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1 Department of Biology and
2 Howard Hughes Medical Institute, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, USA
Reprint requests to: Robert T. Sauer, Department of Biology, Massachusetts Institute of Technology, Cambridge, MA 02139, USA
(RECEIVED October 18, 2002; FINAL REVISION February 5, 2003; ACCEPTED February 6, 2003)
Article and publication are at http://www.proteinscience.org/cgi/doi/10.1110/ps.0237603.
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Abstract |
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S by ClpX were measured with different protein partners and the kinetics of degradation of ssrA-tagged substrates were determined with both nucleotides. ClpX hydrolyzed ATPS to ADP and thiophosphate at a rate (6/min) significantly slower than ATP hydrolysis (140/min), but the hydrolysis of both nucleotides was increased by ssrA-tagged substrates and decreased by ClpP. KM and kcat for hydrolysis of ATP and ATPS were linearly correlated over a 200-fold range, suggesting that protein partners largely affect kcat rather than nucleotide binding, indicating that most bound ATP leaves the enzyme by hydrolysis rather than dissociation, and placing an upper limit of 
15 µM on KD for both nucleotides. Competition studies with ClpX and fluorescently labeled ADP gave inhibition constants for ATPS (2 µM) and ADP (3 µM) under the reaction conditions used for steady-state kinetics. In the absence of Mg2+, where hydrolysis does not occur, the inhibition constant for ATP (55 µM) was weaker but very similar to the value for ATPS (45 µM). Compared with ATP, ATPS supported slow but roughly comparable rates of ClpXP degradation for two Arc-ssrA substrates and denatured GFP-ssrA, but not of native GFP-ssrA. These results show that the processing of protein substrates by ClpX is closely coupled to the maximum rate of nucleotide hydrolysis. Keywords: AAA+ ATPase; catalyzed protein unfolding; ATP-dependent proteolysis; motor proteins; mantADP
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Introduction |
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Studies of protein unfolding by ClpX and/or degradation by ClpXP have been facilitated by the finding that the C-terminal ssrA-tag sequence (AANDENYALAA) provides a binding site for ClpX, and therefore any ssrA-tagged protein becomes a potential ClpX substrate (Levchenko et al. 1997; Gottesman et al. 1998; Lee et al. 2001; Wah et al. 2002). For example, green fluorescent protein with an ssrA tag (GFP–ssrA) is degraded by ClpXP in an ATP-dependent reaction in which protein denaturation is the rate-determining step (Kim et al. 2000). If the protease active sites of ClpXP are inactivated, then GFP–ssrA is denatured and translocated into the proteolytic chamber, where it is trapped in an unfolded state. Spontaneous denaturation of GFP–ssrA is exceedingly slow compared to ClpX-catalyzed unfolding, suggesting that ClpX-applied forces lower the protein denaturation barrier during the enzymatic reaction (Kim et al. 2000). Similar conclusions emerge from the study of ClpXP degradation of stability variants of ssrA-tagged Arc repressor (R.E. Burton et al. 2001). Although these Arc mutants varied over a 107-fold range in terms of their thermodynamic stabilities and spontaneous unfolding rates, ClpXP degraded the set of Arc proteins at rates differing only by 50%.
Roughly 150 molecules of ATP are hydrolyzed during ClpXP degradation of a single molecule of Arc–ssrA, suggesting that many cycles of ATP binding and hydrolysis are needed for efficient substrate unfolding and/or translocation (R.E. Burton et al. 2001). However, although ClpXP is one of the better characterized ATP-dependent proteases, relatively little is known about the linkage between ATP binding or hydrolysis and the protein unfolding and translocation steps that are required for substrate degradation. In this paper, we have determined the kinetic parameters for steady-state hydrolysis of ATP and ATPS by ClpX, ClpXP, or ClpX plus ssrA-tagged substrates. Depending upon the nucleotide and associated proteins, the maximum ClpX-catalyzed hydrolysis rates were found to vary over a 200-fold range. Covariation of kcat and KM in these experiments suggests that ATP and ATPS have roughly the same affinity for ClpX irrespective of its associated protein partners or substrates. At saturating nucleotide concentrations, ATPS hydrolysis was 20- to 40-fold slower than ATP hydrolysis. Relative to ATP, ATPS supported ClpXP-catalyzed processing and degradation of ssrA-tagged Arc substrates and denatured GFP–ssrA at significantly reduced rates. These results provide evidence for strong linkage between the rate of nucleotide hydrolysis and the rate of ClpXP-catalyzed substrate processing. No degradation of native GFP–ssrA by ClpXP could be detected using ATPS. This result suggests that the maximum rate of nucleotide hydrolysis is also linked to the unfolding force that ClpX can generate, setting a threshold for allowable substrates in terms of protein stability.
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Results |
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SS supports hexamerization of ClpX (Grimaud et al. 1998), promotes ClpX binding to ssrA-tagged substrates (Singh et al. 2000; Wah et al. 2002), and is hydrolyzed by ClpX (Zhou et al. 2001), although experiments demonstrating this last result have not been presented. It is not known whether ClpXP can unfold and degrade substrates in the presence of ATPS. To test directly for hydrolysis of this ATP analog by ClpX, we collected a 31P-NMR spectrum of a sample containing 5 mM ATPS and then added 0.3 µM ClpX6 and collected spectra at intervals over a total of 21 h. Figure 1A
shows the initial and 21-hour spectra, revealing a time-dependent loss of ATPS-specific NMR signals and concomitant appearance of ADP-specific NMR signals. At each time point, the sum of the integrated intensities of the ATPS-specific and ADP-specific NMR signals were relatively constant (Fig. 1B), suggesting that the principal hydrolysis reaction catalyzed by ClpX was cleavage of the ß- phosphodiester bond of ATPS to produce ADP and PO3S, although the NMR signal for the latter species was obscured by the resonance from the -phosphate of ATPS. The hydrolysis of ATPS observed in the presence of ClpX in this experiment was significantly faster than the rate of spontaneous hydrolysis determined under comparable conditions (see below).
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S, the radioactive hydrolysis products produced in the presence of ClpX were characterized by thin-layer chromatography (Fig. 2). In this assay, 35S-labeled ATPS chromatographed as a single radioactive species that barely moved from the origin. Unlabeled ATPS chromatographed at the same position. Upon incubation with ClpX, a faster migrating radioactive species appeared in a time-dependent fashion (Fig. 2). This latter species was identified as PO3S by the criterion that chromatography of unlabeled Na3PO3S, followed by visualization by starch–iodide staining, gave a spot at the same migration position. In this experiment, the ClpX-catalyzed hydrolysis rate was roughly 200-fold faster than spontaneous ATPS hydrolysis. Taken together, the NMR and thin-layer chromatography experiments demonstrate that ClpX catalyzes hydrolysis of ATPS to ADP and PO3S.
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S and ATPS and the thin-layer chromatography assay or by using ATP and a coupled spectrophometric assay. In each case, the resulting data fit well to the Michaelis-Menten equation (Fig. 3), allowing determination of kcat and KM values. For ClpX-mediated hydrolysis of ATPS, kcat was 6.0 x 0.1/min per ClpX6 enzyme and KM was 13 x 1.3 µM. By contrast, at saturating nucleotide concentrations, ClpX-mediated hydrolysis of ATP was roughly 20-fold faster (kcat = 140 x 5/min per ClpX6) but the KM value for ATP (190 x 30 µM) was also almost 15-fold higher.
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S and ATP hydrolysis by ClpX were also measured in the presence of its proteolytic partner, ClpP, or in the presence of ssrA-tagged protein substrates. When ClpP was present, kcat for hydrolysis of ATPS by ClpX was reduced 3.5-fold and kcat for hydrolysis of ATP was reduced 2.3-fold (Fig. 3
; Table 1). Reductions in KM of similar magnitudes were also observed for both nucleotides in these experiments (Fig. 3; Table 1). In the presence of Arc–PL8–ssrA or GFP–ssrA, increases in both kcat and KM for ClpX-catalyzed hydrolysis of ATPS and ATP were observed, albeit to different extents (Fig. 3; Table 1). Changes in the ATPase activity of ClpX in the presence of ClpP- or ssrA-tagged substrates have been reported previously (R.E. Burton et al. 2001; Kim et al. 2001) and provide evidence that the observed nucleotide hydrolysis is catalyzed by ClpX and not by some minor contaminant in the enzyme preparation.
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S was 20- to 40-fold lower than kcat for hydrolysis of ATP. We were initially surprised, however, by the observation that ATPS hydrolysis rates saturated at much lower nucleotide concentrations than ATP hydrolysis rates, as reflected in the 14- to 30-fold lower KM values for ATPS relative to ATP (Table 1). Moreover, for both nucleotides, addition of ClpP- or ssrA-tagged substrates consistently resulted in changes in the same direction and of roughly similar magnitude in both kcat and KM. The plot in Figure 4 shows that KM varies in a near linear fashion with kcat over a 200-fold range. In the standard definition,
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 | (1) |
For kcat and KM values measured for each nucleotide and set of conditions, KM will be a linear function of kcat if the values of ka (the association rate constant) and Kd (the dissociation equilibrium constant) are the same for both nucleotides and do not change with different protein partners. The solid line shown in Figure 4 is a fit (R2 = 0.99) to equation 1 using a ka value of 0.8 x 0.1 µ/M/min (1.3•104/M/s) and a Kd value of 10 x 3 µM. Hence, the experimental results are consistent with a model in which nucleotide binding to ClpX is roughly the same for ATP and for ATPS, irrespective of ClpX’s associated partners or protein substrates.
How reliable are the ka and KD values derived from the analysis presented above? The fitted ka values were 0.8 x 0.2 µ/M/min and 0.2 x 0.1 µ/M/min when the steady-state hydrolysis data were fitted just for ATP or ATPS, respectively. In these individual fits, however, the errors in KD were comparable to, or greater than, the fitted values (7 x 7 µM). This allows us to place an upper limit of 15 µM on the KD’s for ATP and ATPS binding, but these values could be significantly lower. Given ka and having an upper limit for KD places an upper limit on the dissociation rate constant (kd) of 12/min for ATP and 3/min for ATPS. For ATP hydrolysis, this maximum kd value is still significantly less than the measured kcat values, indicating that most ATP molecules that bind to ClpX are committed to hydrolysis.
If the model discussed above for the co-variation of kcat and KM is correct, then binding measurements should yield nucleotide binding affinities of 15 µM or less under the conditions used for the steady-state kinetic measurements. To test this prediction, we first measured equilibrium binding of ClpX to the fluorescent ADP analog mantADP (2'(or-3')-O-(N-methylanthraniloyl)adenosine 5'-diphosphate), and then used competition assays to determine apparent binding affinities of ClpX for ADP and ATPS (Fig. 5). ClpX binding enhanced the fluorescence emission of mantADP by approximately 2.2-fold and caused a small blue shift in the spectrum (Fig. 5A). Data obtained from the titration of ClpX against constant mantADP were fit well by a model in which each ClpX monomer bound one nucleotide with an affinity of 7 x 1 µM. Fits of the data from the competition experiments shown in Figure 5B gave apparent binding constants of 1.6 x 0.4 µM for ATPS and 3.2 x 0.8 µM for unmodified ADP.
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S and ATP to ClpX was measured by competition (Fig. 5C). The inhibition constants for ATP (52 x 13 µM) and ATPS (44 x 5 µM) were within error under these conditions, supporting the idea that ClpX does not discriminate between these nucleotides at the binding step. These experiments do not rule out the possibility that Mg2+ stabilizes ATP binding more than ATPS binding, but, in other work, we have found that ATP and ATPS bound with indistinguishable affinities to a ClpX mutant with undetectable ATPase activity in the presence of Mg2+ (S. Joshi, R.E. Burton, T.A. Baker, and R.T. Sauer, in prep.).
ATPS supports selective protein denaturation and degradation
The ability of ClpX to hydrolyze ATPS provided an opportunity to test whether this nucleotide also enabled ClpX to denature and/or translocate substrates to ClpP for degradation. ATPS supported ClpXP degradation of 35S-labeled Arc–PL8–ssrA (Arc–ssrA with a Pro8 
Leu substitution) as assayed by the release of TCA-soluble radioactivity (see Fig. 6A, inset). Figure 6A shows the observed degradation rate as a function of substrate concentration. The kcat value (0.09 x 0.01/min) for degradation of Arc–PL8–ssrA by ClpXP in the presence of ATPS was roughly 14-fold lower than the value (1.3 x 0.1/min) determined previously in the presence of ATP (R.E. Burton et al. 2001), but KM for this protein substrate was similar in the presence of ATPS (0.9 x 0.1 µM) and ATP (0.7 x 0.1 µM). No degradation was observed in the absence of nucleotide, in the absence of ClpP, or in the absence of ClpX (data not shown). For Arc–PL8–ssrA, the reduction in the degradation rate in the presence of ATPS versus ATP was comparable to the reduction in the rate of nucleotide hydrolysis, suggesting coupling between nucleotide hydrolysis and substrate unfolding and/or translocation. By contrast, no ClpXP degradation of GFP–ssrA was detected in the presence of ATPS (Fig. 6A), even though GFP–ssrA and Arc–PL8–ssrA were degraded at roughly similar rates in the presence of ATP (Kim et al. 2000; R.E. Burton et al. 2001). This result shows that nucleotide hydrolysis per se is not sufficient for ClpXP degradation of all protein substrates.
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S might not support ClpXP-catalyzed degradation of GFP–ssrA because ClpXP•ATPS does not bind this substrate, cannot denature native GFP–ssrA, or cannot translocate unfolded GFP–ssrA into ClpP. To test these possibilities, we acid denatured GFP–ssrA prior to addition of ClpXP and ATPS. As shown in Figure 6B, acid-denatured 35S-labeled GFP–ssrA was degraded by ClpXP in the presence of ATPS, although with a distinct lag of several minutes and a steady-state rate roughly 20-fold slower than the rate determined using ATP. There was a lag time of ~2 min in the appearance of peptides when denatured GFP–ssrA was degraded in the presence of ATPS, which could be modeled by assuming a binding step followed by two consecutive slow steps (e.g., substrate engagement and translocation) with rate constants of 0.2/min prior to rapid peptide bond cleavage. Clearly, however, ATPS supported the binding of denatured GFP–ssrA to ClpX and its translocation into ClpP for degradation. ATPS must also allow the binding of native GFP–ssrA, as the latter protein stimulated hydrolysis of ATPS by ClpX (Fig. 3; Wah et al. 2002). Taken together, these experiments indicate that ATPS is unable to support ClpX-mediated denaturation of native GFP–ssrA.
Does ATPS actually support ClpX-catalyzed denaturation of Arc–PL8–ssrA or does the observed degradation depend upon the spontaneous denaturation and trapping of this substrate by ClpXP? To help address this question, we assayed degradation of the Ile37 Val (Arc–IV37–ssrA) variant of Arc–ssrA (Fig. 6C), which is substantially less stable than Arc–PL8–ssrA (R.E. Burton et al. 2001). The Arc–IV37–ssrA mutant was roughly 13% unfolded under the assay conditions used, compared with 0.6% unfolded for the PL8 mutant. As shown in Figure 6C, both Arc–ssrA variants were degraded by ClpXP•ATPS at similar rates. Because the unfolding and refolding of these variants are highly dynamic processes (Milla et al. 1995; Schildbach et al. 1995), these data suggest either that ClpX can actively unfold these molecules using ATPS or that ClpX traps the unfolded forms of these proteins in an irreversible fast step with some subsequent step such as translocation being rate limiting. We favor the former model but cannot rigorously exclude the latter model. In either case, although ClpXP can clearly degrade some proteins using ATPS hydrolysis as an energy source, its capacity to actively unfold protein substrates is significantly diminished.
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Discussion |
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TOPAbstractIntroductionResultsDiscussionMaterials and methodsReferences |
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S in a reaction that is stimulated by ssrA-tagged substrates and suppressed by ClpP; (2) kcat and KM for hydrolysis of ATP and ATPS co-vary in a fashion expected if the binding of protein partners to ClpX largely affects kcat and not the rate constants or equilibrium constant for nucleotide binding; (3) ClpX binds ATPS and ADP with apparent affinities in the low µM range, under standard assay conditions, and binds ATP and ATPS with essentially identical affinities in the absence of Mg2+; (4) ATPS supports ClpXP-mediated denaturation, translocation, and degradation of some, but not all, protein substrates; and (5) the reduced hydrolysis rate for ATPS compared with ATP correlates with reduced rates of substrate processing by ClpXP. These findings provide a foundation for beginning to understand the mechanism by which ClpX transforms the chemical energy of nucleotide binding and/or hydrolysis into the mechanical forces required to unfold protein substrates and translocate them into ClpP for degradation. ATP-dependent motor proteins, such as myosin and kinesin, also generate force and molecular movement. To maintain tight coupling between ATP consumption and motility, motor proteins interleave the chemical and mechanical steps in the overall reaction cycle (Jencks 1989). With myosin, for example, ATP binding to myosin catalyzes release from the actin filament, which stimulates ATP hydrolysis, which allows actin rebinding, which permits release of inorganic phosphate and ADP, which drives the power stroke, and so on. Kinesin appears to operate by a similar mechanism although conformational changes occur during different parts of the ATPase cycle (Rice et al. 1999). Myosin performs its "power stroke" following release of inorganic phosphate and then resets its conformation when ADP is exchanged for ATP (Suzuki et al. 1998). Kinesin applies its power stroke following ATP binding and resets after phosphate release (Rice et al. 1999).
How similar is ClpX to well-characterized motor proteins such as myosin? For myosin, ATP binding and actin binding are competitive; ATP binding causes actin dissociation. For ClpX, the binding of ATP and of ssrA-tagged substrates are not competitive. Although Km for ATP hydrolysis increases when ssrA-tagged substrates bind ClpX, this increase is modest (twofold) and is caused by the increase in kcat. Moreover, in the presence of ATPS, stable binding of ClpX binds to a peptide with the SsrA tag sequence with a KD (2.5 µM) similar to the KM (1–2 µM) of ClpXP-catalyzed degradation of ssrA-tagged substrates (Wah et al. 2002). Myosin, by itself, is a relatively poor steady-state ATPase (1.2/min) because ADP and inorganic phosphate (Pi) are released very slowly; addition of actin accelerates the release rate enormously and increases ATP hydrolysis to >1000/min. This results in efficient coupling of ATP hydrolysis and actin-mediated force generation. ClpX, by contrast, is a robust ATPase (140/min) even in the absence of substrate proteins, but >100 molecules of ATP are hydrolyzed during degradation of a single molecule of substrate (R.E. Burton et al. 2001). Although these differences do not preclude the possibility that ClpX functions like a molecular motor in some ways, they emphasize that significant differences in the coupling efficiency between the chemical and mechanical cycles must exist.
Although ATPS is hydrolyzed by ClpX, kcat is reduced by 20- to 40-fold compared with ATP hydrolysis. It is not currently known what steps determine kcat for ClpX hydrolysis for either ATP or ATPS. Possibilities include breaking the phosphate anhydride bond, dissociation of inorganic phosphate or PO3S, or a conformational change in ClpX that accompanies one of these steps. Although ADP release could be rate limiting for ATP hydrolysis, another step would then have to become rate limiting for ATPS hydrolysis to explain the different rates of ClpX-catalyzed hydrolysis of these nucleotides.
The significant differences in ClpXP-catalyzed hydrolysis of ATP and ATPS allowed us to probe the coupling between nucleotide hydrolysis and substrate processing rates. The slower hydrolysis of ATPS clearly reduced the substrate processing activity of ClpXP. Arc–PL8–ssrA was degraded roughly 14-fold more slowly by ClpXP when ATPS was substituted for ATP, whereas a 35-fold reduction in hydrolysis rates was observed. Peptide-bond hydrolysis by ClpP is known to be a very fast step in the overall degradation reaction (Grimaud et al. 1998). In principle, the reduction in the rate of substrate processing using ATPS could reflect slowing of just the denaturation rate or just the translocation rate. However, changes in both processing steps seem more probable given that native GFP–ssrA could not be denatured by ClpXP•ATPS but was readily denatured by ClpXP•ATP (Kim et al. 2000). Moreover, degradation of acid-denatured GFP–ssrA by ClpXP•ATPS was also much slower than degradation by ClpXP•ATP. Because the available data suggest that ATP and ATPS have similar affinities for ClpXP, these changes in the protein denaturation and translocation rates are probably caused by changes in kcat for hydrolysis of these respective nucleotides.
Once ATP binds to ClpX, it is largely committed to hydrolysis; the experiments presented here establish an upper limit for ATP loss by dissociation (kd 
12/min) whereas loss via hydrolysis occurs >10-times faster (kcat = 140/min). For ATPS, however, the maximum dissociation rate constant (kd 3/min) could be significant in comparison with the hydrolysis rate constant (6/min). Hence, some bound ATPS might dissociate from the ClpX prior to hydrolysis, potentially reducing the energy available for ClpX to catalyze substrate processing. One possibility is that the sulfur atom on the terminal phosphate of ATPS perturbs the hydrolysis cycle at the point where force is generated. Studies of kinesin (Rice et al. 1999) and myosin (Suzuki et al. 1998) have shown that significant conformational changes occur upon Pi release, generating mechanical force or resetting the conformation of the motor protein. If the ATPS thiophosphate disrupts the timing of unfolding or translocation events, then protein substrates may escape before being committed to unfolding and/or translocation.
GFP–ssrA is more thermodynamically stable than Arc–PL8–ssrA (Kim et al. 2000; R.E. Burton et al. 2001) and the inability of ClpXP•ATPS to denature the former protein suggests that the maximal rate of nucleotide hydrolysis affects the amount of unfolding force that can be applied. If this is true, then the ATPase activities of other Clp/HSP100 family members may be linked to their abilities to denature or disassemble hyper-stable proteins. For the E. coli enzymes, ClpX and ClpA both hydrolyze ATP at rates in excess of 100/min (Singh et al. 1999), whereas HslU (ClpY) and ClpB have basal activities of roughly 30/min (Yoo et al. 1996; Li and Sha 2002). It will be interesting to see whether the latter enzymes are poorer protein unfoldases. Chaperones that do not appear to actively unfold protein substrates have even lower basal ATPase rates. GroEL hydrolyzes ATP at 5/min (Todd et al. 1993). DnaK has a very low activity (0.2/min) that is increased to 10/min by DnaJ and GrpE (Liberek et al. 1991). Analysis of different myosin-II family members suggests that alterations in the ATPase cycle can modulate the maximum motility rate by changing the time that myosin remains strongly bound to actin (Spudich 1994). A similar correspondence may exist for Clp family members, where individual enzymes have evolved ATPase activities that are matched to support their biological functions.
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Materials and methods |
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Protein expression and purification
Arc–PL8–ssrA, Arc–IV37–ssrA, GFP–ssrA, and ClpPHis6 were expressed and purified according to published protocols (R.E. Burton et al. 2001). ClpX was purified from an overexpression strain [HMS174(DE3)/pLysS] carrying the clpX gene on a pET3a vector (Levchenko et al. 1995). Cells were grown at 25°C in TB medium (20 g tryptone, 20 g yeast extract, 8 g NaCl, 2 g Na2HPO4, and 1 g KH2PO4 per L) in a 12-L Microferm fermentor (New Brunswick Scientific) to an OD600 of 2.2. ClpX expression was initiated by addition of IPTG to a final concentration of 0.25 mM. Cells were harvested by centrifugation 2 h after induction and were stored as a frozen suspension in a minimal amount of lysis buffer. The cell paste was thawed and diluted to 7 mL of lysis buffer per gram of wet cells. Cell lysis was achieved by addition of 0.2 mg/mL lysozyme, followed by sonication on ice (3 x 1-min pulses at maximum power). Cell debris was removed by centrifugation at 15,000 rpm for 60 min in a Sorvall S-600 rotor, and the supernatant was centrifuged at 40,000 rpm for 1 1 in a Beckman Ti50.2 rotor. Solid (NH4)2SO4 was added to the cleared lysate to 35% saturation and the mixture was left on ice for 1 h, followed by centrifugation at 40,000 rpm for 1 h. The pellet was resuspended in one-tenth of the original volume of lysis buffer and applied to a 14-mL hydroxyapatite column. The column was washed with 2 volumes of lysis buffer and a linear gradient from lysis buffer to 0.3 M NaHPO4 (pH 7.2) and 5 mM DTT was run. ClpX-containing fractions were identified by SDS-PAGE and concentrated by (NH4)2SO4 precipitation, and the resolubilized pellet was loaded onto a Sephacryl-300 column equilibrated in lysis buffer. ClpX-containing fractions were pooled and loaded onto a Q-Sepharose column, and a linear gradient from 0.1 to 1 M KCl in lysis buffer was run. Fractions containing ClpX were desalted into lysis buffer using a PD-10 column, and aliquots were stored frozen at -80°C.
Nucleotide hydrolysis assays
ATP hydrolysis at 30°C was assayed using a coupled spectrophotometric assay (Nørby 1988) in PD buffer plus 20 U/mL pyruvate kinase, 20 U/mL lactate dehydrogenase, 7.5 mM phosphoenolpyruvate, and 1 mM NADH. KCl was added to each reaction to maintain a constant ionic strength of 48 mM. ATPS hydrolysis at 30°C was measured using trace amounts of 35S-labeled ATPS, followed by thin-layer chromatography. Reactions were identical to the ATP hydrolysis reactions, except the components needed for the coupled assay were omitted. Aliquots were removed at time points and quenched by mixing with 2.5 volumes of 50 mM Tris•Cl (pH 7.5), 100 mM EDTA, 20 mM ATPS, and 20 mM PO3S. One microliter of each quenched sample was spotted onto a plastic-backed PEI-cellulose sheet (EM Science) and chromatographed at 4°C in 1.5 M formic acid and 0.4 M LiCl. Following chromatography, the plates were dried, and the radioactivity in each spot was quantified using a Molecular Dynamics PhosphorImager.
A potential complicating factor in interpreting the ATPase activity of ClpX as a function of nucleotide concentration is that ClpX hexamers are less stable in the absence of nucleotide (Grimaud et al. 1998). However, the plots in Figure 3 were fit well by a simple Michaelis-Menten relationship (r > 0.96), suggesting either that ClpX remains hexameric at the nucleotide concentrations studied or that the ATPase activity of the lower-order oligomers is similar to that of the hexamer. In either case, the precise distribution of ClpX oligomers is not critical for the analysis presented here.
Degradation assays
Degradation of ssrA-tagged substrates at 30°C was performed in PD buffer with 5 mM ATPS and 18 mM KCl to match the ionic strength of the nucleotide hydrolysis experiments. 35S-labeled protein was prepared by growing expression strains in minimal media containing 35S-methionine as described previously (R.E. Burton et al. 2001). The concentrations of Arc–ssrA protein stocks were determined by UV absorbance (
280 = 8250/M/cm), whereas GFP–ssrA concentrations were measured by visible absorbance (500 = 54000/M/cm). Reactions were initiated by adding 35S-labeled substrate to a mixture of ClpX, ClpP, ATPS, and buffer that had been pre-equilibrated for 2 min at 30°C. Aliquots were removed at time points, quenched with 1/4 volume 50% trichloroacetic acid, incubated on ice for 20 min, and then spun at 13,000 rpm for 20 min at 4°C in a Heraeus Biofuge 13 centrifuge. The concentration of TCA-soluble peptides was determined by liquid scintillation counting of the supernatant.
31P-NMR spectroscopy
NMR samples were prepared in 50 mM Tris•Cl (pH 7.5), 200 mM KCl, 5 mM MgCl2, 5 mM DTT, 10 mM sodium 3-trimethylsilylproprionate (TMSP), 10% glycerol, and 10% D2O. Samples (3 mL) were loaded into 10-mm diameter NMR tubes and spectra were taken using a Bruker DMX 500 MHz spectrometer. NMR spectra were recorded at 30°C, and referenced indirectly using the TMSP resonance together with the known frequency ratio of 31P to 1H (Wishart et al. 1995).
ClpX binding to mantADP and competition experiments
ClpX binding to mantADP at 30°C was monitored by the change in fluorescence emission at 445 nm after excitation at 357 nm. For mantADP binding, 30 µL samples were prepared with PD buffer plus 48 mM KCl, 1 µM mantADP, and varying concentrations of ClpX. Competition experiments were performed in PD buffer using 12 µM total ClpX (monomer equivalents), 1 µM mantADP, and varying concentrations of ADP or ATPS. KCl was added to each sample to bring the ionic strength to 48 mM above that of PD buffer alone. Binding experiments in the absence of Mg2+ were performed at 30°C in a buffer containing 25 mM HEPES•KOH (pH 7.6), 5 mM KCl, 5 mM EDTA, 0.032% NP-40, and 10% glycerol.
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References |
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(open circles) and ß (open squares) phosphates of ATP
