The bacterial flagellar switch complex is getting more complex
2008; Springer Nature; Volume: 27; Issue: 7 Linguagem: Inglês
10.1038/emboj.2008.48
ISSN1460-2075
AutoresGalit N Cohen-Ben-Lulu, Noreen R. Francis, Eyal Shimoni, Dror Noy, Yaacov Davidov, K. Nagendra Prasad, Yael Sagi, Gary Cecchini, Rose M. Johnstone, Michael Eisenbach,
Tópico(s)Bacterial Genetics and Biotechnology
ResumoArticle13 March 2008free access The bacterial flagellar switch complex is getting more complex Galit N Cohen-Ben-Lulu Galit N Cohen-Ben-Lulu Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Noreen R Francis Noreen R Francis Department of Biology, Brandeis University, Waltham, MA, USA Search for more papers by this author Eyal Shimoni Eyal Shimoni Electron Microscopy Unit, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Dror Noy Dror Noy Department of Plant Sciences, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Yaacov Davidov Yaacov Davidov Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Krishna Prasad Krishna Prasad Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Yael Sagi Yael Sagi Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Gary Cecchini Gary Cecchini Molecular Biology, VA Medical Center, San Francisco, CA, USA Department of Biochemistry and Biophysics, University of California, San Francisco, CA, USA Search for more papers by this author Rose M Johnstone Rose M Johnstone Department of Biochemistry, McGill University, Montreal, Quebec, Canada Search for more papers by this author Michael Eisenbach Corresponding Author Michael Eisenbach Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Galit N Cohen-Ben-Lulu Galit N Cohen-Ben-Lulu Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Noreen R Francis Noreen R Francis Department of Biology, Brandeis University, Waltham, MA, USA Search for more papers by this author Eyal Shimoni Eyal Shimoni Electron Microscopy Unit, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Dror Noy Dror Noy Department of Plant Sciences, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Yaacov Davidov Yaacov Davidov Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Krishna Prasad Krishna Prasad Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Yael Sagi Yael Sagi Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Gary Cecchini Gary Cecchini Molecular Biology, VA Medical Center, San Francisco, CA, USA Department of Biochemistry and Biophysics, University of California, San Francisco, CA, USA Search for more papers by this author Rose M Johnstone Rose M Johnstone Department of Biochemistry, McGill University, Montreal, Quebec, Canada Search for more papers by this author Michael Eisenbach Corresponding Author Michael Eisenbach Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel Search for more papers by this author Author Information Galit N Cohen-Ben-Lulu1, Noreen R Francis2, Eyal Shimoni3, Dror Noy4, Yaacov Davidov1, Krishna Prasad1, Yael Sagi1, Gary Cecchini5,6, Rose M Johnstone7 and Michael Eisenbach 1 1Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel 2Department of Biology, Brandeis University, Waltham, MA, USA 3Electron Microscopy Unit, The Weizmann Institute of Science, Rehovot, Israel 4Department of Plant Sciences, The Weizmann Institute of Science, Rehovot, Israel 5Molecular Biology, VA Medical Center, San Francisco, CA, USA 6Department of Biochemistry and Biophysics, University of California, San Francisco, CA, USA 7Department of Biochemistry, McGill University, Montreal, Quebec, Canada *Corresponding author. Department of Biological Chemistry, The Weizmann Institute of Science, POB 26, 76100 Rehovot, Israel. Tel.: +972 8 934 3923; Fax: +972 8 947 2722; E-mail: [email protected] The EMBO Journal (2008)27:1134-1144https://doi.org/10.1038/emboj.2008.48 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The mechanism of function of the bacterial flagellar switch, which determines the direction of flagellar rotation and is essential for chemotaxis, has remained an enigma for many years. Here we show that the switch complex associates with the membrane-bound respiratory protein fumarate reductase (FRD). We provide evidence that FRD binds to preparations of isolated switch complexes, forms a 1:1 complex with the switch protein FliG, and that this interaction is required for both flagellar assembly and switching the direction of flagellar rotation. We further show that fumarate, known to be a clockwise/switch factor, affects the direction of flagellar rotation through FRD. These results not only uncover a new component important for switching and flagellar assembly, but they also reveal that FRD, an enzyme known to be primarily expressed and functional under anaerobic conditions in Escherichia coli, nonetheless, has important, unexpected functions under aerobic conditions. Introduction Bacterial chemotaxis is considered by many as the signalling system for which the level of understanding is most advanced (see, for a review, Eisenbach, 2004). The object of chemotactic signalling in bacteria is to change the direction of flagellar rotation in response to stimulation, which results in a change in swimming behaviour and direction (Larsen et al, 1974). The structural element that is responsible for switching the direction of flagellar rotation is the switch—a ring-like complex (termed the C-ring) at the base of the flagellar motor, which is one of nature's most intriguing molecular machines (see, for reviews, Macnab, 1995; Eisenbach and Caplan, 1998). The switch complex is built from multiple units of the proteins FliG, FliM, and FliN. It is striking how little is known about switch function despite (a) the centrality of switching for chemotaxis, (b) the many known aspects of switch structure, (c) the availability of functional switching mutants, and (d) the extensive effort invested in studying the switch (see, for reviews, Eisenbach and Caplan, 1998; Berg, 2003; Kojima and Blair, 2004). It is known that the default direction of rotation (i.e., the direction in the absence of intracellular signalling) is counterclockwise (see, for a review, Eisenbach, 1996), and that phosphorylation-controlled binding of CheY to the switch protein, FliM, enables the switch to shift to clockwise rotation (see, for a review, Eisenbach, 2004). However, the post-binding processes that occur at the switch are still a mystery. Over a decade ago, Marwan et al (1990) found that fumarate is a factor needed for the archeal Halobacterium salinarium to swim back and forth. Subsequently, this dicarboxylate was found to be a clockwise switching factor in Escherichia coli. In cytoplasm-free but CheY-containing cell envelopes, it enabled the flagella to switch their direction of rotation (Barak and Eisenbach, 1992; Barak et al, 1996). By employing mutants having exceptionally low or high intracellular levels of fumarate due to the absence of one of the enzymes that act on fumarate, that is, succinate dehydrogenase (SDH) or fumarase, Prasad et al (1998) and Montrone et al (1998) found that, in intact E. coli cells, fumarate increases both the fraction of time spent in clockwise rotation and switching frequency. These effects were due, in part, to reduction of the standard free energy difference between the clockwise and counterclockwise states of the switch (Prasad et al, 1998). Fumarase, a cytosolic protein, was also shown to be involved in a phosphorylation-independent response to some repellents (Montrone et al, 1996). The effects of fumarate in intact cells of E. coli were independent of the presence of CheY in the cell, indicating that fumarate exerted its action on the switch rather than on CheY (Prasad et al, 1998). In the present communication, we discovered that the enzyme fumarate reductase (FRD) is the target of fumarate at the switch. FRD and its closely related homologue SDH are four-subunit membrane-bound enzymes (see Supplementary Figure s1 and Supplementary data), usually thought to be involved in electron transport reactions (see, for review, Cecchini et al, 2002). It is well documented that FRD functions in anaerobic respiration of E. coli and that SDH is involved in aerobic respiration. We provide evidence for a reversible interaction between FRD (hitherto unknown to be associated with motility or flagella) and FliG of the flagellar switch, and we demonstrate that mutants lacking frd are defective in flagellar assembly and switching and are not responsive to fumarate. Results Fumarate does not bind to any of the known switch proteins We initiated this work by trying to determine whether fumarate binds to the switch complex. We isolated the intact switch complex of E. coli flagella (see Materials and methods and Supplementary Figure s2), incubated it with [14C]fumarate, and separated it from the medium by centrifugation. We detected no binding of [14C]fumarate (assayed in the range of 5–50 μM [14C]fumarate) to the isolated switch complex. We also measured the binding of [14C]fumarate to each of the three purified switch proteins. We used both equilibrium dialysis and centrifugal ultrafiltration, described in Materials and methods and Supplementary data, to measure binding of [14C]fumarate in the range 0.5–10 000 μM to each of the three switch proteins (10–200 μM). No binding was detected. Potential targets of fumarate binding to the flagellar switch The absence of detectable direct binding to any switch protein suggested that another protein may transmit the fumarate effect to the switch. This protein is expected to be membrane-bound because earlier it was shown that fumarate enhances switching even in envelopes devoid of cytoplasm (Barak and Eisenbach, 1992; Barak et al, 1996). Potential candidates for transmitting the fumarate effect to the switch are SDH and FRD—enzymes similar in amino-acid composition, 3D structure (Supplementary Figure s1), cofactors, and mechanisms of function (Cecchini et al, 2002). Both interact with fumarate and are membrane-bound. Thus, we tested the interaction of SDH and FRD with the switch. Evidence consistent with SDH binding to the switch complex was observed following incubation of [3H]SDH with the isolated intact switch complex and separation by centrifugation (Figure 1A; Kd=1.1±0.3 μM; mean±s.e.m.). To reduce nonspecific SDH binding, the experiment (detailed in Materials and methods) was carried out in the presence of at least 40-fold excess BSA, high salt concentration, and detergent. Insufficient amounts of the switch complex made direct binding measurements with an irrelevant protein impractical. Figure 1.SDH and FRD binding to the switch complex. (A) Concentration-dependent SDH binding to the isolated switch complex. See Materials and methods for details. The free SDH concentration was calculated by subtracting the concentration of bound SDH from the added SDH concentration. The data shown are normalized and derived from three different switch batches. The concentration of the switch was 0.1 mg protein/ml. The connecting line between the experimental points is a theoretical fit, performed by Origin computing program according to the Hill model (R2=0.90). (B) Protein A-gold particles bound to FRD in isolated extended flagellar basal bodies. The preparation of extended basal bodies was preincubated sequentially with FRD, anti-FRD antibody, and Protein A-gold. The number of gold particles counted was 1985. A total of 1265 basal bodies were counted; 715 were labelled at C-ring, M-ring, or between these rings, 87 were nonspecifically labelled in other parts of the basal body, and 463 were not labelled. The average number of gold particles associated with a basal body was 2.5 (the most common being 2 or 3 particles/basal body; range=1–10 particles/basal body). For the negative control with boiled FRD (not shown in the figure), a total of 382 basal bodies were counted: 3 (0.8%) were labelled at C- or M-ring, 4 (1%) were nonspecifically labelled in other parts of the basal body, and 375 (98%) were not labelled. Similar numbers were observed with BSA substituting for boiled FRD. (C) Image of an isolated extended flagellar basal body, treated as in (B) (bar=10 nm). (D) Protein A-gold particles bound to FRD in isolated basal bodies lacking the structure of the C-ring. Gold particles counted were 1619. Of the 1679 basal bodies counted, 1156 were labelled at the M-ring, 155 were nonspecifically labelled in other parts of the basal body, and 368 were not labelled. The average number of gold particles associated with such a basal body was 1.3 (the most common being 1 particle/body; range=1–6 particles/basal body). (E) Image of an isolated basal body lacking the structure of the C-ring, treated as in (B) (bar=10 nm). Download figure Download PowerPoint To examine FRD binding to the switch, we employed immuno-electron microscopy, which requires much less material and does not involve chemical modification of the enzyme. We isolated flagellar hook basal bodies (termed 'extended basal bodies'; Francis et al, 1994) and allowed them to bind FRD. (The basal body is the rotor of the flagellar motor, connected to the flagellar filament by means of a hook. It consists of a rod surrounded by coaxial rings: the M ring, a doublet ring embedded in the cytoplasmic membrane; the P ring, embedded in the peptidoglycan layer; and the L ring, embedded in the outer membrane (Figure 1E). Extended basal bodies also contain the C-ring (Figure 1C).) BSA was used as a control both for nonspecific binding/adsorption of the antibodies to the basal bodies and detection of protein A-Gold trapping by the basal body particles in the labelling protocol. Denatured FRD was used as a negative control. After binding, all the samples were incubated sequentially with anti-FRD antibody and protein A-Gold. FRD was localized to the C-ring (i.e., the switch complex) and, to a much lesser extent, to the M-ring (Figure 1B and C). Neither BSA nor denatured FRD was similarly localized. Thus, both SDH and FRD appear to bind to the switch complex. To substantiate FRD binding to the M-ring, we assessed by immuno-electron microscopy its binding to a preparation of basal bodies that lack the C-ring structure and do not contain the switch proteins FliM and FliN (Francis et al, 1994). However, they do contain FliG, as ∼80% of the basal bodies are labelled following anti-FliG immunolabelling (Supplementary Figure s3) and as previously demonstrated (Francis et al, 1992, 1994). FRD binding to this preparation was evident (Figure 1D and E). The observation that both the C-ring and M-ring bound FRD raised the possibility that the site of FRD and, probably, SDH binding was FliG. This is because FliG is the only switch protein that was present in both basal-body preparations and because it appears to be associated with both the C-ring and M-ring (Oosawa et al, 1994; Thomas et al, 2001). To explore this possibility, we employed three independent experimental approaches. (a) Following incubation with His-tagged FliG, biotinylated SDH and FRD were brought down by streptavidin–agarose beads and probed for the presence of FliG by western blotting with anti-His-tagged FliG antibody. His-tagged FliG colocalized with either SDH or FRD (Figure 2A). (b) Real-time surface plasmon resonance analysis showed that both purified SDH (Figure 2B) and FRD (Figure 2C), when individually immobilized onto the sensor surface, bound His-tagged FliG in a concentration-dependent manner. The dissociation rate constants, obtained on the basis of the Langmuir 1:1 bimolecular kinetic model, were similar for SDH and FRD (2 × 10−3 s−1), as were the association rate constants (3000 and 5000 M−1 s−1 for SDH and FRD, respectively) and the resulting dissociation constants (Kd=0.6±0.1 and 0.4±0.3 μM; mean±s.d. for SDH and FRD, respectively). Earlier tests showed that immobilized SDH bound neither FliM nor FliN (Prasad, 2001). The similarity between the Kd value obtained in this experiment for SDH binding to FliG and the Kd value for SDH binding to the isolated switch complex in Figure 1A strongly supports the argument that the SDH–switch binding in Figure 1A was specific. (c) Analytical ultracentrifugation was used to assess complex formation between His-tagged FliG and FRD. The shift in the location of the major protein peak from the molar mass of FRD to that of a 1:1 FliG–FRD complex when His-tagged FliG was present (Figure 2D) confirmed the formation of a complex and identified its stoichiometry. Figure 2.Binding of purified His-tagged FliG to purified FRD or SDH. (A) Pull-down assay. FliG was incubated with biotinylated FRD (lane 1), biotinylated SDH (lane 2), or, as a negative control, with neither FRD nor SDH (lane 3), precipitated with streptavidin–agarose beads, and probed with an anti-His-tagged FliG antibody. (B, C) Real-time interaction of His-tagged FliG with biotinylated SDH (B) or biotinylated FRD (C), measured by surface plasmon resonance. RU, resonance units. The values shown are after subtraction of the value of the empty channel. The kinetic and equilibrium constants, shown in the text, were obtained by a global fit, using the Langmuir 1:1 bimolecular kinetic model (A+B⇔AB; fit quality: χ2=5 and 11 for the SDH and FRD curves, respectively). (D) Analytical ultracentrifugation of the FliG-FRD complex. The c(M) distribution is a variant of the distribution of Lamm equation solutions using Sedfit (see Supplementary data). Three samples were run: FRD alone (2 μM; solid line), His-tagged FliG alone (16 μM; inset), and both together (2 μM each; dotted line). The major peak observed in the His-tagged FliG and FRD mixture (dotted line) was at 150 kDa (corresponding to a 1:1 FliG:FRD complex; 158 kDa theoretical value, calculated from the amino-acid composition). The changes of the sedimentation rates of the lesser peaks of FRD were minor in the presence of His-tagged FliG. The major peak of His-tagged FliG (inset) was at 32 kDa (37 kDa theoretical); the low peak at 75 kDa probably reflects a FliG dimer. The peaks of FRD (solid line) correspond, in the order of decreasing abundance, to FRD (134 kDa, consisting of all four subunits FrdA, FrdB, FrdC, and FrdD; 121 kDa theoretical), FrdA (71 kDa; 66 kDa theoretical), and FrdB (23 kDa; 27 kDa theoretical). FrdC and FrdD are not observed in the figure, as expected, due to their extreme hydrophobicity (Cecchini et al, 2002), which precludes them from the solution (either sedimenting to the bottom or adhering to the wall of the tube). Download figure Download PowerPoint The seminal issues, however, are whether this binding has physiological significance, and whether either or both SDH and FRD are responsible for the action of fumarate on the switch. FRD but not SDH affects the functions of FliG FliG is central to the function of the flagellar motor. It is known to be directly involved in motor assembly, torque generation, and switching (see, for a review, Kojima and Blair, 2004). Consequently, defects in this protein may lead to loss of flagella, paralysis, or biased direction of flagellar rotation. Assuming that the interaction of FliG with FRD or SDH is required for its function, absence of FRD or SDH may lead to any one of these phenotypes. Therefore, we prepared three deletion mutants: a Δfrd mutant, deleted for the genes encoding all the subunits of FRD; a Δsdh mutant in which two of the four genes encoding SDH were deleted, resulting in complete absence of SDH (Montrone, 1996; Prasad et al, 1998); and a double Δfrd Δsdh mutant. The Δsdh mutant did not differ from its wild-type parent with respect to motility (data not shown), whereas, strikingly, the Δfrd mutant and the double mutant were barely motile. As shown for the Δfrd mutant (Figure 3A), many cells did not swim at all, others swam more slowly than usual and, in most of these latter cases, the movement was wobbly. This behaviour resulted from a decrease in the number of flagella (Figure 3B and C). The wild-type parent had a median of 5 flagella/cell, but the Δfrd mutant had a median of only 1 flagellum/cell, with many cells having no flagella at all. Similar data (not shown) were obtained for the double mutant. To verify that the observed phenotype was due to the absence of FRD, we complemented the frd deletion with a plasmid, producing a single copy of FRD under its native promoter (pEWF1). The plasmid restored, at least partially, the number of flagella (median of 3 flagella/cell; Figure 3B and C) and increased the fraction of motile cells (Figure 3A). As the frd deletion did not affect the expression level of FliG, as evident from western blots with anti-FliG antibody (Supplementary Figure s4), the results suggest that FRD is required for normal flagellar assembly. Figure 3.Effects of frd and sdh deletions on swimming, assembly of flagella, and switching the direction of flagellar rotation. (A) Percentage of motile cells. Swimming cells were video-recorded and the fraction of motile cells was determined blindly. Values shown are the mean±s.e.m. of three experiments, 100–200 cells for each strain in each experiment. The asterisk indicates a statistically significant difference from the other columns (P<0.01 and P<0.05 for wild-type and Δfrd+pEWF1 columns, respectively; one-way ANOVA plus Tukey–Kramer tests). (B) Typical micrographs of strains RP437 (a; bar=1 μm), RP437Δfrd (b; 0.5 μm), and RP437Δfrd containing pEWF1 (c; 1 μm). (C) Distribution of the number of flagella per cell. Cells were negatively stained with uranyl acetate and photographed using a transmission electron microscope. The number of flagella of ∼200 cells of each strain were counted without prior knowledge of the strain being counted. The difference in the number of flagella between the strains was very significant (P<0.001; Krusal–Wallis test). Open columns, strain RP437; solid columns, RP437Δfrd; hatched columns, RP437Δfrd containing pEWF1. Download figure Download PowerPoint FRD deletion could potentially reduce the energy level and elevate the fumarate level in the cell, contributing to the observed phenotypes of the Δfrd mutant. No evidence for these scenarios was found. We measured the intracellular ATP concentration and found it to be similar in all the strains used, with a variation of ∼10% from the value of 2.5±0.1 mM measured for the wild-type strain. (It should be mentioned, however, that although removal of FRD did not have a significant effect on cellular ATP levels, its absence might have other, unexplored effects on cellular metabolism.) The possible contribution of an elevated fumarate level was ruled out earlier by showing that, in cells deleted of the genes encoding fumarase, elevated cellular fumarate has no effect on the motility level (Montrone et al, 1998; Prasad et al, 1998). To examine the effect of frd deletion on flagellar rotation of those cells that had residual flagella, we tethered them through a flagellum to a glass slide and monitored the rotation of the cells (Silverman and Simon, 1974). The tethered Δfrd cells rarely switched their direction of rotation, making a reversal once every 2.5 min on average (Table I, line 2) and rotating counterclockwise 99.9% of the time (line 1). The addition of the strong repellent benzoate increased the probability of clockwise rotation in both the Δfrd strain and its wild-type parent (line 3 versus line 1). However, in the Δfrd strain, the fraction of responsive cells was smaller (line 5), and the level of clockwise rotation achieved after this stimulation was significantly lower than that in the wild type (line 3). In contrast, the switching frequency was higher than that in the wild type (line 4), possibly reflecting futile attempts to switch to clockwise rotation. These results are consistent with the clockwise-promoting effect of fumarate in the intact cells of E. coli (Montrone et al, 1998; Prasad et al, 1998) and with the possibility, investigated below, that FRD is the protein that transmits the fumarate effect to the switch. Table 1. Comparison between wild-type and Δfrd strains with respect to the parameters of flagellar rotation Parameter testeda Wild type Δfrd P-value for the difference between the strainsb Before stimulation 1. Time fraction spent in clockwise rotation±s.e.m. (%) 2.0±0.9 0.1±0.1 0.25 2. Switching frequency±s.e.m. (min−1) 2.1±0.5 0.4±0.2 <0.009 After stimulation 3. Time fraction spent in clockwise rotation±s.e.m. (%) 70±4 13±4 <0.0001 4. Switching frequency±s.e.m. (min−1) 0.7±0.2 16±2 <0.0001 5. Fraction of responsive cells (%) 96 75 0.001 a The rotation of 46–59 tethered cells was monitored for periods of 40 s before and subsequent to stimulation with the repellent benzoate (50 mM). b Calculated by the Mann–Whitney test, except for the fraction of responsive cells, which was calculated by the Fisher's exact test. FRD transmits the effect of fumarate to the flagellar switch To examine this possibility, we employed two approaches. First, we studied the binding of [14C]fumarate to a preparation of M-rings associated with FliG (Figure 4A and B)—the minimal structure shown to bind FRD (Figure 1D and E)—in the presence and absence of FRD. The presence of FRD significantly increased the binding of fumarate to this preparation (Figure 4C). Next, we measured the effect of fumarate on flagellar rotation using tethered, cytoplasm-free envelopes containing His-tagged CheY. In intact bacteria, fumarate mainly increases the fraction of time spent in clockwise rotation (Montrone et al, 1998; Prasad et al, 1998). In envelopes, fumarate enables switching: tethered envelopes require this dicarboxylate to switch from one direction of rotation to the other (Barak and Eisenbach, 1992; Barak et al, 1996). If fumarate binding to the switch complex depends on FRD, envelopes prepared from the Δfrd mutant should be incapable of switching either in the presence or absence of fumarate. The results confirmed this prediction. Although fumarate was present, none of the Δfrd envelopes switched directions, rotating exclusively in one direction, mostly counterclockwise. In contrast, wild-type envelopes showed switching (Figure 4D). These results (Figure 4C and D), taken together with the observation that Δsdh mutant envelopes continue to switch when fumarate is added (Barak et al, 1996), suggest that, FRD, but not SDH, transmits the fumarate effect to the switch and that FRD may be a functional component of the switching mechanism of E. coli. Figure 4.FRD transmits the effect of fumarate to FliG at the switch–motor complex. (A) Electron micrographs of purified M-rings. Bar=50 nm. (B) SDS gel of the M-rings (composed of the protein FliF), demonstrating the co-occurrence of FliG. (C) Binding of fumarate to the M-ring in the presence and absence of FRD. Fumarate binding was measured with 50 μM [14C]fumarate and 20 μM FRD. The concentration of the M-ring proteins was 10 μM. The data shown are the means (±s.e.m.) of 3–4 repetitions. The asterisk indicates a statistically significant difference (P<0.04, unpaired t-test). (D) Effect of frd deletion on the rotation of tethered, CheY-containing, cytoplasm-free envelopes in the presence of fumarate at room temperature. The figure shows the distribution of envelopes according to their direction of rotation. Open columns, counterclockwise-rotating cells; hatched columns, clockwise-rotating cells; solid column, switching cells. The numbers of monitored envelopes were 115 and 39 for the wild-type strain and the Δfrd mutant, respectively. Download figure Download PowerPoint Discussion In this study, we have made three important discoveries, none of which were predictable on the basis of pre-existing knowledge. First, we demonstrated that FRD is important for the function of the switch and appears to form a dynamic complex with it. This conclusion is based on the observations that (a) absence of FRD results in a sharp decrease in the number of flagella per cell and, consequently, in cell motility (Figure 3), without affecting the level of FliG in the cell (Supplementary Figure s4); (b) the ability of existing flagella to switch rotation to the clockwise direction is much reduced in the absence of FRD (Table I); and (c) FRD forms a 1:1 complex with the switch protein FliG (Figures 1 and 2). These observations thus demonstrate that FRD is involved in both flagellar assembly and switching to clockwise rotation. Second, we identified FRD as the target binding site of fumarate at the switch, enabling fumarate to exert its stimulating effect on clockwise switching (Figure 4). Third, we have demonstrated that FRD has an important role under aerobic conditions in E. coli, unassociated with ATP production. All the results shown were obtained under aerobic conditions, where FRD is minimally expressed (∼4% of the expression level compared to anaerobic conditions; Jones and Gunsalus, 1985) and, according to the current understanding, it has no essential function (Cecchini et al, 2002). Furthermore, although FRD, similar
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