![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
|
|
|
|
|
||
Vol. 18, Issue 7, 2728-2734, July 2007
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Howard Hughes Medical Institute and Department of Biochemistry and Biophysics, University of California at San Francisco, San Francisco, CA 94158
Submitted March 1, 2007;
Revised April 25, 2007;
Accepted May 3, 2007
Monitoring Editor: Reid Gilmore
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
). One of the highly conserved features of the targeting machinery is that SRP requires an RNA subunit to function (Walter and Blobel, 1982). This study elaborates the mechanism by which the SRP RNA subunit contributes to the protein targeting reaction.
In the first step of cotranslational protein targeting, the SRP binds to the signal sequence of a nascent polypeptide chain emerging from the ribosome (Walter et al., 1981; Keenan et al., 2001; Halic et al., 2004). The resulting SRP-ribosome-nascent chain complex then binds to the SRP receptor (SR), which resides at the target membrane (Gilmore et al., 1982a,b). The SRP and the SR associate with each other through related GTPase modules, but only when GTP-bound (Miller et al., 1993; Egea et al., 2004; Focia et al., 2004). After transfer of the ribosome to the protein translocation channel (translocon), the SRP and the SR hydrolyze their respective bound GTPs, which causes them to dissociate, and allows for a new round of targeting (Connolly et al., 1991). The SRP and the SR reciprocally activate the other's GTPase and are therefore GTPase-activating proteins (GAPs) for each other (Powers and Walter, 1995).
Although SRP-dependent protein targeting is conserved in all organisms, the prokaryotic system has the fewest components and is therefore the simplest (Poritz et al., 1990; Larsen and Zwieb, 1993). In Escherichia coli, the SRP consists of a single protein, Ffh, and a small RNA, the 4.5S RNA (Poritz et al., 1990). Ffh consists of two domains: the M domain, which contains both the binding site for signal sequences and the 4.5S RNA, and the NG domain, which includes the GTPase module that binds the SRP receptor FtsY. A flexible linker joins the two domains of Ffh (Keenan et al., 1998; Egea et al., 2005). In E. coli, Ffh, FtsY, and the 4.5S RNA are all essential genes.
The SRP RNA is an almost universally conserved component of the SRP targeting system. The only known exception is chloroplast SRP, which lacks an SRP RNA but also functions in a different, posttranslational mode by binding to proteins entering the chloroplast from the cytosol and targeting them to the thylakoid membrane. In E. coli, the 4.5S RNA has two known biochemical activities: to catalyze the interaction of Ffh and FtsY, accelerating both the on and off rate of complex formation by over two orders of magnitude without changing the KD, and to enhance the GTPase activity of the Ffh·FtsY complex by about sevenfold (this RNA-dependent stimulation of GTPase activity is distinct from the much larger GTPase stimulation caused directly by the interaction of Ffh and FtsY; Peluso et al., 2000, 2001). Thus, the 4.5S RNA represents an amazing example of an RNA that modulates the behavior of proteins. However, the relationship between the biochemically defined activities of the 4.5S RNA and the universal requirement for an RNA in cotranslational protein targeting is unknown.
A clue to the role of the 4.5S RNA comes from structural work, which shows that the signal sequence-binding pocket (a deep groove lined by hydrophobic amino acids) is spatially contiguous with the 4.5S RNA (Keenan et al., 1998; Batey et al., 2000). Moreover, the hydrophobic groove can assume multiple conformations, indicating that its open state may not be stable in an aqueous environment but closes so that hydrophobic residues can pack against each other and thus be shielded from water (Rosendal et al., 2003). Because this conformational variability occurs in close juxtaposition to the 4.5S RNA, it is plausible that the RNA might monitor the occupancy of the signal sequence-binding pocket and transmit that information to the NG domain, thereby modulating its interaction with FtsY. However, it has remained unresolved how the 4.5S RNA might be linked to these conformational dynamics.
Solution studies looking at the conformation of Ffh suggested that in the absence of the 4.5S RNA, the M domain may occlude the binding site for FtsY. 4.5S RNA binding to Ffh may relieve this inhibition to promote complex formation (Buskiewicz et al., 2005a,b; Halic et al., 2006a,b; Mainprize et al., 2006; Schaffitzel et al., 2006). Although this model provides a plausible mechanism for 4.5S RNA activity, it would suggest that the role of 4.5S RNA is limited to mediate a single Ffh activation step during SRP assembly. Once bound, it would be an inert bystander that would not contribute actively to regulation of the SRP cycle, as there is currently no evidence that 4.5S RNA dissociates from Ffh after binding.
Here, we describe a reverse-genetic and biochemical analysis designed to ask if and how the 4.5S RNA facilitates communication between the M and NG domains of Ffh. We describe mutants in Ffh that bind 4.5S RNA normally and interact normally with FtsY when the 4.5S RNA is absent, but impair the rate of association or GTP hydrolysis when the 4.5S RNA is present. These mutations map to regions of Ffh that undergo major conformational rearrangements during the targeting cycle, suggesting that the 4.5S RNA coordinates the steps of the targeting reaction.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
. Mutations in Ffh were introduced using the QuickChange Mutagenesis protocol (Stratagene, La Jolla, CA). For all in vitro experiments, a truncated form of FtsY (aa 47–497) was used, as previously described (Powers and Walter, 1997). In all cases, assays were performed at 25°C in 50 mM HEPES, pH 7.5, 150 mM potassium acetate, 2 mM magnesium acetate, 0.01% Nikkol, and 2 mM dithiothreitol. The NG domain of Ffh was generated by limited digestion as described (Zopf et al., 1993). The M domain was removed by flowing the digested protein over SP Sepharose, and the NG domain was further purified by gel filtration on Superdex 75.
Fluorescence-binding Assays
Fluorescence-binding experiments were performed as described (Peluso et al., 2000). Rapid reactions [wild type (wt) and Ffh(L301P) + RNA] were performed on a stopped flow fluorimeter (KinTek). Slow reactions were performed on an SLM fluorimeter. For on rates, data were fit to a single exponential and observed rate constants were plotted as a function of concentration. Rate constants were calculated using the following equation: kobs = kon * [Ffh] + koff. Off rates were calculated by preforming complexes of 2 µM of each Ffh (±4.5S RNA) and FtsY and then trapping dissociated components with an excess of GDP. Curves were fit to a single exponential function.
GTPase Assays
Assays were performed as described (Peluso et al., 2001) with slight modifications. To calculate the basal activities of Ffh mutants, trace amounts of [32P]GTP were added to varying concentrations of Ffh and reactions were followed to completion. The data were fit to a single exponential equation to calculate the kobs. In contrast, we used a multiple-turnover regime to measure the stimulated GTPase activity of the Ffh variants. A fixed concentration of Ffh (0.1 µM for Ffh(wt) + 4.5S RNA and other fast reactions and 0.5 µM for Ffh(wt) – 4.5S RNA and other slow reactions) was used with varying concentrations of FtsY. The initial linear portion of the reaction was followed, corrected for the contribution of basal hydrolysis from FtsY (<20% of total hydrolysis observed), and fit to the following equation: turnovers/complex = kobs * time.
Gel Shift Assays
Gel shift analysis of binding of the 4.5S RNA to Ffh mutants was carried out by mixing trace amounts of 32P-end–labeled 4.5S RNA with 0.25 µM cold 4.5S RNA and 0.25 µM Ffh (cold RNA was important to help prevent aggregation of Ffh and RNA in the well) in assay buffer supplemented with 10% glycerol. This mixture was separated in TAE buffer (40 mM Tris-acetate, 20 mM sodium acetate, and 1 mM EDTA) supplemented with 2.5 mM magnesium acetate on a 7% (29:1 acrylamide:bisacrylamide) polyacrylamide gel.
Biotinylation Experiments
Wam121 cells [MC4100 ara+ ffh::kan attB::(OriR6K PBAD-ffh tet); Phillips and Silhavy, 1992] were transformed with plasmid pHP44 (pBR322-acrR'acrA acrB576-PSBT; Tian and Beckwith, 2002) and pHDB7 (pACYC184-Ffh; Lee and Bernstein, 2001) or variants in which point mutations were introduced. Transformants were selected on plates in the presence of arabinose and were then grown overnight at 37°C in liquid medium in the absence of arabinose. Cells were then diluted back and harvested during log phase as described (Tian and Beckwith, 2002). Western blots were performed using streptavidin-horseradish peroxidase (HRP) conjugate (Amersham). HRP was inactivated with azide, and the blots were reprobed with an antibody to Ffh (Poritz et al., 1990) and visualized using chemiluminescence.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
,b). To test this possibility, we generated the NG domain of Ffh (Ffh-NG) by removal of the M domain by V8 protease digestion (Figure 1A). Next, we monitored the kinetics of complex formation by two independent assays: 1) measuring the change in intrinsic tryptophan fluorescence of FtsY (Jagath et al., 2000; Peluso et al., 2000) and 2) measuring the stimulation of GTP hydrolysis (Peluso et al., 2001). In both assays, Ffh-NG behaved as full-length Ffh in the absence of 4.5S RNA and by contrast to full-length Ffh could not be accelerated by addition of the RNA (Figure 1, B–E).
|
; Peluso et al., 2000). We used this fluorescence shift to measure binding of Ffh-NG to FtsY. The rate of Ffh-NG·FtsY complex formation (kon = 99 M–1s–1; Figure 1, C and D) was within error from that of full-length Ffh in the absence of the 4.5S RNA (kon = 180 M–1s–1) and over 500 times slower than full-length Ffh in the presence of the SRP RNA (kon = 5.7 x 104 M–1s–1; Peluso et al., 2000).
Monitoring stimulation of GTP hydrolysis after Ffh-NG·FtsY complex formation yielded similar results. The stimulated GTPase reaction measures the entire interaction cycle of Ffh and FtsY. At low concentrations of FtsY the GTP hydrolysis rate is primarily dependent on the rate of binding of Ffh and FtsY, whereas at higher concentrations the catalytic step of GTP hydrolysis becomes rate limiting (Peluso et al., 2001). Thus, the kmax is a direct measure of the catalytic rate, and the K1/2 depends on both the binding rate and the kmax. As the 4.5S RNA affects both the binding rate and the rate of catalysis, we observe an increase in the kmax as well as a decrease in the K1/2 upon addition of the 4.5S RNA (Figure 1E; cf. 
and 
). When we compared the stimulated GTPase activity of Ffh-NG to that of full-length Ffh, we saw that both the kmax and the K1/2 were nearly identical to that measured for full-length Ffh in the absence of the 4.5S RNA (Figure 1E, Table 1). Furthermore, addition of 4.5S RNA in 10-fold excess of the 0.5 µM KD for the 4.5S RNA and Ffh-NG Buskiewicz et al., 2005a) had no affect on either parameter of the GTPase reaction for Ffh-NG. These results indicate that the Ffh M domain is required for 4.5S RNA to accelerate Ffh·FtsY complex formation. 4.5S RNA bound to the M domain therefore actively enhances the rate of complex formation, rather than the RNA-free M domain slowing it down.
|
Through this analysis, we identified four point mutations in Ffh (L301P, L303D, L350D, and L354D) that impair the activity of 4.5S RNA in interesting ways. All four Ffh mutants bind to the 4.5S RNA (Figure 2A) and hydrolyze GTP in a manner indistinguishable from wild-type Ffh (the measurement refers to basal GTPase in the absence of FtsY; Figure 2B).
|
). By contrast, the binding rates were only modestly affected in the absence of the 4.5S RNA (a 2- to 10-fold decrease; Figure 3B, ).
|
|
), mutations that affect the activity of the 4.5S RNA should affect the on and off rates of complex formation to the same degree. To test this prediction, we determined the off rates of the Ffh mutants in the presence of the 4.5S RNA by following tryptophan fluorescence after addition of GDP to trap Ffh and FtsY in their dissociated, GDP-bound states. As shown in Figure 3, C and D, all three Ffh mutants bound to 4.5S RNA showed dissociation rates comparable to those of complexes containing wild-type Ffh lacking 4.5S RNA (Figure 3D, ). In summary, these data demonstrate that the mutations L303D, L350D, and L354D abrogate the activity of 4.5S RNA to catalyze complex formation between Ffh and FtsY but, importantly, do not impair the affinity with which Ffh and FtsY interact (Figure 3D, KDs 
).
Ffh Mutations L301P and L303D Diminish the Rate of Stimulated GTP Hydrolysis in the Ffh·FtsY Complex
In addition to catalyzing Ffh·FtsY complex formation, the 4.5S RNA also stimulates the rate of GTP hydrolysis by the complex (kmax), albeit to a lesser extent (Peluso et al., 2001). All four Ffh mutants analyzed here showed some reduction of RNA stimulation of GTPase activity by the complex. Ffh(L350D) and Ffh(L354D) had less than a twofold reduction of maximal GTP hydrolysis activity in the presence of the 4.5S RNA (Figure 4, C and D, Table 1). The Ffh(L303D) and Ffh(L301P)·FtsY complex showed kmax levels that were reduced to near the level of the wild-type Ffh·FtsY complex lacking 4.5S RNA (Figure 4, A, B, and D, Table 1). In contrast to the other three mutants, Ffh(L301P) binds to FtsY with normal rates (Figure 3B and Table 2). Although the 4.5S RNA stimulation of Ffh·FtsY GTPase activity is small, the effects of the mutations were consistent across at least three independent experiments and two independent preparations of protein for each Ffh variant.
|
Mutations That Impair Either Activity of the 4.5S RNA Lead to In Vivo Protein Targeting Defects
Previously it was unknown how the biochemical activities of the 4.5S RNA related to the cellular function of the 4.5S RNA in cotranslational protein targeting. Having identified Ffh mutants that specifically impair the activities of the 4.5S RNA, we were able to assess the importance of these activities for protein targeting in vivo without directly perturbing the 4.5S RNA. To do this, we expressed the Ffh mutants in cells that harbor a reporter for cotranslational protein targeting and conditionally express wild-type Ffh.
To monitor cotranslational protein targeting, we used the elegant system developed by Tian and Beckwith (2002) in which the multispanning membrane protein, AcrB is fused to the Proprionibacterium shermanii transcarboxylase (PSBT) biotinylation domain. When this fusion protein is targeted to the membrane by SRP, the PSBT is targeted to the periplasm, where it is not biotinylated (Figure 5A). However, if the SRP-targeting system is defective, the protein accumulates in the cytoplasm and is biotinylated. We introduced plasmids to direct the expression of either wild-type Ffh or Ffh bearing the L301P, L303D, L350D, and L354D mutations into E. coli cells harboring the AcrB-PSBT fusion and in which the sole genomic copy of Ffh was conditionally expressed by the presence of arabinose (Phillips and Silhavy, 1992).
|
). Reduced levels of active SRP in combination with overexpression of an SRP substrate such as AcrB, however, can lead to pronounced synthetically toxic effects (Ulbrandt et al., 1997; Bernstein and Hyndman, 2001). Western blotting of cell extracts with an Ffh-specific antibody showed that all Ffh variants were expressed to comparable levels (Figure 5B, bottom). Taken together, these data indicate that both biochemical activities of the 4.5S RNA—stimulating Ffh·FtsY complex formation and stimulating GTPase activity in the complex—are critical for protein targeting in vivo.
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
A striking feature of the mutations characterized here is that all four are found in positions of Ffh that are conformationally dynamic, based on comparison of available crystal structures (Figure 6). In particular, L350 and L354 are both part of a short helix in the finger loop of the M domain. In one crystal structure of the M domain (Thermus aquaticus; Keenan et al., 1998; Figure 6A, top), the residues line the top of one side of the signal sequence-binding groove. By contrast, in another crystal structure (Sulfolobus solfataricus; Rosendal et al., 2003), Figure 6B, top) the residues point away from the signal sequence-binding pocket. It is interesting to note that in the Keenan structure, two adjacent M domains in the crystal lattice intertwine, such that hydrophobic residues from one M domain partially occupy the signal sequence-binding groove of the other. Thus, the conformational differences in the two M domain structures may represent a change in conformation from a closed state (T. aquaticus structure) to an open state (S. solfataricus structure) that occurs when the signal sequence-binding pocket becomes occupied. This view suggests that the activity of the SRP RNA may be dependent on the occupancy of the signal sequence-binding groove. For example, binding of the SRP to a signal sequence may enhance the activity of the SRP RNA, giving cargo bound SRP a kinetic advantage to interact with FtsY.
|
(the metazoan ortholog of FtsY) was observed, indicating that binding to the SR induces a conformational change (Halic et al., 2006b). Solution studies further support the notion that complex formation between the SRP and SR induce a conformational change in this region (Buskiewicz et al., 2005a; Spanggord et al., 2005). Taken together with our results, these data indicate that a specific conformation of the linker is required for GTPase stimulation by the 4.5S RNA and that factors affecting the relative conformation of the M and NG domains of Ffh could affect the rate of Ffh·FtsY dissociation driven by GTP hydrolysis. For example, a ribosome bound to SRP may inhibit the conformation required for RNA-dependent GTPase stimulation and thus delay Ffh·FtsY dissociation until the ribosome has been transferred to the translocon.
Significantly, we found that the two activities of the 4.5S RNA, to promote complex formation and to promote GTP hydrolysis, are differentially sensitive to the mutations in Ffh that mimic the absence of 4.5S RNA in vitro. Although the two mutations in the signal sequence-binding domain, L350D and L354D, primarily affect 4.5S RNA stimulation of binding rate to FtsY, L301P primarily affects 4.5S RNA stimulation of GTPase activity, and L303D causes dramatic reductions in both binding rate and GTPase activity. Although these distinctions are not absolute, the differences contrast with previously reported mutations in the tetraloop region of the 4.5S RNA that act similarly to the L303D mutation and compromise both the rate of complex formation and the maximal rate of GTP hydrolysis in the complex (Jagath et al., 2001; Siu et al., 2006). Thus, the data presented here demonstrate that the two activities can be differentially affected. Therefore the activities of the 4.5S RNA to promote complex formation and disassembly could, in principle, be differentially regulated, consistent with the models presented above.
Taken together, the mutations identified in this study support the model that the SRP RNA links the major conformational changes in the signal sequence-binding subunit of the SRP to the interaction cycle of the SRP and the SR. Such molecular communication within SRP provides an attractive mechanism for coordination of the interaction of the SRP and the SR with ribosome recruitment and transfer to the translocon and an explanation for the centrality of the SRP RNA to efficient protein targeting.
|
ACKNOWLEDGMENTS |
|---|
|
Footnotes |
|---|
Address correspondence to: Peter Walter (pwalter{at}biochem.ucsf.edu)
Abbreviations used: SRP, signal recognition particle; SR, SRP receptor.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Bernstein, H. D., and Hyndman, J. B. (2001). Physiological basis for conservation of the signal recognition particle targeting pathway in Escherichia coli. J. Bacteriol 183, 2187–2197.
Buskiewicz, I., Kubarenko, A., Peske, F., Rodnina, M. V., and Wintermeyer, W. (2005a). Domain rearrangement of SRP protein Ffh upon binding 4.5S RNA and the SRP receptor FtsY. RNA 11, 947–957.
Buskiewicz, I., Peske, F., Wieden, H. J., Gryczynski, I., Rodnina, M. V., and Wintermeyer, W. (2005b). Conformations of the signal recognition particle protein Ffh from Escherichia coli as determined by FRET. J. Mol. Biol 351, 417–430.[CrossRef][Medline]
Connolly, T., Rapiejko, P. J., and Gilmore, R. (1991). Requirement of GTP hydrolysis for dissociation of the signal recognition particle from its receptor. Science 252, 1171–1173.
Egea, P. F., Shan, S. O., Napetschnig, J., Savage, D. F., Walter, P., and Stroud, R. M. (2004). Substrate twinning activates the signal recognition particle and its receptor. Nature 427, 215–221.[CrossRef][Medline]
Egea, P. F., Stroud, R. M., and Walter, P. (2005). Targeting proteins to membranes: structure of the signal recognition particle. Curr. Opin. Struct. Biol 15, 213–220.[CrossRef][Medline]
Focia, P. J., Shepotinovskaya, I. V., Seidler, J. A., and Freymann, D. M. (2004). Heterodimeric GTPase core of the SRP targeting complex. Science 303, 373–377.
Gilmore, R., Blobel, G., and Walter, P. (1982a). Protein translocation across the endoplasmic reticulum. I. Detection in the microsomal membrane of a receptor for the signal recognition particle. J. Cell Biol 95, 463–469.
Gilmore, R., Walter, P., and Blobel, G. (1982b). Protein translocation across the endoplasmic reticulum. II. Isolation and characterization of the signal recognition particle receptor. J. Cell Biol 95, 470–477.
Halic, M., Becker, T., Pool, M. R., Spahn, C. M., Grassucci, R. A., Frank, J., and Beckmann, R. (2004). Structure of the signal recognition particle interacting with the elongation-arrested ribosome. Nature 427, 808–814.[CrossRef][Medline]
Halic, M., Blau, M., Becker, T., Mielke, T., Pool, M. R., Wild, K., Sinning, I., and Beckmann, R. (2006a). Following the signal sequence from ribosomal tunnel exit to signal recognition particle. Nature 444, 507–511.[CrossRef][Medline]
Halic, M., Gartmann, M., Schlenker, O., Mielke, T., Pool, M. R., Sinning, I., and Beckmann, R. (2006b). Signal recognition particle receptor exposes the ribosomal translocon binding site. Science 312, 745–747.
Jagath, J. R., Matassova, N. B., de Leeuw, E., Warnecke, J. M., Lentzen, G., Rodnina, M. V., Luirink, J., and Wintermeyer, W. (2001). Important role of the tetraloop region of 4.5S RNA in SRP binding to its receptor FtsY. RNA 7, 293–301.[Abstract]
Jagath, J. R., Rodnina, M. V., and Wintermeyer, W. (2000). Conformational changes in the bacterial SRP receptor FtsY upon binding of guanine nucleotides and SRP. J. Mol. Biol 295, 745–753.[CrossRef][Medline]
Keenan, R. J., Freymann, D. M., Stroud, R. M., and Walter, P. (2001). The signal recognition particle. Annu. Rev. Biochem 70, 755–775.[CrossRef][Medline]
Keenan, R. J., Freymann, D. M., Walter, P., and Stroud, R. M. (1998). Crystal structure of the signal sequence binding subunit of the signal recognition particle. Cell 94, 181–191.[CrossRef][Medline]
Larsen, N., and Zwieb, C. (1993). The signal recognition particle database (SRPDB). Nucleic Acids Res 21, 3019–3020.
Lee, H. C., and Bernstein, H. D. (2001). The targeting pathway of Escherichia coli presecretory and integral membrane proteins is specified by the hydrophobicity of the targeting signal. Proc. Natl. Acad Sci. USA 98, 3471–3476.
Mainprize, I. L., Beniac, D. R., Falkovskaia, E., Cleverley, R. M., Gierasch, L. M., Ottensmeyer, F. P., and Andrews, D. W. (2006). The structure of Escherichia coli signal recognition particle revealed by scanning transmission electron microscopy. Mol. Biol. Cell 17, 5063–5074.
Miller, J. D., Wilhelm, H., Gierasch, L., Gilmore, R., and Walter, P. (1993). GTP binding and hydrolysis by the signal recognition particle during initiation of protein translocation. Nature 366, 351–354.[CrossRef][Medline]
Peluso, P., Herschlag, D., Nock, S., Freymann, D. M., Johnson, A. E., and Walter, P. (2000). Role of 4.5S RNA in assembly of the bacterial signal recognition particle with its receptor. Science 288, 1640–1643.
Peluso, P., Shan, S. O., Nock, S., Herschlag, D., and Walter, P. (2001). Role of SRP RNA in the GTPase cycles of Ffh and FtsY. Biochemistry 40, 15224–15233.[CrossRef][Medline]
Pettersen, E. F., Goddard, T. D., Huang, C. C., Couch, G. S., Greenblatt, D. M., Meng, E. C., and Ferrin, T. E. (2004). UCSF Chimera-a visualization system for exploratory research and analysis. J. Comput. Chem 25, 1605–1612.[CrossRef][Medline]
Phillips, G. J., and Silhavy, T. J. (1992). The E. coli ffh gene is necessary for viability and efficient protein export. Nature 359, 744–746.[CrossRef][Medline]
Poritz, M. A., Bernstein, H. D., Strub, K., Zopf, D., Wilhelm, H., and Walter, P. (1990). An E. coli ribonucleoprotein containing 4.5S RNA resembles mammalian signal recognition particle. Science 250, 1111–1117.
Powers, T., and Walter, P. (1995). Reciprocal stimulation of GTP hydrolysis by two directly interacting GTPases. Science 269, 1422–1424.
Powers, T., and Walter, P. (1997). Co-translational protein targeting catalyzed by the Escherichia coli signal recognition particle and its receptor. EMBO J 16, 4880–4886.[CrossRef][Medline]
Rosendal, K. R., Wild, K., Montoya, G., and Sinning, I. (2003). Crystal structure of the complete core of archaeal signal recognition particle and implications for interdomain communication. Proc. Natl. Acad. Sci. USA 100, 14701–14706.
Schaffitzel, C., Oswald, M., Berger, I., Ishikawa, T., Abrahams, J. P., Koerten, H. K., Koning, R. I., and Ban, N. (2006). Structure of the E. coli signal recognition particle bound to a translating ribosome. Nature 444, 503–506.[CrossRef][Medline]
Shan, S. O., and Walter, P. (2003). Induced nucleotide specificity in a GTPase. Proc. Natl. Acad. Sci. USA 100, 4480–4485.
Siu, F. Y., Spanggord, R. J., and Doudna, J. A. (2006). SRP RNA provides the physiologically essential GTPase activation function in cotranslational protein targeting. RNA 2, 240–250.
Spanggord, R. J., Siu, F., Ke, A., and Doudna, J. A. (2005). RNA-mediated interaction between the peptide-binding and GTPase domains of the signal recognition particle. Nat. Struct. Mol. Biol 12, 1116–1122.[CrossRef][Medline]
Tian, H., and Beckwith, J. (2002). Genetic screen yields mutations in genes encoding all known components of the Escherichia coli signal recognition particle pathway. J. Bacteriol 184, 111–118.
Ulbrandt, N. D., Newitt, J. A., and Bernstein, H. D. (1997). The E. coli signal recognition particle is required for the insertion of a subset of inner membrane proteins. Cell 88, 187–196.[CrossRef][Medline]
Walter, P., and Blobel, G. (1982). Signal recognition particle contains a 7S RNA essential for protein translocation across the endoplasmic reticulum. Nature 299, 691–698.[CrossRef][Medline]
Walter, P., Ibrahimi, I., and Blobel, G. (1981). Translocation of proteins across the endoplasmic reticulum. I. Signal recognition protein (SRP) binds to in-vitro-assembled polysomes synthesizing secretory protein. J. Cell Biol 91, 545–550.
Zopf, D., Bernstein, H. D., and Walter, P. (1993). GTPase domain of the 54-kD subunit of the mammalian signal recognition particle is required for protein translocation but not for signal sequence binding. J. Cell Biol 120, 1113–1121.
This article has been cited by other articles:
|
|
 |
|
  T. Hainzl, S. Huang, and A. E. Sauer-Eriksson Interaction of signal-recognition particle 54 GTPase domain and signal-recognition particle RNA in the free signal-recognition particle PNAS, September 18, 2007; 104(38): 14911 - 14916. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
: Ffh NG – 4.5S RNA). The inset graph is magnified to show the slow reactions. (E) Stimulated GTP hydrolysis reactions for Ffh·FtsY and Ffh-NG·FtsY. Hydrolysis rates (per complex, per second) are plotted as a function of concentration of FtsY (
: Ffh NG + 4.5S RNA).
), Ffh(L350D) (
), and Ffh(L354D) (
). Reactions were performed with trace [32P]GTP and varying amounts of Ffh. Curves (solid line Ffh(wt), dashed lines for Ffh mutants) were fit to the equation kobs = kcat * [Ffh]/(KM + [Ffh]). Values of kcat are as follows: Ffh(wt), 0.070 min–1; Ffh(L301P), 0.077 min–1; Ffh(L303D), 0.092 min–1; Ffh(L350D), 0.10 min–1; and Ffh(L354D), 0.10 min–1.
