Introduction

RNase E is the main endoribonuclease in Escherichia coli, known for its role in RNA processing and mRNA degradation^1-3. It is an essential protein^4,5, and homolog proteins are found across many bacterial species^6-8. The essentiality stems from the N-terminal domain (NTD) or the catalytic domain⁹. The NTD is followed by a membrane targeting sequence (MTS) and the C-terminal domain (CTD) or macromolecular interaction domain², where RhlB (a DEAD-box RNA helicase), PNPase (a 3’→5’ exonuclease), and enolase (a glycolytic enzyme) bind to form the RNA degradosome complex⁶. The MTS forms an amphipathic α helix, responsible for the localization of RNase E in the inner membrane^10,11. Interestingly, the membrane localization of RNase E and the presence of CTD are not essential in E. coli nor are conserved across bacterial species, in contrast to the broad conservation of NTD across bacteria as well as chloroplast^7,12,13. This raises a question about the role of membrane localization and CTD in the in vivo function of RNase E.

E. coli strains with cytoplasmic RNase E (due to the removal of MTS) are viable although they grow slower than the wild-type (WT) cells^10,14. In vitro studies showed that membrane binding of RNase E does not necessarily increase its enzymatic rates^10,14. However, membrane localization is likely important for gene regulation in vivo because RNase E becomes sequestered from the cytoplasmic pool of mRNAs, giving mRNAs time for translation. This idea is supported by our recent observation that the membrane-bound RNase E limits the degradation of nascent mRNAs while cytoplasmic RNase E (ΔMTS) can degrade nascent mRNAs during transcription¹⁵. We found that transcripts encoding membrane proteins can be an exception to this rule, in that they can experience co-transcriptional degradation assisted by transertion effect¹⁵. These findings agree with results from a genome-wide study, indicating that the membrane localization of RNase E (but not the cytoplasmic localization of RNase E) allows for differential regulation of mRNA stability for genes encoding cytoplasmic proteins versus inner membrane proteins in E. coli¹⁶.

Previous studies have reported evidence that E. coli RNase E can localize in the cytoplasm—for example, when cells were grown anaerobically¹⁷ or when membrane fluidity was reduced by changes in lipid composition¹⁸. These findings imply that RNase E can dissociate from the membrane; however, the origin of weak membrane binding remains unknown.

Across bacteria, several species within α-proteobacteria have cytoplasmic RNase E¹⁹ while other species have membrane-bound RNase E. Among these, B. subtilis RNase Y (a functional homolog of RNase E) associates with the membrane via a transmembrane motif²⁰, instead of an amphipathic motif used by E. coli and other γ-proteobacteria⁷. Considering different types of membrane-binding motifs that evolution has introduced, E. coli RNase E may possibly be engineered with a transmembrane motif. Such a mutant will serve as a useful model for investigating the impact of membrane-binding motifs on the localization, diffusion, and activity of RNase E.

Lastly, the CTD of E. coli RNase E is a natively unstructured region²¹ that has been shown to contribute to the enzymatic activity happening at the NTD^15,22-24. We recently showed that this is likely due to an internal allosteric effect within RNase E, instead of the roles played by additional factors bound at the CTD, such as RhlB and PNPase¹⁵. Interestingly, CTD is not conserved across bacteria⁷, and its contribution to the structural stability and membrane binding affinity of RNase E remains unknown.

In this study, we quantified the membrane binding percentage (MB%) of RNase E in E. coli cells using single-molecule microscopy and investigated how its membrane association affects its diffusion as well as its function in mRNA degradation. We examined the effects of different membrane-binding motifs, including the original MTS and transmembrane segments derived from LacY, on the localization and diffusion of RNase E, in the presence and absence of CTD. Our work provides new insights into the spatial organization of RNase E in bacteria and highlights the importance of MTS and CTD in regulating the localization, diffusion, and function of RNase E. Our findings offer potential avenues for modulating RNase E’s subcellular localization and diffusion dynamics and hence its activity, for various applications.

Results

Membrane binding percentage (MB%) of RNase E

Fluorescence microscopy studies have shown that RNase E is localized to the membrane in E. coli^10,11,16, but the percentage of molecules bound to the membrane has not been quantitatively examined in live cells. To analyze RNase E localization and dynamics at the single-molecule level, we fused RNase E with a photo-convertible fluorescent protein, mEos3.2²⁵ and imaged individual RNase E molecules over time in two dimensions (Fig. 1A). The location of fluorescent molecules was identified in each frame and linked to form tracks using the open-source software u-track²⁶ (Fig. 1A). The subcellular locations were also calculated relative to the cell boundaries identified from bright-field images using another open-source image analysis package Oufti²⁷ (Fig 1B). To combine data from many cells, molecular positions along the short and long axes of a cell were normalized to the cell width and cell length for xNorm and yNorm, respectively (Fig. 1C). Based on the yNorm, molecules in the cylindrical part of cells were selected, and their xNorm values were used to obtain an xNorm histogram. Hereinafter, we will focus on the xNorm histogram to compare the membrane enrichment of proteins.

Figure 1:

The xNorm histogram of RNase E shows two peaks corresponding to each side edge of the membrane (Fig. 1D and Fig. S1), very similar to the xNorm histogram of LacY, obtained by imaging LacY-mEos3.2 using the same method. LacY is a membrane channel for lactose and is composed of 12 transmembrane segments²⁸. It is expected to be inserted into the inner membrane during translation^29-31, such that all imaged LacY is expected to be localized in the inner membrane³².

We further quantified the percentage of molecules bound to the membrane (membrane-binding percentage or MB%) from the xNorm histogram. To do this, we developed a mathematical model based on a 2D projection of molecules randomly distributed on the surface of or inside a cylinder. The model includes imaging effects that affect the shape of xNorm histograms: the localization error, limited focal depth of the quasi-TIRF illumination we used, and the location of the inner membrane relative to the cell boundary (Fig. S2). The model fitting was performed using the Markov-Chain Monte Carlo algorithm (MCMC). For validation, we applied the model to xNorm histograms of LacY and LacZ for complete membrane binding or complete cytoplasmic localization, respectively. The MB% of LacY was 99% with a 95% confidence interval of [96%, 100%], and LacZ showed MB% of 3.4% [0.2%, 7.3%] (Fig. 1E-F, Fig. S3A). Both MB% agree with the expectation for membrane and cytoplasmic proteins. For RNase E, we found MB% of 93% [91%, 96%], suggesting that most of the RNase E molecules are localized in the inner membrane (Fig. 1E).

Factors affecting slow diffusion of RNase E

The membrane localization of RNase E likely limits its interaction with mRNAs in the cytoplasm. We measured how fast RNase E moves in the membrane by analyzing the trajectories of individual RNase E-mEos3.2 imaged at 21.7 ms acquisition interval (Fig. 1A). We calculated the diffusion coefficient D by fitting the mean-squared displacement (MSD) of trajectories with MSD = 4Dτ + b, where τ is lag time and b is a combination of dynamic and static localization error³³ (Supplementary Information). We obtained D of 0.0184 ± 0.0002 μm²/s (mean ± SEM) for RNase E (Fig. 2A). As a comparison, RNase E’s D was larger than the low limit of D our microscope can measure from stationary, surface-immobilized mEos3.2, i.e. D = 0.0020 ± 0.0001 μm²/s (Fig. S3A, Supplementary Discussion). Also, RNase E’s D was comparable to that of its RNA substrates, i.e. mRNAs bound with ribosomes, estimated to be D = 0.015 μm²/s based on the diffusion of ribosomal protein L1 (Fig. S4C). We investigated the origin of RNase E’s slow mobility by testing the effect of (i) mRNA substrates, (ii) RNA degradosome components, and (iii) membrane attachment on the diffusion of RNase E.

Figure 2:

To test the effect of mRNA substrates on the diffusion of RNase E, we treated cells with rifampicin, which blocks transcription initiation thus depleting cellular mRNAs. In rifampicin-treated cells, D of RNase E was 0.0270 ± 0.0003 μm²/s, a 1.47-times increase from untreated cells (Fig. 2B). We note that LacY also exhibited a statistically significant increase in D in rifampicin-treated cells with a 1.25-times increase from untreated cells (Fig. 2C). Although this increase is relatively small, it was unexpected because LacY is not an RNA-binding protein. The increase in D of LacY in rifampicin-treated cells likely results from the depletion of mRNAs near the membrane (e.g., the mRNAs undergoing transertion), which could otherwise hinder the diffusion of membrane proteins^34,35.

The fact that RNase E exhibits less than a two-fold increase in D upon rifampicin treatment is surprising. In the case of the ribosome, the diffusion of its large subunit L1 protein was 7 times faster in rifampicin-treated cells (Fig. 2D), consistent with previous reports^36,37. This big change can be explained by the fact that ribosomes form polysomes, such that the effective mass of L1 protein in non-treated cells would be 2 or more times larger than that in rifampicin-treated cells (where it remains as a free subunit). In the case of RNase E, it forms the RNA degradosome complex, whose mass can be from 450 kDa⁶ to 2.3 MDa depending on the partial or full occupancy of the canonically known component proteins (RhlB, PNPase, enolase)^2,38-41 (Supplementary Discussion). In either case, the RNase E complex is smaller than the 70S ribosome of ∼2.5 MDa⁴². If RNase E interacts with an mRNA with n ribosomes, the total mass of the RNA degradosome complex would increase by (n + 1) fold or more. Thus, a substantial increase in D of RNase E is expected upon mRNA depletion.

To explain the marginal increase in RNase E’s diffusion upon rifampicin treatment, we considered two possibilities; it is possible that only a small percentage of RNase E interacts with mRNAs and/or RNase E interacts with mRNA very briefly, unlike ribosomes spending an order of 10-100 s in a polysome state for translation elongation⁴³. When we increased the polysome pool of the cellular mRNA by treating cells with a translation elongation inhibitor chloramphenicol⁴⁴ or by overexpressing lacZ mRNA from a high-copy plasmid, D of RNase E remained unchanged from untreated cells (Fig. S4D). This result rules out the possibility that only a small percentage of RNase E interacts with mRNAs and weighs in favor of the scenario that RNase E interacts with mRNAs briefly.

Next, we investigated the effect of membrane association on the diffusion of RNase E by comparing D of the WT RNase E with a cytoplasmic RNase E mutant lacking the MTS segment. The WT RNase E and RNase E ΔMTS have a similar molecular mass since MTS is only 15 residues of 1000 residues in a monomer RNase E (Fig. 2E and Fig. S5). Therefore, the difference in D should be attributed to their subcellular localizations (Fig. 2F; membrane vs cytoplasm) instead of the mass. We found that the cytoplasmic RNase E diffuses ∼5.5 times faster than the membrane-bound, WT RNase E (Fig. 2G). A similar increase (∼5.3 times) was observed when we examined the CTD truncation mutants, membrane-bound RNase E (1-592) and cytoplasmic RNase E (1-529), which have similar mass but different localizations (Fig. 2F-G).

The CTD region is where the RNA degradosome proteins (enolase, RhlB, and PNPase) bind to create the large RNA degradosome complex (Fig. 2E). To test how these proteins affect the diffusion of RNase E, we measured the diffusion of RNase E in strains where rhlB or pnp was deleted. D increased 1.08 and 1.04 times for ΔrhlB and Δpnp, respectively. The increase was very minute (especially for Δpnp), considering that the mass of the RNA degradosome is expected to decrease by 15% and 40% for rhlB and pnp deletions, respectively (Fig. 2H-I). This finding possibly suggests complex stoichiometry in the RNA degradosome (see Discussions). Despite this ambiguity, a significant 4.04 times increase in D was observed in RNase E ΔCTD, or RNase E (1-592), where the CTD of RNase E was deleted to eliminate interactions with all RNA degradosome components while keeping the membrane localization (Fig. 2H). In this mutant, the mass of the RNase E complex is expected to decrease to 396 kDa (Fig. 2I). Altogether, these results suggest that the slow diffusion of RNase E is largely affected by the membrane localization and the formation of the massive RNA degradosome and less by interaction with mRNA substrates, unlike other RNA-interacting proteins, such as ribosomes.

Diffusion and localization of MTS and transmembrane segments

Unlike RNase E in E. coli, RNase Y, a functional homolog of RNase E in B. subtilis, is localized to the membrane via a transmembrane domain. Any differences between a peripheral (like MTS of E. coli’s RNase E) and a transmembrane motif on membrane localization and mobility of RNase E, remain unclear. To address this question, we created RNase E mutants that contain a transmembrane domain in place of the MTS.

Before creating the mutants, we characterized individual short membrane-binding motifs by fusing them with mEos3.2. LacY in its native form has 12 transmembrane segments that are spatially arranged into two groups of 6 transmembrane segments²⁸. We were able to express the first two transmembrane segments (LacY(1-73) or LacY2), the first six transmembrane segments (LacY(1-192) or LacY6), and the original LacY (LacY12) fused to mEos3.2 from the IPTG-inducible promoter on the chromosome. We then imaged their membrane localization and diffusivity (Fig. 3A). The MTS segment and LacY-derived transmembrane segments showed a strong membrane enrichment (Fig. 3B). In terms of diffusion, LacY2 and LacY6 diffused faster than the MTS segment (Fig. 3C). This is remarkable because their molecular masses (and thus size) are expected to be larger than that of MTS (Fig. 3D). The Saffman-Delbruck diffusion model states that the diffusion coefficient of membrane proteins decreases logarithmically as the radius of the embedded part increases given the same membrane environment⁴⁵. Our data suggest that the two types of membrane-binding motifs (MTS and LacY2) experience different membrane environments, possibly due to the way they interact with the lipids.

Figure 3:

To further explore protein-membrane interactions, we conducted all-atom molecular dynamics simulations of MTS and LacY2 interacting with the E. coli membrane using the NAMD software⁴⁶ (Fig. 3E; see Supplementary Information). In the simulations, protein motion was calculated for 1 μs. Although the D values were larger than the experimental values (possibly due to the absence of mEos3.2 in the model), MTS again diffused slower than LacY2 (Fig. 3F and Fig. S6A). By calculating membrane-protein interaction energies, we found that MTS-membrane interaction is more stable than LacY2-membrane interaction (Fig. S6B). These results suggest that the diffusion of MTS, located on the surface of the membrane, is slower than that of LacY2 due to stronger interaction with lipid head groups.

RNase E mutants carrying a transmembrane motif

Since LacY2 and LacY6 showed a strong membrane enrichment similar to LacY12 (Fig. 3B), we replaced MTS in RNase E with LacY2, LacY6, and LacY12 in the presence or absence of CTD (Fig. 4A-B and Fig. S5). All the chimeric RNase E mutants were expressed from the native locus on the chromosome as the only copy of RNase E and had mEos3.2 fused at the C terminus for imaging. The strains did not show noticeable differences in growth rate compared to the WT strain (Table S3), suggesting that the RNase E mutants were functionally active.

Figure 4:

xNorm histograms of ΔCTD mutants indicated membrane localization similar to LacY (Fig. 4C). However, mutants containing CTD showed noticeable cytoplasmic subpopulations when LacY2 and LacY6 were used in place of MTS (Fig. 4D). The mathematical model fitting of xNorm histograms suggested that MB% of RNase E-LacY2-CTD and RNase E-LacY6-CTD to be 69% [66%, 73%] and 86% [84%, 90%], respectively (Fig. 4F). We note that the non-perfect membrane localization was observed only in the mutants containing CTD and the same protein without CTD showed MB% of 100% (Fig. 4E-F, Fig. S7A-B). This observation suggests that CTD contributes to the unstable membrane binding of RNase E mutants. Such a difference between the presence and absence of CTD was not observed in the chimera based on LacY12 (Fig. 4E-F), possibly due to a stable membrane insertion of LacY12.

Membrane lipid composition is affected by the central metabolism⁴⁷ and hence, may be different when cells are grown in a different carbon source. To test the possibility that MB% of RNase E changes in different media (due to changes in the membrane property), we imaged RNase E from cells grown in M9 succinate (instead of M9 glycerol with casamino acids and thiamine, CAAT; see Table S3 for growth rate difference). We found that MB% of RNase E-LacY2-CTD reduced even lower to 43% [41%, 46%] (Fig. S7C).

This large reduction contrasted with the small MB% change observed in WT RNase E (from 93% to 92%) and LacY2 (from 99.6% to 100%) when cells were grown in M9 succinate, suggesting that RNase E-LacY2-CTD may exhibit a broad MB% range across different growth conditions (Fig. S7C-D).

Next, we examined whether the diffusion coefficients of chimeric RNase E mutants differed based on the type of membrane motifs they had. For example, MTS diffused slower than LacY2, despite being smaller in size (Fig. 3C-D). From this, we expected the chimeric RNase E with LacY2 to diffuse faster than the RNase E with MTS. Indeed, in the absence of CTD, we found that LacY2-based RNase E diffuses 1.33 times faster than MTS-based RNase E (Fig. 4G). In the presence of CTD, LacY2 and LacY6-based RNase E diffused 3.33 and 1.14 times faster than the MTS-based RNase E counterpart (i.e. the WT RNase E). In this case (with CTD), the large increase in diffusion is also affected by the presence of cytoplasmic population (∼31% for LacY2 and ∼14% for LacY6; Fig. 4H) which diffuses faster than the membrane-bound population (possibly about 5 times faster, based on Fig. 2G). Taken together, our data suggest that transmembrane motifs can facilitate the diffusion of RNase E and allow for dissociation from the membrane in the presence of CTD.

Functional consequence of subcellular localization and diffusion of RNase E

To check the functional consequence of cytoplasmic localization of RNase E, we measured lacZ mRNA degradation in various RNase E mutants presented in this study. Recently, we developed an assay measuring co-transcriptional and post-transcriptional mRNA degradation rates of lacZ by inducing its transcription for only 75 s to capture the degradation of nascent mRNA¹⁵. We found that the rate of co-transcriptional degradation (k_d1) of lacZ mRNA is about 10 times slower than that of post-transcriptional degradation (k_d2) in WT cells. We found that k_d1 increases about 3 fold in cells expressing RNase E ΔMTS, suggesting that cytoplasmic RNase E can freely diffuse and degrade nascent mRNAs in the cytoplasm, in contrast to the membrane-bound RNase E¹⁵. This result indicates that the chimeric RNase E that showed cytoplasmic subpopulation (Fig. 4F) may exhibit larger k_d1 than other RNase E variants.

We repeated this assay in a strain expressing the WT RNase E fused with mEos3.2. Note that this strain is different from the one we used for imaging because a monocistronic lacZ gene is needed to aid data interpretation¹⁵. The relative abundances of 5’ lacZ mRNA (Z5) before 3’ lacZ mRNA (Z3) level increases (between ∼100 and 210 s) were constant, confirming negligible co-transcriptional degradation of lacZ mRNA¹⁵ (Fig. 5A). However, when we examined lacZ mRNA degradation in cells expressing RNase E-LacY2-CTD (69 MB%), the Z5 levels at the same time points exhibited a downward trend (Fig. 5B). Its k_d1 was larger than any other RNase E mutants and close to what was observed in RNase E ΔMTS¹⁵ (Fig. 5C). RNase E-LacY6-CTD also showed some cytoplasmic population (86 MB%) but did not exhibit a significant increase in k_d1 (Fig. 5C). The result from RNase E-LacY2-CTD suggests that the cytoplasmic subpopulation of RNase E is functional and may affect the degradation pattern of mRNAs globally.

Figure 5:

In terms of the post-transcriptional mRNA degradation rate (k_d2), we tested if the slow diffusion of RNase E in the membrane affects k_d2. In the presence of CTD, k_d2 did not vary much among different membrane motifs (MTS and LacY; Fig. 5D). For example, even though the LacY12 motif slows down the diffusion of RNase E, k_d2 was similar to that of WT RNase E (Fig. 5D). This might be due to differences in RNase E copy numbers. Specifically, the LacY12 version of RNase E might be more abundant than the WT version because RNase E expression level is affected by its ability to regulate its own transcript levels⁴⁸. The higher expression level might compensate for the slow diffusion to give a similar k_d2. However, when we examined the over-expression of WT RNase E, we observed that k_d1 and k_d2 remained unaffected (Fig. S8A). These results suggest that k_d2 is not limited by the copy number and diffusion of membrane-bound RNase E in the presence of CTD.

In the absence of CTD, k_d2 was, in general, lower than those measured from RNase E variants including CTD (Fig. 5D), indicating the importance of CTD in RNase E’s catalytic activity. Similar results have been reported earlier for RNase E based on MTS^15,22-24. Among different membrane motifs for RNase E ΔCTD, we noticed that the LacY2 version, which diffuses faster than the MTS version (Fig. 4G), does not necessarily yield higher k_d2 (Fig. 5D). Plus, LacY6 and LacY12 versions showed even lower k_d2 than MTS-based RNase E ΔCTD. Overall, the slow k_d2 coming from the absence of CTD could not be rescued by faster diffusion of RNase E, but it can be worsened by the large and slow membrane motifs. The presence of CTD appears to buffer the effect of membrane motifs on the catalytic activity of RNase E (k_d2).

Discussion

Our study establishes the membrane enrichment and slow diffusion of RNase E in E. coli. This supports the notion that sequestration of RNase E on the membrane confers the spatial and temporal separation between synthesis and decay of mRNAs¹⁵. Furthermore, the processing of rRNA^49-53 and tRNA^54,55 and small RNA-based gene regulations^56,57 mediated by RNase E likely take place on the membrane.

For the WT RNase E, our analysis showed MB% of 93%, close to 91% observed using immunogold labeling and freeze-fracture method in a previous study⁵⁸. Because the membrane localization of RNase E was shown to be affected by membrane lipid composition¹⁸, we tested if the MB% of WT RNase E changes in different growth conditions. When cells were grown in M9 with succinate as a sole carbon source without CAAT, different from the media used in all our measurements (M9 glycerol with CAAT), the MTS segment exhibited MB% reduction from 100% to 83% while the LacY2 segment maintained ∼100% (Fig. S3E, S3G, S6D), suggesting that MTS can lose membrane binding affinity in this condition. This result may be consistent with the fact that amphipathic motifs tend to have a weaker association with the membrane than transmembrane motifs due to peripheral interaction with the membrane⁵⁹. For example, an amphipathic motif within MinD was observed to be completely cytoplasmic when the motif alone was expressed with a GFP fusion in E. coli⁶⁰. MinD becomes a membrane protein as a dimer, suggesting that the dimer formation is needed for strong membrane association⁶⁰. Similarly, the nature of tetramer formation may ensure membrane localization of RNase E when a single MTS exhibits weak membrane association. The MB% of WT RNase E was marginally affected by the growth media (Fig. S3F-G).

While individual transmembrane motifs exhibited stronger membrane association than amphipathic motif MTS, we found LacY2-based RNase E can lose membrane binding affinity, unlike MTS-based RNase E. This was observed only in the presence of the CTD of RNase E, suggesting that in addition to the membrane binding motif, the CTD also contributes to the membrane binding affinity. This idea is also supported by MB% results of 93% for WT RNase E (containing CTD) vs 100% for its ΔCTD version, RNase E (1-592). Similarly, the chimeric RNase E with LacY6 and CTD showed MB% of 86% while its ΔCTD version showed 100% (Fig 4E-F). Functionally, our mRNA degradation assay supported the idea that the CTD might have an allosteric effect on the NTD²⁴, affecting mRNA degradation rates. This was true regardless of the subcellular localization of RNase E: in the membrane, RNase E (1-592) showed lower k_d2 than WT RNase E, and in the cytoplasm, RNase E (1-529) showed lower k_d2 than RNase E ΔMTS¹⁵. Future analysis of the full RNase E structure and conformational dynamics would help address remaining questions on the role of CTD on MB% and the function of RNase E.

D of RNase E we measured in E. coli (0.0183 ± 0.0002 μm²/s) is comparable to those measured in other bacterial species. For example, in Caulobacter crescentus, D of its cytoplasmic RNase E was shown to be about 0.03 μm²/s⁶¹. The diffusion of RNase Y in B. subtilis was found in either a slow (0.031 μm²/s) or fast (0.3 μm²/s) state⁶². The authors showed that the slow population represents RNase Y binding to mRNA and/or the putative RNA degradosome while the fast population is freely diffusing RNase Y. We note that our data do not show two distinct diffusion states for RNase E in E. coli (Fig. S4B), possibly suggesting stable formation of RNA degradosome complexes in E. coli, in contrast to the dynamic formation observed in B. subtilis.

Diffusion is affected by the size of the particle in a given medium⁴⁵, and it has been used to identify different size forms of a protein, due to biochemical interactions or complex formation^36,63. Related, the diffusion of RNase E in ΔrhlB and Δpnp backgrounds can be used to understand the stoichiometry of the RNA degradosome complex. We observed that the WT RNase E (and thus RNA degradosome) in ΔrhlB diffuses faster than that in Δpnp strain. This result is inconsistent with the expectation that the size of RNA degradosome should decrease more by the absence of PNPase than that of RhlB. This expectation assumes that all binding sites for RhlB and PNPase in RNase E are occupied. However, it has been proposed that a PNPase (trimer) may interact with more than one RNase E monomer^2,41. It indicates that the stoichiometry between PNPase and RNase E might be lower than expected, yielding a smaller mass reduction in the RNA degradosome when PNPase is absent.

RNA substrates interacting with RNase E can also increase the effective mass of RNase E and lower its D value. However, when cellular mRNAs were depleted by rifampicin treatment, we found that RNase E diffusion increased less than that of other RNA-binding molecules related to transcription and translation. For example, the large and small ribosomal subunits^36,37,64, tRNA⁶⁵, and RNA polymerase⁴⁴ showed a large (about 10-20 fold) increase in D upon rifampicin treatment. Interestingly, Hfq, an RNA chaperon involved in small RNA regulation together with RNase E, showed a moderate increase (∼2 fold) in D when RNA is depleted by rifampicin⁶⁶. We note that our result is consistent with previous studies that examined the effect of rifampicin on RNase E diffusion in E. coli^11,67 as well as RNase E in C. crescentus and RNase Y in B. subtilis^62,68. These findings likely imply that RNase E (and RNase Y) interacts with its substrate for a short time duration, different from other RNA-binding proteins related to transcription and translation.

Lastly, the slow diffusion of MTS in comparison to LacY2 and LacY6 suggests that MTS is not an efficient membrane-binding motif for diffusion in the membrane. While our molecular dynamics simulations demonstrated that it originates from stronger interaction energy between protein and the membrane, whether this can be generalized to any other peripheral and integral membrane motifs remains to be tested. We speculate that this can be a general phenomenon as peripheral motifs interact with lipids orthogonally while integral membrane motifs interact with the lipids in parallel and can diffuse together with them. Diffusion of membrane-bound molecules contains information about protein-lipid contacts and membrane dynamics⁶⁹. Future studies may help address the distinct diffusion dynamics of MTS and transmembrane motifs.

Acknowledgements

We thank Drs. Mark Arbing, Agamemnon Carpousis, Johan Elf, and Christine Jacobs-Wagner for strains. We thank Maggie Liu, Kavya Vaidya, and Zach Wang for their contributions in the early phase of this work and the members of Kim lab for critical reading of the manuscript. This work was supported by the NSF Center for Physics of Living Cells (1430124), NSF Science and Technology Center for Quantitative Cell Biology (2243257), NIH (R35GM143203; R24GM145965), and Searle Scholars Program.

Additional information

Author contributions

L.T. and S.K. designed the research; L.T. and Y.W. performed imaging and analysis; L.T. and Se.K. measured mRNA lifetimes; LT, Se.K., and S.K. performed genetics; L.T., Y.W., and J.W. developed data analysis methods; S. performed all-atom MD simulations under supervision of E.T.; L.T. and S.K. wrote the manuscript with input from all authors. S.K. supervised the study.

Data/Code

https://github.com/sjkimlab/Code_Publication/tree/master/RNaseE%202024

Additional files

Supplemental Information