). The N-terminal domain
73 (NTD) harbors the Primary Binding Site (PBS), which mediates Rho's recognition of and binding
74 to *Rut* sites in nascent transcripts. The C-terminal domain (CTD) contains motifs essential for
75 hexamer cohesion, ATP hydrolysis, and RNA translocation, which occurs via the Secondary
76 Binding Site (SBS) loops within the central Rho channel. The CTD is highly conserved, while the
77 NTD is more variable and often contains large polypeptide insertions (Figure 1A). These
78 insertions are particularly common in *Actinomycetes* and *Bacteroidota* (D'Heygere et al. 2013;
79 Moreira et al. 2024) where they likely aid Rho in recognizing genuine *Rut* sites (Nowatzke et
80 al. 1997; D'Heygere et al. 2015; Simon et al. 2021). In some species, the NTD insertion domain
81 is intrinsically disordered, facilitating either liquid-liquid phase separation (Krypotou et al.
82 2023) or prion-like aggregation (Yuan and Hochschild 2017) of Rho.
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A recent cryo-electron microscopy (cryoEM) study of *E. coli*'s Rho bound to the
83 prototypical λtR1 terminator RNA provides structural insights into how Rho's composite,
84 crown-like PBS recognizes and binds a *Rut* site (Figure 1B) (Molodtsov et al. 2023). Each Rho
85 subunit engages five consecutive single-stranded nucleotides (nt) separated by unbound
86 linkers of 4-5 nt. These contacts extend beyond the 5'YC-binding PBS clefts (where Y
87 represents a U or C residue) previously identified in crystal structures of *E. coli*'s Rho (Bogden
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et al. 1999; Skordalakes and Berger 2003). Although only two of these PBS clefts appear to be
occupied by YC dinucleotides in the cryoEM structure (Figure 1B, PBS clefts in purple with
bound dinucleotides in green), the nucleotide identities should be considered tentative given
the resolution of the structure (4.27 Å).

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Alignment of the six λtR1 RNA segments bound to the Rho subunits provides a
tentative RNA consensus sequence for PBS interaction, with A and C residues preferred at
most positions (Figure 1C, top). Analysis of the hundreds of putative Rut sites recently
identified in the E. coli genome by Helicase-SELEX (Figure 1E) (Delaleau et al. 2022) yields a
similar, albeit shorter consensus sequence (Figure 1C, bottom), possibly reflecting PBS
preferences of individual subunits and the minimal specificity of inter-subunit RNA linkers.

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Overall, these findings reinforce the long-standing view that *Rut* sites are difficult to
define, aside from their preferential localization in C>G-biased regions with limited secondary
structure stability (Alifano et al. 1991; Peters et al. 2012; Nadiras et al. 2018a; Delaleau et al.
2022). The optimal *Rut* site length appears to be in the 60-90 nt range (Hart and Roberts 1994;
Walmacq et al. 2004; Molodtsov et al. 2023), though shorter (~30 nt) RNA sequences or
hairpin-like structures can still bind Rho tightly (Schneider et al. 1993) or elicit Rho activity
(Delaleau et al. 2022). This suggests that not all PBS clefts need to be occupied or bridged
uniformly, to trigger Rho's catalytic isomerization. Due to these cryptic features, predicting
*Rut* sites (and by extension, Rho-dependent terminators) is challenging, requiring cautious use
of specialized algorithms (Nadiras et al. 2018a; Di Salvo et al. 2019). Nonetheless, Helicase-
SELEX screening suggests that *Rut* sites occur only at discrete locations (Figure 1E), collectively
covering less than 5% of the *E. coli* genome (Delaleau et al. 2022). This implies that *Rut* site
recognition by Rho is highly specific, positioning it as a key regulatory checkpoint in the RDTT
pathway (see below for further discussion).

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Extensive structural and biophysical data confirm that the Rho hexamer alternates
between open (Skordalakes and Berger 2003; Skordalakes et al. 2005; Canals et al. 2010; Hao
et al. 2021a; Said et al. 2021; Saridakis et al. 2022) and closed (Thomsen and Berger 2009;
Thomsen et al. 2016; Molodtsov et al. 2023) ring conformations. The open-ring form
predominates when Rho is free of RNA or during PBS-mediated RNA loading (Skordalakes and
Berger 2003; Canals et al. 2010). In this state, the corkscrewed subunit arrangement is
catalytically inactive but guides the RNA chain towards the central ring channel. Upon entry into the central cavity, RNA interacts with Rho's SBS loops, triggering ring closure and catalytic activation (Figure 1D) (Thomsen and Berger 2009; Thomsen et al. 2016). This conformational transition is rate limiting in vitro (Kim and Patel 2001; Walmacq et al. 2004) and serves as another important regulatory checkpoint, for instance modulated by elongation factor NusG (Lawson et al. 2018) or blocked by the drug Bicyclomycin (Lawson et al. 2016).

Once activated, Rho translocates along the RNA in the 5'→3' direction in an ATP-dependent manner (Steinmetz and Platt 1994; Schwartz et al. 2007; Schwartz et al. 2009), moving rapidly (Gocheva et al. 2015) but with moderate processivity (Walmacq et al. 2004; Soares et al. 2014). During translocation, SBS-RNA interactions are remodeled while the *Rut* site remains bound to the PBS (Figure 2A) (Steinmetz and Platt 1994; Koslover et al. 2012; Gocheva et al. 2015). This "tethered tracking" mechanism (Steinmetz and Platt 1994) allows Rho to catch up with the RNA polymerase (RNAP), typically at transcriptional pause sites (Richardson and Richardson 1996; Guerin et al. 1998; Hollands et al. 2014). Once in contact with the RNAP, the Rho motor either fully dissociates the TEC or only disengages the RNA from the TEC while the RNAP remains bound to the DNA template (Song et al. 2022). This "catch-up" model is backed by a wealth of studies (Table 1), including kinetic coupling experiments with Rho and RNAP velocity mutants (Jin et al. 1992; Shashni et al. 2012), and a recent cryoEM reconstruction of the *E. coli* pre-termination complex where Rho interacts with RNAP through its CTD face (Figure 2B, in gray) (Molodtsov et al. 2023). Elongation factor NusG stabilizes this pre-termination complex by linking Rho and RNAP (Figure 2B), potentially aiding termination by acting as a stator for the Rho motor that restricts its rotation with respect to RNAP (Molodtsov et al. 2023). The organization of this "catch-up" pre-termination complex is consistent with previous structures of binary Rho-RNA (Thomsen and Berger 2009; Thomsen et al. 2016) and Rho-NusG (Lawson et al. 2018) complexes.

Distinct cryo-EM reconstructions of the *E. coli* pre-termination complex (Hao et al. 2021a; Said et al. 2021) support an alternative "stand-by" model in which an open, catalytically inactive Rho hexamer is pre-bound to RNAP in an inverted CTD→NTD face orientation (Figure 2C, D). In this model, Rho binds RNAP in non-coding or untranslated regions of nascent transcripts, scans the growing RNA chain from this RNAP-bound position, and triggers termination by allosteric inactivation and destabilization of the pre-termination complex upon
detecting a *Rut* site (Epshtein et al. 2010; Hao et al. 2021a; Said et al. 2021). Although the RNA trajectory cannot be definitely determined from the cryo-EM reconstructions, it was proposed that the RNA chain might be guided by the transcription factor NusA to contact Rho's SBS before the PBS, in a reverse order when compared to the traditional "catch-up" model (Hao et al. 2021a; Hao et al. 2021b). While this proposal remains subject to debate (Table 1), RNA is not required for the "stand-by" Rho-RNAP assembly, which is stabilized by NusA and, to a lesser extent, by NusG (Figure 2D) (Hao et al. 2021a; Hao et al. 2021b; Said et al. 2021).

Which model best describes RDTT is still debated, as there are pros and cons for each proposed mechanism (Hao et al. 2021b; Rashid and Berger 2023), while an alternative perspective incorporates “stand-by” complexes into the classical “catch-up” scheme (see Table 1 and below). One appealing feature of the “stand-by” model is that Rho is co-localized with the nascent transcript through its interaction with RNAP, providing a straightforward way to segregate genuine RNA targets from the bulk of cellular RNAs, which are RNAP-free (Hao et al. 2021a; Hao et al. 2021b; Said et al. 2021). However, target discrimination or subcellular localization could also be achieved by other mechanisms, such as liquid-liquid phase separation (Nandana and Schrader 2021), protein filament polymerization (Mance et al. 2024), or topological constraints imposed on the nucleoid by various DNA- and RNA-binding proteins (Hustmyer and Landick 2024). For example, the Rho factor of *Bacteroides thetaiotaomicron*, contains an unusually large intrinsically disordered region (IDR) inserted within its NTD (Figure 1A), which promotes the formation of Rho condensates independently of RNAP presence (Krypotou et al. 2023). These Rho condensates are detectable *in vivo* by immunofluorescence, unless the IDR is deleted, highlighting their importance for the bacterium's fitness. Alternatively, Rho polymerization may regulate the spatial distribution of inactive versus active Rho factors, similar to the mechanism observed for other transcription factors (Mance et al. 2024). A recent work suggests that *E. coli*'s Rho can indeed oligomerize into long filaments under certain conditions, including cellular stresses (Wang et al. 2025). Further investigations using advanced super-resolution microscopy approaches will be necessary to determine the subcellular oligomeric states of Rho and whether all Rho particles, or only spatially defined subsets, actively participate in transcription termination.

Intriguingly, interactions that stabilize the "stand-by" pre-termination complex (Figure 2D) include Rho contacts to a lineage-specific insertion sequence in the β subunit of RNAP
181 (Said et al. 2021; Simon et al. 2021). While some of these interactions are supported by genetic
182 or biochemical evidence (Epshtein et al. 2010; Hao et al. 2021a; Said et al. 2021), it remained
183 unclear whether "stand-by" pre-termination complexes could form in species distant from E.
184 coli (Simon et al. 2021). Recent cryo-EM reconstructions of the pre-termination complex from
185 Thermus thermophilus (Murayama et al. 2023) provide some insight. In this species, the Rho
186 factor binds the TEC in a configuration that closely resembles the "catch-up" pre-termination
187 complex observed in E. coli (Figure 2B). However, 3D classification of the cryo-EM particles
188 revealed an alternative binding mode where the NTD face of the Rho hexamer contacts RNAP
189 in a "stand-by"-like manner. Nonetheless, this configuration differs from the "stand-by" model
190 as Rho adopts a closed-ring conformation while RNAP is in an apo-like rather than TEC-like
191 state. Biochemical probing of the Rho-RNAP interfaces suggested that the "catch-up" complex
192 is the genuine pre-termination state, while the "stand-by"-like complex may be an off-
193 pathway intermediate (Murayama et al. 2023).

194 Recent work by S. Hohng and colleagues provides key insights that may help reconcile
195 the "catch-up" vs "stand-by" views. Using single-molecule fluorescence assays, these authors
196 demonstrated that Rho could engage in both pathways in vitro (Song et al. 2022). Across all
197 terminators tested, the "catch-up" RDTT pathway, leading to RNA release and subsequent 1-
198 dimensional sliding of RNAP on DNA, was the most frequently utilized and the fastest (Song et
199 al. 2023). In contrast, the "stand-by" RDTT route was the slowest and, intriguingly, the only
200 pathway in which termination efficiency positively correlated with the duration of RNAP
201 pausing along DNA (Song et al. 2023). This finding, together with the inactive (open)
202 conformation of the Rho hexamer in the "stand-by" complex (Figure 2D), may reflect the time
203 needed for the complex to transition into a productive state. This suggests that the "stand-
204 by" complex could represent either a very early intermediate or an off-pathway state,
205 preceding Rho's engagement in the active RDTT pathway. Such inactive Rho association with
206 the TEC may explain the previously noted similarity between Rho and RNAP distributions along
207 the E. coli genome, as observed in ChIP-chip analyses (Mooney et al. 2009). Notably, "catch-
208 up" Rho-RNA complexes (Figure 2A) are unlikely to be efficiently cross-linked to DNA and,
209 thus, may go undetected in ChIP-chip experiments. These observations raise the possibility
210 that two Rho recruitment pathways coexist: one in which Rho diffuses freely in solution before
211 engaging the transcript, and another in which Rho is pre-bound to RNAP and subsequently
transfers to the nascent RNA, ultimately triggering TEC inactivation and RDTT. Evidence
supporting the coexistence of two distinct Rho recruitment pathways *in vivo* comes from
examining how Rho and NusG alterations impact RDTT profiles at the genome-wide level (see
section below).

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217 RDTT IN GLOBAL TRANSCRIPTION MANAGEMENT

Transcriptome analyses comparing the RNA profiles of untreated cells with those of cells
undergoing Rho inhibition or depletion have enabled the mapping of the RDTT landscape in
key model bacteria. These studies have uncovered significant differences in how various
bacteria rely on Rho to terminate transcription at the end of genes. For instance, while only
about 20% of *E. coli* genes carry Rho-dependent terminators at their 3' ends (Peters et al.
2012), the vast majority of genes in Mycobacteria have their transcripts terminated by Rho
(Botella et al. 2017; Ahmad et al. 2023). However, a common finding across all studied
organisms is the central role played by RDTT in suppressing pervasive antisense transcription
within coding regions (Nicolas et al. 2012; Peters et al. 2012; Leela et al. 2013; Mader et al.
2016; Raghunathan et al. 2018). This activity is thought to help prevent the formation of
deleterious transcriptional R-loops upstream of RNAPs (Leela et al. 2013; Raghunathan et al.
2018; Pandiyan et al. 2024) and to facilitate the removal of stalled TECs, which can otherwise
create conflicts with DNA replication and lead to DNA breaks (Washburn and Gottesman 2010;
Dutta et al. 2011; Jain et al. 2019). One striking finding from early transcriptomic studies was
that Rho, assisted by NusA and NusG, silences the expression of horizontally acquired A/T-rich
regions in the *E. coli* genome (Cardinale et al. 2008). Among Rho-silenced targets are toxic
prophage genes (Cardinale et al. 2008; Menouni et al. 2013; Hafeezunnisa et al. 2021;
Baniulyte and Wade 2025), which partially accounts for the essentiality of NusG and Rho in *E.
coli* (Cardinale et al. 2008). This elevated termination activity in A/T-rich regions was initially
attributed to the presence of codons with a lower adaptation index, which cause translational
stalling and thereby promote RDTT (see below) (Cardinale et al. 2008).

An alternative scenario emerged from the finding that Rho-mediated silencing sites
coincide with regions bound by the nucleoid-structuring protein H-NS (Saxena and
Gowrishankar 2011; Peters et al. 2012). H-NS, a highly abundant DNA-binding protein, plays a
central role in silencing horizontally acquired genes (Lucchini et al. 2006; Navarre et al. 2006).
It binds to A/T-rich DNA sequences, forming nucleoprotein filaments that block RNAP access
to promoters over extended regions (Lucchini et al. 2006; Landick et al. 2015). This repression
not only prevents transcription from regular promoters but also silences the multitude of
spurious intragenic promoters that arise due to the high A/T content of these regions (Singh
et al. 2014; Lamberte et al. 2017). The presence of Rho-dependent terminators within H-NS-
bound regions suggests that H-NS-mediated repression of transcription initiation is
incomplete, and that Rho is required to terminate elongating transcripts that occasionally
escape this control. This functional cooperation between H-NS and Rho was recently
demonstrated in *Salmonella*, where certain *rho* mutations or NusG depletion result in the
constitutive activation of H-NS-silenced pathogenicity islands (Bossi et al. 2019). In the case of
*Salmonella* Pathogenicity Island 1 (SPI-1), read-through transcripts originating several
hundred base pairs upstream of the master regulator gene *hilD*, relieve H-NS-mediated
repression without actually reaching the *hilD* promoter. Remarkably, repression is restored by
inserting a strong gyrase site (SGS) within the intervening sequence (Figueroa-Bossi et al.
2024). This led to a model in which transcription destabilizes H-NS:DNA filaments at a distance
via the generation of positive DNA supercoiling (Figure 3). Drawing from *in vitro* evidence that
H-NS can constrain negative supercoils (Tupper et al. 1994), the model proposes that H-NS
filaments *in vivo* form on negatively supercoiled pletonemic DNA, with H-NS bridging the
opposing arms of the pletoneme. Rotational diffusion of positive supercoils generated by
neighboring transcription (Liu and Wang 1987; Wu et al. 1988) may cause the H-NS-bound
plectoneme to “unroll”, disrupting H-NS bridges and leading to transient filament disassembly.
After the passage of RNAP, the filament can rapidly reassemble, aided by the accumulation of
negative supercoils behind the moving transcription complex (Figure 3). This dynamic scenario
aligns with ChIP data showing that substantial amounts of H-NS remain bound to SPI-1 even
when the island is actively transcribed (Figueroa-Bossi et al. 2022; Figueroa-Bossi et al. 2024).

A major conclusion from these studies is that, *in vivo*, H-NS alone is insufficient to halt
elongating transcription complexes and instead relies on Rho to perform this function. H-
NS:DNA filaments have been shown to prolong RNAP pausing (Kotlajich et al. 2015; Boudreau
et al. 2018), creating a window for NusG-mediated recruitment of Rho. This recruitment must
occur early during elongation, before the surge of positive supercoiling leads to filament
273 destabilization. Interestingly, a Rho mutant defective in *Rut* site recognition (Y80C) was found
274 to retain full capacity to support H-NS-mediated gene silencing (Bossi et al. 2019). This
275 observation suggests that RDTT at H-NS-repressed loci occurs predominantly via the “stand-
276 by” pathway, which is expected to be less dependent on strong RNA binding and more
277 influenced by the duration of RNAP pausing (Song et al. 2023).

278 Genome-wide studies in *E. coli* have hinted at the presence of Rho-dependent
279 terminators in the 5'UTRs of various genes (Peters et al. 2009; Sedlyarova et al. 2016;
280 Sedlyarova et al. 2017; Adams et al. 2021), implying that Rho could play a significant role in
281 the biogenesis of non-coding RNAs and/or in transcription attenuation mechanisms. This
282 hypothesis is further supported by a recent map of putative *Rut* sites generated using
283 Helicase-SELEX (Delaleau et al. 2022), a mapping approach that is not biased by the extensive
284 posttranscriptional 3'-end processing of Rho-dependent transcripts (Dar and Sorek 2018).
285 Alongside the frequent occurrence of RDTT in horizontally acquired portions of the *E. coli*
286 genome, these observations suggest that RDTT could serve as a major adaptive mechanism,
287 enhancing bacterial fitness to environmental fluctuations. Genome-wide studies of other
288 bacteria have since revealed that these features are not exclusive to *E. coli* (D'Halluin et al.
289 2023; Petroni et al. 2023). Moreover, individual cases of mechanisms of conditional Rho-
290 dependent regulation have been identified across various species (Table 2). In the following
291 section, we briefly present these mechanisms and highlight the most recent discoveries. For a
292 more comprehensive discussion of previously identified mechanisms, readers may refer to
293 other recent reviews (Kriner et al. 2016; Bastet et al. 2018; Roberts 2019; Turnbough 2019;
294 Bossi et al. 2020).

295

296 RDTT IN THE REGULATION OF SPECIFIC GENOMIC LOCI

297 *Mechanisms of conditional Rho activity*

298 Conditional RDTT requires the direct or indirect regulation of Rho activity by an inducer at one
299 or more stages of the RDTT pathway. These stages include Rho loading onto RNA, catalytic
300 isomerization, translocation along RNA, formation of the pre-termination complex with RNAP,
301 and Rho-mediated disassembly of the transcription complex.
Among these possibilities, the regulation of Rho loading onto RNA is perhaps the best documented. In the bacteria where transcription and translation are coupled (Woodgate and Zenkin 2023), actively translating ribosomes protect nascent mRNAs by physically obstructing Rho's access to intragenic *Rut* sites (**Figure 4A**). Factors that uncouple translation from transcription expose intragenic *Rut* sites, leading to RDTT-induced silencing of the corresponding gene(s) and downstream genes of the same operon. This process is called transcriptional polarity (Richardson et al. 1975; Ratner 1976; Wek et al. 1987). Ribosome-targeting antibiotics (Zhu et al. 2019) or the depletion of translation elongation factors (Elgamal et al. 2016) promote ribosome stalling, thereby enhancing Rho-mediated transcriptional polarity. Similarly, conditions that impair or reduce the frequency of translation initiation also induce polarity effects (de Smit et al. 2009).

This principle is widely exploited in conditional silencing mechanisms. For instance, sRNA binding to 5' UTRs (Bossi et al. 2012) or riboswitch-induced restructuring of mRNA leader sequences (Bastet et al. 2017) sequester the ribosome binding site (RBS) within an RNA duplex, preventing ribosome association and promoting Rho-dependent polarity (**Figure 4B,C**; see examples in **Table 2**). In *E. coli*, RNAP pausing near the RBS and start codon in the 5'UTR (Larson et al. 2014), likely facilitates sRNA binding or mRNA leader restructuring before the ribosome associates with the nascent transcript (Chauvier et al. 2019; Chauvier et al. 2021). A similar attenuation system was discovered recently in *Vibrio cholerae* where the sRNA inducer, OppZ, is the last product of its own operon, *oppABCDFZ* (**Table 2**) (Hoyos et al. 2020). OppZ binding to the intergenic region between *oppA* and *oppB* blocks *oppB* translation exposing an intragenic Rho-dependent terminator that silences downstream genes via a negative feedback loop (Hoyos et al. 2020). In all these cases, the combination of transcriptional and translational repression ensures that the inducer triggers robust gene/operon silencing.

Riboswitches and sRNAs can also regulate RDTT more directly by sequestering *Rut* sites within secondary RNA structures or RNA duplexes, preventing Rho from loading onto the nascent RNA (**Figure 4D,E**). One likely case leads to transcription antitermination and the stimulation of *rpoS* expression upon binding of sRNA DsrA, ArcZ, or RprA to the 5'UTR of *rpoS* (Sedlyarova et al. 2016) at a site overlapping with a putative *Rut* site mapped by helicase-SELEX (Delaleau et al. 2022). While this steric interference mechanism has been proposed in several other instances (**Table 2**), the impact of the inducer on Rho's affinity for its RNA target
has rarely been tested. However, a recent study demonstrated that temperature specifically modulates Rho binding to the mRNA leader sequences of genes and operons involved in the E. coli's cold shock response, including cspA, cspB, cspG, and nsrR-rnr-yjfH1 (Delaleau et al. 2024). At 37°C, Rho efficiently binds these mRNA leaders, contributing to the repression of cold shock gene expression (Figure 5, top). Upon cold shock, the mRNA leaders adopt "closed" conformations that are less favorable to Rho binding while exposing RBS sequences, thereby preventing RDTT and optimizing the translation of the cold shock proteins (Figure 5, middle). During cold acclimation, the cold shock proteins provide negative feedback regulation by binding the mRNA leaders once they reach a sufficient concentration, switching them back to conformations more favorable for RDTT (Figure 5, bottom) (Delaleau et al. 2024). This discovery challenges the long-held view that the cold shock response is primarily a posttranscriptional program (Zhang et al. 2018). Instead, it highlights RDTT as an integral component of a complex regulatory circuit that ensures precise control of gene expression (Figure 5) (Delaleau et al. 2024).

Translation of a small upstream open reading frame (uORF) provides another mechanism for conditional RDTT regulation (Table 2). A classic example is the regulation of the tryptophanase operon (tnaCAB) via translation of the tnaC uORF (Figure 6A) (Stewart et al. 1986). At high L-tryptophan concentration, tryptophan invades the ribosomal exit tunnel, inducing ribosome stalling at the tnaC termination codon and preventing cleavage and release of the tnaC leader peptide (Gong et al. 2001; van der Stel et al. 2021). The stalled ribosome then blocks Rho from accessing an adjacent Rut site in the tnaC-tnaA intergenic region, allowing expression of the downstream tnaA (tryptophanase) and tnaB (tryptophan transporter) genes (Stewart et al. 1986; Gish and Yanofsky 1995). A similar uORF-dependent attenuation-by-RDTT mechanism regulates the expression of the ornithine decarboxylase gene (speF) in E. coli and Salmonella in response to ornithine availability (Table 2) (Ben-Zvi et al. 2019; Herrero Del Valle et al. 2020). However, other instances of uORF-dependent attenuation-by-RDTT deviate from this paradigm and instead activate rather than inhibit RDTT in response to the inducer (Table 2). In the cruR-bfrG-bp2921 operon of Bordetella pertussis, high copper ion concentrations trigger ribosome release from the cruR uORF where it otherwise masks an intragenic Rut site (Figure 6B) (Roy et al. 2022). Copper thus exposes the Rut site within cruR, activating RDTT-mediated attenuation of the downstream bfrG gene,
364 which encodes a putative Cu importer, likely contributing to Cu homeostasis. The molecular
365 mechanisms underlying ribosome stalling by the CruR peptide and ribosome release by Cu
366 ions remain unclear but appear to involve two conserved motifs in the peptide: CXXC (middle)
367 and RAPP (C-terminal) (Roy et al. 2022).

368 The mechanisms by which uORFs govern Rho-dependent attenuation in the corA,
369 mgtA, and mgtC genes, involved in magnesium homeostasis in Salmonella (Table 2), have
370 been studied extensively (reviewed in: (Kriner et al. 2016; Bossi et al. 2020)). In these cases,
371 Mg2+ concentration modulates uORF translation efficiency, which in turn influences how the
372 downstream mRNA leader sequence folds during transcription (Kriner and Groisman 2015;
373 Gall et al. 2016; Kriner and Groisman 2017). Under low Mg2+ conditions, ribosome stalling in
374 the uORF promotes the formation of an RDTT-resistant mRNA conformation, enabling the
375 expression of downstream Mg2+ importer genes. Conversely, high Mg2+ levels enhance uORF
376 translation and drive the mRNA leader into an RDTT-prone conformation promoting
377 transcriptional attenuation. For the mgtC attenuator, however, mRNA leader restructuring
378 does not directly regulate Rho access to the Rut site. Instead, it controls a RARE (Rho-
379 antagonizing RNA element) motif in the nascent transcript, which can trap the Rho-RNA
380 complex in an inactive state (Sevostyanova and Groisman 2015). When uORF translation is
381 inefficient, RARE remains single-stranded and inhibits Rho, allowing mgtCBR expression. In
382 contrast, when uORF translation is efficient, RARE is sequestered in a stem-loop structure,
383 preventing its inhibitory effect and facilitating RDTT-mediated attenuation of mgtCBR. Given
384 the growing number of newly identified uORFs (Orr et al. 2020; Adams et al. 2021), particularly
385 in taxa such as Mycobacteria where RDTT also plays a critical role (Botella et al. 2017; Ahmad
386 et al. 2023), these known cases of uORF-dependent attenuation-by-RDTT may represent just
387 the tip of the iceberg.

388 RDTT can also be regulated stochastically, enabling bimodal expression of fitness or
389 virulence factors while allowing bacterial pathogens to evade host immune responses through
390 phase variation and phenotypic heterogeneity (Bidnenko and Bidnenko 2018). Notable
391 examples include invertible DNA elements upstream from fimbriae operons in E. coli (Hinde
392 et al. 2005) and flagella operons in Clostridioides difficile (Warren Norris et al. 2024). These
393 invertible elements encode Rut site(s) in only one orientation, enabling the operons to switch
394 between ON and OFF states depending on the stochastic activity of recombinases (Table 2).
Additionally, stochastic read-through of Rho-dependent terminators provides another mechanism for bimodal gene expression. This has been observed in *Salmonella*, where it regulates virulence genes (Figueroa-Bossi et al. 2022) and in *B. subtilis* where it influences motility and cell differentiation genes (Bidnenko and Bidnenko 2018).

### Protein factors that contribute to regulate RDTT

Besides H-NS, whose functional interaction with Rho was discussed earlier, various proteins regulate RDTT at specific terminators or under particular conditions such as stress responses or the hijacking of bacterial transcription by bacteriophages (Table 3). These protein factors and their roles in RDTT have been studied primarily for *E. coli* and may not be universally conserved across bacteria (Reay et al. 2004; Kalarickal et al. 2010).

In *E. coli*, two major Rho partners are transcription elongation factors NusA and NusG (Table 3). Both factors can bind Rho and RNAP within pre-termination complexes (see above and Figure 2B, D) (Hao et al. 2021a; Said et al. 2021; Molodtsov et al. 2023) and regulate RDTT at specific subsets of terminators in *E. coli* (Peters et al. 2012; Qayyum et al. 2016) and *Salmonella* (Carlomagno and Nappo 2003; Bossi et al. 2019). Notably, NusG enhances RDTT at suboptimal *Rut* sequences with little or no C>G bias (Peters et al. 2012; Delaleau et al. 2022). In contrast, NusA inhibits Rho's ATPase activity (Schmidt and Chamberlin 1984) and typically shifts the termination window downstream while reducing RDTT efficiency in vitro (Lau and Roberts 1985; Chen and Richardson 1987; Sigmund and Morgan 1988; Burns et al. 1998; Qayyum et al. 2016; Simon et al. 2021; Zhu et al. 2022), although a stimulatory effect has been reported in a few cases (Kainz and Gourse 1998; Hao et al. 2021a). Reasons for this discrepancy remain unclear.

NusA and NusG also participate in multipartite antitermination complexes that protect the transcription of ribosomal *rrn* operons (which are untranslated) and lambdoid phage early genes from Rho action (Table 3) (Santangelo and Artsimovitch 2011; Wang and Artsimovitch 2020). These *rrn* and λN antitermination complexes share several core components (NusA/G/B/E) but also include distinct factors (S4 and SuhB versus λN) that collectively transform the TEC into a highly processive, RDTT-resistant form (Figure 7A) (Krupp et al. 2019;
424 Huang et al. 2020). A key step in the assembly of these complexes is the recruitment of NusB/E
425 by a *boxA* sequence in the nascent RNA (**Figure 7B**) (Nodwell and Greenblatt 1993).

426 Interestingly, however, *boxA* recognition by Nus factors can sometimes promote
427 rather than inhibit RDTT. For example, recruitment of a NusB/A complex by an intragenic *boxA*
428 sequence enhances RDTT at a suboptimal terminator in the *hisG* gene of *Salmonella*
429 (Carlomagno and Nappo 2003). Similarly, a *boxA* sequence in the 5' leader of the *suhB* mRNA
430 in *E. coli* and *Salmonella* (**Figure 7B**) cooperates with NusB/E to trigger RDTT within the *suhB*
431 coding sequence (Baniulyte et al. 2017). Formation of a *boxA*-NusB/E complex near the RBS
432 appears to inhibit *suhB* translation through a cryptic, non-steric occlusion mechanism, thereby
433 promoting Rho-dependent polarity. Computational analyses suggest that *boxA*-mediated
434 regulation of RDTT is phylogenetically widespread, extending beyond these few reported
435 cases (Baniulyte et al. 2017).

436 Several antitermination proteins can independently shield the TEC from RDTT using
437 distinct molecular strategies. For instance, the NusG paralog RfaH recognizes 12 nt-long *ops*
438 (operon polarity suppressor) non-template DNA sequences located in the 5'UTRs of a select
439 group of long, xenogeneic operons encoding fitness and virulence factors in *E. coli*, *Salmonella*,
440 and possibly other Enterobacteriaceae (Wang and Artsimovitch 2020). The *ops* sequence
441 induces hyper-pausing of RNAP, which facilitates RfaH recruitment and its structural transition
442 from an auto-inhibited to an active conformation. Once active, RfaH binds both RNAP and the
443 ribosome, reinforcing transcription-translation coupling throughout the coding regions of the
444 *ops*-bearing operons (Molodtsov et al. 2024; Zuber et al. 2024). Lambdoid phages employ Q
445 antitermination proteins, which recognize *qut* (Q utilization) DNA sequences near the
446 promoter to associate with σ-dependent paused TECs. Once loaded, these proteins remain
447 persistently associated with the TEC, ensuring the uninterrupted transcription of the phage
448 late genes (**Table 3**) (Roberts et al. 2008). The Q protein of phage 21 (Q21) alone forms a strong
449 antitermination complex with the TEC (Shi et al. 2019; Yin et al. 2019) whereas the non-
450 homologous Qλ and Q82 proteins require NusA as a cofactor (Yang et al. 1989; Shankar et al.
451 2007; Yin et al. 2022). Nonetheless, structural analyses of Q21 (Shi et al. 2019; Yin et al. 2019)
452 and Qλ (Yin et al. 2022) antitermination complexes suggest a shared mechanism: the Q
453 protein forms a torus-like structure that extends and narrows the RNA exit channel of RNAP.
454 This steric alteration prevents the formation of pause or termination-inducing RNA hairpins
within the channel and likely interferes with Rho-mediated TEC disruption (Yang and Roberts 1989; Yarnell and Roberts 1992; Shankar et al. 2007), though this hypothesis requires further validation. Additionally, the UvrABD complex, which facilitates transcription-coupled repair (TCR) utilizes RNAP as a binding platform (Bharati et al. 2022). Indirect evidence suggests that UvrABD competes with Rho and NusG for the same RNAP binding surface, thereby promoting antitermination during TCR (Martinez et al. 2022).

Several antitermination proteins inhibit Rho directly rather than targeting RNAP (Table 3). One example is the Psu protein from phage P4, which supports late gene transcription in helper phage P2 by preventing Rho-dependent polarity (Linderoth and Calendar 1991). Recent cryo-EM reconstructions of the Rho-Psu complex suggest a mechanism in which Psu dimers bridge and lock two Rho hexamers in tail-to-tail, inactive open-ring conformation (Figure 8A) (Gjorgjevik et al. 2025). Interestingly, the helical pitch of the Rho rings in this complex is higher than in crystal structures of Psu-free, open Rho hexamers (Skordalakes and Berger 2003; Skordalakes et al. 2005), allowing for the oligomeric expansion of Rho rings in a subset of the cryoEM particles (Figure 8A) (Gjorgjevik et al. 2025). This mechanism contrasts with that of the Sm-like protein Rof (aka YaeO), an endogenous Rho inhibitor that protects *E. coli* from oxidative stress (Kawamura et al. 2005) and is required for virulence and host invasion in *Salmonella* (Zhang et al. 2024). Rof molecules bind to the protomer interfaces of the Rho hexamer (one Rof per interface), collectively locking Rho in an inactive, open Rho ring configuration (Figure 8B) (Said et al. 2024; Zhang et al. 2024). Unlike Psu, binding of Rof does not appear to extend the helical pitch of the Rho ring. Additionally, bound Rof molecules partially occlude Rho's PBS, impairing its ability to interact with RNA or NusA/G-containing TECs *in vitro* (Said et al. 2024). Another recently discovered Rho inhibitor is the stress response kinase YihE (Wang et al. 2022). YihE expression is stimulated during the conjugative pilus expression (Cpx) stress response, where it helps protect *E. coli* from oxidative damage and lipid peroxidation. A low-resolution crystal structure of the Rho-YihE complex suggests that a single YihE molecule binds the PBS of the leftmost Rho subunit at the gap of an open Rho hexamer (Wang et al. 2022). However, whether this interaction inactivates Rho by preventing ring closure or by interfering with other steps of the RDTT pathway remains unclear. Notably, the structural data for the Rho-YihE complex have not been deposited by the authors (only
that for a structure of Rho alone under PDB code 7X2R), which raises some doubts about the findings.

Another intriguing endogenous Rho interactor and inhibitor is the RNA chaperone Hfq (Rabhi et al. 2011a), which stabilizes sRNA:mRNA complexes during sRNA-mediated gene silencing (Katsuya-Gaviria et al. 2022). Hfq forms a relatively stable binary complex with Rho ($K_d$ ~40 nM) but effective inhibition of Rho activity requires the simultaneous interaction of both proteins with their RNA target (Rabhi et al. 2011a). While Hfq's antitermination activity has been observed in *E. coli* using a plasmid reporter system carrying the prototypical $\lambda$tR1 terminator (Rabhi et al. 2011a), its physiological antitermination targets remain to be firmly identified. A strong candidate is the Rho-dependent terminator in the 5'UTR of the *rpoS* stress response gene, which is inactivated when sRNA DsrA, ArcZ, or RprA binds to the *rpoS* mRNA leader sequence in an Hfq-dependent manner (Sedlyarova et al. 2016; Nadiras et al. 2018b). Virulence factors subject to opposing regulation by Rho and Hfq are also compelling possibilities (Rabhi et al. 2011a). Finally, a recent study reporting Hfq's association with hundreds of nascent transcripts in *Pseudomonas aeruginosa* further supports its potential role as a Rho interactor (Kambara et al. 2018).

Other Rho interactors have been identified by mass spectrometry (Table 3) (Butland et al. 2005) but their direct interactions with Rho or their effects on RDTT remain unverified. Additionally, Rho has been identified via mass spectrometry in a large c-di-GMP-dependent protein complex upon overexpression and pull-down of the oxygen sensors DosC and DosP (Tuckerman et al. 2011). Intriguingly, the complex also contained the RNA degradosome components PNPase, enolase and RNAse E, providing the first direct evidence of a condition-dependent link between Rho and the RNA degradosome in *E. coli*. In contrast, in *Rhodobacter capsulatus*, Rho appears to associate more stably with the RNA degradosome (Jager et al. 2001), though its proportion within the complex fluctuates depending on oxygen availability during growth (Jager et al. 2004).

### Accessory nucleic acid sequences that regulate RDTT

Accessory DNA/RNA sequences or motifs play a crucial role in regulating RDTT, ensuring that this regulation remains confined to specific regions of the bacterial or phage genome. For
example, the presence of a *cut* or *ops* sequence in the DNA template is essential for recruiting
the antitermination factors Q and RfaH, respectively (see section above and **Table 3**).
Accessory RNA sequences influence co-transcriptional RNA folding and modulate Rho access
to *Rut* sites (**Figure 4B-D**) or, in the case of the RARE sequence, drive the catalytic
isomerization of the Rho-RNA complex (Sevostyanova and Groisman 2015). *BoxA* sequences
within the nascent transcript recruit Nus factors either stimulating or inhibiting RDTT,
depending on the genomic context (see section above). Similarly, a *boxB* sequence aids in
assembling the λN antitermination complex by recruiting the λN protein (Vieu and Rahmouni
2004) whereas comparable promoter-proximal *boxB* sequences play a much less significant
role in *rrn* antitermination in *E. coli* (Warner et al. 2023).

Regulation of RDTT can also occur through the binding of accessory RNA sequences to
RNAP. For example, promoter-proximal sequences known as *put* sites, encoded by the phage
HK022 genome interact with RNAP to transform the TEC into a termination-resistant form
(King and Weisberg 2003). Cryo-EM reconstructions of *put*RNA-TEC complexes suggest that
this interaction prevents "swiveling", a conformational shift in RNAP that facilitates
transcriptional pausing (Hwang et al. 2022). However, whether this anti-pausing activity fully
accounts for *put*RNA-mediated inhibition of RDTT (King and Weisberg 2003) remains unclear.
More recently, natural RNAP-binding aptamers (RAPs) have been identified in *E. coli* through
genomic SELEX, including a subset of inhibitory RAPs (iRAPs) that promote RDTT (Sedlyarova
et al. 2017). The authors have proposed that iRAP binding to RNAP hampers the progression
of the trailing ribosome, uncoupling translation from transcription thereby eliciting Rho-
dependent polarity (Sedlyarova et al. 2017). However, the subsequent discovery of iRAPs in
non-coding, antisense regions (Magan et al. 2019) suggests that they may trigger RDTT
through a different yet-undetermined mechanism. To date, the physiological roles of RAPs
and iRAPs, as well as their modes of engagement—whether constitutive or conditional- remain
largely unknown.

541

542 TWEAKING RHO-DEPENDENT ATTENUATION FOR SYNTHETIC BIOLOGY

Despite the key role of RDTT in the regulation of bacterial gene expression, its application in
synthetic biology has been limited. However, certain *rho* mutants evolved under specific stress
conditions, such as heat stress (Kishimoto et al. 2010; Deatherage et al. 2017), ethanol exposure (Freddolino et al. 2012; Haft et al. 2014) or carbon source limitation (Conrad et al. 2009; Worthan et al. 2024), may hold potential for bioproduction processes or other biotechnological applications. Notably the rho R109H mutation transforms Rho into a pH-dependent enzyme within the physiological pH range (Worthan et al. 2024), offering a unique opportunity for pH-responsive genetic control.

The tryptophan-responsive tnaC attenuator of the tnaCAB operon (Figure 6A) is a rare example of Rho-dependent regulatory element integrated into synthetic biological circuits. Its dynamic response (Wang et al. 2020a) and orthogonal control can be fine-tuned, with the latter enhanced in nusB-depleted strains, where cross-regulation by other amino acids and natural metabolites is reduced (Sherman et al. 2021). The tnaC attenuator has been used to detect and enhance L-tryptophan production and to optimize metabolic fluxes where L-tryptophan serves as an intermediate (Fang et al. 2016; Liu et al. 2017; Chen et al. 2019; Wang et al. 2020b; Gwon et al. 2021). For example, in a synthetic E. coli co-culture platform designed for tryptamine bio-production, the tetracycline resistance gene tetA was placed under tnaC control (Wang et al. 2020b). The tnaC-tetA sensor selectively promotes the growth of the subpopulation of cells that produce high amounts of L-tryptophan, the tryptamine precursor, in the presence of tetracycline. In another application, an amber UAG stop codon has been introduced into tnaC to tailor its sensing capability to an unnatural amino acid and its corresponding, synthetic aminoacyl-tRNA synthetase (Liu et al. 2011). Similarly, engineering the 5'UTR upstream from tnaC enables the attenuator to respond to RNA duplex formation with a complementary RNA, modulating RBS accessibility and downstream transcriptional output (Mihailovic et al. 2018).

The growing repertoire of natural Rho-dependent attenuators (Table 2) provides new opportunities for developing synthetic regulatory devices responsive to diverse signal inputs. Another promising avenue is the *de novo* engineering of synthetic RDTT attenuators. Certain synthetic aptamers and riboswitches contain C-rich sequences that can be modified into *Rut*-like switches enabling ligand-dependent Rho recruitment (Boudvillain et al., unpublished results) in a way reminiscent of the CsrA-responsive *pgaA* attenuator (Table 2) (Figueroa-Bossi et al. 2014). Additionally, switchable *Rut* motifs can be evolved *de novo* using a modified Helicase-SELEX approach. By applying alternating rounds of selection and counter-
576 selection -depending on the presence and absence of a target ligand- this strategy yields RNA
577 sequences that regulate Rho helicase activity in a ligand-dependent manner (Boudvillain et al.
578 2021). As a proof or principle, this approach has been successfully used to evolve synthetic
579 serotonin-responsive *Rut* sequences capable of modulating Rho activity both *in vitro* and *in
vivo* (Delaleau et al. 2022).
580

581

582 CONCLUSIONS AND OPEN QUESTIONS

583 Paraphrasing the title of a previous review of RDTT (Banerjee et al. 2006), and despite
584 significant advances since its publication, many questions remain regarding the mechanisms
585 and physiological roles of Rho. While both *in vitro* (Figure 2) (Hao et al. 2021a; Said et al. 2021;
586 Song et al. 2022; Molodtsov et al. 2023; Murayama et al. 2023; Song et al. 2023) and *in vivo
587* (Shashni et al. 2014; Bossi et al. 2019) studies support the existence of parallel RDTT pathways,
588 their specific mechanisms and biological relevance require further investigation. Is there a
589 pathway dominant under all conditions, or do different environmental or cellular contexts
590 favor distinct RDTT mechanisms? Is the RDTT pathway for a given terminator permanently
591 defined, or can it change dynamically in response to variable or inducible factor(s)? More
592 broadly, are parallel RDTT pathways a common feature across Rho-bearing species or do they
593 represent exceptions rather than the rule?

594 RDTT is traditionally regarded as the primary mechanism for monitoring translation-
595 transcription coupling. However, recent findings suggest that this coupling may be less
596 prevalent than previously assumed (Woodgate and Zenkin 2023) or that mRNA can be
597 significantly "looped out" between RNAP and the ribosome, making it accessible to Rho
598 (Qureshi and Duss 2024). If this is the case, how is spurious RDTT prevented? Is Rho activity
599 spatially confined to specific subcellular or nucleoid regions? Could some RAPs function
600 similarly to *put* sequences, shielding RNAP from Rho-dependent termination? Furthermore,
601 how is the action of numerous modulators of RDTT (Table 3) coordinated to ensure a precise
602 spatiotemporal regulation of Rho?

603 Recent findings suggest that *E. coli* Rho can undergo reversible inactivation through
604 the formation of filamentous Rho hyper-oligomers, particularly under stress conditions that
605 disrupt translation-transcription coupling such as exposure to translation inhibitors (Wang et
606 al. 2025). However, do the Rho-containing cellular aggregates observed in presence of these
607 inhibitors truly represent inactive Rho filaments, given that similar inhibitory conditions have
608 been shown to stimulate rather than suppress RDTT (Zhu et al. 2019)? Alternatively, could
609 oligomeric Rho filaments serve other functions, such as facilitating the subcellular clustering
610 of Rho activity, a phenomenon recently observed for other filament-forming transcription
611 factors (Mance et al. 2024)?

612 Many Rho-regulated processes have now been identified and recognized to be critical
613 for bacterial adaptation and fitness (Table 2). However, our understanding remains heavily
614 biased towards the E. coli model. While RDTT continues to be intensely studied in this
615 organism, research is expanding to uncover Rho's functions and mechanisms across the
616 bacterial kingdom. Exciting discoveries suggest that RDTT can take diverse forms in
617 phylogenetically distant bacteria, often influenced by taxon-specific variations in the Rho
618 polypeptide (D'Heygere et al. 2015; Yuan and Hochschild 2017; Simon et al. 2021; Krypotou et
619 al. 2023; Warren Norris et al. 2024). These variations extend to the regulation of Rho activity,
620 which can occur through mechanisms such as liquid-liquid phase separation (Krypotou et al.
621 2023), β-amyloid aggregation (Yuan and Hochschild 2017), interaction with endogenous
622 inhibitors (Zhang et al. 2024), RNA oxidation (Han et al. 2022), or growth-dependent
623 fluctuations in intracellular Rho levels (Italiani et al. 2002; Silva et al. 2019; Bidnenko et al.
624 2024). What other regulatory surprises will emerge as RDTT is studied across the bacteria
625 diversity? How does Rho function in extremophiles? Beyond bacterial fitness and adaptation,
626 what roles does RDTT play in bacterial development programs or microbial community
627 interactions?

628 Sophisticated experimental tools will be essential to tackle these timely questions.
629 Genome-wide mapping of putative *Rut* sites is now feasible with Helicase-SELEX (Delaleau et
630 al. 2022), which has already led to new discoveries such as Rho's involvement in the cold shock
631 response of *E. coli* and *Salmonella* (Delaleau et al. 2024). Cryo-EM will likely continue to
632 provide valuable structural insights into RDTT, potentially extending to cellular ultrastructure
633 imaging through cryo-electron tomography in the near future. Additionally, super-resolution
634 fluorescence microscopy approaches hold great promise for tracking single Rho particles—and
635 their potential complexes with RNAP or Nus factors— in real time, assessing their subcellular
636 distributions and velocities, and probing RDTT mechanisms in live cells. While super-resolution
637 live-cell imaging of RNAP and NusG has been achieved (El Sayyed et al. 2024), extending this
638 approach to Rho will require the development of an effective non-disruptive method for
639 fluorescently labeling intracellular Rho molecules.

640

641 ACKNOWLEDGMENTS

642 This work was undertaken in partial fulfilment of the requirements for the doctoral degree of
643 T.D.D. and was supported by grants from the French Agence Nationale de la Recherche to
644 M.B. (ANR-19-CE44-0009-01, and ANR-22-CE44-0017). J.C.B. is a recipient of a PhD fellowship
645 from the Université d'Orléans. L.B. and N.F-B. acknowledge the support of the French National
646 Research Council (CNRS).
<table><caption>Table 1: Comparison of the catch-up and stand-by models (not all relevant references are cited, for brevity)</caption><thead><tr><th></th><th>Catch-up model</th><th>Stand-by model</th></tr></thead><tbody><tr><th>Pros</th><td><ul><li><strong>Historical model</strong> supported by extensive biochemical evidence (Jin et al. 1992; Steinmetz and Platt 1994; Richardson and Richardson 1996; Xu et al. 2002; Park and Roberts 2006; Schwartz et al. 2009; Kalyani et al. 2011; Rabhi et al. 2011b), single-molecule manipulation studies (Koslover et al. 2012; Gocheva et al. 2015), and structural analyses (Skordalakes and Berger 2003; Thomsen and Berger 2009; Lawson et al. 2018; Molodtsov et al. 2023)</li><li><strong>Consistent with in vitro enzymatic properties of Rho</strong>, including RNA-stimulated ATP hydrolysis (Lowery-Goldhammer and Richardson 1974) and ATP-dependent, 5'→3' directional translocation along RNA leading to disruption of RNA-DNA duplexes (Brennan et al. 1987; Walmacq et al. 2004; Walmacq et al. 2006) or biotin-streptavidin complexes (Schwartz et al. 2007).</li><li><strong>Genetic evidence for kinetic coupling</strong>: Slow RNAP mutants suppress RDTT defects caused by rho and nusG mutations or by inhibitors such as Psu and Bycyclomycin (Shashni et al. 2012), supporting the model of kinetic coupling between Rho and RNAP translocations (Jin et al. 1992).</li><li><strong>CryoEM reconstructions</strong> of pre-termination complexes are compatible with this model (Molodtsov et al. 2023; Murayama et al. 2023); the nucleic-acid substrates and assembly strategy used in these studies have been validated through biochemical RDTT assays.</li></ul></td><td><ul><li><strong>Recently proposed model based on new evidence</strong> from a biochemical study (Epshtein et al. 2010) and cryoEM reconstructions of pre-termination complexes (Hao et al. 2021a; Said et al. 2021) containing NusA and NusG.</li><li><strong>Colocalization of Rho and RNAP</strong> along the <em>E. coli</em> genome revealed by ChIP-seq experiments (Mooney et al. 2009)</li><li><strong>Paralell RDTT pathway(s)</strong> supported by single-molecule fluorescence studies (Song et al. 2022; Song et al. 2023)</li></ul></td></tr><tr><th>Cons</th><td><ul><li><strong>Cofactor missing in cryoEM structures</strong>: <em>E. coli</em> 'catch-up' pre-termination complexes characterized by CryoEM (Molodtsov et al. 2023) did not include NusA.</li></ul></td><td><ul><li><strong>Biochemical validation missing</strong>: The reconstituted "stand-by" complexes on DNA bubble substrates (Hao et al. 2021a; Said et al. 2021) have not been shown to support RDTT.</li><li><strong>Inactive Rho</strong>: In the 'stand-by' complexes, the Rho hexamer adopts an open, catalytically inactive conformation and binds RNAP in an orientation incompatible with its translocase activity (Figure 2D), suggesting the captured state may be off-pathway or represent an early, pre-productive intermediate.</li><li><strong>Off-pathway evidence</strong>: Biochemical characterization of analogous complexes from <em>T. thermophilus</em> support an off-pathway interpretation (Murayama et al. 2023).</li><li><strong>Uncertain RNA trajectory</strong>: The proposed inverted RNA contact sequence (SBS→PBS) (Hao et al. 2021a) lacks strong experimental validation and may instead reflect off-pathway RNA sequestration by NusA.</li><li><strong>Steric hindrance at the PBS</strong>: Limited PBS accessibility in the stand-by' complexes (Figure 2D) raises questions about <em>Rut</em> site recognition, especially in cases of conditional RDTT requiring RNA structural remodeling.</li></ul></td></tr></tbody></table>
<table><caption>Table 2: genes and operons regulated by Rho in a conditional manner</caption><thead><tr><th>Type</th><th>Regulated locus</th><th>Species</th><th>Metabolic pathway</th><th>Inducer(s)</th><th>Inducer effect</th><th>Mechanism</th><th>Year</th></tr></thead><tbody><tr><td rowspan="10">sRNA-dependent</td><td>chiPQ</td><td>Salmonella</td><td>Carbon utilization</td><td>sRNA ChiX</td><td>Termination</td><td>Inhibition of translation upon ChiX pairing to <i>chiP</i> leader unmasks intragenic terminator in <i>chiP</i> CDS (Bossi et al. 2012)</td><td>2012</td></tr><tr><td>galKM</td><td>E. coli</td><td>Galactose catabolism</td><td>sRNA Spot 42 (aka spf)</td><td>Termination</td><td>sRNA binding inhibits galK translation and activates RDTT; exact mechanism still unclear but likely involves translation uncoupling (Jeon et al. 2022)</td><td>2015</td></tr><tr><td>oppABCDFZ</td><td>Vibrio cholerae</td><td>Oligopeptide uptake</td><td>sRNA OppZ</td><td>Termination</td><td>OppZ binding to the <i>oppAB</i> intergenic region inhibits <i>oppB</i> translation and unmasks an intragenic terminator in <i>oppB</i> (Hoyos et al. 2020)</td><td>2020</td></tr><tr><td>rpoS</td><td>E. coli</td><td>General stress response</td><td>sRNAs DsrA, ArcZ, rprA</td><td>Antitermination</td><td>rpoS transcription is stimulated by sRNA binding to the <i>rpoS</i> leader stimulates (Sedlyarova et al. 2016; Nadiras et al. 2018b) at a site overlapping with a putative <i>Rut</i> site (Delaleau et al. 2022)</td><td>2016</td></tr><tr><td>astE</td><td>E. coli</td><td>Arginine catabolism</td><td>sRNAs DsrA, ArcZ</td><td>Antitermination</td><td>Putative ArcZ and DsrA binding sites in <i>astE</i> leader (Sedlyarova et al. 2016); Exact mechanism undetermined</td><td>2016</td></tr><tr><td>fadE</td><td>E. coli</td><td>Carbon source utilization</td><td>sRNAs DsrA, ArcZ</td><td>Antitermination</td><td>Putative ArcZ binding site in <i>fadE</i> leader (Sedlyarova et al. 2016); Exact mechanism undetermined</td><td>2016</td></tr><tr><td>yqeC</td><td>E. coli</td><td>Unknown</td><td>sRNAs DsrA, ArcZ</td><td>Antitermination</td><td>Putative ArcZ and DsrA binding sites in <i>yqeC</i> leader (Sedlyarova et al. 2016); Exact mechanism undetermined</td><td>2016</td></tr><tr><td>rho</td><td>Salmonella</td><td>RDTT regulation</td><td>sRNA SraL</td><td>Antitermination</td><td>SraL binding site overlaps with a putative <i>Rut</i> site suggesting a binding competition mechanism (Silva et al. 2019)</td><td>2019</td></tr><tr><td>CRISP-2</td><td>Pseudomonas aeruginosa</td><td>Adaptive immunity</td><td>sRNA PhrS</td><td>Antitermination</td><td>Exact mechanism undetermined (Lin et al. 2019)</td><td>2019</td></tr><tr><td>trpE</td><td>E. coli</td><td>Tryptophan biosynthesis</td><td>sRNA RydC</td><td>Unknown</td><td>Predicted from H-SELEX screening, putative <i>Rut</i> site in <i>trpE</i> CDS (Delaleau et al. 2022); awaits experimental validation</td><td>2022</td></tr><tr><td rowspan="4">Riboswitch</td><td>ribB</td><td>E. coli</td><td>Riboflavin biosynthesis</td><td>FMN (flavin mononucleotide)</td><td>Termination</td><td>Restructuring of <i>ribB</i> leader by FMN likely unmasks <i>Rut</i> site otherwise sequestered in alternative structure (Hollands et al. 2012; Bastet et al. 2017)</td><td>2012</td></tr><tr><td>btuB</td><td>E. coli</td><td>Cobalamin (Vitamin B12) transport</td><td>Adenosylcobalamin</td><td>Termination</td><td>Exact mechanism undetermined but RDTT requires upstream section of the <i>btuB</i> CDS (Nadiras et al. 2018a; Bastet et al. 2024); <i>btuB</i> is also regulated by sRNAs OmrA/B in a Rho-independent manner (Bastet et al. 2024)</td><td>2024</td></tr><tr><td>lysC</td><td>E. coli</td><td>Lysine biosynthesis</td><td>Lysine</td><td>Termination</td><td>Restructuring of <i>lysC</i> leader by Lysine unmasks <i>Rut</i> site otherwise sequestered in alternative structure (Sedlyarova et al. 2016; Bastet et al. 2017; Ghosh et al. 2024)</td><td>2017</td></tr><tr><td>thiBPQ</td><td>E. coli</td><td>Thiamine uptake</td><td>TPP (thiamin pyrophosphate)</td><td>Termination</td><td>Exact mechanism undetermined; putative <i>Rut</i> site in <i>thiB</i> leader (Bastet et al. 2017; Delaleau et al. 2022)</td><td>2017</td></tr></tbody></table>
<table><tbody><tr><td></td><td>thiCEFSGH</td><td>E. coli</td><td>Vitamin B1 biosynthesis</td><td>TPP (thiamin pyrophosphate)</td><td>Termination</td><td>Restructuring of thiC leader by FMN likely unmasks <i>Rut</i> site otherwise sequestered in alternative structure (Bastet et al. 2017; Chauvier et al. 2017)</td><td>2017</td></tr><tr><td></td><td>thiMD</td><td>E. coli</td><td>Vitamin B1 biosynthesis</td><td>TPP (thiamin pyrophosphate)</td><td>Termination</td><td>Restructuring of thiM leader by TPP inhibits translation and unmasks intragenic <i>Rut</i> site in thiM (Bastet et al. 2017; Delaleau et al. 2022)</td><td>2017</td></tr><tr><td></td><td>ribM</td><td>Corynebacterium glutamicum</td><td>Riboflavin transport</td><td>FMN (flavin mononucleotide)</td><td>Termination</td><td>Exact mechanism undetermined but likely involves RDTT and mRNA decay (Takemoto et al. 2015)</td><td>2015</td></tr><tr><td></td><td>mntP</td><td>E. coli</td><td>Manganese homeostasis</td><td>Mn<sup>2+</sup> ion</td><td>Antitermination</td><td>Binding of Mn<sup>2+</sup> to the riboswitch restores translation-transcription coupling and expression of the MntP efflux pump (Prakash et al. 2024). Evidence for <i>Rut</i> site in upstream part of mntP CDS (Delaleau et al. 2022).</td><td>2024</td></tr><tr><td rowspan="6">Riboswitch-like</td><td>trpEDCFBA</td><td>B. subtilis</td><td>Tryptophan biosynthesis</td><td>TRAP protein</td><td>Termination</td><td>TRAP binding to the trpE leader inhibits translation and unmasks intragenic terminator(s) (Yakhnin et al. 2001)</td><td>2001</td></tr><tr><td>pgaABCD</td><td>E. coli</td><td>Biofilm formation</td><td>CsrA protein</td><td>Termination</td><td>Restructuring of pgaA leader by CsrA unmasks <i>Rut</i> site otherwise sequestered in a RNA hairpin (Figueroa-Bossi et al. 2014)</td><td>2014</td></tr><tr><td>tkt-gap-lpp0152-lpp0151</td><td>Legionella pneumophila</td><td>Pyruvate metabolism</td><td>CsrA protein</td><td>Antitermination</td><td>CsrA binding to an AGGA motif in the gap CDS inhibits RDTT in vitro, presumably upon RNA restructuring (Sahr et al. 2017). This effect correlates with a lower amount of gap RNA in csaR- vs csaR+ strains, although Rho involvement in vivo was not tested formally (Sahr et al. 2017).</td><td>2017</td></tr><tr><td>ABUW_1645, ABUW_1959, ABUW_2818</td><td>Acinetobacter baumannii</td><td>Phase variation & virulence</td><td>CsrA protein</td><td>Antitermination</td><td>CsrA inhibits RDTT by binding to a GGA-rich region of the ABUW_1645 mRNA leader, which also contains a <i>Rut</i> site. The ATPase activity of Rho is inhibited in vitro by CsrA binding to ABUW_1645 mRNA leader (Singh et al. 2025).</td><td>2025</td></tr><tr><td>cspA/B/G</td><td>E. coli (cspA/B/G) and Salmonella (CspA)</td><td>Cold stress response</td><td>Cold temperature<br>CspA protein</td><td>Antitermination<br>Termination</td><td>Restructuring of cspA/B/G leader at low temperature blocks access to <i>Rut</i> site; CspA binding to the mRNA leader unmasks the <i>Rut</i> site (Delaleau et al. 2024)</td><td>2024</td></tr><tr><td>nsrR-rnr-yjfH1</td><td>E. coli</td><td>Cold stress response</td><td>Cold temperature<br>CspA protein</td><td>Antitermination<br>Termination</td><td>Restructuring of nsrR-rnr-yjfH1 leader at low temperature blocks access to <i>Rut</i> site; CspA binding to the mRNA leader unmasks the <i>Rut</i> site (Delaleau et al. 2024)</td><td>2024</td></tr><tr><td></td><td>cspl</td><td>E. coli</td><td>Cold stress response</td><td>Cold temperature</td><td>Antitermination</td><td>Exact mechanism undetermined; Putative <i>Rut</i> site upstream from promoter (Delaleau et al. 2024)</td><td>2024</td></tr><tr><td></td><td>DeaD</td><td>E. coli</td><td>Cold stress response</td><td>Cold temperature</td><td>Antitermination</td><td>Exact mechanism undetermined (Delaleau et al. 2024); <i>Rut</i> site identified in 5'UTR (Ojha and Jain 2020)</td><td>2024</td></tr><tr><td>uORF-mediated</td><td>tnaCAB</td><td>E. coli</td><td>Tryptophan catabolism</td><td>Tryptophan</td><td>Antitermination</td><td>Tryptophan binding to the ribosome prevents its release from the last codon of the tnaC uORF, thereby shielding an adjacent <i>Rut</i> site (van der Stel et al. 2021)</td><td>1985</td></tr></tbody></table>
<table><tr><td></td><td>mgtA</td><td>Salmonella</td><td>Magnesium homeostasis</td><td>Mg<sup>2+</sup> ion</td><td>Termination</td><td>Magnesium stimulates translation of uORF mgtL, which promotes restructuring of the mgtA leader, unmasks the Rut site, and stimulates RNAP pausing (Hollands et al. 2012; Hollands et al. 2014; Gall et al. 2016); May not be conserved in E. coli (Bastet et al. 2017)</td><td>2012</td></tr><tr><td></td><td>corA</td><td>Salmonella and E. coli</td><td>Magnesium homeostasis</td><td>Mg<sup>2+</sup> ion</td><td>Termination</td><td>Magnesium stimulates translation of corL uORF and restructuring of corA leader, which unmasks Rut site and stimulates RNAP pausing (Kriner and Groisman 2015; Kriner and Groisman 2017; Vezina Bedard et al. 2024); putative Rut site in the corA leader of E. coli (Delaleau et al. 2022; Vezina Bedard et al. 2024)</td><td>2015</td></tr><tr><td></td><td>mgtCBR</td><td>Salmonella</td><td>Magnesium homeostasis</td><td>Mg<sup>2+</sup> ion</td><td>Termination</td><td>Magnesium stimulates translation of uORF mgtM, which promotes restructuring of the mgtC leader and sequestering of the RARE antitermination sequence that otherwise inhibits the Rho-Rut complex (Sevostyanova and Groisman 2015)</td><td>2015</td></tr><tr><td></td><td>speF</td><td>E. coli and Salmonella</td><td>Polyamine homeostasis</td><td>Ornithine</td><td>Antitermination</td><td>During translation of the uORF speFL, ornithine invades the ribosomal exit channel and stalls the ribosome on the termination codon, which masks a Rut site and allows expression of the ornithine decarboxylase SpeF (Ben-Zvi et al. 2019; Herrero Del Valle et al. 2020)</td><td>2020</td></tr><tr><td></td><td>cruR-bfrG-bp2921</td><td>Bordetella pertussis</td><td>Copper homeostasis</td><td>Copper</td><td>Termination</td><td>Copper binding to the CruR leader peptide triggers release of stalled ribosome otherwise shielding an intragenic Rut site in the uORF (Roy et al. 2022)</td><td>2022</td></tr><tr><td></td><td>hisGDCBHAFI</td><td>E. coli and Salmonella</td><td>Histidine biosynthesis</td><td>Charged tRNA<sup>His</sup></td><td>Termination</td><td>Shortage of charged tRNA<sup>His</sup> slows translation of His-rich hisL uORF, inducing an antitermination conformation of the mRNA leader. Attenuation may combine intrinsic termination (Frunzio et al. 1981) and RDTT (Lawther and Hatfield 1978). Intragenic RDTT sites have been identified in the hisGDC cistrons of Salmonella (Alifano et al. 1991; Rivellini et al. 1991)</td><td>1978</td></tr><tr><td rowspan="3">uORF-like</td><td>tufB</td><td>Salmonella</td><td>Translation regulation</td><td>Tu protein</td><td>Termination</td><td>Tu promotes rapid displacement of the elongating ribosome and folding of the tufB leader that unmasks a Rut site. Ribosome traffic jam in absence of Tu favors formation of the antitermination conformation (Brandis et al. 2016)</td><td>2016</td></tr><tr><td>mdtJI</td><td>E. coli</td><td>Polyamine homeostasis</td><td>Spermidine</td><td>Antitermination</td><td>Expression of mdtJI is regulated by Rho, the translation of an uORF, and spermidine but the exact mechanism is undetermined (Adams et al. 2021)</td><td>2021</td></tr><tr><td>topAI (yjhX)</td><td>E. coli</td><td>Type II toxin-antitoxin system</td><td>Antibiotics</td><td>Termination</td><td>Antibiotic-induced ribosome stalling unmasks intragenic terminator within topAI. Regulation also depends on upstream uORF toiL (Baniulyte and Wade 2025). Putative Rut sites in both topAI and toiL (Delaleau et al. 2022). Exact mechanism remains to be determined.</td><td>2024</td></tr></table>
<table><tr><td>iRAP</td><td>nadD</td><td>E. coli</td><td>NAD+/NADP+ biosynthesis</td><td>Unknown, related to growth phase or oxidative stress</td><td>Termination</td><td>iRAP sequence in CDS promotes ribosome stalling and unmasking of intragenic terminator (Sedlyarova et al. 2017); mechanism of conditional Rho activation not determined.</td><td>2017</td></tr><tr><td rowspan="2">Invertible DNA</td><td>fimAICDFGH operon</td><td>E. coli</td><td>Phase variation & virulence</td><td>DNA inversion by FimB & FimE recombinases (expression is growth- & condition-dependent)</td><td>Termination</td><td>The DNA switch in the OFF orientation enables RDTT (Hinde et al. 2005)</td><td>2002</td></tr><tr><td>flgB operon</td><td>Clostridioides difficile</td><td>Phase variation & virulence</td><td>DNA inversion mediated by random activation of recombination?</td><td>Termination</td><td>The DNA switch in the OFF orientation enables RDTT; Exact mechanism undetermined (Warren Norris et al. 2024)</td><td>2020</td></tr></table>
<table><caption>Table 3: Proteins that regulate RDTT</caption><thead><tr><th>Category</th><th>Protein</th><th>Main features and role(s) in RDTT</th></tr></thead><tbody><tr><td rowspan="6">Transcription elongation factors</td><td>NusG</td><td>NTD binds RNAP while CTD binds ribosome or Rho in mutually exclusive manner. Increases RNAP processivity except at RDTT terminators (Ray-Soni et al. 2016; Yakhnin et al. 2023). Stimulates RDTT in untranslated regions, notably at suboptimal <i>Rut</i> sites, by increasing the rate of catalytic activation of the Rho-RNA complex (Lawson et al. 2018). Constituent of λ and <i>rrn</i> antitermination complexes (Santangelo and Artsimovitch 2011; Huang et al. 2020). Universally conserved, NusG has idiosyncratic features in some bacteria, such as <i>M. tuberculosis</i> (Kalarickal et al. 2010) or <i>Thermus thermophilus</i> (Reay et al. 2004) where it does not bind Rho.</td></tr><tr><td>NusA</td><td>NTD binds the RNA exit channel of RNAP and nascent RNA structures within it to enhance RNAP pausing or intrinsic termination (Ray-Soni et al. 2016). CTD binds Rho (Hao et al. 2021a; Said et al. 2021). Role in RDTT is debated, likely context-dependent (Burns et al. 1998; Carlomagno and Nappo 2003; Ray-Soni et al. 2016). Proposed to inhibit RDTT by binding RNA at sites overlapping with <i>Rut</i> sites (Qayyum et al. 2016) or to stimulate RDTT (Cardinale et al. 2008) by slowing down RNAP (Ray-Soni et al. 2016) or stabilizing the 'Stand-by' pre-termination complex (Fig. 2D) (Hao et al. 2021a; Said et al. 2021). Constituent of λ and <i>rrn</i> antitermination complexes (Santangelo and Artsimovitch 2011; Huang et al. 2020). A <i>nusA</i> deletion mutant is viable provided that Rho activity is reduced (Zheng and Friedman 1994).</td></tr><tr><td>RfaH</td><td>NusG paralog and transcription antitermination factor operating at a subset of horizontally acquired operons in <i>Enterobacteriaceae</i>. Prevents Rho-dependent polarity in specific operons containing an <i>ops</i> sequence by excluding NusG, altering the RNAP conformation, and facilitating ribosome recruitment (Wang and Artsimovitch 2020; Molodtsov et al. 2024; Zuber et al. 2024).</td></tr><tr><td>NusB</td><td>Constituent of λ and <i>rrn</i> antitermination complexes (Santangelo and Artsimovitch 2011; Huang et al. 2020). Enhances RDTT at some terminators in a <i>boxA</i>-dependent manner (Carlomagno and Nappo 2003; Baniulyte et al. 2017).</td></tr><tr><td>GreA</td><td>Prevents formation of long-lived backtracked TECs by stimulating cleavage of 2-3 nt-long extruded RNA 3'end. Cooperates with Rho to remove arrested TECs and avoid genotoxic collisions with the replisome (Dutta et al. 2011). Putative Rho interaction detected by mass spectrometry (Butland et al. 2005).</td></tr><tr><td>GreB</td><td>Reactivates backtracked TECs by stimulating cleavage of up to 18 nt-long extruded RNA 3'end. Cooperates with Rho to remove arrested TECs and avoid genotoxic collisions with the replisome (Dutta et al. 2011). Stimulates RDTT with arrested, backtracked TECs <i>in vitro</i> (Dutta et al. 2008).</td></tr><tr><td rowspan="6">Ribosomal proteins</td><td>L3 (RplC)</td><td rowspan="6">50S subunit constituents. Putative, individual interactions with Rho detected by mass spectrometry (Butland et al. 2005). Effects on RDTT are unknown.</td></tr><tr><td>L4 (RplD)</td></tr><tr><td>L7 (RplL)</td></tr><tr><td>S2 (RpsB)</td></tr><tr><td>S10 (NusE)</td></tr><tr><td>S4</td></tr><tr><td>Inositol monophosphatase</td><td>SuhB</td><td>30S subunit constituent. Binds NusG (Burmann et al. 2010) and regulates RDTT by promoting transcription-translation coupling through its interaction with NusG (Woodgate and Zenkin 2023). Constituent of λ and <i>rrn</i> antitermination complexes (Santangelo and Artsimovitch 2011; Huang et al. 2020).</td></tr><tr><td></td><td></td><td>30S subunit constituent. Constituent of the <i>rrn</i> antitermination complex (Santangelo and Artsimovitch 2011; Huang et al. 2020).</td></tr><tr><td></td><td></td><td>May contribute to rRNA and ribosome maturation independently from its monophosphatase activity (Singh et al. 2016). Constituent of the <i>rrn</i> antitermination complex (Singh et al. 2016; Huang et al. 2020). May be more important for ribosome maturation than for antitermination <i>per se</i> (Singh et al. 2016).</td></tr></tbody></table>
<table><tbody><tr><td rowspan="5">RNA binding and processing proteins</td><td>Hfq</td><td>Ring-shaped RNA chaperone, involved in gene silencing where it stabilizes sRNA-mRNA duplexes (Katsuya-Gaviria et al. 2022). Hfq can stably bind Rho and trigger antitermination in vitro and in E. coli cells with a model system (Rabhi et al. 2011a). Interaction with RNA is required for antitermination. Physiological targets of Hfq-dependent antitermination are unidentified but may include the general stress regulator rpoS gene (Sedlyarova et al. 2016).</td></tr><tr><td>CsrA</td><td>Regulates RDTT in the 5'UTR of the pgaABCD operon of E. coli by preventing formation of a hairpin like-structure blocking the Rut site (Figueroa-Bossi et al. 2014).</td></tr><tr><td>CspA</td><td>Major cold shock protein. Regulates RDTT-dependent expression of cold shock proteins (Figure 5) (Delaleau et al. 2024).</td></tr><tr><td>Rnase E</td><td>Major component of RNA degradosome. Rho associates with RNase E in Rhodobacter capsulatus (Jager et al. 2001; Jager et al. 2004) and under specific growth conditions in E. coli (Tuckerman et al. 2011).</td></tr><tr><td>YgjH</td><td>Putative tRNA binding protein. Reduces RDTT in in vivo reporter screen based on sgrS terminator (Morita et al. 2022).</td></tr><tr><td rowspan="2">DNA binding proteins</td><td>H-NS</td><td>Filament-forming NAP. Binds A/T-rich DNA and works synergistically with Rho to silence specific loci, including xenogeneic regions of E. coli and Salmonella genomes (Peters et al. 2012; Bossi et al. 2019).</td></tr><tr><td>UvrABD complex</td><td>Involved in SOS-mediated transcription antitermination during TCR, possibly because of overlapping, mutually exclusive Rho/NusG and UvrABD binding sites on RNAP (Martinez et al. 2022).</td></tr><tr><td rowspan="2">Direct, endogenous Rho inhibitors</td><td>Rof (YaeO)</td><td>Direct Rho interactor and inhibitor (Pichoff et al. 1998). Protects E. coli from oxidative stress (Kawamura et al. 2005) and is required for virulence gene expression and host cell invasion in Salmonella (Zhang et al. 2024). CryoEM structures of Rof-Rho complexes show Rof molecules bound at Rho protomer interfaces where they partly occlude the PBS and lock Rho in an inactive, open ring conformation (Said et al. 2024; Zhang et al. 2024). YaeO from V. cholerae may operate by a distinct mechanism disrupting the oligomeric state of V. cholerae Rho (Pal et al. 2019).</td></tr><tr><td>YihE</td><td>Stress response kinase. Direct Rho interactor and antagonist in the cell envelope stress response (Wang et al. 2022).</td></tr><tr><td rowspan="3">Phage proteins</td><td>λN</td><td>Allows expression of phage λ early genes. Intrinsically unstructured (Krupp et al. 2019). Constituent of λ antitermination complex (Santangelo and Artsimovitch 2011; Wang and Artsimovitch 2020). Invades RNAP hybrid channel in cryoEM structure of the antitermination complex (Krupp et al. 2019).</td></tr><tr><td>Qλ, Q21, Q82</td><td>Allow expression of phage late genes. Qλ, Q21, and Q82 are functional (but not sequence) antitermination homologs from lambdoid phages. Q protein loads onto promoter-proximal, σ-dependent paused TEC using a Q-binding element (aka <i>qut</i>) in the DNA template (Roberts et al. 2008). Loaded Q extends and narrows the RNA exit channel of RNAP, making it highly processive and less prone to pausing or termination (Shi et al. 2019; Yin et al. 2019; Yin et al. 2022). Qλ is a proven polarity suppressor (Forbes and Herskowitz 1982). NusA stabilizes Qλ and Q82 antitermination complexes and helps protect the Q82 complex against Rho in vitro (Yang and Roberts 1989; Shankar et al. 2007).</td></tr><tr><td>Psu</td><td>Bacteriophage P4 capsid decorating protein. Binds and inactivates Rho (Linderoth et al. 1997) as a dimer (Ranjan et al. 2013; Gjorgjevikj et al. 2025). Rho's ATPase activity is affected while its RNA binding capacity is not (Pani et al. 2006). Recent cryoEM structures of the Psu-Rho complex supports that Psu constrains the Rho ring into an open, inactive conformation allowing its hyper-oligomerization (Gjorgjevikj et al. 2025).</td></tr><tr><td rowspan="4">Putative Rho interactors detected by mass spectrometry (Butland et al. 2005)</td><td>Hda</td><td>AAA+ ATPase. Inhibits reinitiation of DNA replication. Effect on RDTT is unknown.</td></tr><tr><td>ClpA</td><td>AAA+ ATPase. Component of the ClpAP and ClpAXP protease complexes. Effect on RDTT is unknown.</td></tr><tr><td>ClpP</td><td>Serine protease. Component of ClpAP, ClpAPX and ClpXP protease complexes. Effect on RDTT is unknown.</td></tr><tr><td>SspB</td><td>ClpXP protease specificity-enhancing factor. Effect on RDTT is unknown.</td></tr><tr><td></td><td>CspC</td><td>Cold shock protein. Promotes transcriptional read through at the sgrS locus, which is regulated by Rho inhibitors Rof and bicyclomycin (Morita et al. 2022) and contains a candidate Rut site (Delaleau et al. 2022). Effect of CspC on RDTT has not been investigated.</td></tr></tbody></table>
<table><tr><td>CspD</td><td>Cold shock protein. Promotes transcriptional read through at the sgrS locus, which is regulated by Rho inhibitors Rof and bicyclomycin (Morita et al. 2022) and contains a candidate Rut site (Delaleau et al. 2022). Effect of CspD on RDTT has not been investigated.</td></tr><tr><td>CspE</td><td>Cold shock protein. Effect on RDTT is unknown.</td></tr><tr><td>DnaJ</td><td>Cochaperone of Hsp70 chaperone; binds σ32 factor. Effect on RDTT is unknown.</td></tr><tr><td>DxS</td><td>1-deoxy-D-xylulose-5-phosphate synthase. Effect on RDTT is unknown.</td></tr><tr><td>FabI</td><td>Enoyl-[acyl-carrier-protein] reductase. Effect on RDTT is unknown.</td></tr><tr><td>FtsA</td><td>Essential cell division protein. Overexpression of Rho inhibitor protein Rof suppresses temperature-sensitive mutations in ftsA (Pichoff et al. 1998). Effect on RDTT is unknown.</td></tr><tr><td>GcpE</td><td>(E)-4-hydroxy-3-methylbut-2-enyl-diphosphate synthase. Effect on RDTT is unknown.</td></tr><tr><td>GyrA</td><td>DNA gyrase subunit A. Effect on RDTT is unknown.</td></tr><tr><td>LpxD</td><td>UDP-3-O-(3-hydroxymyristoyl)glucosamine N-acyltransferase. Effect on RDTT is unknown.</td></tr><tr><td>MalT</td><td>DNA-binding transcriptional activator. Effect on RDTT is unknown.</td></tr><tr><td>MreB</td><td>Cytoskeletal protein. Effect on RDTT is unknown.</td></tr><tr><td>PlsB</td><td>Glycerol-3-phosphate 1-O-acyltransferase. Effect on RDTT is unknown.</td></tr><tr><td>PrsA</td><td>Ribose-phosphate diphosphokinase. Effect on RDTT is unknown.</td></tr><tr><td>PyrH</td><td>UMP kinase. Effect on RDTT is unknown.</td></tr><tr><td>GsP</td><td>Glutathionylspermidine amidase/synthetase. Effect on RDTT is unknown.</td></tr><tr><td>MenF</td><td>Isochorismate synthase. Effect on RDTT is unknown.</td></tr><tr><td>Tu</td><td>Translation elongation factor. Regulates its own expression in Salmonella by an attenuation-by-RDTT mechanism (Brandis et al. 2016). Effect on RDTT is otherwise unknown.</td></tr><tr><td>YhbY</td><td>Ribosome assembly factor. Effect on RDTT is unknown.</td></tr><tr><td>RnpA</td><td>RNase P subunit. Effect on RDTT is unknown.</td></tr><tr><td>RuvB</td><td>Holliday junction branch migration protein. Effect on RDTT is unknown.</td></tr><tr><td>SecA</td><td>Protein translocation ATPase. Effect on RDTT is unknown.</td></tr><tr><td>SelB</td><td>Specialized translation factor. Effect on RDTT is unknown.</td></tr></table>
FIGURE LEGENDS

**Figure 1:** Transcription termination factor Rho and its interaction with RNA. (A) Diagram of the primary structure of Rho. (B) Structure of the closed Rho hexamer from *Escherichia coli* bound to λtR1 RNA (PDBs 8E3H and 8E6W). RNA is in red, with dinucleotides in YC-binding pockets shown in green. The YC-binding clefts (PBS) are in magenta and the SBS residues in orange. (C) Consensus RNA binding motifs deduced from (Top) the alignment of the λtR1 RNA segments bound to the Rho hexamer shown in panel B or (bottom) differential MEME enrichment of the 2511 *E. coli* putative *Rut* peaks consistently detected by Helicase-SELEX versus 3000 *Rut*-less control sequences (1315 sites; E-value: 5.8 x 10<sup>-30</sup>) (Delaleau et al. 2022). (D) Diagram depicting the trajectory of RNA along the crown-like PBS site of Rho, its entry in the central Rho channel, and its interaction with the SBS loops, which triggers the closure and catalytic activation of the Rho hexamer. (E) Helicase-SELEX with Rho revealed putative *Rut* sites along the *E. coli* genome, visualized as discrete sequence enrichment peaks (blue) in this representative region of the (+) strand. Approximately one-third of the putative *Rut* sites are located upstream of or within genes (Delaleau et al. 2022), representing primary candidate sites for conditional RDTT.

**Figure 2:** Models of Rho-dependent termination of transcription (RDTT). (A) Diagram of the "catch-up" RDTT model and (B) cryoEM reconstruction of the "catch-up" pre-termination complex (PDBs 8E3F, 8E3H, and 8E6W). (C) Diagram of the "stand-by" RDTT model and (D) cryoEM reconstruction of the "stand-by" pre-termination complex (PDB 6Z9P). Note that the Rho hexamer is in an open, inactive configuration in the "stand-by" complex. To better visualize the orientation of the Rho hexamer relative to RNAP, Rho's NTD and CTD segments are shown in magenta and gray, respectively, in all panels.

**Figure 3:** Model for the role of RDTT in H-NS-mediated gene silencing. H-NS:DNA filaments form on negatively supercoiled DNA and constrain negative supercoils (Tupper et al. 1994). H-NS represses transcription initiation from multiple promoters within the filament, including spurious intragenic promoters. Transcripts from promoters that escape repression are intercepted and terminated by Rho, aided by NusG (top). This function is crucial for maintaining the silenced state. If Rho or NusG is defective, RNAP invades the H-NS:DNA filament, generating positive DNA supercoiling as it translocates along the DNA (Wu et al.
1988). As positive and negative supercoils neutralize each other, the resulting loss of negative superhelicity destabilizes the H-NS:DNA complex, leading to transient filament disassembly (bottom). The filament rapidly reassembles after RNAP passage, driven by the surge of negative supercoils accumulating behind the elongating RNAP.

**Figure 4:** Regulation of RDTT by blocking of the *Rut* site. (A) Translation-transcription coupling protects intragenic *Rut* sites from Rho action. (B) Binding of a sRNA to the mRNA leader at/near the RBS prevents translation and unmasks intragenic *Rut* site(s) leading to RDTT. (C) Restructuring of the mRNA leader upon ligand binding to its cognate riboswitch masks the RBS, allowing RDTT from intragenic *Rut* sites. Inhibition of RDTT may also stem from sequestering of the *Rut* site in (D) a secondary RNA structure or (E) a sRNA-mRNA duplex, or (F) from blocking of the *Rut* site by a RNA-binding protein (hypothetical case not discussed in main text).

**Figure 5:** Diagram depicting the regulation of cold shock genes during the cold shock. At 37°C (top), the mRNA leaders adopt OFF conformations that favor RDTT and disfavor translation, inhibiting expression of the cold shock proteins (csp). Temperature-dependent restructuring of the mRNA leaders upon cold shock (middle) has an opposite effect, masking the *Rut* sites and unmasking the RBSs, to allow csp expression. During cold acclimation (bottom), the concentrations of csp proteins increase until they become sufficient for the csp proteins to bind their mRNA leaders and promote their restructuring into OFF conformations favoring RDTT and disfavoring translation.

**Figure 6:** uORF-dependent attenuation-by-RDTT. (A) Diagram depicting the regulation of the *E. coli tnaCAB* operon by tryptophan. At high concentration, tryptophan prevents release of the ribosome from the termination codon, shielding the downstream *Rut* site and allowing expression of *tnaAB*. (B) Diagram depicting the regulation of the *B. pertussis tnaCAB cruR-bfrG-bp2921* operon by Cu ions. The ribosome is stalled on the *cruR* stop codon in absence of Cu ions, shielding the intragenic *Rut* site and allowing expression of the putative Cu importer *bfrG*. In presence of Cu ions, the ribosome is efficiently released from the *cruR* stop codon, allowing RDTT and silencing of *bfrG* (Roy et al. 2022).
**Figure 7:** CryoEM reconstructions of the λN (PDB 6GOV) and *rnn* (PDB 6TQO) antitermination complexes show how the antitermination factors bind RNAP and the nascent transcript to shield them efficiently from Rho action (compare with Figure 2B, D). The complexes also sequester NusG-CTD (by NusE) and prevent RNAP pausing or backtracking (Krupp et al. 2019; Huang et al. 2020).

**Figure 8:** Cryo-EM reconstructions of the (A) Rho-Psu complex (PDB 9GCT) and (B) Rho-Rof complex (PDB 8W8D) show how the two Rho inhibitors use distinct strategies to keep Rho into an inactive configuration. Rho's NTD residues are colored in magenta.
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<figure><img src="image_2.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_3.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_4.png" /><figcaption>D</figcaption></figure>

<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /><figcaption>NTD face (top view)</figcaption></figure>

<figure><img src="image_7.png" /><figcaption>Helicase-SELEX</figcaption></figure>
<figure><img src="image_8.png" /><figcaption>‘Catch-up’ model</figcaption></figure>

<figure><img src="image_9.png" /><figcaption>Alternative
‘Stand-by’ model</figcaption></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>
<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>
A

<figure><img src="image_14.png" /></figure>

Expression

<figure><img src="image_15.png" /></figure>

B

<figure><img src="image_16.png" /></figure>

- sRNA

<figure><img src="image_17.png" /></figure>

C

<figure><img src="image_18.png" /></figure>

- Ligand

D

<figure><img src="image_19.png" /></figure>

E

<figure><img src="image_20.png" /></figure>

F

<figure><img src="image_21.png" /></figure>

+ sRNA

<figure><img src="image_22.png" /></figure>

Termination

+ Ligand

<figure><img src="image_23.png" /></figure>
## HOST TEMPERATURE

### Cold shock gene silencing

<figure><img src="image_24.png" /></figure>

<figure><img src="image_25.png" /></figure>

<figure><img src="image_26.png" /></figure>

<figure><img src="image_27.png" /></figure>

<figure><img src="image_28.png" /></figure>

## COLD SHOCK

### Expression of cold-shock proteins

<figure><img src="image_29.png" /></figure>

<figure><img src="image_30.png" /></figure>

<figure><img src="image_31.png" /></figure>

<figure><img src="image_32.png" /></figure>

Time

<figure><img src="image_33.png" /></figure>

<figure><img src="image_34.png" /></figure>

## COLD ACCLIMATION

### Cold shock gene silencing

<figure><img src="image_35.png" /></figure>

<figure><img src="image_36.png" /></figure>

<figure><img src="image_37.png" /></figure>

<figure><img src="image_38.png" /></figure>

Time

<figure><img src="image_39.png" /></figure>
A

<figure><img src="image_40.png" /><figcaption>Low [Trp]:</figcaption></figure>

<figure><img src="image_41.png" /></figure>

B

<figure><img src="image_42.png" /><figcaption>High [Cu]:</figcaption></figure>

<figure><img src="image_43.png" /></figure>
<figure><img src="image_44.png" /><figcaption>A</figcaption></figure>

λN antitermination
complex

<figure><img src="image_45.png" /></figure>

<figure><img src="image_46.png" /></figure>

rrn antitermination
complex

<figure><img src="image_47.png" /><figcaption>B</figcaption></figure>
A

<figure><img src="image_48.png" /></figure>

B

<figure><img src="image_49.png" /><figcaption>Rho:Rof complex</figcaption></figure>
<figure><img src="image_50.png" /></figure>

RNA

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# Rho-dependent transcription termination: mechanisms and roles in bacterial fitness and adaptation to environmental changes

Thuy Duong Do, Nara Figueroa-Bossi, Johnathan C Black, et al.

RNA published online June 3, 2025

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Published online June 3, 2025 in advance of the print journal.

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