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

For example, in Escherichia coli, ClpA is responsible, either directly or indirectly via the adaptor protein ClpS, for recognition of substrates such as SsrA-tagged proteins or N-end rul

1 Zentrum fu¨r Molekulare Biologie Heidelberg, Universita¨t Heidelberg, Heidelberg, Germany

2 Department of Biochemistry, La Trobe University, Melbourne, Australia

3 Institut fu¨r Biologie, Freie Universita¨t Berlin, Berlin, Germany

4 MPI fu¨r Entwicklungsbiologie, Tubingen, Germany

The AAA+ superfamily [1] is an extensive group of

proteins involved in a broad range of biological

func-tions Its members are present in all kingdoms of life

and often play a crucial role in cell maintenance In

bacteria, 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 of proteins, including large macromolecular complexes [3] Although several of these proteins share consider-able sequence similarity, they demonstrate distinct substrate specificity For example, in Escherichia coli, ClpA is responsible, either directly or indirectly via the adaptor protein ClpS, for recognition of substrates such as SsrA-tagged proteins or N-end rule substrates

Keywords

AAA+; binding; ClpA; SsrA; unfolding

Correspondence

D A Dougan, Department of Biochemistry,

La Trobe University, Melbourne 3086,

Australia

Fax: +61 3 9479 2467

Tel: +61 3 9479 3276

E-mail: d.dougan@latrobe.edu.au

B Bukau, Zentrum fu¨r Molekulare Biologie

Heidelberg, Universita¨t Heidelberg, INF 282,

Heidelberg D-69120, Germany

Fax: +49 6221 54 5894

Tel: +49 6221 54 6795

E-mail: bukau@zmbh.uni-heidelberg.de

*These authors contributed equally to this

work

(Received 22 November 2007, revised 10

January 2008, accepted 14 January 2008)

doi:10.1111/j.1742-4658.2008.06304.x

Protein degradation in the cytosol of Escherichia coli is carried out by a variety of different proteolytic machines, including ClpAP The ClpA com-ponent is a hexameric AAA+ (ATPase associated with various cellular activities) chaperone that utilizes the energy of ATP to control substrate recognition and unfolding The precise role of the N-domains of ClpA in this process, however, remains elusive Here, we have analysed the role of five highly conserved basic residues in the N-domain of ClpA by monitor-ing the bindmonitor-ing, unfoldmonitor-ing 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 of ClpA (H94, R86 and R100) did not alter substrate degradation In contrast mutation of two conserved arginine residues (R90 and R131), flanking a putative peptide-binding groove within the N-domain of ClpA, specifically compromised the ability of ClpA to unfold and degrade selected substrates but did not prevent substrate recognition, ClpS-mediated substrate delivery

or ClpP binding In contrast, a highly conserved tyrosine residue lining the central pore of the ClpA hexamer was essential for the degradation of all substrate 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, both

of which are required for efficient unfolding of some protein substrates

Abbreviations

AAA+, ATPase associated with various cellular activities; FITC, fluorescein isothiocyanate; GFP, green fluorescent protein; kR, lambda repressor.

[4,5] Once recognized, these substrates are unfolded

by 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 are

degraded into short peptides

AAA+ proteins usually contain an N-terminal

domain (N-domain) that serves as a docking site for

various adaptor proteins [6–10] ClpA consists of three

domains: an N-domain and two ATP-binding domains

referred 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-domains

might regulate ClpA function is controversial, although

it has been proposed that the N-domain controls

bind-ing of ClpA to ClpP [14] Interestbind-ingly, there is also

considerable debate regarding the role of the ClpB

N-domain (which shares a common fold with the

N-domain of ClpA) in substrate selection [15–17] One

difficulty in understanding the role of the N-domain of

ClpA stems from the variety of activities exhibited by

various DNClpA constructs tested, each containing

different lengths of ‘linker’ residues that connect the

N-domain to the D1 domain In order to avoid the

potential problems associated with ‘ragged’ ends of

DNClpA, we chose to create several single and double

point mutations within the N-domain to probe

N-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), located

within the N-domain of ClpA, that dramatically alters

the ability of ClpA to degrade selected substrates This

element, although dispensable for the recognition of

the SsrA tag per se, modulates the binding, unfolding

and subsequent degradation of SsrA-tagged protein

substrates We propose that this element plays an

important role in the binding and subsequent release

of substrates, by triggering ‘local’ unfolding of the

sub-strate We speculate that the ATP-dependent global

unfolding of some protein substrates is initiated

through productive binding to the substrate via two

elements in ClpA, one in the N-domain and the other

in the pore of the ClpA hexamer In the case of short

unstructured peptides or unfolded proteins such as

casein, binding to the tyrosine residues in the

hexamer-ic pore of ClpA is sufficient for substrate translocation

to occur; however, in other cases such as SsrA-tagged

protein substrates, binding at both sites is required for

translocation-mediated global unfolding to proceed

efficiently

Results

Two conserved arginine residues (R90 and R131) within the N-domain are required for full ClpA function

We were interested to understand how substrates are recognized and subsequently unfolded by ClpA As mutation of the tyrosine residue located in the pore has been demonstrated to inhibit degradation of all substrates tested [18], we postulated that substrate dis-crimination must arise from an alternative region within ClpA Based on previous findings showing that deletion of the N-domain of ClpA dramatically reduced the rate of degradation of GFP–ssrA and to a lesser extent casein [8,11], we speculated that the N-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 of the N-domains in substrate recognition, we compared the amino acid sequences of this region in several AAA+ proteins (Fig 1) From this analysis, we noted

a high occurrence of conserved basic residues distrib-uted throughout the domain, several of which (R86, R90, H94, R100 and R131) flanked a hydrophobic groove (Fig 2A) To test the role of these basic resi-dues, we constructed a number of single (R86A, R90A and R131A) and double (H94A⁄ R100A and R90A⁄ R131A) point mutations in the N-domain of ClpA (Fig 2A)

First, we compared the degradation of SsrA-tagged GFP 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 mutant H94A⁄ R100A (Fig 2B, filled diamonds) was unaf-fected In contrast the rate of ClpP-mediated degrada-tion by the single mutants R90A (Fig 2B, open diamonds) and R131A (Fig 2B, open triangles) was reduced approximately threefold when compared to wild-type ClpA (Fig 2B, open circles) Interestingly, when we combined these two single point mutants to create the double mutant R90A⁄ R131A (herein referred

to as RR⁄ AA), the degradation of GFP–ssrA was reduced dramatically (Fig 2B, filled circles) Although these mutant proteins exhibited different abilities with regard to mediation of GFP–ssrA degradation (Fig 2B), the basal ATPase activity was not affected (Fig 3E, compare lanes 1 and 4) Given that the ATPase activity of ClpA is dependent on its oligomeri-zation [19], as the nucleotide is bound between two adjacent subunits, this result suggests that the overall hexameric structure of RR⁄ AA was maintained

To determine whether this dramatic change in

ClpP-mediated degradation of GFP–ssrA by RR⁄ AA was

simply due to an inability to bind ClpP, we performed

co-immunoprecipitation experiments using a-ClpP

anti-serum (Fig 2C) The co-immunoprecipitation of ClpA

with the a-ClpP antiserum was specific, as recovery of

ClpA 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 not

show any change in ClpP interaction when compared to

wild-type ClpA (Fig 2C, lane 3), as determined by

quantification of ClpA amounts after

co-immunopre-cipitation (Fig 2C, lower panel), suggesting that the

overall 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-type

ClpA behaviour (data not shown)

An alternative explanation for the lack of GFP–ssrA

degradation exhibited by RR⁄ AA could be that the

N-domain was structurally compromised as a result of

mutations in this region To confirm that neither the

N-domain structure nor the overall structure of these

mutant proteins were adversely affected, we tested the

degradation of a model N-end rule substrate,

FR-lin-ker–GFP [20] The ClpP-mediated degradation of this

substrate class requires specific interaction between

ClpS and the N-domain of ClpA Consequently,

dra-matic changes to the structure of the N-domain of

ClpA would inhibit ClpS binding and thereby

ClpS-dependent degradation of this substrate As expected,

the ClpP-mediated degradation of FR-linker–GFP

by wild-type ClpA required the addition of ClpS

(Fig 2D) Importantly, like wild-type ClpA, RR⁄ AA was also able to support the ClpS-dependent degrad-ation of FR-linker–GFP (Fig 2D), demonstrating a functional interaction between ClpS and the N-domain

of RR⁄ AA, and this result suggests that neither the local nor the overall structure of the RR⁄ AA mutant was compromised

Mutation of the conserved arginine residues has only a moderate effect on degradation of short unstructured peptides and the model unfolded protein, casein

To determine whether RR⁄ AA also demonstrated an inability to degrade other known ClpAP substrates, we examined the ClpP-dependent degradation of several model ClpA substrates, including the N-terminal domain of the k repressor fused to the SsrA tag (kR– ssrA) [21], two short peptides, and the model unfolded protein a-casein [22] As for GFP–ssrA, the rate of fluorescein-labelled kR–ssrA degradation mediated by

RR⁄ AA was dramatically reduced when compared to wild-type ClpA (Fig 3A, filled circles and open circles) Interestingly, the rate of RR⁄ AA-mediated degradation was not significantly altered for an SsrA-tagged peptide (Fig 3B), indicating that recognition of the SsrA tag is not affected by RR⁄ AA Moreover, two other unfolded substrates, a-casein (Fig 3C) and

a 21-amino-acid polypeptide derived from r32 (a loosely folded protein) [23] (Fig 3D), were also degraded by RR⁄ AA with similar kinetics to wild-type ClpA, in a ClpP-dependent manner In contrast to the

Fig 1 Multiple sequence alignment of the N-domain of bacterial ClpA homologues and E coli ClpB Amino acid sequences of the N-domain

of 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 together with the amino acid sequence of the N-domain of E coli ClpB (P63284) Conserved hydrophobic residues are highlighted in grey, conserved basic residues are highlighted in blue, and conserved acidic residues are highlighted in red Amino acid numbering corresponds to the ClpA sequence from E coli Residues chosen for mutation are indicated by asterisks.

substrate-dependent degradation exhibited by RR⁄ AA,

the ClpA pore mutant (Y259A), which is unable to

translocate GFP–ssrA [18], prevented the degradation

of both kR–ssrA (Fig 3A, open diamonds) and

fluo-rescein isothiocyanate-labelled casein (FITC-casein;

data not shown) Consistent with these results, casein

stimulated the ATPase activity of both wild-type ClpA

and RR⁄ AA (Fig 3E), while, in contrast, SsrA-tagged

GFP only stimulated the ATPase activity of wild-type

ClpA (Fig 3E) Together, these data suggest that

RR⁄ AA has a reduced ability to initiate unfolding of

more tightly folded proteins, but retains full ability to

translocate short unstructured peptides and model

unfolded proteins into the ClpP chamber for degrada-tion

RR⁄ AA delays the release and subsequent unfolding of certain protein substrates Before testing the unfolding activity of RR⁄ AA, we wished to compare the ability of the RR⁄ AA mutant

to bind to the various substrates tested To do this, we constructed a ClpA variant in which the glutamic acid residue within the Walker B motif of each AAA domain (E286, E565) was changed to alanine This double Walker B mutant (herein referred to as dWB)

Fig 2 Two conserved arginine residues flanking a hydrophobic groove are essential for 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 and stick (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 of wild-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 separaco-immunoprecipita-tion 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 ClpP was determined from quantification of three independent experiments Error bars represent the standard error of the mean A non-specific protein 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).

and the corresponding mutant in RR⁄ AA (referred to

as RR⁄ dWB) were used to monitor substrate binding

as determined by co-elution of substrate–ClpA

complexes during gel filtration Initially, we tested the

ability 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 of

ATP and dWB (Fig 4A, open triangles) or RR⁄ dWB

(Fig 4A, filled diamonds), the FITC-casein peak

shifted and formed two new peaks, the largest of

which co-eluted with the ClpA hexamer (Fig 4A, grey

block) Quantification of this peak indicated that

approximately 30 and 40 pmol of FITC-casein were

bound 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 or

RR⁄ dWB As controls, each substrate (in the absence

of ClpA) was also separated by gel filtration and the amount of substrate was quantified in the hexamer region of the gel filtration profile (Fig 4B,C, lane 1) Similarly, as a further control, each substrate in the presence of dWB (Fig 4B,C, lane 2) or RR⁄ dWB (Fig 4B,C, lane 4) was also quantified after separation

by gel filtration in the absence of ATP These controls demonstrated a strict requirement for ATP in the interaction between dWB ClpA and each substrate tested Interestingly, under the same conditions, although very little change in the binding of FITC-casein was observed, approximately threefold more

Fig 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 l M ) was monitored in the absence of ClpA (ClpP) and the presence of wild-type (ClpA) or mutant (RR ⁄ AA) proteins (C) Time course of a-casein degradation by ClpA or RR ⁄ AA in the presence of ClpP (D) ClpP-mediated degradation of a short unstructured peptide derived from r 32 (QRKLFFNLEKTKQRLGWFNQC) by RR ⁄ AA is not compromised ClpP-mediated degradation of the peptide (50 l M ) was monitored over time

in the presence of wild-type ClpA (open circles) or RR ⁄ AA (filled circles) The amount of peptide remaining was determined by quantification

of 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 bars represent the standard error of the mean.

SsrA-tagged substrate co-eluted with RR⁄ dWB when

compared to dWB ClpA (Fig 4B,C) These data are

consistent with the notion that RR⁄ AA is able to bind

each substrate but exhibits a change in the release of

some substrates (e.g kR–ssrA) This lack of release is

expected to hinder unfolding and ultimately reduce

degradation of the substrate

To further test the possibility that RR⁄ AA has a

compromised 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, filled

circles) was strongly compromised Surprisingly, the

kinetics of unfolding by RR⁄ AA measured using the

GroEL trap were slower than expected As this

method does not directly measure the change in sub-strate conformation and may be affected by rapid refolding of the substrate, we chose to validate this finding using a more sensitive and direct approach Thus, hydrogen–deuterium exchange was used to mea-sure the unfolding of GFP–ssrA in the presence and absence of either wild-type ClpA or RR⁄ AA Follow-ing incubation of GFP–ssrA (28 954 Da) in deuterated buffer, the mass of GFP–ssrA rapidly increased to

29 034 Da within the first 5 min of the experiment This change in mass occurred in the absence (data not shown) and the presence of wild-type or mutant ClpA (indicated by the hash symbol, #, in Fig 5B,C), and resulted from the rapid exchange of 80 accessible amide protons In the absence of ClpA, the remaining amide protons within the protected core did not exchange over a period of 2 h (data not shown) In the

Fig 4 Mutations in the N-domain do not prevent substrate interaction (A) FITC-casein (500 pmol) was separated by gel filtration in the presence of 2 m M ATP (open circles), 160 pmol dWB ClpA6plus 2 m M ATP (open triangles) or 160 pmol RR ⁄ dWB ClpA 6 plus 2 m M 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 ClpA 6 eluted 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 ClpA 6 without (lane 2) or with addition of 2 m M ATP (lane 3), or in the presence of 160 pmol RR ⁄ dWB ClpA6without (lane 4) or with addition of 2 m M ATP (lane 5) as described in (A) (C) GFP–ssrA (990 pmol) was separated by gel filtration without the addition of ATP (lane 1), in the presence of 160 pmol dWB ClpA6without (lane 2) or with addition of 2 m M ATP (lane 3), or in the presence of 160 pmol RR ⁄ dWB ClpA 6 without (lane 4) or with addition of 2 m M ATP (lane 5) as described in (A).

presence of ClpA and ATP, we observed a further

peak (Fig 5B, asterisk), which arises from

incorpora-tion of deuterium into the core region of GFP–ssrA as

a result of its unfolding With time, the relative

amount 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 of

RR⁄ 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 after

30 min (Fig 5C, asterisk) Taken together, these data

suggest that the change in degradation of GFP–ssrA

mediated by RR⁄ AA stems from a delayed release of

substrate, which results in reduced unfolding of the

substrate

Discussion

As for most AAA+ proteases, ClpA utilizes the

hydrolysis of ATP to drive substrate unfolding and

translocation into the associated peptidase (ClpP) To

date, however, the role of the N-domains in this

pro-cess has not been well defined as several conflicting

roles have been proposed Despite this, one aspect of

the N-domain function is unambiguous – it is essential

for ClpS binding and hence the delivery of N-end rule

substrates to ClpAP Currently, much of our

mecha-nistic understanding of the ClpAP machine is based

largely on the use of model proteins such as casein and

GFP–ssrA Previous studies have demonstrated that a

ring of tyrosine residues located in the pore of the

ClpA hexamer is essential for the translocation and

degradation of all substrates [18] In contrast, various

N-domain deletions of ClpA have exhibited differing

affects on substrate degradation [8,11,12], which may simply result from reduced ClpP interaction [14] In order to better understand N-domain function, we analysed in detail both the sequence and three-dimen-sional structure of the ClpA N-domain [25]

In this study, we have identified an element within the N-domain of ClpA (composed of two conserved basic 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 the hexameric pore tyrosine residue (which abolishes degradation of all substrates examined), the RR⁄ AA mutant alters the unfolding of certain substrate types For example, SsrA-tagged proteins are bound by

RR⁄ 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 observed for the model unfolded protein casein or short peptide substrates (including an SsrA-tagged peptide) as deter-mined by rapid degradation of these peptides (Fig 3B,D) RR⁄ AA also exhibited a reduced rate of GFP–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 SsrA tag does not bind to the N-domain of ClpA, and sug-gest that these basic residues influence substrate release from the N-domain, which in turn allows substrate unfolding to proceed

Importantly, in contrast to previous studies on the N-domain of ClpA (which examined the effect of removing the entire domain and resulted in dramatic affects on ATPase activity or ClpP binding [14,26]), our site-directed mutagenesis approach has allowed us

Fig 5 Mutations in the N-domain reduce substrate unfolding Unfolding of GFP–ssrA was monitored (A) in the presence of the GroEL trap D87K upon the addition of ClpA (open circles) or RR ⁄ AA (closed circles), (B)

by hydrogen–deuterium exchange in D 2 O buffer in the presence of ClpA and ATP, or (C) by hydrogen–deuterium exchange in

D2O buffer in the presence of RR ⁄ AA and ATP (D) The relative amount of ‘unfolded’ GFP–ssrA (29 130 Da) was determined in the presence of wild-type ClpA (open circles) or RR ⁄ AA (filled circles).

to specifically probe N-domain function The RR⁄ AA

mutant not only displays normal basal ATPase activity

but also retains the ability to interact with both ClpS

and ClpP and most importantly is able to translocate

short peptides and unfolded protein substrates into the

ClpP chamber for degradation Despite these normal

activities, RR⁄ AA shows a dramatic decrease in the

ability to degrade SsrA-tagged proteins (Figs 2 and 3)

Nevertheless, this defect does not result from a simple

inability to interact with the SsrA tag (as demonstrated

by efficient degradation of the SsrA peptide by

RR⁄ AA and stable binding of two SsrA-tagged

pro-teins to RR⁄ dWB) In contrast, this region appears to

be involved in a more general binding task, which is

consistent with previous findings [18,27], and moreover

may modulate the ability of ClpA to bind, unfold and

ultimately degrade substrates such as GFP–ssrA

Interestingly, ClpS was observed to interact directly

with R131 in a ClpS–ClpA complex [28] However,

this interaction is not required for ClpS-mediated

action [25], and hence most likely mimics a substrate

interaction, suggesting that R131 may interact directly

with some substrates Nevertheless, using an in vitro

crosslinking approach [29], which permits the detection

of dynamic interactions, we did not observe an

inter-action between various N-domain residues (His94,

Leu109 and Val134) located in close proximity to R90

and R131 and a substrate (data not shown)

Impor-tantly, these variants were able to crosslink to ClpS

and mediated the degradation of GFP–ssrA and

FR-linker–GFP by ClpP (data not shown) Thus it remains

unclear whether these arginine residues are directly

involved in substrate binding Of note, although the

N-domains of ClpA and p97 are not structurally

related, mutations in several basic residues (R95G,

R155C, R155H) within the N-domain of p97 have

been implicated in the inclusion body myopathy

asso-ciated with Paget’s disease of bone and

fronto-tempo-ral dementia [30] Interestingly, in the crystal structure

of p97, these basic residues are not surface-exposed

but face the AAA domain, in close proximity to the

Walker A motif Hence, it is appealing to speculate

that the arginine residues in ClpA do not regulate

sub-strate unfolding directly through interaction with the

substrate, but instead coordinate substrate

bind-ing⁄ release via an interaction elsewhere in ClpA

Inter-estingly, although RR⁄ AA has a dramatic inhibitory

effect on the degradation of SsrA-tagged GFP, it does

not affect binding or delivery of the model N-end rule

substrate (FR-linker–GFP) by ClpS (Fig 2), which

suggests one of two possibilities Firstly, that the defect

in RR⁄ AA-mediated unfolding is not dependent on

the 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) This can be understood by examining the N- and C-termi-nal structures of GFP The first 11 N-termiC-termi-nal residues

of GFP form an a-helix, which leads into a parallel b-sheet In contrast, the last 12 amino acids of GFP form a b-strand leading into an anti-parallel b-sheet [31] In the case of GFP–ssrA, release of this substrate from ClpA may be more effective than from RR⁄ AA, allowing ClpA to perform the ATP-dependent unfold-ing step more efficiently Interestunfold-ingly, it has been pro-posed [32,33] that an a-helix is more easily unfolded than a b-sheet Therefore, these data support the idea that RR⁄ AA has reduced ability to trigger local unfolding of a substrate (at the N- or C-terminus) and are consistent with the idea that local unfolding by the N-domain may be required before global unfolding of the substrate can proceed, as has been suggested for the AAA+ protein PAN [34] Alternatively, different ClpA substrates may utilize various recogni-tion⁄ unfolding pathways – some that require these arginine 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 the N-domain In this case, it is appealing to speculate that ClpS itself may act as the second binding site required for unfolding, thereby replacing a need for the N-domain

Experimental procedures

Proteins

ClpA, ClpP and GFP–ssrA were over-expressed from an isopropyl thio-b-d-galactoside-inducible plasmid and puri-fied from the claripuri-fied lysates as described previously [8] All ClpA mutant proteins were purified as for wild-type ClpA kR–ssrA was labelled with fluorescein as described previously [21] Purification of FR-linker–GFP was per-formed as previously described [20] FITC-casein was obtained from Sigma (St Louis, MO, USA) All proteins were > 95% pure as determined by Coomassie-stained SDS–PAGE Protein concentrations were determined using

a Bradford assay system (Bio-Rad, Munich, Germany) using BSA purchased from Pierce (Rockford, IL, USA) as

a standard, and refer to the protomer

Unfolding and protein degradation assays

GFP–ssrA degradation was monitored by changes in fluorescence (excitation at 400 nm and emission at

510 nm) Degradation of fluorescein-labelled kR–ssrA and

were performed in the presence of GroEL trap D87K as

previously described [24] Non-fluorescent degradation

assays (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 ClpA

proteins) and 1 lm ClpP were used Samples were removed

from 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 and

peptide substrates by 16.5% Tris⁄ Tricine SDS–PAGE

Proteins were visualized by Coomassie brilliant blue

stain-ing When required, protein bands were quantified using

geleval1.21 (FrogDance Software, Dundee, UK)

Analysis of ClpA–ClpP complexes by

co-immunoprecipitation

To assess ClpA–ClpP complex formation, wild-type or

mutant ClpA (1 lm) and ATPcS (2 mm) were preincubated

in co-immunoprecipitation buffer (50 mm Tris⁄ HCl pH 7.5,

300 mm NaCl, 40 mm Mg-acetate, 10% glycerol) at room

temperature for 2 min prior to the addition of ClpP (1 lm)

After a further 3 min incubation at room temperature, the

protein samples were mixed by end-over-end rotation for

1 h at 4C with Protein A–Sepharose obtained from Sigma

(St Louis, MO, USA) containing pre-bound antibodies

against E coli ClpP Following removal of the unbound

fraction, the Protein A–Sepharose beads were washed three

times with ice-cold co-immunoprecipitation buffer

contain-ing 10 mm ATP, then bound proteins were eluted with

50 mm glycine, pH 2.5 Proteins were separated by 10%

SDS–PAGE and detected by Coomassie brilliant blue

staining

Gel filtration and substrate binding

Gel filtration was carried out at 4C using a Superose 6

column (GE Healthcare, Uppsala, Sweden) in buffer

con-taining 20 mm Tris⁄ HCl pH 7.4, 100 mm KCl, 40 mm

NaCl, 10 mm MgCl2, 5 mm dithiothreitol, 0.1 mm EDTA,

5% glycerol with or without 2 mm ATP Fractions of

250 lL were collected in a 96-well plate and samples

analy-sed by fluorescence and 15% SDS–PAGE

ATPase assay

The 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,

10.5 gÆL ammonium molybdate in 1 N HCl) and 100 lL

of 34% citrate

Hydrogen–deuterium exchange and mass spectrometry

GFP–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 the exchange reaction (t = 0 min); for the control experiment, equal volumes of non-deuterated buffer were added At indicated time points, samples were removed from the reac-tion The hydrogen–deuterium exchange was stopped by rapidly lowering the pH to 2.4 at 4C All subsequent steps were carried out on ice to minimize back exchange The pro-teins were separated on a micro-C4RP column connected to

an ESI-QTOF mass spectrometer (Applied Biosystems, Foster City, CA, USA) using an acetonitrile gradient

Acknowledgements

We thank E Weber-Ban (Eidgeno¨ssiche Technische Hochschule Zurich) for providing fluorescently labelled kR–ssrA This research was supported by the Austra-lian Research Council Discovery Project scheme (DP0450051), the Deutsche Forschungsgemeinschaft priority program ‘Proteolysis in Prokaryotes: Protein Quality Control and Regulatory Principles’ and Aus-tralian Research Council QEII Fellowships to D.A.D and K.N.T

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