Báo cáo khoa học: Conserved residues in the N-domain of the AAA+ chaperone ClpA regulate substrate recognition and unfolding pdf

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Conserved residues in the N-domain of the AAA+chaperone ClpA regulate substrate recognition andunfoldingAnnette H. Erbse1,*, Judith N. Wagner1,*, Kaye N. Truscott2, Sukhdeep K. Spall2, Janine Kirstein1,3,Kornelius Zeth4,Ku¨rsad Turgay1,3, Axel Mogk1, Bernd Bukau1and David A. Dougan1,21 Zentrum fu¨r Molekulare Biologie Heidelberg, Universita¨t Heidelberg, Heidelberg, Germany2 Department of Biochemistry, La Trobe University, Melbourne, Australia3 Institut fu¨r Biologie, Freie Universita¨t Berlin, Berlin, Germany4 MPI fu¨r Entwicklungsbiologie, Tubingen, GermanyThe AAA+ superfamily [1] is an extensive group ofproteins involved in a broad range of biological func-tions. Its members are present in all kingdoms of lifeand often play a crucial role in cell maintenance. Inbacteria, several AAA+ proteins (e.g. ClpA, ClpB,ClpX, HslU and Lon) are central to the protein qual-ity-control network [2]. They employ a common mech-anism, involving the binding and hydrolysis of ATP,to mediate the unfolding ⁄ disassembly of a variety ofproteins, including large macromolecular complexes[3]. Although several of these proteins share consider-able sequence similarity, they demonstrate distinctsubstrate specificity. For example, in Escherichia coli,ClpA is responsible, either directly or indirectly via theadaptor protein ClpS, for recognition of substratessuch as SsrA-tagged proteins or N-end rule substratesKeywordsAAA+; binding; ClpA; SsrA; unfoldingCorrespondenceD. A. Dougan, Department of Biochemistry,La Trobe University, Melbourne 3086,AustraliaFax: +61 3 9479 2467Tel: +61 3 9479 3276E-mail: d.dougan@latrobe.edu.auB. Bukau, Zentrum fu¨r Molekulare BiologieHeidelberg, Universita¨t Heidelberg, INF 282,Heidelberg D-69120, GermanyFax: +49 6221 54 5894Tel: +49 6221 54 6795E-mail: bukau@zmbh.uni-heidelberg.de*These authors contributed equally to thiswork(Received 22 November 2007, revised 10January 2008, accepted 14 January 2008)doi:10.1111/j.1742-4658.2008.06304.xProtein degradation in the cytosol of Escherichia coli is carried out by avariety of different proteolytic machines, including ClpAP. The ClpA com-ponent is a hexameric AAA+ (ATPase associated with various cellularactivities) chaperone that utilizes the energy of ATP to control substraterecognition and unfolding. The precise role of the N-domains of ClpA inthis process, however, remains elusive. Here, we have analysed the role offive highly conserved basic residues in the N-domain of ClpA by monitor-ing the binding, unfolding and degradation of several different substrates,including short unstructured peptides, tagged and untagged proteins. Inter-estingly, mutation of three of these basic residues within the N-domain ofClpA (H94, R86 and R100) did not alter substrate degradation. In contrastmutation of two conserved arginine residues (R90 and R131), flanking aputative peptide-binding groove within the N-domain of ClpA, specificallycompromised the ability of ClpA to unfold and degrade selected substratesbut did not prevent substrate recognition, ClpS-mediated substrate deliveryor ClpP binding. In contrast, a highly conserved tyrosine residue lining thecentral pore of the ClpA hexamer was essential for the degradation of allsubstrate types analysed, including both folded and unstructured proteins.Taken together, these data suggest that ClpA utilizes two structural ele-ments, one in the N-domain and the other in the pore of the hexamer, bothof which are required for efficient unfolding of some protein substrates.AbbreviationsAAA+, ATPase associated with various cellular activities; FITC, fluorescein isothiocyanate; GFP, green fluorescent protein; kR, lambdarepressor.1400 FEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBS[4,5]. Once recognized, these substrates are unfoldedby the AAA+ protein, in an ATP-dependent manner,and translocated through the central pore of the oligo-mer into the associated ClpP peptidase, where they aredegraded into short peptides.AAA+ proteins usually contain an N-terminaldomain (N-domain) that serves as a docking site forvarious adaptor proteins [6–10]. ClpA consists of threedomains: an N-domain and two ATP-binding domainsreferred to as the D1 and D2 domains. Interestingly,deletion of the N-domain from ClpA not only abol-ishes binding of the adaptor protein, ClpS, but addi-tionally modulates ClpA substrate specificity [8,11–13].This change in substrate specificity is poorly under-stood, and the mechanism by which the N-domainsmight regulate ClpA function is controversial, althoughit has been proposed that the N-domain controls bind-ing of ClpA to ClpP [14]. Interestingly, there is alsoconsiderable debate regarding the role of the ClpBN-domain (which shares a common fold with theN-domain of ClpA) in substrate selection [15–17]. Onedifficulty in understanding the role of the N-domain ofClpA stems from the variety of activities exhibited byvarious DNClpA constructs tested, each containingdifferent lengths of ‘linker’ residues that connect theN-domain to the D1 domain. In order to avoid thepotential problems associated with ‘ragged’ ends ofDNClpA, we chose to create several single and doublepoint mutations within the N-domain to probeN-domain function.Here, using mutational analysis, we report the iden-tification of a structural element composed of con-served basic amino acids (R90 and R131), locatedwithin the N-domain of ClpA, that dramatically altersthe ability of ClpA to degrade selected substrates. Thiselement, although dispensable for the recognition ofthe SsrA tag per se, modulates the binding, unfoldingand subsequent degradation of SsrA-tagged proteinsubstrates. We propose that this element plays animportant role in the binding and subsequent releaseof substrates, by triggering ‘local’ unfolding of the sub-strate. We speculate that the ATP-dependent globalunfolding of some protein substrates is initiatedthrough productive binding to the substrate via twoelements in ClpA, one in the N-domain and the otherin the pore of the ClpA hexamer. In the case of shortunstructured peptides or unfolded proteins such ascasein, binding to the tyrosine residues in the hexamer-ic pore of ClpA is sufficient for substrate translocationto occur; however, in other cases such as SsrA-taggedprotein substrates, binding at both sites is required fortranslocation-mediated global unfolding to proceedefficiently.ResultsTwo conserved arginine residues (R90 and R131)within the N-domain are required for full ClpAfunctionWe were interested to understand how substrates arerecognized and subsequently unfolded by ClpA. Asmutation of the tyrosine residue located in the porehas been demonstrated to inhibit degradation of allsubstrates tested [18], we postulated that substrate dis-crimination must arise from an alternative regionwithin ClpA. Based on previous findings showing thatdeletion of the N-domain of ClpA dramaticallyreduced the rate of degradation of GFP–ssrA and to alesser extent casein [8,11], we speculated that theN-domain facilitates an early binding step, contribut-ing to specific recognition of substrates such as SsrA-tagged proteins. In order to further study the role ofthe N-domains in substrate recognition, we comparedthe amino acid sequences of this region in severalAAA+ proteins (Fig. 1). From this analysis, we noteda high occurrence of conserved basic residues distrib-uted throughout the domain, several of which (R86,R90, H94, R100 and R131) flanked a hydrophobicgroove (Fig. 2A). To test the role of these basic resi-dues, we constructed a number of single (R86A, R90Aand R131A) and double (H94A ⁄ R100A andR90A ⁄ R131A) point mutations in the N-domain ofClpA (Fig. 2A).First, we compared the degradation of SsrA-taggedGFP by wild-type and mutant ClpAP complexes(Fig. 2B). The ClpP-dependent degradation of GFP–ssrA mediated by either the single mutant R86A(Fig. 2B, open inverted triangles) or the double mutantH94A ⁄ R100A (Fig. 2B, filled diamonds) was unaf-fected. In contrast the rate of ClpP-mediated degrada-tion by the single mutants R90A (Fig. 2B, opendiamonds) and R131A (Fig. 2B, open triangles) wasreduced approximately threefold when compared towild-type ClpA (Fig. 2B, open circles). Interestingly,when we combined these two single point mutants tocreate the double mutant R90A ⁄ R131A (herein referredto as RR ⁄ AA), the degradation of GFP–ssrA wasreduced dramatically (Fig. 2B, filled circles). Althoughthese mutant proteins exhibited different abilities withregard to mediation of GFP–ssrA degradation(Fig. 2B), the basal ATPase activity was not affected(Fig. 3E, compare lanes 1 and 4). Given that theATPase activity of ClpA is dependent on its oligomeri-zation [19], as the nucleotide is bound between twoadjacent subunits, this result suggests that the overallhexameric structure of RR ⁄ AA was maintained.A. H. Erbse et al. Substrate recognition and unfolding by ClpAFEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBS 1401To determine whether this dramatic change in ClpP-mediated degradation of GFP–ssrA by RR ⁄ AA wassimply due to an inability to bind ClpP, we performedco-immunoprecipitation experiments using a-ClpP anti-serum (Fig. 2C). The co-immunoprecipitation of ClpAwith the a-ClpP antiserum was specific, as recovery ofClpA required the addition of both ATPcS (a non-hydrolysable analogue of ATP) and ClpP (Fig. 2C,lane 3). Importantly, RR ⁄ AA (Fig. 2C, lane 4) did notshow any change in ClpP interaction when compared towild-type ClpA (Fig. 2C, lane 3), as determined byquantification of ClpA amounts after co-immunopre-cipitation (Fig. 2C, lower panel), suggesting that theoverall structure of the RR ⁄ AA mutant is not compro-mised. Likewise, the other N-domain mutants tested(i.e. R86A and H94A ⁄ R100A) also exhibited wild-typeClpA behaviour (data not shown).An alternative explanation for the lack of GFP–ssrAdegradation exhibited by RR ⁄ AA could be that theN-domain was structurally compromised as a result ofmutations in this region. To confirm that neither theN-domain structure nor the overall structure of thesemutant proteins were adversely affected, we tested thedegradation of a model N-end rule substrate, FR-lin-ker–GFP [20]. The ClpP-mediated degradation of thissubstrate class requires specific interaction betweenClpS and the N-domain of ClpA. Consequently, dra-matic changes to the structure of the N-domain ofClpA would inhibit ClpS binding and thereby ClpS-dependent degradation of this substrate. As expected,the ClpP-mediated degradation of FR-linker–GFPby wild-type ClpA required the addition of ClpS(Fig. 2D). Importantly, like wild-type ClpA, RR ⁄ AAwas also able to support the ClpS-dependent degrad-ation of FR-linker–GFP (Fig. 2D), demonstrating afunctional interaction between ClpS and the N-domainof RR ⁄ AA, and this result suggests that neither thelocal nor the overall structure of the RR ⁄ AA mutantwas compromised.Mutation of the conserved arginine residues hasonly a moderate effect on degradation of shortunstructured peptides and the model unfoldedprotein, caseinTo determine whether RR ⁄ AA also demonstrated aninability to degrade other known ClpAP substrates, weexamined the ClpP-dependent degradation of severalmodel ClpA substrates, including the N-terminaldomain of the k repressor fused to the SsrA tag (kR–ssrA) [21], two short peptides, and the model unfoldedprotein a-casein [22]. As for GFP–ssrA, the rate offluorescein-labelled kR–ssrA degradation mediated byRR ⁄ AA was dramatically reduced when compared towild-type ClpA (Fig. 3A, filled circles and opencircles). Interestingly, the rate of RR ⁄ AA-mediateddegradation was not significantly altered for an SsrA-tagged peptide (Fig. 3B), indicating that recognition ofthe SsrA tag is not affected by RR ⁄ AA. Moreover,two other unfolded substrates, a-casein (Fig. 3C) anda 21-amino-acid polypeptide derived from r32(aloosely folded protein) [23] (Fig. 3D), were alsodegraded by RR ⁄ AA with similar kinetics to wild-typeClpA, in a ClpP-dependent manner. In contrast to theFig. 1. Multiple sequence alignment of the N-domain of bacterial ClpA homologues and E. coli ClpB. Amino acid sequences of the N-domainof ClpA from E. coli (P0ABH9), V. cholera (Q9KSW2), P. aeruginosa (Q9I0L8), X. fastidosa (Q87DL7), B. japonicum (Q89JW6), C. crescentus(Q9A5H9), N. meningitidis (Q9JZZ6), D. radiodurans (Q9RWS7), C. acetobutylicum (Q97I30) and H. pylori (O24875) were aligned togetherwith the amino acid sequence of the N-domain of E. coli ClpB (P63284). Conserved hydrophobic residues are highlighted in grey, conservedbasic residues are highlighted in blue, and conserved acidic residues are highlighted in red. Amino acid numbering corresponds to the ClpAsequence from E. coli. Residues chosen for mutation are indicated by asterisks.Substrate recognition and unfolding by ClpA A. H. Erbse et al.1402 FEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBSsubstrate-dependent degradation exhibited by RR ⁄ AA,the ClpA pore mutant (Y259A), which is unable totranslocate GFP–ssrA [18], prevented the degradationof both kR–ssrA (Fig. 3A, open diamonds) and fluo-rescein isothiocyanate-labelled casein (FITC-casein;data not shown). Consistent with these results, caseinstimulated the ATPase activity of both wild-type ClpAand RR ⁄ AA (Fig. 3E), while, in contrast, SsrA-taggedGFP only stimulated the ATPase activity of wild-typeClpA (Fig. 3E). Together, these data suggest thatRR ⁄ AA has a reduced ability to initiate unfolding ofmore tightly folded proteins, but retains full ability totranslocate short unstructured peptides and modelunfolded proteins into the ClpP chamber for degrada-tion.RR ⁄ AA delays the release and subsequentunfolding of certain protein substratesBefore testing the unfolding activity of RR ⁄ AA, wewished to compare the ability of the RR ⁄ AA mutantto bind to the various substrates tested. To do this, weconstructed a ClpA variant in which the glutamic acidresidue within the Walker B motif of each AAAdomain (E286, E565) was changed to alanine. Thisdouble Walker B mutant (herein referred to as dWB)Fig. 2. Two conserved arginine residues flanking a hydrophobic groove are essential for N-domain function. (A) Structure of the ClpA N-domain. ClpA is shown as a ribbon diagram (dark grey), and the side chains of R86, R90, H94, R100 and R131 are represented as a ball andstick (blue) flanking the putative peptide-binding groove (orange). The surface of the N-domain is shaded light grey, and R86, R90, H94,R100 and R131 are highlighted in blue. (B) The ClpP-mediated degradation of GFP–ssrA was monitored by fluorescence in the presence ofwild-type ClpA (open circles), R86A (inverted open triangles), H94A ⁄ R100A (filled diamonds), R90A (open diamonds), R131A (open triangles)and RR ⁄ AA (filled circles). (C) The interaction between wild-type ClpA (lane 3) or RR ⁄ AA (lane 4) with ClpP, assessed by co-immunoprecipita-tion using a-ClpP antiserum, was visualized by staining of the protein bands using Coomassie brilliant blue following separation by SDS–PAGE. In the absence of added ATPcS (lane 1) or ClpP (lane 2), ClpA was not co-precipitated. The relative amount of ClpA binding to ClpPwas determined from quantification of three independent experiments. Error bars represent the standard error of the mean. A non-specificprotein band is indicated by an asterisk. (D) The functional interaction between ClpS and ClpA (wild-type and RR ⁄ AA) was observed by moni-toring the ClpS-dependent degradation of FR-linker–GFP (in the presence of ClpP).A. H. Erbse et al. Substrate recognition and unfolding by ClpAFEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBS 1403and the corresponding mutant in RR⁄ AA (referred toas RR ⁄ dWB) were used to monitor substrate bindingas determined by co-elution of substrate–ClpAcomplexes during gel filtration. Initially, we tested theability of dWB and RR ⁄ dWB to interact with FITC-casein. As a control, in the absence of ClpA,FITC-casein eluted in a single peak at 21.5 mL(Fig. 4A, open circles). However, upon addition ofATP and dWB (Fig. 4A, open triangles) or RR ⁄ dWB(Fig. 4A, filled diamonds), the FITC-casein peakshifted and formed two new peaks, the largest ofwhich co-eluted with the ClpA hexamer (Fig. 4A, greyblock). Quantification of this peak indicated thatapproximately 30 and 40 pmol of FITC-casein werebound to the hexamers of dWB and RR ⁄ dWB respec-tively. Next we compared the ability of kR–ssrA(Fig. 4B) and GFP–ssrA (Fig. 4C) to bind to dWB orRR ⁄ dWB. As controls, each substrate (in the absenceof ClpA) was also separated by gel filtration and theamount of substrate was quantified in the hexamerregion of the gel filtration profile (Fig. 4B,C, lane 1).Similarly, as a further control, each substrate in thepresence of dWB (Fig. 4B,C, lane 2) or RR ⁄ dWB(Fig. 4B,C, lane 4) was also quantified after separationby gel filtration in the absence of ATP. These controlsdemonstrated a strict requirement for ATP in theinteraction between dWB ClpA and each substratetested. Interestingly, under the same conditions,although very little change in the binding of FITC-casein was observed, approximately threefold moreFig. 3. RR ⁄ AA exhibits different abilities with regard to degradation of various ClpA substrates. (A) ClpP-mediated degradation of fluores-cein-labelled kR–ssrA by ClpA (open circles), RR ⁄ AA (closed circles) and Y259A (open diamonds) was monitored by an increase in fluores-cence (excitation at 490 nm and emission at 520 nm). (B) ClpP-mediated degradation of a SsrA tagged peptide (50 lM) was monitored inthe absence of ClpA (ClpP) and the presence of wild-type (ClpA) or mutant (RR ⁄ AA) proteins. (C) Time course of a-casein degradation byClpA or RR ⁄ AA in the presence of ClpP. (D) ClpP-mediated degradation of a short unstructured peptide derived from r32(QRKLFFNLEKTKQRLGWFNQC) by RR ⁄ AA is not compromised. ClpP-mediated degradation of the peptide (50 lM) was monitored over timein the presence of wild-type ClpA (open circles) or RR ⁄ AA (filled circles). The amount of peptide remaining was determined by quantificationof the Coomassie-stained band following separation of the proteins by Tris ⁄ Tricine SDS–PAGE. (E) The ATPase activity of wild-type ClpA(lanes 1–3), RR ⁄ AA (lanes 4–6) and Y259A (lanes 7–9) was determined either in the absence of substrate (white bars; lanes 1, 4 and 7,respectively) or in the presence of GFP–ssrA (grey bars; lanes 2, 5 and 8, respectively) or a-casein (black bars; lanes 3, 6 and 9, respec-tively). The ATPase activity (relative to ClpA in the absence of substrate) was determined from three independent experiments. Error barsrepresent the standard error of the mean.Substrate recognition and unfolding by ClpA A. H. Erbse et al.1404 FEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBSSsrA-tagged substrate co-eluted with RR ⁄ dWB whencompared to dWB ClpA (Fig. 4B,C). These data areconsistent with the notion that RR ⁄ AA is able to bindeach substrate but exhibits a change in the release ofsome substrates (e.g. kR–ssrA). This lack of release isexpected to hinder unfolding and ultimately reducedegradation of the substrate.To further test the possibility that RR ⁄ AA has acompromised unfolding activity, we compared the abil-ity of wild-type ClpA and RR ⁄ AA to unfold SsrA-tagged GFP in the presence of the GroEL trap [24].As expected wild-type ClpA, in the absence of ClpP,was able to unfold GFP–ssrA (Fig. 5A, open circles)but the unfolding ability of RR ⁄ AA (Fig. 5A, filledcircles) was strongly compromised. Surprisingly, thekinetics of unfolding by RR ⁄ AA measured using theGroEL trap were slower than expected. As thismethod does not directly measure the change in sub-strate conformation and may be affected by rapidrefolding of the substrate, we chose to validate thisfinding using a more sensitive and direct approach.Thus, hydrogen–deuterium exchange was used to mea-sure the unfolding of GFP–ssrA in the presence andabsence of either wild-type ClpA or RR⁄ AA. Follow-ing incubation of GFP–ssrA (28 954 Da) in deuteratedbuffer, the mass of GFP–ssrA rapidly increased to29 034 Da within the first 5 min of the experiment.This change in mass occurred in the absence (data notshown) and the presence of wild-type or mutant ClpA(indicated by the hash symbol, #, in Fig. 5B,C), andresulted from the rapid exchange of 80 accessibleamide protons. In the absence of ClpA, the remainingamide protons within the protected core did notexchange over a period of 2 h (data not shown). In theFig. 4. Mutations in the N-domain do not prevent substrate interaction. (A) FITC-casein (500 pmol) was separated by gel filtration in thepresence of 2 mM ATP (open circles), 160 pmol dWB ClpA6plus 2 mM ATP (open triangles) or 160 pmol RR ⁄ dWB ClpA6plus 2 mM ATP(filled diamonds) as described in Experimental procedures. The molecular mass standards thyroglobulin (669 kDa), ferritin (440 kDa), aldolase(232 kDa) and ovalbumin (43 kDa) eluted as indicated by the arrows labelled 669, 440, 232 and 43 respectively. The position at which ClpA6eluted is indicated with an arrow labelled ClpA6. The amount of casein bound was calculated from the peak elution (boxed in grey) that co-eluted with ClpA6. (B) Fluorescein-labelled kR–ssrA (450 pmol) was separated by gel filtration without the addition of ATP (white bar, lane 1),in the presence of 160 pmol dWB ClpA6without (lane 2) or with addition of 2 mM ATP (lane 3), or in the presence of 160 pmol RR ⁄ dWBClpA6without (lane 4) or with addition of 2 mM ATP (lane 5) as described in (A). (C) GFP–ssrA (990 pmol) was separated by gel filtrationwithout the addition of ATP (lane 1), in the presence of 160 pmol dWB ClpA6without (lane 2) or with addition of 2 mM ATP (lane 3), or inthe presence of 160 pmol RR ⁄ dWB ClpA6without (lane 4) or with addition of 2 mM ATP (lane 5) as described in (A).A. H. Erbse et al. Substrate recognition and unfolding by ClpAFEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBS 1405presence of ClpA and ATP, we observed a furtherpeak (Fig. 5B, asterisk), which arises from incorpora-tion of deuterium into the core region of GFP–ssrA asa result of its unfolding. With time, the relativeamount of this heavier species (29 130 Da) increased,reflecting complete unfolding of all GFP–ssrA by ClpA(Fig. 5D, open circles). In contrast, the rate ofRR ⁄ AA-mediated unfolding (Fig. 5D, filled circles)was significantly slower than that of wild-type ClpA,with more than half of the GFP–ssrA still folded after30 min (Fig. 5C, asterisk). Taken together, these datasuggest that the change in degradation of GFP–ssrAmediated by RR⁄ AA stems from a delayed release ofsubstrate, which results in reduced unfolding of thesubstrate.DiscussionAs for most AAA+ proteases, ClpA utilizes thehydrolysis of ATP to drive substrate unfolding andtranslocation into the associated peptidase (ClpP). Todate, however, the role of the N-domains in this pro-cess has not been well defined as several conflictingroles have been proposed. Despite this, one aspect ofthe N-domain function is unambiguous – it is essentialfor ClpS binding and hence the delivery of N-end rulesubstrates to ClpAP. Currently, much of our mecha-nistic understanding of the ClpAP machine is basedlargely on the use of model proteins such as casein andGFP–ssrA. Previous studies have demonstrated that aring of tyrosine residues located in the pore of theClpA hexamer is essential for the translocation anddegradation of all substrates [18]. In contrast, variousN-domain deletions of ClpA have exhibited differingaffects on substrate degradation [8,11,12], which maysimply result from reduced ClpP interaction [14]. Inorder to better understand N-domain function, weanalysed in detail both the sequence and three-dimen-sional structure of the ClpA N-domain [25].In this study, we have identified an element withinthe N-domain of ClpA (composed of two conservedbasic residues, R90 and R131) that flanks a hydropho-bic groove. This element, via an unknown mechanism,contributes to the dynamic nature of substrate inter-action with ClpA. In contrast to mutation of thehexameric pore tyrosine residue (which abolishesdegradation of all substrates examined), the RR ⁄ AAmutant alters the unfolding of certain substrate types.For example, SsrA-tagged proteins are bound byRR ⁄ AA but release of the substrate is inhibited(Fig. 4). This slow substrate release appears to be spe-cific for SsrA-tagged proteins and was not observedfor the model unfolded protein casein or short peptidesubstrates (including an SsrA-tagged peptide) as deter-mined by rapid degradation of these peptides(Fig. 3B,D). RR ⁄ AA also exhibited a reduced rate ofGFP–ssrA unfolding as measured by hydrogen–deute-rium exchange or in the presence of the GroEL trap(Fig. 5). Collectively, these data confirm that the SsrAtag does not bind to the N-domain of ClpA, and sug-gest that these basic residues influence substrate releasefrom the N-domain, which in turn allows substrateunfolding to proceed.Importantly, in contrast to previous studies on theN-domain of ClpA (which examined the effect ofremoving the entire domain and resulted in dramaticaffects on ATPase activity or ClpP binding [14,26]),our site-directed mutagenesis approach has allowed usFig. 5. Mutations in the N-domain reducesubstrate unfolding. Unfolding of GFP–ssrAwas monitored (A) in the presence of theGroEL trap D87K upon the addition of ClpA(open circles) or RR ⁄ AA (closed circles), (B)by hydrogen–deuterium exchange in D2Obuffer in the presence of ClpA and ATP, or(C) by hydrogen–deuterium exchange inD2O buffer in the presence of RR ⁄ AA andATP. (D) The relative amount of ‘unfolded’GFP–ssrA (29 130 Da) was determined inthe presence of wild-type ClpA (opencircles) or RR ⁄ AA (filled circles).Substrate recognition and unfolding by ClpA A. H. Erbse et al.1406 FEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBSto specifically probe N-domain function. The RR ⁄ AAmutant not only displays normal basal ATPase activitybut also retains the ability to interact with both ClpSand ClpP and most importantly is able to translocateshort peptides and unfolded protein substrates into theClpP chamber for degradation. Despite these normalactivities, RR ⁄ AA shows a dramatic decrease in theability to degrade SsrA-tagged proteins (Figs 2 and 3).Nevertheless, this defect does not result from a simpleinability to interact with the SsrA tag (as demonstratedby efficient degradation of the SsrA peptide byRR ⁄ AA and stable binding of two SsrA-tagged pro-teins to RR⁄ dWB). In contrast, this region appears tobe involved in a more general binding task, which isconsistent with previous findings [18,27], and moreovermay modulate the ability of ClpA to bind, unfold andultimately degrade substrates such as GFP–ssrA.Interestingly, ClpS was observed to interact directlywith R131 in a ClpS–ClpA complex [28]. However,this interaction is not required for ClpS-mediatedaction [25], and hence most likely mimics a substrateinteraction, suggesting that R131 may interact directlywith some substrates. Nevertheless, using an in vitrocrosslinking approach [29], which permits the detectionof dynamic interactions, we did not observe an inter-action between various N-domain residues (His94,Leu109 and Val134) located in close proximity to R90and R131 and a substrate (data not shown). Impor-tantly, these variants were able to crosslink to ClpSand mediated the degradation of GFP–ssrA and FR-linker–GFP by ClpP (data not shown). Thus it remainsunclear whether these arginine residues are directlyinvolved in substrate binding. Of note, although theN-domains of ClpA and p97 are not structurallyrelated, mutations in several basic residues (R95G,R155C, R155H) within the N-domain of p97 havebeen implicated in the inclusion body myopathy asso-ciated with Paget’s disease of bone and fronto-tempo-ral dementia [30]. Interestingly, in the crystal structureof p97, these basic residues are not surface-exposedbut face the AAA domain, in close proximity to theWalker A motif. Hence, it is appealing to speculatethat the arginine residues in ClpA do not regulate sub-strate unfolding directly through interaction with thesubstrate, but instead coordinate substrate bind-ing ⁄ release via an interaction elsewhere in ClpA. Inter-estingly, although RR ⁄ AA has a dramatic inhibitoryeffect on the degradation of SsrA-tagged GFP, it doesnot affect binding or delivery of the model N-end rulesubstrate (FR-linker–GFP) by ClpS (Fig. 2), whichsuggests one of two possibilities. Firstly, that the defectin RR ⁄ AA-mediated unfolding is not dependent onthe global thermodynamic stability of the substrate,but rather correlates with local unfolding of the sub-strate (i.e. unfolding of the N- or C-terminus). Thiscan be understood by examining the N- and C-termi-nal structures of GFP. The first 11 N-terminal residuesof GFP form an a-helix, which leads into a parallelb-sheet. In contrast, the last 12 amino acids of GFPform a b-strand leading into an anti-parallel b-sheet[31]. In the case of GFP–ssrA, release of this substratefrom ClpA may be more effective than from RR ⁄ AA,allowing ClpA to perform the ATP-dependent unfold-ing step more efficiently. Interestingly, it has been pro-posed [32,33] that an a-helix is more easily unfoldedthan a b-sheet. Therefore, these data support the ideathat RR ⁄ AA has reduced ability to trigger localunfolding of a substrate (at the N- or C-terminus) andare consistent with the idea that local unfolding by theN-domain may be required before global unfolding ofthe substrate can proceed, as has been suggested forthe AAA+ protein PAN [34]. Alternatively, differentClpA substrates may utilize various recogni-tion ⁄ unfolding pathways – some that require thesearginine residues in the N-domain (e.g. GFP–ssrA)and other that do not (e.g. N-end rule substrates,delivered by ClpS). Therefore, ClpS-delivered sub-strates may bypass the need for these residues in theN-domain. In this case, it is appealing to speculatethat ClpS itself may act as the second binding siterequired for unfolding, thereby replacing a need forthe N-domain.Experimental proceduresProteinsClpA, ClpP and GFP–ssrA were over-expressed from anisopropyl thio-b-d-galactoside-inducible plasmid and puri-fied from the clarified lysates as described previously [8].All ClpA mutant proteins were purified as for wild-typeClpA. kR–ssrA was labelled with fluorescein as describedpreviously [21]. Purification of FR-linker–GFP was per-formed as previously described [20]. FITC-casein wasobtained from Sigma (St Louis, MO, USA). All proteinswere > 95% pure as determined by Coomassie-stainedSDS–PAGE. Protein concentrations were determined usinga Bradford assay system (Bio-Rad, Munich, Germany)using BSA purchased from Pierce (Rockford, IL, USA) asa standard, and refer to the protomer.Unfolding and protein degradation assaysGFP–ssrA degradation was monitored by changes influorescence (excitation at 400 nm and emission at510 nm). Degradation of fluorescein-labelled kR–ssrA andA. H. Erbse et al. Substrate recognition and unfolding by ClpAFEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBS 1407FITC-labelled a-casein was monitored by changes in fluo-rescence (excitation at 490 nm and emission at 520 nm)using a Perkin-Elmer fluorescence spectrometer LS50B(Waltham, MA, USA). The reactions were carried out asdescribed previously [24]. Unfolding assays for GFP–ssrAwere performed in the presence of GroEL trap D87K aspreviously described [24]. Non-fluorescent degradationassays (GFP–ssrA, a-casein, FR-linker–GFP and peptides)were preformed as previously described [8,20]. Unless other-wise stated, 1 lm ClpA (wild-type and all mutant ClpAproteins) and 1 lm ClpP were used. Samples were removedfrom the reactions at the indicated time points and degra-dation was stopped by the addition of sample buffer. Pro-tein substrates were separated by 15% SDS–PAGE andpeptide substrates by 16.5% Tris ⁄ Tricine SDS–PAGE.Proteins were visualized by Coomassie brilliant blue stain-ing. When required, protein bands were quantified usinggeleval1.21 (FrogDance Software, Dundee, UK).Analysis of ClpA–ClpP complexes byco-immunoprecipitationTo assess ClpA–ClpP complex formation, wild-type ormutant ClpA (1 lm) and ATPcS(2mm ) were preincubatedin co-immunoprecipitation buffer (50 mm Tris ⁄ HCl pH 7.5,300 mm NaCl, 40 mm Mg-acetate, 10% glycerol) at roomtemperature for 2 min prior to the addition of ClpP (1 lm).After a further 3 min incubation at room temperature, theprotein samples were mixed by end-over-end rotation for1 h at 4 °C with Protein A–Sepharose obtained from Sigma(St Louis, MO, USA) containing pre-bound antibodiesagainst E. coli ClpP. Following removal of the unboundfraction, the Protein A–Sepharose beads were washed threetimes with ice-cold co-immunoprecipitation buffer contain-ing 10 mm ATP, then bound proteins were eluted with50 mm glycine, pH 2.5. Proteins were separated by 10%SDS–PAGE and detected by Coomassie brilliant bluestaining.Gel filtration and substrate bindingGel filtration was carried out at 4 °C using a Superose 6column (GE Healthcare, Uppsala, Sweden) in buffer con-taining 20 mm Tris ⁄ HCl pH 7.4, 100 mm KCl, 40 mmNaCl, 10 mm MgCl2,5mm dithiothreitol, 0.1 mm EDTA,5% glycerol with or without 2 mm ATP. Fractions of250 lL were collected in a 96-well plate and samples analy-sed by fluorescence and 15% SDS–PAGE.ATPase assayThe ATPase activity of wild-type and mutant ClpA(0.5 lm) was measured at 660 nm, in degradation buffer(25 mm Tris ⁄ HCl pH 7.5, 100 mm NaCl, 100 mm KCl,20 mm MgCl2, 0.05% Triton X-100, 10% glycerol) in theabsence or presence of unlabelled substrate (5 lm). Thereaction was started by the addition of 2 mm ATP andstopped by the addition of 800 lL malachite green solution(0.034% malachite green, 0.1% Triton X-100 and10.5 gÆ L)1ammonium molybdate in 1 N HCl) and 100 lLof 34% citrate.Hydrogen–deuterium exchange and massspectrometryGFP–ssrA (2 lm) was diluted 75· into deuterated buffer(50 mm Tris ⁄ HCl pH 7.5, 300 mm NaCl, 10% glycerol,0.5 mm dithiothreitol, 10 mm ATP). Where appropriate,ClpA or RR ⁄ AA (2 lm) was added at the start of theexchange reaction (t = 0 min); for the control experiment,equal volumes of non-deuterated buffer were added. Atindicated time points, samples were removed from the reac-tion. The hydrogen–deuterium exchange was stopped byrapidly lowering the pH to 2.4 at 4 °C. All subsequent stepswere carried out on ice to minimize back exchange. The pro-teins were separated on a micro-C4RP column connected toan ESI-QTOF mass spectrometer (Applied Biosystems,Foster City, CA, USA) using an acetonitrile gradient.AcknowledgementsWe thank E. Weber-Ban (Eidgeno¨ssiche TechnischeHochschule Zurich) for providing fluorescently labelledkR–ssrA. This research was supported by the Austra-lian Research Council Discovery Project scheme(DP0450051), the Deutsche Forschungsgemeinschaftpriority program ‘Proteolysis in Prokaryotes: ProteinQuality Control and Regulatory Principles’ and Aus-tralian Research Council QEII Fellowships to D.A.Dand K.N.T.References1 Neuwald AF, Aravind L, Spouge JL & Koonin EV(1999) AAA+: a class of chaperone-like ATPases asso-ciated with the assembly, operation, and disassembly ofprotein complexes. Genome Res 9, 27–43.2 Dougan DA, Mogk A & Bukau B (2002) Protein fold-ing and degradation in bacteria: to degrade or not todegrade? That is the question. 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Iwakura M & Matouschek A (2001) ATP-dependent proteases degrade their substrates by processively unraveling them from the degradation signal Mol Cell 7, 627–637 34 Navon A & Goldberg AL (2001) Proteins are unfolded on the surface of the ATPase ring before transport into the proteasome Mol Cell 8, 1339–1349 FEBS Journal 275 (2008) 1400–1410 ª 2008 The Authors Journal compilation ª 2008 FEBS . productive binding to the substrate via twoelements in ClpA, one in the N-domain and the other in the pore of the ClpA hexamer. In the case of shortunstructured. with the N-domain of ClpA) in substrate selection [15–17]. Onedifficulty in understanding the role of the N-domain of ClpA stems from the variety of activities
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