Role of communication between subunits and enzymes in
ClpXP-mediated substrate unfolding and degradation
by
Shilpa Arun Joshi
A.B., Molecular and Cell Biology
University of California at Berkeley, 1997
Submitted to the Department of Biology in Partial Fulfillment
of the Requirements for the Degree of
Doctor of Philosophy in Biology
at the
Massachusetts Institute of Technology
February 2004
 2004 Shilpa A. Joshi. All rights reserved.
The author hereby grants MIT permission to reproduce and to distribute publicly paper and
electronic copies of this thesis document in whole or in part.
Signature of Author............................................................................................................................
Department of Biology
February 2, 2004
Certified by........................................................................................................................................
Robert T. Sauer, Salvador E. Luria Professor of Biology
Thesis Supervisor
Accepted by.......................................................................................................................................
Alan D. Grossman, Professor of Biology
Co-Chair, Biology Graduate Committee
Role of communication between subunits and enzymes in
ClpXP-mediated substrate unfolding and degradation
by
Shilpa Arun Joshi
Submitted to the Department of Biology on February 2, 2004 in Partial
Fulfillment of the Requirements for the Degree of Doctor of Philosophy
ABSTRACT
Chaperones and proteases play important roles in quality control by helping proteins fold,
by dismantling hyper-stable complexes, and by degrading unwanted proteins. The highly
conserved AAA+ Clp/Hsp100 proteins are ATPases which function as disassembly chaperones
as well as essential components of energy-dependent proteases. For example, the ClpX ATPase
disassembles macromolecular complexes and combines with the ClpP peptidase to form ClpXP,
a molecular machine with structural and functional similarity to the eukaryotic 26S proteasome.
ClpXP consists of hexameric ClpX rings stacked coaxially against the double-ring ClpP14
peptidase. ClpXP's peptidase active sites reside in a sequestered chamber accessible through a
narrow channel which excludes native, folded substrates. ClpX binds specific substrates, unfolds
them in a reaction requiring ATP hydrolysis, and then translocates them into ClpP for
degradation. When ClpP is absent, ClpX unfolds and releases specific substrates, an activity that
can disassemble otherwise stable complexes.
Although ClpX-mediated substrate unfolding and ClpXP-mediated degradation have
been studied extensively, the role of communication between subunits within a ClpX hexamer or
between enzymes in ClpXP has not been addressed. I initially screened numerous ClpX point
mutants for their ability to support host-cell lysis by bacteriophage Mu, and then purified several
mutants for functional characterization in vitro. Three of these mutants contained substitutions at
intersubunit interfaces; these variants bound substrate well, but displayed unusual changes in
ATP hydrolysis in response to substrate binding and had low protein unfolding activity. These
results suggest that communication across these subunit-subunit interfaces coordinates and
regulates nucleotide hydrolysis in response to substrate binding and is critical for efficient
substrate unfolding. I also discovered that ClpX and ClpP communicate with each other during
substrate processing and degradation. ClpX binds ClpP more strongly during substrate unfolding
than during translocation, by a mechanism that is linked to the ATP hydrolysis rate. Interaction
between ClpX and DFP-modified ClpP, mimicking a peptide-cleavage intermediate, is
strengthened significantly suggesting that ClpP also communicates its functional state to ClpX.
Overall, this work suggests that communication within the ClpX hexamer and between enzymes
in ClpXP modulates and coordinates substrate processing.
Thesis Supervisor: Robert T. Sauer
Title: Salvador E. Luria Professor of Biology
2
Acknowledgements
I thank Bob and Tania, the best advisors I could have hoped to find. I have learned a lot from
you two and have grown a great deal as a result. Thanks for taking the job of training us
seriously, making time for everyone even when you are busy, and setting high standards for all
tasks. I cannot say strongly enough how much I have valued these traits, along with your humor,
youthful enthusiasm, and love of experiments. Thanks for everything!
Thanks also to the other faculty on my thesis committee (Alan Grossman, David Jeruzalmi, and
Graham Walker) and other people in MIT Biology. I have appreciated my committee's
comments and discussion and thank all of you for taking time to be on my committee. I
generally thank all members of the Sauer and Baker laboratories for daily discussion, help, and
insights. I particularly thank Randy Burton, Peter Chivers, Ehud Gazit, Wali Karzai, and David
Wah for teaching me many things through the years.
I am indebted immensely to my parents, Scott, and the rest of my family for their love, support,
and encouragement. Aiee and Baba, you are the best parents and the kindest people I have ever
met. Thanks for your inspiring courage, making time to listen, and never telling me I am foolish
when I try new things. Scott, thanks for your questioning way, sensitivity, and endless jokes.
You have brightened these days. All four grandparents, you are truly inspiring in how you
strived for self-improvement and tried to better the larger world. Dada and Tai, thanks for
instilling in me a sense of competition and watching me even when you would have rather done
something else. Benevie and Reshma, thanks for the lively discussions. Sachin, Sonal,
Akhilesh, and Priya, thanks for being young and always providing a different perspective on life.
I finally acknowledge other friends, both new and old, who have made life as wonderful as it is
for me. Thanks to H. Schwartz, Audrey Chang, Vaishali and Sanjay Dixit, Janice Lee, Sheila
Menzies, and Ryan McQuade for the companionship, laughs, and memories. Thanks also to
Thea Buckingham, Amber Habib, Irit Pandey, Shandev Rai, Adrian Recinos, Aparna Vadlamani,
and Elinor Velasquez for being there through the years. My life would be grayer had I not had
the privilege of meeting all of you.
3
TABLE OF CONTENTS
Abstract
2
Acknowledgements
3
Chapter one:
5
Chapter two:
Clp/Hsp100 proteins: Disassembly chaperones and protease
complexes vital to cellular quality control
Figure 1
Table 1
Figure 2
9
10
23
C-terminal domain mutations in ClpX uncouple substrate
binding from an engagement step required for unfolding
(Shilpa A. Joshi, Tania A. Baker, Robert T. Sauer, 2003.
Molecular Microbiology, 48, pp. 67-76.)
64
Figure 1
Figure 2
Figure 3
Figure 4
Figure 5
Figure 6
Figure 7
69
71
75
77
79
81
87
Chapter three: Communication between ClpX and ClpP during substrate
processing and degradation
(Shilpa A. Joshi, Greg L. Hersch, Tania A. Baker, Robert T. Sauer,
2004. Submitted, Nature Structural and Molecular Biology.)
Figure 1 (work of G.L.H.)
Figure 2
Figure 3
Figure 4
Figure 5
Figure 6
101
107
109
111
115
117
121
Biographical note:
137
4
CHAPTER ONE
Clp/Hsp100 proteins: Disassembly chaperones and
protease complexes vital to cellular quality control
5
Proteins carry out critical macromolecular functions in every living cell. In order to work
properly, however, proteins must be correctly folded. Within the crowded cellular environment,
there is ample opportunity for proteins to become misfolded or aggregated. It is not surprising,
therefore, that cells have evolved molecular systems to help proteins fold properly and to help
take apart aggregates. Hsp70 and Hsp60 proteins help proteins fold and much is known about
the activities of these protein families. Dismantling of protein aggregates is carried out by
members of the more recently discovered Clp/Hsp100 family.
Hsp70 and Hsp60 chaperone systems
Classical
chaperone
systems,
like
DnaK/DnaJ
(Hsp70/Hsp40)
and
GroEL/GroES
(Hsp60/Hsp10), function to assist protein folding. Prokaryotic DnaK/DnaJ and GroEL/GroES
are heat shock proteins which are transcriptionally upregulated (Cowing et al., 1985; Yamamori
and Yura, 1980) in response to diverse kinds of cellular stress in addition to high temperature
(VanBogelen et al., 1987). Escherichia coli DnaK and GroEL, which are the main chaperone
components of each system, bind hydrophobic sequences (Houry et al., 1999; Rudiger et al.,
1997) presumably exposed because a given protein is unfolded or improperly folded. DnaK
tends to bind extended stretches of amino acids, while GroEL binds more compact shapes (Flynn
et al., 1993; Hayer-Hartl et al., 1994; Landry et al., 1992). The DnaK system is believed
important for nascent polypeptide folding and for protein translocation across membranes
(Frydman et al., 1994; Gaitanaris et al., 1994; Langer et al., 1992a; Petit et al., 1994; Wild et al.,
1992; Wild et al., 1996), whereas the GroEL system is thought by some to be more important for
the refolding of misfolded proteins (Gordon et al., 1994; Viitanen et al., 1992). Substrates bound
by the DnaK system are maintained in a soluble folding-competent state by repeated cycles of
6
binding to the exposed hydrophobic patches and release (Buchberger et al., 1996; Szabo et al.,
1994). In contrast, GroEL is believed to help bound proteins fold by providing a protected
environment within an encapsulated chamber where aggregation cannot occur (Fenton and
Horwich, 1997; Ranson et al., 1995; Walter et al., 1996; Zahn et al., 1996).
ATP binding and hydrolysis regulates the reaction cycles of both systems. In E. coli, the DnaK
system involves a nucleotide-exchange factor, GrpE, in addition to the co-chaperone DnaJ,
whereas GroEL functions with its co-chaperone, GroES (Friedman et al., 1984; Georgopoulos et
al., 1973; Georgopoulos et al., 1972; Sternberg, 1973). DnaKATP binds substrate proteins
weakly, while DnaKADP assumes a different conformation which binds substrates more tightly
(Palleros et al., 1993; Palleros et al., 1994; Pierpaoli et al., 1997). DnaJ stimulates DnaK’s
ATPase activity (McCarty et al., 1995; Szabo et al., 1994), and GrpE binding causes ADP
release from DnaKADPsubstrate (Szabo et al., 1994). In one model, a DnaKATPsubstrate
complex binds DnaJ and is converted into DnaKADPsubstrate, whereas in another model this
same complex is formed after DnaJsubstrate binds DnaKATP (Fink, 1999). GrpE then causes
ADP to be released from DnaKADPsubstrate, and subsequent ATP binding causes both a
conformational change and substrate release (Buchberger et al., 1995; Palleros et al., 1993;
Theyssen et al., 1996). In addition to being regulated by ATP binding and hydrolysis, GroEL’s
reaction cycle is modulated by GroES binding.
GroEL is composed of two back-to-back
heptameric rings (Braig et al., 1994), which form two cylindrical chambers, each with a single
opening at its end. GroES binds to the end of one ring, closing off that GroEL chamber from the
cellular environment (Langer et al., 1992b; Xu et al., 1997). Substrate proteins originally bind to
hydrophobic surfaces in a GroEL chamber (Braig et al., 1994; Braig et al., 1993; Fenton et al.,
7
1994), and only have a chance to refold when the inner surfaces of the GroEL cavity become
hydrophilic as a result of a dramatic conformational change after GroES has bound (Chen et al.,
1994; Roseman et al., 1996; Xu et al., 1997). Differences in affinity between GroES and the
different nucleotide-bound GroEL forms (Jackson et al., 1993) and negative allostery in the
system (Hayer-Hartl et al., 1995; Todd et al., 1994; Xu et al., 1997) drive the reaction cycle
forward.
Functional overview of the Clp/Hsp100 protein family
Cellular roles.
Unlike the Hsp70 and Hsp60 chaperone systems (Goloubinoff et al., 1989;
Schroder et al., 1993; Veinger et al., 1998), the Clp/Hsp100 protein family’s main function is to
disassemble macromolecular complexes or disaggregate protein aggregates after they have
formed. Clp/Hsp100 proteins take apart some multimers by unfolding individual components
(Burton et al., 2001a; Kim et al., 2000; Weber-Ban et al., 1999). Some Clp proteins also
function as part of ATP-dependent proteases (Figure 1) (Gottesman et al., 1997a; Gottesman et
al., 1997b; Wickner et al., 1999). The first energy-dependent protease was discovered in E. coli.
It was known that when ATP generation was disrupted, protein degradation in vivo decreased by
90-95% (Goldberg and St John, 1976).
Soon thereafter, an ATP-dependent protease, Lon
(protease La), was isolated and identified (Charette et al., 1981; Chung and Goldberg, 1981;
Swamy and Goldberg, 1981). However, casein, a model substrate, was still found to be degraded
in lysates from cells with a disrupted lon gene (Maurizi et al., 1985). Using casein degradation
as an assay, the ATP-dependent protease, ClpAP, was discovered (Katayama et al., 1988;
Katayama-Fujimura et al., 1987).
ClpA was shown to be an ATPase, whereas ClpP had
peptidase activity.
8
Figure 1. Some Clp proteins have dual functions.
E. coli
ClpX/ClpA/HslU
Disassembled/
disaggregated
substrate
Substrate
E. coli
ClpXP/ClpAP/HslUV
protease
Degraded
substrate
Table 1. Clp proteins support various biological processes
Representative processes
Developmental
stage/process
Organism
Stress
tolerance
A. thaliana
ClpB (heat)
B. subtilis
ClpP
ClpX; ClpP
(sporulation)
B. thuringiensis
ClpP1
ClpC/ClpP2
(sporulation)
L. lactis
Virulence
process
ClpP
ClpP1
ClpP
ClpP2
ClpB
ClpC
ClpE
ClpC/ClpE/
ClpP
M. leprae
ClpC
M. tuberculosis
ClpC
ClpP (biofilm
formation)
P. fluorescens
S. cerevisiae
Hsp104
(heat)
S. typhimurium
ClpP
S. hyodysenteriae
ClpC
S. aureus
ClpX
S. pneumoniae
ClpC (heat)
Y. enterocolitica
ClpC
ClpC/ClpE
ClpP (mycelium
differentiation)
S. coelicolor
Synechococcus sp.
PCC 7942
Uptake
of DNA
ClpX
(bacteriophage
Mu cell-lysis)
ClpA/ClpB/
HslU (heat);
ClpX/ClpP
(many)
ClpP (heat,
pH)
L. donovani
L. monocytogenes
Cell
motility
ClpP
C. crescentus
E. coli
Cell
division
ClpB/ClpP1
(thermal);
ClpP1 (UVB light)
ClpP
ClpC
Since the discovery of E. coli ClpA, ATPases of the Clp/Hsp100 family have been found to be
highly conserved and to exist in all biological kingdoms (Gottesman et al., 1990b; Maurizi et al.,
1990b; Neuwald et al., 1999; Schirmer et al., 1996; Squires and Squires, 1992). Known Clp
ATPases include ClpA, ClpB, ClpC, ClpE, ClpX, and ClpY. Many of these ATPases associate
with peptidases to form energy-dependent proteases. Clp/Hsp100 family ATPases and proteases
have critical roles in a diverse variety of processes in many different prokaryotic and eukaryotic
organisms (Table 1). For example, Clp proteins are key parts of stress tolerance systems.
Lactococcus lactis ClpP is involved in tolerance to heat and pH changes, and cells lacking ClpP
are much less viable (Frees and Ingmer, 1999). Synechococcus sp. PCC 7942, a cyanobacterial
strain, requires ClpB for survival after heat stress (Eriksson and Clarke, 1996) and ClpP1 for
long-term acclimation to different stresses, particularly cold temperatures and exposure to UV-B
light (Clarke et al., 1998; Porankiewicz et al., 1998). Heat stress tolerance in Arabidopsis
thaliana and Saccharomyces cerevisiae require ClpB or Hsp104, respectively (Hong and
Vierling, 2000; Sanchez and Lindquist, 1990). In Bacillus subtilis, Listeria monocytogenes, and
Streptococcus pneumoniae, Clp proteins function in stress tolerance and in other processes. In B.
subtilis, ClpP is important for tolerance to heat, salt, oxidative stress, exposure to ethanol and
puromycin, and glucose and oxygen deprivation (Gerth et al., 1998; Kruger et al., 1994; Msadek
et al., 1998; Volker et al., 1994). Additionally, ClpP is vital for competence development
(Turgay et al., 1998), sporulation (Nanamiya et al., 2000), correct cell morphology and motility
(Gerth et al., 1998), and ClpX and ClpP are critical for proper developmental regulation of
transcription (Nakano et al., 2001; Nakano et al., 2003). L. monocytogenes ClpC is involved in
stress tolerance (Rouquette et al., 1996) and is important for virulence, as are ClpE and ClpP
(Gaillot et al., 2000; Nair et al., 1999; Rouquette et al., 1998). ClpE is also important for cell
11
division in L. monocytogenes (Nair et al., 1999). ClpC in S. pneumoniae is involved in heat
tolerance, cell division, autolysis, genetic transformability, ability to adhere to host cells
(Charpentier et al., 2000), and virulence, as is ClpE (Lau et al., 2001; Polissi et al., 1998).
Streptomyces coelicolor ClpP is important for the differentiation of mycelium (de Crecy-Lagard
et al., 1999), whereas Synechococcus sp. PCC 7942 ClpP null cells are filamentous and grow
slowly (Clarke et al., 1998). Virulence roles have been ascribed for Salmonella typhimurium
ClpP (Hensel et al., 1995), Yersinia enterocolitica ClpP (Pederson et al., 1997), Staphylococcus
aureus ClpX (Mei et al., 1997), Serpulina hyodysenteriae ClpC (ter Huurne et al., 1994),
Mycobacterium leprae ClpC (Misra et al., 1996), Mycobacterium tuberculosis ClpC (Misra et
al., 1996), and Leishmania donovani ClpB (Hubel et al., 1997; Krobitsch and Clos, 1999).
Bacillus thuringiensis ClpC and ClpP2 are involved in sporulation, ClpP2 is important for
motility, and ClpP1 is critical for cell division and response to low temperatures (Fedhila et al.,
2002). Finally, Caulobacter crescentus ClpXP is needed for progression through the cell cycle
and for viability (Jenal and Fuchs, 1998; Jenal and Shapiro, 1996); Pseudomonas fluorescens
ClpP is needed for biofilm formation (O'Toole and Kolter, 1998); Caenorhabditis elegans OOC5, a putative Clp protein, is necessary for mitotic spindle orientation and polarity in embryos
(Basham and Rose, 1999; Basham and Rose, 2001); and the Chlamydomonas plastid ClpP is
essential for correct chloroplast function and for resulting algal growth (Huang et al., 1994).
Biochemical studies of Clp/Hsp100 ATPases and proteases have focused most intensively on the
E. coli enzymes. E. coli contains four Clp ATPases (ClpX (Gottesman et al., 1993; Wojtkowiak
et al., 1993), HslU (Chuang et al., 1993; Rohrwild et al., 1996), ClpA (Katayama et al., 1988),
and ClpB (Kitagawa et al., 1991)) and three of these proteins also form bipartite protease
12
complexes with either the ClpP (Katayama et al., 1988; Maurizi et al., 1990a) or HslV (Chuang
et al., 1993; Rohrwild et al., 1996) peptidases. ClpX or ClpA interact with ClpP to form the
ClpXP or ClpAP proteases, respectively, whereas HslU joins HslV to form the HslUV protease.
In genetic studies, E. coli Clp proteins often function to combat cellular stress. ClpX-defective
cells survive less well after UV irradiation (Neher et al., 2003a) and cannot be lysed by
bacteriophage Mu (Mhammedi-Alaoui et al., 1994). Additionally, both ClpX-defective and
ClpP-defective cells are less viable during lengthened stationary phase (Weichart et al., 2003),
die after UV exposure under amino acid deprivation (Burger et al., 2002), are poorly mutable by
UV light (Frank et al., 1996), are defective in acquiring type IA and IB restriction systems
(Makovets et al., 1998), and do not support restriction alleviation of the type IA system after
DNA damage (Makovets et al., 1999). Cells lacking HslU are inhibited for growth at high
temperatures and are smaller in size at low temperatures (Katayama et al., 1996).
ClpA-
defective cells have growth defects at 46 °C (Thomas and Baneyx, 1998), but not at temperatures
between 25 °C and 42 °C (Katayama et al., 1988). ClpB-defective cells also show problems
with thermal stress, as they grow slowly at 44 °C and survive poorly at 50 °C (Squires et al.,
1991). ClpB-defective and ClpP-defective cells also fail to grow with more than 0.5% SDS,
whereas wild-type cells grow in the presence of 20-fold higher detergent concentrations
(Rajagopal et al., 2002). Furthermore, ClpP-defective cells show delayed recovery after repeated
cycles of carbon starvation and growth (Damerau and St John, 1993), and HslV-minus cells are
lethal under conditions where there is an altered amount of signal recognition particle present
(Bernstein and Hyndman, 2001). Finally, recent studies suggest that ClpXP may be important in
response to oxidative stress and to changes between aerobic and anaerobic conditions (Flynn et
al., 2003).
13
E. coli Clp proteins function as disassembly chaperones and proteases.
ClpB is believed to act
only as a disassembly chaperone, whereas HslU, ClpA, and ClpX have additional roles as part of
HslUV, ClpAP, or ClpXP, three of E. coli’s five known ATP-dependent proteases (Gottesman,
1996; Schirmer et al., 1996; Wickner et al., 1999).
ClpB is particularly important for
solubilizing large protein aggregates (Diamant et al., 2000; Laskowska et al., 1996; Thomas and
Baneyx, 2000), a function which also requires the DnaK chaperone system (Zolkiewski, 1999).
ClpB is thought to act first in aggregate disassembly, exposing hydrophobic patches which are
then bound by the DnaK system (Goloubinoff et al., 1999; Weibezahn et al., 2003). Aggregated
proteins which have small Hsps bound also are disaggregated by ClpB with DnaK/DnaJ/GrpE
(Mogk et al., 2003a). This ClpB/DnaK system acts more specifically when it disassembles
dimers of TrfA, plasmid RK2’s replication initiation protein, to active monomers (Konieczny
and Liberek, 2002).
In contrast to ClpB’s general disassembly of large protein aggregates, known chaperone
activities for the remaining Clp proteins are far more substrate specific. HslU disaggregates
SulA, an inhibitor of cell division, both in vivo and in vitro (Seong et al., 2000). ClpA activates
P1 RepA DNA-binding activity by converting inactive RepA dimers into monomers (Wickner et
al., 1994). ClpX disassembles three known substrates. In two cases, TrfA (Konieczny and
Helinski, 1997) and O (Wawrzynow et al., 1995), ClpX disassembles dimers into monomers.
In the third case, disassembly of the MuA4-DNA complex, an interesting phenotype results if
ClpX is not present to take apart this protein-DNA complex (Kruklitis et al., 1996; Levchenko et
al., 1995; Mhammedi-Alaoui et al., 1994). Namely, bacteriophage Mu cannot replicate in or
14
lyse bacterial cells lacking functional ClpX because failure to disassemble the MuA4-DNA
complex prevents replication of Mu DNA. The MuA4-DNA complex is an intermediate in a
recombination reaction catalyzed by MuA (Mu transposase). The timing of ClpX action is
controlled exquisitely by MuB, a protein which stimulates MuA’s catalysis of recombination
events (Baker et al., 1991).
MuB and ClpX bind overlapping parts of MuA’s C-terminus
(Levchenko et al., 1997). ClpX commences disassembly only after recombination is completed
and MuB has dissociated because ClpX-mediated disassembly requires recognition of MuA’s
last ten amino acids (Levchenko et al., 1997).
Like the proteasome, Clp proteases are compartmentalized, barrel-shaped machines with their
peptidase active sites sequestered in an internal chamber accessible only by passage through
narrow axial portals (Larsen and Finley, 1997). Therefore, native proteins must be denatured
and translocated into the proteolytic chamber prior to degradation. The list of substrates for
degradation by HslUV, ClpAP, and ClpXP is growing steadily. HslUV degrades SulA in vivo
and in vitro (Kanemori et al., 1999; Seong et al., 1999; Wu et al., 1999) and more generally
degrades misfolded proteins (Missiakas et al., 1996). ClpAP degrades itself (Gottesman et al.,
1990a), ClpA-lacZ fusion proteins (Gottesman et al., 1990a), P1 RepA (Wickner et al., 1994),
HemA (glutamyl-tRNA reductase) (Wang et al., 1999), some carbon starvation proteins
(Damerau and St John, 1993), and heat-aggregated proteins (Dougan et al., 2002). ClpXP
degrades O (Gottesman et al., 1993; Wojtkowiak et al., 1993), s (RpoS) (Schweder et al.,
1996), UmuD´ in a UmuD/UmuD´ heterodimer (Frank et al., 1996), Mu vir repressors
(Mhammedi-Alaoui et al., 1994; Welty et al., 1997), P1 Phd (Lehnherr and Yarmolinsky, 1995),
the HsdR subunit of EcoKI (Doronina and Murray, 2001), and Dps (Flynn et al., 2003; Stephani
15
et al., 2003). Recently, more than 50 new ClpXP substrates were identified (Flynn et al., 2003),
including transcription factors, ribosomal proteins, stress response proteins, and metabolic
enzymes. Additionally, ClpXP and ClpAP process HlyC in the presence of HlyA (GuzmanVerri et al., 2001) and degrade ssrA-tagged proteins (Gottesman et al., 1998). The latter proteins
contain an 11-residue C-terminal ssrA degradation tag that is added to nascent chains on stalled
ribosomes via a cotranslational process (Karzai et al., 2000; Keiler et al., 1996). ClpXP and
ClpAP degradation of ssrA-tagged proteins is part of a generalized protein quality control
system.
Structural overview of E. coli Clp/Hsp100 proteins
AAA+ protein family functions.
The Clp/Hsp100 protein family is part of the AAA+ ATPase
family, a large group of structurally related proteins which are involved in a diverse variety of
cellular activities, including membrane fusion, proteolysis, DNA replication and repair,
recombination, and transcription (Neuwald et al., 1999). Most AAA+ ATPases use nucleotide
binding or hydrolysis to exert mechanical force, thereby remodeling macromolecular complexes
and assisting in their assembly, disassembly, or activation (Neuwald et al., 1999; Ogura and
Wilkinson, 2001). An ancient family, AAA+ ATPases are found in all three divisions of life
(bacteria, archaeabacteria, and eukaryotes) (Neuwald et al., 1999), underscoring the functional
importance of AAA+ proteins. Two subgroups, AAA proteins and Replication Factor C (RFC)related clamp-loader proteins, are present in all organisms. AAA proteins are a group of related
chaperones which function in diverse processes (Confalonieri and Duguet, 1995; Patel and
Latterich, 1998; Swaffield et al., 1995). One AAA subfamily, p97/Cdc48/VCP, is found in some
bacteria (like M. tuberculosis) as well as in all archaeabacteria and eukaryotes (Neuwald et al.,
16
1999).
p97/Cdc48 is important for Golgi, endoplasmic reticulum, and nuclear envelope
membrane fusion events (Acharya et al., 1995; Latterich et al., 1995; Rabouille et al., 1995) and
for endoplasmic reticulum-associated degradation (Kopito, 1997; Rabinovich et al., 2002).
Other important AAA proteins include the 19S regulatory component ATPases of the 26S
proteasome (Baumeister and Lupas, 1997; Baumeister et al., 1998) and the N-ethylmaleimidesensitive fusion protein (NSF) (Hay and Scheller, 1997; Weber et al., 1998), which is critical for
SNARE-mediated membrane fusion events (Hanson et al., 1997; May et al., 2001; Sollner et al.,
1993).
RFC-related clamp-loader proteins are the most conserved AAA+ proteins involved in DNA
replication, but other less conserved AAA+ proteins are also involved in replication and repair.
The five S. cerevisiae RFC proteins which comprise the heteropentameric clamp-loader complex
are AAA+ proteins (Cullmann et al., 1995; Neuwald et al., 1999). PCNA, the trimeric ringshaped processivity factor, is loaded onto DNA by the RFC clamp loader to initiate DNA
synthesis (Kelman and O'Donnell, 1995; Krishna et al., 1994). An analogous system, involving
the -complex clamp loader and -clamp processivity factor, exists in E. coli (Jeruzalmi et al.,
2001; Kong et al., 1992). Four of the five essential -complex components (Pritchard et al.,
2000; Stewart et al., 2001; Studwell and O'Donnell, 1990) are AAA+ proteins (Neuwald et al.,
1999), though only one type () is capable of interacting with nucleotide (Xiao et al., 1995).
Less conserved AAA+ proteins are involved in DNA replication initiation and repair. Initiation
of DNA replication is mediated in S. cerevisiae and other eukaryotes by sequential assembly of
different protein complexes at origins of replication (Aparicio et al., 1997; Bell and Stillman,
1992; Diffley and Cocker, 1992; Donovan et al., 1997; Tanaka et al., 1997). A hexameric ORC
17
(Bell et al., 1993), composed of Orc1-Orc6, binds the origin first and then recruits Cdc6 (Liang
et al., 1995), which in turn recruits the heterohexameric MCM (Chong et al., 1996) protein
complex (Aparicio et al., 1997; Coleman et al., 1996; Tanaka et al., 1997).
Subsequent
phosphorylation of MCM complex components leads to replication initiation (Bousset and
Diffley, 1998; Lei et al., 1997; Santocanale and Diffley, 1998). Orc1, Orc4, Orc5, Cdc6, and all
six MCM proteins are AAA+ family members (Neuwald et al., 1999). In E. coli, the AAA+
DnaA protein, which is conserved in all bacteria (Neuwald et al., 1999), binds oriC replication
origins in a manner functionally similar to eukaryotic ORC (Baker et al., 1986; Fuller et al.,
1984). The AAA+ Rad17 (Neuwald et al., 1999) in Schizosaccharomyces pombe and humans is
a DNA-damage checkpoint protein with homology to RFC1 (of the eukaryotic clamp loader) (alKhodairy et al., 1994; Enoch et al., 1992; Freire et al., 1998; Griffiths et al., 1995). Human
Rad17 associates with RFC2-RFC5 and a PCNA-like clamp composed of three checkpoint
proteins (Caspari et al., 2000; Lindsey-Boltz et al., 2001), suggesting a role in repair of lesions
for this clamp loader and associated clamp (Kelly and Brown, 2000; O'Connell et al., 2000;
Venclovas and Thelen, 2000).
AAA+ proteins also have interesting roles in recombination and transcription. The AAA+ RuvB
helicase is found in all bacteria except the extreme thermophile, Aquifex aeolicus (Neuwald et
al., 1999). RuvB promotes DNA branch migration during recombination at Holliday junctions
(Rice et al., 1997; West, 1997). Two hexameric RuvB rings sandwich RuvA (Iwasaki et al.,
1989) and a channel passing through the complex is used presumably for DNA threading (Yu et
al., 1997) during branch migration. Bacterial NtrC protein is a AAA+ transcriptional enhancer
which helps regulate the expression of genes involved in nitrogen metabolism (Magasanik,
18
1989). Using energy from ATP hydrolysis, NtrC helps convert a closed complex of promoterbound RpoN(54)-RNA polymerase to an active open complex (Hwang et al., 1999; Porter et al.,
1993; Wedel and Kustu, 1995; Wyman et al., 1997). NtrC is found in all bacterial species which
contain RpoN (Neuwald et al., 1999).
AAA+ family structural characteristics.
The AAA+ ATPase family belongs to a large group,
the P-loop-type NTPases, which have nucleotide-binding sites defined by classical Walker-A and
Walker-B motifs (Gorbalenya and Koonin, 1989; Koonin, 1993b; Saraste et al., 1990; Walker et
al., 1982). AAA+ proteins further belong to a subgroup, the motif C (or sensor I)-containing
ATPases (Koonin, 1993a; Neuwald et al., 1999). Like other P-loop-type NTPases, AAA+
proteins contain a conserved core subdomain with an ,-fold, generally similar to the RecA
ATP-binding domain (Hubbard et al., 1997; Neuwald et al., 1999; Story and Steitz, 1992). This
subdomain, typically of 200-250 residues, contains the Walker-A and Walker-B motifs in a
structure which contains a five-stranded -sheet with two or three -helices on either side
(Confalonieri and Duguet, 1995; Neuwald et al., 1999; Patel and Latterich, 1998). The AAA+
module, however, is comprised of a larger structural unit, which includes a smaller
predominantly -helical C-terminal subdomain (Neuwald et al., 1999). Nucleotide is bound
between the large and small subdomains (Lupas and Martin, 2002). Amino acids from both
subdomains of a AAA+ module, and sometimes residues from an adjacent subunit of a
multimeric AAA+ protein, generally contact ATP/ADP. Some AAA+ proteins neither bind nor
hydrolyze ATP. For instance, ´ of the E. coli -complex clamp loader has a disrupted P-loop
motif (Guenther et al., 1997), and at least one of the AAA+ clamp-loader subunits in most
bacteria also have disrupted P-loops (Neuwald et al., 1999).
19
In addition to the Walker motifs, AAA+ proteins contain a sensor-I motif and often contain
other motifs called RFC boxes because they were initially identified in RFC-related proteins
(Cullmann et al., 1995; Guenther et al., 1997; Neuwald et al., 1999). For example, the RFC boxII, box-VII, and sensor-II motifs are conserved to different extents in different AAA+ proteins
(Neuwald et al., 1999). In contrast, AAA proteins usually do not contain a sensor-II motif, but
instead contain a conserved 19-residue motif called the SRH (for second region of homology)
(Karata et al., 1999; Karata et al., 2001; Neuwald et al., 1999). AAA proteins thus constitute a
distinct subgroup of the AAA+ protein family. In ATPases containing Walker motifs, it appears
that ATP is hydrolyzed by an activated water molecule which attacks the -phosphate of the
bound ATP (Karata et al., 2001; Ogura and Wilkinson, 2001). Negative charges in the pentacoordinate transition state are stabilized in part by a magnesium ion. Amino acids from different
conserved motifs stabilize interactions with phosphate groups or the magnesium ion, or are
involved in activating the water by extracting a proton. The Walker-A motif (P loop) is Gly(X)4-Gly-Lys-Thr, where X is any amino acid (Ogura and Wilkinson, 2001; Saraste et al., 1990;
Walker et al., 1982). The P-loop lysine interacts with the - and -phosphates of bound ATP,
while the threonine helps bind the magnesium. Walker B is (H)4-Asp-Glu-(X)2, where H is a
hydrophobic amino acid and X is any residue.
The aspartate also helps coordinate the
magnesium ion, whereas glutamate is often the residue which activates a water molecule for
attack. The sensor-I motif in the large ,-subdomain typically contains a conserved asparagine
or threonine residue important for sensing whether the ATP -phosphate is present (Guenther et
al., 1997; Koonin, 1993a), whereas the RFC box-II motif appears to be involved in recognition
of the ATP adenine moiety (Guenther et al., 1997; Lenzen et al., 1998). Both the box-VII and
20
sensor-II motifs are found in the small C-terminal subdomain and both typically contain a highly
conserved arginine. The box-VII arginine, however, is believed to sense nucleotide binding in
an adjacent subunit, whereas the sensor-II arginine is important for sensing whether the terminal
-phosphate of the ATP/ADP bound to the same subunit is present (Lenzen et al., 1998; Neuwald
et al., 1999; Yu et al., 1998).
Domain structure of E. coli Clp proteins.
ClpX (Kim and Kim, 2003), HslU (Bochtler et al.,
2000; Sousa et al., 2000; Wang et al., 2001), ClpA (Guo et al., 2002b), and ClpB (Lee et al.,
2003) all form hexameric rings with a central axial channel (Beuron et al., 1998; Ortega et al.,
2000). ClpX and HslU contain a single AAA+ module, whereas ClpA and ClpB contain two
modules (Figure 2).
The ClpX and HslU ,-subdomains are homologous to the ,-
subdomains in the second AAA+ module of ClpA and ClpB (Gottesman et al., 1993; Schirmer et
al., 1996). Whereas ClpA’s first ,-subdomain (NBD1) is important for hexamerization and its
second ,-subdomain (NBD2) is critical for ATP hydrolysis activity (Singh and Maurizi, 1994),
the opposite is true for ClpB’s NBD1 and NBD2 (Barnett and Zolkiewski, 2002; Mogk et al.,
2003b). In addition, ClpX contains a unique N-terminal domain (Donaldson et al., 2003), HslU
contains an intermediate domain (I-domain) between the Walker-A and –B motifs in its AAA+
module (Schirmer et al., 1996), and ClpA and ClpB both have their own conserved N-terminal
domains (Guo et al., 2002a; Li and Sha, 2003; Zeth et al., 2002). ClpB also contains a coiledcoil intermediate domain (I-domain) near the junction between its AAA+ modules (Lee et al.,
2003; Schirmer et al., 1996). ClpX and ClpA, which both interact with the ClpP peptidase, also
share a conserved tripeptide motif required for binding ClpP in a surface loop (ClpP-loop) in
their homologous ,-subdomains (Kim et al., 2001; Singh et al., 2001).
21
Interestingly,
alternative translational initiation sites (Seol et al., 1994; Squires et al., 1991; Woo et al., 1992)
in both the clpA and clpB genes result in synthesis of some ClpA and ClpB proteins lacking the
conserved N-terminal domain.
The subfamily-specific domains in the Clp/Hsp100 proteins may help mediate recognition of
substrate proteins. ClpX and ClpXP bind substrates either directly, usually via N- or C-terminal
peptide sequences (Flynn et al., 2003), or indirectly, via interactions with adaptor proteins which
themselves bind substrates. Known ClpX adaptors include SspB (Levchenko et al., 2000), RssB
(Zhou et al., 2001), and UmuD (Neher et al., 2003b). ClpX’s N-domain is important for
processing O and MuA, but not ssrA-tagged substrates (Ortega et al., 2000; Singh et al., 2001;
Wojtyra et al., 2003). Likewise, delivery of ssrA-tagged substrates by SspB requires this Ndomain (Dougan et al., 2003; Wojtyra et al., 2003). HslU’s I-domain is involved in recognition
of some substrates but not others (Song et al., 2000). Like ClpXP, ClpAP also has an adaptor
protein, ClpS (Dougan et al., 2002), which binds its N-domain (Dougan et al., 2002; Guo et al.,
2002a; Zeth et al., 2002). In the presence of ClpP, a ClpA protein lacking the N-domain no
longer degrades casein (Seol et al., 1994; Singh et al., 2001) or RepA (Lo et al., 2001), but still
degrades ssrA-tagged substrates (Dougan et al., 2002; Lo et al., 2001; Singh et al., 2001). ClpB
lacking its N-domain is defective for disaggregation of casein (Barnett et al., 2000; Park et al.,
1993) and insulin (Park et al., 1993), but still can disaggregate heat-aggregated MDH and
luciferase (Mogk et al., 2003b). Interestingly, specific ClpB N-domain point mutants have been
constructed which bind but no longer disaggregate heat-treated luciferase (Liu et al., 2002),
suggesting a role for this domain beyond substrate binding. Consistent with this model, the I-
22
Figure 2. Domain structure of E. coli Clp proteins
ClpA:
N-domain
ATPase 1
ATPase 2
Cdom.
ClpP-loop
ClpB:
N-domain
ATPase 1
ATPase 2
Cdom.
ATPase 2
Cdom.
I-domain
ClpX:
Ndom.
ClpP-loop
HslU/ClpY:
ATPase 2
I-domain
Cdom.
24
domain of ClpB has been suggested to be critical for substrate binding (Lee et al., 2003; Mogk et
al., 2003b).
Substrate unfolding and degradation by ClpXP and ClpAP
ClpX and ClpA either unfold proteins or disassemble macromolecular complexes on their own,
or act as the unfoldase components within the ClpXP or ClpAP proteases (Gottesman et al.,
1997a; Gottesman et al., 1997b; Wickner et al., 1999). Protein unfolding and translocation
provides a mechanism for substrate disassembly, as well as a mechanism to degrade substrates in
association with a compartmentalized peptidase. In the former case, the substrate is released to
the cellular environment, whereas, in the latter case, the substrate is degraded. ClpXP and
ClpAP are composed of hexameric ClpX or ClpA rings stacked coaxially against the double-ring
ClpP14 peptidase (Beuron et al., 1998; Grimaud et al., 1998; Kessel et al., 1995; Ortega et al.,
2000). The two heptameric ClpP rings are assembled face-to-face with the peptidase active sites
at the internal ClpP ring interface (Wang et al., 1997). A channel or pore passes through the
ClpX or ClpA hexamers into the degradation chamber of ClpP. This channel is too narrow to
permit the passage of folded proteins (Wang et al., 1997) which must first be denatured and then
translocated into ClpP prior to cleavage into peptides. Electron microscopy studies show that the
channels in ClpXP and ClpAP are used for substrate translocation (Ishikawa et al., 2001; Ortega
et al., 2002; Ortega et al., 2000).
ClpX and ClpA are responsible for substrate recognition by ClpXP and ClpAP. Generally, ClpX
and ClpA bind specific peptide sequences at the N- or C-termini of substrate proteins (Flynn et
al., 2003; Gonciarz-Swiatek et al., 1999; Gottesman et al., 1990a; Gottesman et al., 1998;
25
Hoskins et al., 2000a; Levchenko et al., 1997). Sometimes, however, these sequences are hidden
and require other events, like proteolysis, in order to be accessible. For instance, intact LexA is
not a ClpXP substrate, but its N-terminal domain becomes a substrate after it has been separated
from LexA’s C-terminal domain by auto-proteolytic cleavage (Neher et al., 2003a). In other
instances, adaptors are required for degradation. RpoS (s) is a substrate for ClpXP when it is
presented by a phosphorylated form of RssB, an adaptor protein (Hengge-Aronis, 2002; Pratt and
Silhavy, 1996; Zhou et al., 2001). During stationary phase, however, RssB is not phosphorylated
and no longer binds s. Stable s is then free to activate transcription of genes needed in
stationary phase. Adaptor proteins also can improve the recognition of substrates which can be
bound on their own. For example, SspB improves recognition by ClpX of ssrA-tagged substrates
(Levchenko et al., 2000). A further level of control exists in this instance because SspB binding
to the ssrA tag prevents ClpA binding, and thus directs ssrA-tagged substrates to ClpX (Flynn et
al., 2001; Gottesman et al., 1998). The adaptor, ClpS, acts in yet another way as it both redirects
ClpA to act on aggregated proteins and inhibits ClpA from binding and degrading ssrA-tagged
substrates (Dougan et al., 2002).
After ClpX or ClpA bind native substrate proteins, they unfold the proteins and either release
them or translocate them to ClpP for degradation. Polypeptide cleavage by ClpP and subsequent
peptide release occur very quickly (Thompson and Maurizi, 1994; Thompson et al., 1994), so
either unfolding or translocation theoretically is rate limiting. Recent studies show that either
step can be rate limiting in ClpXP degradation, depending on the substrate (Kenniston et al.,
2003). ATP hydrolysis is needed for both unfolding and translocation (Burton et al., 2003;
Hoskins et al., 2000b; Kenniston et al., 2003; Kim et al., 2000; Singh et al., 2000). While
26
ClpXP unfolds proteins, it hydrolyzes ATP at a constant rate but spends more time (and more
ATP) when unfolding a more stable substrate (Kenniston et al., 2003). After substrates are
bound via recognition tags, they move into the Clp protein channels in a directional way with
regions close to the tags entering the channels first (Reid et al., 2001). In one model, Clp
ATPases denature substrates by pulling them into the translocation channel by their recognition
tags (Kenniston et al., 2003; Lee et al., 2001; Matouschek, 2003). Both ClpX and ClpA globally
unfold substrate proteins (Burton et al., 2001a; Kim et al., 2000; Singh et al., 2000; Weber-Ban
et al., 1999), if ClpP is not present, and release them in nonnative forms (Fenton et al., 1994;
Kim et al., 2000; Singh et al., 2000; Weber-Ban et al., 1999). Only proteins with recognition
tags are denatured, however, even when these proteins are in multimers with untagged proteins
(Burton et al., 2001a; Burton et al., 2001b; Gonzalez et al., 2000).
Role of communication in substrate unfolding and degradation by ClpXP
Relatively little is known about the extent and importance of inter-subunit communication, either
within a ClpX hexamer during substrate processing or between ClpX and ClpP during processing
and degradation. To date, studies with E. coli ClpA, S. cerevisiae Hsp104, and Mus musculus
p97 suggest that interdomain communication between parts of each protein’s two AAA+
modules is important for protein function. In ClpA, ATP binding or hydrolysis by the first
nucleotide-binding domain (NBD1) is needed for communication to NBD2, and this signaling is
critical for chaperone activity (Pak et al., 1999). Hsp104 uses interdomain communication in
several ways. The six NBD1 (or NBD2) portions communicate with each other, as each domain
shows cooperative ATP hydrolysis kinetics, and NBD2 communicates its nucleotide-binding
state to NBD1 within the same Hsp104 monomer (Hattendorf and Lindquist, 2002).
27
Functionally, communication upon peptide binding between NBD2’s -helical C-terminal
subdomain and NBD1 is needed for chaperone activity (Cashikar et al., 2002).
A
conformational change in Hsp104’s I-domain propagates the substrate-binding signal to NBD1,
and NBD1 hydrolyzes more ATP in response.
Like Hsp104, p97 has interdomain
communication between its substrate-binding domain and its NBD (NBD2) which is important
for ATP hydrolysis (DeLaBarre and Brunger, 2003). Communication also occurs in HslUV and
the 26S proteasome between the ATPase and peptidase components. HslV’s channel into its
peptidase active sites is open (Bochtler et al., 1997), unlike the channel into the S. cerevisiae 20S
proteasome (Groll et al., 1997). In order for HslV to degrade substrates, however, its peptidase
sites require activation by HslU, which is mediated by HslV binding HslU’s C-termini
(Ramachandran et al., 2002; Seong et al., 2002; Sousa et al., 2002; Sousa et al., 2000). In
contrast, the 19S regulatory particle physically opens up the channel into the 20S proteasome,
allowing the already active peptidase sites access to translocated substrates (Groll et al., 2000).
In both cases, communication between the ATPases and peptidases is needed for proteolysis.
My efforts have been directed toward addressing these questions about communication for ClpX
and ClpXP. For ClpX, I have found that intersubunit communication between the -helical Cterminal subdomain of one subunit and the adjacent ,-subdomain of another subunit appears to
be critical for proper response to substrate binding and for resulting unfolding activity (Chapter
two).
Amino acids in the conserved sensor-II helix are important for mediating this
communication and help to connect nucleotide-binding sites in adjacent subunits of the ClpX
hexamer.
To test the extent of communication between ClpX and ClpP in ClpXP during
substrate processing and degradation, I developed a solution assay to monitor the affinity
28
between both enzymes in the working protease. These results show that ClpX and ClpP interact
more tightly during substrate denaturation than translocation, revealing that ClpX communicates
its protein-processing task to ClpP (Chapter three). The apparent affinity between ClpP and
ClpX strengthens significantly when ClpP’s active sites are modified and mimic intermediates
during peptide-bond hydrolysis. Thus, ClpP also communicates to ClpX. Intersubunit contacts
in a ClpX hexamer and some aspect of ATP cycling also affect interactions with ClpP. Overall,
these studies demonstrate that communication between subunits within a ClpX hexamer and
between enzymes in the ClpXP protease helps modulate activity and can be required for
substrate processing activity.
29
References
Acharya, U., Jacobs, R., Peters, J.M., Watson, N., Farquhar, M.G., and Malhotra, V. (1995). The
formation of Golgi stacks from vesiculated Golgi membranes requires two distinct fusion
events. Cell 82, 895-904.
al-Khodairy, F., Fotou, E., Sheldrick, K.S., Griffiths, D.J., Lehmann, A.R., and Carr, A.M.
(1994). Identification and characterization of new elements involved in checkpoint and
feedback controls in fission yeast. Mol Biol Cell 5, 147-160.
Aparicio, O.M., Weinstein, D.M., and Bell, S.P. (1997). Components and dynamics of DNA
replication complexes in S. cerevisiae: redistribution of MCM proteins and Cdc45p
during S phase. Cell 91, 59-69.
Baker, T.A., Mizuuchi, M., and Mizuuchi, K. (1991). MuB protein allosterically activates strand
transfer by the transposase of phage Mu. Cell 65, 1003-1013.
Baker, T.A., Sekimizu, K., Funnell, B.E., and Kornberg, A. (1986). Extensive unwinding of the
plasmid template during staged enzymatic initiation of DNA replication from the origin
of the Escherichia coli chromosome. Cell 45, 53-64.
Barnett, M.E., Zolkiewska, A., and Zolkiewski, M. (2000). Structure and activity of ClpB from
Escherichia coli. Role of the amino- and -carboxyl-terminal domains. J Biol Chem 275,
37565-37571.
Barnett, M.E., and Zolkiewski, M. (2002). Site-directed mutagenesis of conserved charged
amino acid residues in ClpB from Escherichia coli. Biochemistry 41, 11277-11283.
Basham, S.E., and Rose, L.S. (1999). Mutations in ooc-5 and ooc-3 disrupt oocyte formation and
the reestablishment of asymmetric PAR protein localization in two-cell Caenorhabditis
elegans embryos. Dev Biol 215, 253-263.
30
Basham, S.E., and Rose, L.S. (2001). The Caenorhabditis elegans polarity gene ooc-5 encodes a
Torsin-related protein of the AAA ATPase superfamily. Development 128, 4645-4656.
Baumeister, W., and Lupas, A. (1997). The proteasome. Curr Opin Struct Biol 7, 273-278.
Baumeister, W., Walz, J., Zuhl, F., and Seemuller, E. (1998). The proteasome: paradigm of a
self-compartmentalizing protease. Cell 92, 367-380.
Bell, S.P., Kobayashi, R., and Stillman, B. (1993). Yeast origin recognition complex functions in
transcription silencing and DNA replication. Science 262, 1844-1849.
Bell, S.P., and Stillman, B. (1992). ATP-dependent recognition of eukaryotic origins of DNA
replication by a multiprotein complex. Nature 357, 128-134.
Bernstein, H.D., and Hyndman, J.B. (2001). Physiological basis for conservation of the signal
recognition particle targeting pathway in Escherichia coli. J Bacteriol 183, 2187-2197.
Beuron, F., Maurizi, M.R., Belnap, D.M., Kocsis, E., Booy, F.P., Kessel, M., and Steven, A.C.
(1998). At sixes and sevens: characterization of the symmetry mismatch of the ClpAP
chaperone-assisted protease. J Struct Biol 123, 248-259.
Bochtler, M., Ditzel, L., Groll, M., and Huber, R. (1997). Crystal structure of heat shock locus V
(HslV) from Escherichia coli. Proc Natl Acad Sci USA 94, 6070-6074.
Bochtler, M., Hartmann, C., Song, H.K., Bourenkov, G.P., Bartunik, H.D., and Huber, R. (2000).
The structures of HsIU and the ATP-dependent protease HsIU-HsIV. Nature 403, 800805.
Bousset, K., and Diffley, J.F. (1998). The Cdc7 protein kinase is required for origin firing during
S phase. Genes Dev 12, 480-490.
31
Braig, K., Otwinowski, Z., Hegde, R., Boisvert, D.C., Joachimiak, A., Horwich, A.L., and Sigler,
P.B. (1994). The crystal structure of the bacterial chaperonin GroEL at 2.8 Å. Nature
371, 578-586.
Braig, K., Simon, M., Furuya, F., Hainfeld, J.F., and Horwich, A.L. (1993). A polypeptide bound
by the chaperonin GroEL is localized within a central cavity. Proc Natl Acad Sci USA 90,
3978-3982.
Buchberger, A., Schroder, H., Hesterkamp, T., Schonfeld, H.J., and Bukau, B. (1996). Substrate
shuttling between the DnaK and GroEL systems indicates a chaperone network
promoting protein folding. J Mol Biol 261, 328-333.
Buchberger, A., Theyssen, H., Schroder, H., McCarty, J.S., Virgallita, G., Milkereit, P.,
Reinstein, J., and Bukau, B. (1995). Nucleotide-induced conformational changes in the
ATPase and substrate binding domains of the DnaK chaperone provide evidence for
interdomain communication. J Biol Chem 270, 16903-16910.
Burger, A., Raymer, J., and Bockrath, R. (2002). DNA damage-processing in E. coli: on-going
protein synthesis is required for fixation of UV-induced lethality and mutation. DNA
Repair (Amst) 1, 821-831.
Burton, B.M., Williams, T.L., and Baker, T.A. (2001a). ClpX-mediated remodeling of Mu
transpososomes: selective unfolding of subunits destabilizes the entire complex. Mol Cell
8, 449-454.
Burton, R.E., Baker, T.A., and Sauer, R.T. (2003). Energy-dependent degradation: Linkage
between ClpX-catalyzed nucleotide hydrolysis and protein-substrate processing. Protein
Sci 12, 893-902.
32
Burton, R.E., Siddiqui, S.M., Kim, Y.I., Baker, T.A., and Sauer, R.T. (2001b). Effects of protein
stability and structure on substrate processing by the ClpXP unfolding and degradation
machine. EMBO J 20, 3092-3100.
Cashikar, A.G., Schirmer, E.C., Hattendorf, D.A., Glover, J.R., Ramakrishnan, M.S., Ware,
D.M., and Lindquist, S.L. (2002). Defining a pathway of communication from the Cterminal peptide binding domain to the N-terminal ATPase domain in a AAA protein.
Mol Cell 9, 751-760.
Caspari, T., Dahlen, M., Kanter-Smoler, G., Lindsay, H.D., Hofmann, K., Papadimitriou, K.,
Sunnerhagen, P., and Carr, A.M. (2000). Characterization of Schizosaccharomyces
pombe Hus1: a PCNA-related protein that associates with Rad1 and Rad9. Mol Cell Biol
20, 1254-1262.
Charette, M.F., Henderson, G.W., and Markovitz, A. (1981). ATP hydrolysis-dependent protease
activity of the Lon (CapR) protein of Escherichia coli K-12. Proc Natl Acad Sci USA 78,
4728-4732.
Charpentier, E., Novak, R., and Tuomanen, E. (2000). Regulation of growth inhibition at high
temperature, autolysis, transformation and adherence in Streptococcus pneumoniae by
ClpC. Mol Microbiol 37, 717-726.
Chen, S., Roseman, A.M., Hunter, A.S., Wood, S.P., Burston, S.G., Ranson, N.A., Clarke, A.R.,
and Saibil, H.R. (1994). Location of a folding protein and shape changes in GroELGroES complexes imaged by cryo-electron microscopy. Nature 371, 261-264.
Chong, J.P., Thommes, P., and Blow, J.J. (1996). The role of MCM/P1 proteins in the licensing
of DNA replication. Trends Biochem Sci 21, 102-106.
33
Chuang, S.E., Burland, V., Plunkett, G., 3rd, Daniels, D.L., and Blattner, F.R. (1993). Sequence
analysis of four new heat-shock genes constituting the hslTS/ibpAB and hslVU operons in
Escherichia coli. Gene 134, 1-6.
Chung, C.H., and Goldberg, A.L. (1981). The product of the lon (capR) gene in Escherichia coli
is the ATP-dependent protease, protease La. Proc Natl Acad Sci USA 78, 4931-4935.
Clarke, A.K., Schelin, J., and Porankiewicz, J. (1998). Inactivation of the clpP1 gene for the
proteolytic subunit of the ATP-dependent Clp protease in the cyanobacterium
Synechococcus limits growth and light acclimation. Plant Mol Biol 37, 791-801.
Coleman, T.R., Carpenter, P.B., and Dunphy, W.G. (1996). The Xenopus Cdc6 protein is
essential for the initiation of a single round of DNA replication in cell-free extracts. Cell
87, 53-63.
Confalonieri, F., and Duguet, M. (1995). A 200-amino acid ATPase module in search of a basic
function. Bioessays 17, 639-650.
Cowing, D.W., Bardwell, J.C., Craig, E.A., Woolford, C., Hendrix, R.W., and Gross, C.A.
(1985). Consensus sequence for Escherichia coli heat shock gene promoters. Proc Natl
Acad Sci USA 82, 2679-2683.
Cullmann, G., Fien, K., Kobayashi, R., and Stillman, B. (1995). Characterization of the five
replication factor C genes of Saccharomyces cerevisiae. Mol Cell Biol 15, 4661-4671.
Damerau, K., and St John, A.C. (1993). Role of Clp protease subunits in degradation of carbon
starvation proteins in Escherichia coli. J Bacteriol 175, 53-63.
de Crecy-Lagard, V., Servant-Moisson, P., Viala, J., Grandvalet, C., and Mazodier, P. (1999).
Alteration of the synthesis of the Clp ATP-dependent protease affects morphological and
physiological differentiation in Streptomyces. Mol Microbiol 32, 505-517.
34
DeLaBarre, B., and Brunger, A.T. (2003). Complete structure of p97/valosin-containing protein
reveals communication between nucleotide domains. Nat Struct Biol 10, 856-863.
Diamant, S., Ben-Zvi, A.P., Bukau, B., and Goloubinoff, P. (2000). Size-dependent
disaggregation of stable protein aggregates by the DnaK chaperone machinery. J Biol
Chem 275, 21107-21113.
Diffley, J.F., and Cocker, J.H. (1992). Protein-DNA interactions at a yeast replication origin.
Nature 357, 169-172.
Donaldson, L.W., Wojtyra, U., and Houry, W.A. (2003). Solution structure of the dimeric zinc
binding domain of the chaperone ClpX. J Biol Chem 278, 48991-48996.
Donovan, S., Harwood, J., Drury, L.S., and Diffley, J.F. (1997). Cdc6p-dependent loading of
MCM proteins onto pre-replicative chromatin in budding yeast. Proc Natl Acad Sci USA
94, 5611-5616.
Doronina, V.A., and Murray, N.E. (2001). The proteolytic control of restriction activity in
Escherichia coli K-12. Mol Microbiol 39, 416-428.
Dougan, D.A., Reid, B.G., Horwich, A.L., and Bukau, B. (2002). ClpS, a substrate modulator of
the ClpAP machine. Mol Cell 9, 673-683.
Dougan, D.A., Weber-Ban, E., and Bukau, B. (2003). Targeted delivery of an ssrA-tagged
substrate by the adaptor protein SspB to its cognate AAA+ protein ClpX. Mol Cell 12,
373-380.
Enoch, T., Carr, A.M., and Nurse, P. (1992). Fission yeast genes involved in coupling mitosis to
completion of DNA replication. Genes Dev 6, 2035-2046.
35
Eriksson, M.J., and Clarke, A.K. (1996). The heat shock protein ClpB mediates the development
of thermotolerance in the cyanobacterium Synechococcus sp. strain PCC 7942. J
Bacteriol 178, 4839-4846.
Fedhila, S., Msadek, T., Nel, P., and Lereclus, D. (2002). Distinct clpP genes control specific
adaptive responses in Bacillus thuringiensis. J Bacteriol 184, 5554-5562.
Fenton, W.A., and Horwich, A.L. (1997). GroEL-mediated protein folding. Protein Sci 6, 743760.
Fenton, W.A., Kashi, Y., Furtak, K., and Horwich, A.L. (1994). Residues in chaperonin GroEL
required for polypeptide binding and release. Nature 371, 614-619.
Fink, A.L. (1999). Chaperone-mediated protein folding. Physiol Rev 79, 425-449.
Flynn, G.C., Beckers, C.J., Baase, W.A., and Dahlquist, F.W. (1993). Individual subunits of
bacterial luciferase are molten globules and interact with molecular chaperones. Proc
Natl Acad Sci USA 90, 10826-10830.
Flynn, J.M., Levchenko, I., Seidel, M., Wickner, S.H., Sauer, R.T., and Baker, T.A. (2001).
Overlapping recognition determinants within the ssrA degradation tag allow modulation
of proteolysis. Proc Natl Acad Sci USA 98, 10584-10589.
Flynn, J.M., Neher, S.B., Kim, Y.I., Sauer, R.T., and Baker, T.A. (2003). Proteomic discovery of
cellular substrates of the ClpXP protease reveals five classes of ClpX-recognition signals.
Mol Cell 11, 671-683.
Frank, E.G., Ennis, D.G., Gonzalez, M., Levine, A.S., and Woodgate, R. (1996). Regulation of
SOS mutagenesis by proteolysis. Proc Natl Acad Sci USA 93, 10291-10296.
Frees, D., and Ingmer, H. (1999). ClpP participates in the degradation of misfolded protein in
Lactococcus lactis. Mol Microbiol 31, 79-87.
36
Freire, R., Murguia, J.R., Tarsounas, M., Lowndes, N.F., Moens, P.B., and Jackson, S.P. (1998).
Human and mouse homologs of Schizosaccharomyces pombe Rad1+ and Saccharomyces
cerevisiae RAD17: linkage to checkpoint control and mammalian meiosis. Genes Dev 12,
2560-2573.
Friedman, D.I., Olson, E.R., Georgopoulos, C., Tilly, K., Herskowitz, I., and Banuett, F. (1984).
Interactions of bacteriophage and host macromolecules in the growth of bacteriophage
lambda. Microbiol Rev 48, 299-325.
Frydman, J., Nimmesgern, E., Ohtsuka, K., and Hartl, F.U. (1994). Folding of nascent
polypeptide chains in a high molecular mass assembly with molecular chaperones.
Nature 370, 111-117.
Fuller, R.S., Funnell, B.E., and Kornberg, A. (1984). The DnaA protein complex with the E. coli
chromosomal replication origin (oriC) and other DNA sites. Cell 38, 889-900.
Gaillot, O., Pellegrini, E., Bregenholt, S., Nair, S., and Berche, P. (2000). The ClpP serine
protease is essential for the intracellular parasitism and virulence of Listeria
monocytogenes. Mol Microbiol 35, 1286-1294.
Gaitanaris, G.A., Vysokanov, A., Hung, S.C., Gottesman, M.E., and Gragerov, A. (1994).
Successive action of Escherichia coli chaperones in vivo. Mol Microbiol 14, 861-869.
Georgopoulos, C.P., Hendrix, R.W., Casjens, S.R., and Kaiser, A.D. (1973). Host participation in
bacteriophage lambda head assembly. J Mol Biol 76, 45-60.
Georgopoulos, C.P., Hendrix, R.W., Kaiser, A.D., and Wood, W.B. (1972). Role of the host cell
in bacteriophage morphogenesis: effects of a bacterial mutation on T4 head assembly.
Nat New Biol 239, 38-41.
37
Gerth, U., Kruger, E., Derre, I., Msadek, T., and Hecker, M. (1998). Stress induction of the
Bacillus subtilis clpP gene encoding a homologue of the proteolytic component of the
Clp protease and the involvement of ClpP and ClpX in stress tolerance. Mol Microbiol
28, 787-802.
Goldberg, A.L., and St John, A.C. (1976). Intracellular protein degradation in mammalian and
bacterial cells: Part 2. Annu Rev Biochem 45, 747-803.
Goloubinoff, P., Christeller, J.T., Gatenby, A.A., and Lorimer, G.H. (1989). Reconstitution of
active dimeric ribulose bisphosphate carboxylase from an unfolded state depends on two
chaperonin proteins and Mg-ATP. Nature 342, 884-889.
Goloubinoff, P., Mogk, A., Zvi, A.P., Tomoyasu, T., and Bukau, B. (1999). Sequential
mechanism of solubilization and refolding of stable protein aggregates by a bichaperone
network. Proc Natl Acad Sci USA 96, 13732-13737.
Gonciarz-Swiatek, M., Wawrzynow, A., Um, S. J., Learn, B.A., McMacken, R., Kelley, W.L.,
Georgopoulos, C., Sliekers, O., and Zylicz, M. (1999). Recognition, targeting, and
hydrolysis of the O replication protein by the ClpP/ClpX protease. J Biol Chem 274,
13999-14005.
Gonzalez, M., Rasulova, F., Maurizi, M.R., and Woodgate, R. (2000). Subunit-specific
degradation of the UmuD/D´ heterodimer by the ClpXP protease: the role of trans
recognition in UmuD´ stability. EMBO J 19, 5251-5258.
Gorbalenya, A.E., and Koonin, E.V. (1989). Viral proteins containing the purine NTP-binding
sequence pattern. Nucleic Acids Res 17, 8413-8440.
38
Gordon, C.L., Sather, S.K., Casjens, S., and King, J. (1994). Selective in vivo rescue by
GroEL/ES of thermolabile folding intermediates to phage P22 structural proteins. J Biol
Chem 269, 27941-27951.
Gottesman, S. (1996). Proteases and their targets in Escherichia coli. Annu Rev Genet 30, 465506.
Gottesman, S., Clark, W.P., de Crecy-Lagard, V., and Maurizi, M.R. (1993). ClpX, an alternative
subunit for the ATP-dependent Clp protease of Escherichia coli. Sequence and in vivo
activities. J Biol Chem 268, 22618-22626.
Gottesman, S., Clark, W.P., and Maurizi, M.R. (1990a). The ATP-dependent Clp protease of
Escherichia coli. Sequence of clpA and identification of a Clp-specific substrate. J Biol
Chem 265, 7886-7893.
Gottesman, S., Maurizi, M.R., and Wickner, S. (1997a). Regulatory subunits of energydependent proteases. Cell 91, 435-438.
Gottesman, S., Roche, E., Zhou, Y., and Sauer, R.T. (1998). The ClpXP and ClpAP proteases
degrade proteins with carboxy-terminal peptide tails added by the SsrA-tagging system.
Genes Dev 12, 1338-1347.
Gottesman, S., Squires, C., Pichersky, E., Carrington, M., Hobbs, M., Mattick, J.S., Dalrymple,
B., Kuramitsu, H., Shiroza, T., Foster, T., Clark, W.P., Ross, B., Squires, C.L., and
Maurizi, M.R. (1990b). Conservation of the regulatory subunit for the Clp ATPdependent protease in prokaryotes and eukaryotes. Proc Natl Acad Sci USA 87, 35133517.
Gottesman, S., Wickner, S., and Maurizi, M.R. (1997b). Protein quality control: triage by
chaperones and proteases. Genes Dev 11, 815-823.
39
Griffiths, D.J., Barbet, N.C., McCready, S., Lehmann, A.R., and Carr, A.M. (1995). Fission yeast
rad17: a homologue of budding yeast RAD24 that shares regions of sequence similarity
with DNA polymerase accessory proteins. EMBO J 14, 5812-5823.
Grimaud, R., Kessel, M., Beuron, F., Steven, A.C., and Maurizi, M.R. (1998). Enzymatic and
structural similarities between the Escherichia coli ATP-dependent proteases, ClpXP and
ClpAP. J Biol Chem 273, 12476-12481.
Groll, M., Bajorek, M., Kohler, A., Moroder, L., Rubin, D.M., Huber, R., Glickman, M.H., and
Finley, D. (2000). A gated channel into the proteasome core particle. Nat Struct Biol 7,
1062-1067.
Groll, M., Ditzel, L., Lowe, J., Stock, D., Bochtler, M., Bartunik, H.D., and Huber, R. (1997).
Structure of 20S proteasome from yeast at 2.4 Å resolution. Nature 386, 463-471.
Guenther, B., Onrust, R., Sali, A., O'Donnell, M., and Kuriyan, J. (1997). Crystal structure of the
´ subunit of the clamp-loader complex of E. coli DNA polymerase III. Cell 91, 335-345.
Guo, F., Esser, L., Singh, S.K., Maurizi, M.R., and Xia, D. (2002a). Crystal structure of the
heterodimeric complex of the adaptor, ClpS, with the N-domain of the AAA+ chaperone,
ClpA. J Biol Chem 277, 46753-46762.
Guo, F., Maurizi, M.R., Esser, L., and Xia, D. (2002b). Crystal structure of ClpA, an Hsp100
chaperone and regulator of ClpAP protease. J Biol Chem 277, 46743-46752.
Guzman-Verri, C., Chaves-Olarte, E., Garcia, F., Arvidson, S., and Moreno, E. (2001). In vivo
proteolytic degradation of the Escherichia coli acyltransferase HlyC. J Biol Chem 276,
16660-16666.
Hanson, P.I., Heuser, J.E., and Jahn, R. (1997). Neurotransmitter release - four years of SNARE
complexes. Curr Opin Neurobiol 7, 310-315.
40
Hattendorf, D.A., and Lindquist, S.L. (2002). Cooperative kinetics of both Hsp104 ATPase
domains and interdomain communication revealed by AAA sensor-1 mutants. EMBO J
21, 12-21.
Hay, J.C., and Scheller, R.H. (1997). SNAREs and NSF in targeted membrane fusion. Curr Opin
Cell Biol 9, 505-512.
Hayer-Hartl, M.K., Ewbank, J.J., Creighton, T.E., and Hartl, F.U. (1994). Conformational
specificity of the chaperonin GroEL for the compact folding intermediates of lactalbumin. EMBO J 13, 3192-3202.
Hayer-Hartl, M.K., Martin, J., and Hartl, F.U. (1995). Asymmetrical interaction of GroEL and
GroES in the ATPase cycle of assisted protein folding. Science 269, 836-841.
Hengge-Aronis, R. (2002). Signal transduction and regulatory mechanisms involved in control of
the s (RpoS) subunit of RNA polymerase. Microbiol Mol Biol Rev 66, 373-395, table of
contents.
Hensel, M., Shea, J.E., Gleeson, C., Jones, M.D., Dalton, E., and Holden, D.W. (1995).
Simultaneous identification of bacterial virulence genes by negative selection. Science
269, 400-403.
Hong, S.W., and Vierling, E. (2000). Mutants of Arabidopsis thaliana defective in the
acquisition of tolerance to high temperature stress. Proc Natl Acad Sci USA 97, 43924397.
Hoskins, J.R., Kim, S.Y., and Wickner, S. (2000a). Substrate recognition by the ClpA chaperone
component of ClpAP protease. J Biol Chem 275, 35361-35367.
41
Hoskins, J.R., Singh, S.K., Maurizi, M.R., and Wickner, S. (2000b). Protein binding and
unfolding by the chaperone ClpA and degradation by the protease ClpAP. Proc Natl
Acad Sci USA 97, 8892-8897.
Houry, W.A., Frishman, D., Eckerskorn, C., Lottspeich, F., and Hartl, F.U. (1999). Identification
of in vivo substrates of the chaperonin GroEL. Nature 402, 147-154.
Huang, C., Wang, S., Chen, L., Lemieux, C., Otis, C., Turmel, M., and Liu, X.Q. (1994). The
Chlamydomonas chloroplast clpP gene contains translated large insertion sequences and
is essential for cell growth. Mol Gen Genet 244, 151-159.
Hubbard, T.J., Murzin, A.G., Brenner, S.E., and Chothia, C. (1997). SCOP: a structural
classification of proteins database. Nucleic Acids Res 25, 236-239.
Hubel, A., Krobitsch, S., Horauf, A., and Clos, J. (1997). Leishmania major Hsp100 is required
chiefly in the mammalian stage of the parasite. Mol Cell Biol 17, 5987-5995.
Hwang, I., Thorgeirsson, T., Lee, J., Kustu, S., and Shin, Y.K. (1999). Physical evidence for a
phosphorylation-dependent conformational change in the enhancer-binding protein NtrC.
Proc Natl Acad Sci USA 96, 4880-4885.
Ishikawa, T., Beuron, F., Kessel, M., Wickner, S., Maurizi, M.R., and Steven, A.C. (2001).
Translocation pathway of protein substrates in ClpAP protease. Proc Natl Acad Sci USA
98, 4328-4333.
Iwasaki, H., Shiba, T., Nakata, A., and Shinagawa, H. (1989). Involvement in DNA repair of the
ruvA gene of Escherichia coli. Mol Gen Genet 219, 328-331.
Jackson, G.S., Staniforth, R.A., Halsall, D.J., Atkinson, T., Holbrook, J.J., Clarke, A.R., and
Burston, S.G. (1993). Binding and hydrolysis of nucleotides in the chaperonin catalytic
42
cycle: implications for the mechanism of assisted protein folding. Biochemistry 32, 25542563.
Jenal, U., and Fuchs, T. (1998). An essential protease involved in bacterial cell-cycle control.
EMBO J 17, 5658-5669.
Jenal, U., and Shapiro, L. (1996). Cell cycle-controlled proteolysis of a flagellar motor protein
that is asymmetrically distributed in the Caulobacter predivisional cell. EMBO J 15,
2393-2406.
Jeruzalmi, D., O'Donnell, M., and Kuriyan, J. (2001). Crystal structure of the processivity clamp
loader gamma () complex of E. coli DNA polymerase III. Cell 106, 429-441.
Kanemori, M., Yanagi, H., and Yura, T. (1999). The ATP-dependent HslVU/ClpQY protease
participates in turnover of cell division inhibitor SulA in Escherichia coli. J Bacteriol
181, 3674-3680.
Karata, K., Inagawa, T., Wilkinson, A.J., Tatsuta, T., and Ogura, T. (1999). Dissecting the role
of a conserved motif (the second region of homology) in the AAA family of ATPases.
Site-directed mutagenesis of the ATP-dependent protease FtsH. J Biol Chem 274, 2622526232.
Karata, K., Verma, C.S., Wilkinson, A.J., and Ogura, T. (2001). Probing the mechanism of ATP
hydrolysis and substrate translocation in the AAA protease FtsH by modelling and
mutagenesis. Mol Microbiol 39, 890-903.
Karzai, A.W., Roche, E.D., and Sauer, R.T. (2000). The SsrA-SmpB system for protein tagging,
directed degradation and ribosome rescue. Nat Struct Biol 7, 449-455.
Katayama, T., Kubota, T., Takata, M., Akimitsu, N., and Sekimizu, K. (1996). Disruption of the
hslU gene, which encodes an ATPase subunit of the eukaryotic 26S proteasome homolog
43
in Escherichia coli, suppresses the temperature-sensitive dnaA46 mutation. Biochem
Biophys Res Commun 229, 219-224.
Katayama, Y., Gottesman, S., Pumphrey, J., Rudikoff, S., Clark, W.P., and Maurizi, M.R.
(1988). The two-component, ATP-dependent Clp protease of Escherichia coli.
Purification, cloning, and mutational analysis of the ATP-binding component. J Biol
Chem 263, 15226-15236.
Katayama-Fujimura, Y., Gottesman, S., and Maurizi, M.R. (1987). A multiple-component, ATPdependent protease from Escherichia coli. J Biol Chem 262, 4477-4485.
Keiler, K.C., Waller, P.R., and Sauer, R.T. (1996). Role of a peptide tagging system in
degradation of proteins synthesized from damaged messenger RNA. Science 271, 990993.
Kelly, T.J., and Brown, G.W. (2000). Regulation of chromosome replication. Annu Rev Biochem
69, 829-880.
Kelman, Z., and O'Donnell, M. (1995). DNA polymerase III holoenzyme: structure and function
of a chromosomal replicating machine. Annu Rev Biochem 64, 171-200.
Kenniston, J.A., Baker, T.A., Fernandez, J.M., and Sauer, R.T. (2003). Linkage between ATP
consumption and mechanical unfolding during the protein processing reactions of an
AAA+ degradation machine. Cell 114, 511-520.
Kessel, M., Maurizi, M.R., Kim, B., Kocsis, E., Trus, B.L., Singh, S.K., and Steven, A.C. (1995).
Homology in structural organization between E. coli ClpAP protease and the eukaryotic
26S proteasome. J Mol Biol 250, 587-594.
Kim, D.Y., and Kim, K.K. (2003). Crystal Structure of ClpX Molecular Chaperone from
Helicobacter pylori. J Biol Chem 278, 50664-50670.
44
Kim, Y.I., Burton, R.E., Burton, B.M., Sauer, R.T., and Baker, T.A. (2000). Dynamics of
substrate denaturation and translocation by the ClpXP degradation machine. Mol Cell 5,
639-648.
Kim, Y.I., Levchenko, I., Fraczkowska, K., Woodruff, R.V., Sauer, R.T., and Baker, T.A.
(2001). Molecular determinants of complex formation between Clp/Hsp100 ATPases and
the ClpP peptidase. Nat Struct Biol 8, 230-233.
Kitagawa, M., Wada, C., Yoshioka, S., and Yura, T. (1991). Expression of ClpB, an analog of
the ATP-dependent protease regulatory subunit in Escherichia coli, is controlled by a
heat shock sigma factor (32). J Bacteriol 173, 4247-4253.
Kong, X.P., Onrust, R., O'Donnell, M., and Kuriyan, J. (1992). Three-dimensional structure of
the  subunit of E. coli DNA polymerase III holoenzyme: a sliding DNA clamp. Cell 69,
425-437.
Konieczny, I., and Helinski, D.R. (1997). The replication initiation protein of the broad-hostrange plasmid RK2 is activated by the ClpX chaperone. Proc Natl Acad Sci USA 94,
14378-14382.
Konieczny, I., and Liberek, K. (2002). Cooperative action of Escherichia coli ClpB protein and
DnaK chaperone in the activation of a replication initiation protein. J Biol Chem 277,
18483-18488.
Koonin, E.V. (1993a). A common set of conserved motifs in a vast variety of putative nucleic
acid-dependent ATPases including MCM proteins involved in the initiation of eukaryotic
DNA replication. Nucleic Acids Res 21, 2541-2547.
Koonin, E.V. (1993b). A superfamily of ATPases with diverse functions containing either
classical or deviant ATP-binding motif. J Mol Biol 229, 1165-1174.
45
Kopito, R.R. (1997). ER quality control: the cytoplasmic connection. Cell 88, 427-430.
Krishna, T.S., Kong, X.P., Gary, S., Burgers, P.M., and Kuriyan, J. (1994). Crystal structure of
the eukaryotic DNA polymerase processivity factor PCNA. Cell 79, 1233-1243.
Krobitsch, S., and Clos, J. (1999). A novel role for 100 kD heat shock proteins in the parasite
Leishmania donovani. Cell Stress Chaperones 4, 191-198.
Kruger, E., Volker, U., and Hecker, M. (1994). Stress induction of clpC in Bacillus subtilis and
its involvement in stress tolerance. J Bacteriol 176, 3360-3367.
Kruklitis, R., Welty, D.J., and Nakai, H. (1996). ClpX protein of Escherichia coli activates
bacteriophage Mu transposase in the strand transfer complex for initiation of Mu DNA
synthesis. EMBO J 15, 935-944.
Landry, S.J., Jordan, R., McMacken, R., and Gierasch, L.M. (1992). Different conformations for
the same polypeptide bound to chaperones DnaK and GroEL. Nature 355, 455-457.
Langer, T., Lu, C., Echols, H., Flanagan, J., Hayer, M.K., and Hartl, F.U. (1992a). Successive
action of DnaK, DnaJ and GroEL along the pathway of chaperone-mediated protein
folding. Nature 356, 683-689.
Langer, T., Pfeifer, G., Martin, J., Baumeister, W., and Hartl, F.U. (1992b). Chaperoninmediated protein folding: GroES binds to one end of the GroEL cylinder, which
accommodates the protein substrate within its central cavity. EMBO J 11, 4757-4765.
Larsen, C.N., and Finley, D. (1997). Protein translocation channels in the proteasome and other
proteases. Cell 91, 431-434.
Laskowska, E., Kuczynska-Wisnik, D., Skorko-Glonek, J., and Taylor, A. (1996). Degradation
by proteases Lon, Clp and HtrA, of Escherichia coli proteins aggregated in vivo by heat
shock; HtrA protease action in vivo and in vitro. Mol Microbiol 22, 555-571.
46
Latterich, M., Frohlich, K.U., and Schekman, R. (1995). Membrane fusion and the cell cycle:
Cdc48p participates in the fusion of ER membranes. Cell 82, 885-893.
Lau, G.W., Haataja, S., Lonetto, M., Kensit, S.E., Marra, A., Bryant, A.P., McDevitt, D.,
Morrison, D.A., and Holden, D.W. (2001). A functional genomic analysis of type 3
Streptococcus pneumoniae virulence. Mol Microbiol 40, 555-571.
Lee, C., Schwartz, M.P., Prakash, S., Iwakura, M., and Matouschek, A. (2001). ATP-dependent
proteases degrade their substrates by processively unraveling them from the degradation
signal. Mol Cell 7, 627-637.
Lee, S., Sowa, M.E., Watanabe, Y.H., Sigler, P.B., Chiu, W., Yoshida, M., and Tsai, F.T. (2003).
The structure of ClpB: a molecular chaperone that rescues proteins from an aggregated
state. Cell 115, 229-240.
Lehnherr, H., and Yarmolinsky, M.B. (1995). Addiction protein Phd of plasmid prophage P1 is a
substrate of the ClpXP serine protease of Escherichia coli. Proc Natl Acad Sci USA 92,
3274-3277.
Lei, M., Kawasaki, Y., Young, M.R., Kihara, M., Sugino, A., and Tye, B.K. (1997). MCM2 is a
target of regulation by Cdc7-Dbf4 during the initiation of DNA synthesis. Genes Dev 11,
3365-3374.
Lenzen, C.U., Steinmann, D., Whiteheart, S.W., and Weis, W.I. (1998). Crystal structure of the
hexamerization domain of N-ethylmaleimide-sensitive fusion protein. Cell 94, 525-536.
Levchenko, I., Luo, L., and Baker, T.A. (1995). Disassembly of the Mu transposase tetramer by
the ClpX chaperone. Genes Dev 9, 2399-2408.
Levchenko, I., Seidel, M., Sauer, R.T., and Baker, T.A. (2000). A specificity-enhancing factor
for the ClpXP degradation machine. Science 289, 2354-2356.
47
Levchenko, I., Yamauchi, M., and Baker, T.A. (1997). ClpX and MuB interact with overlapping
regions of Mu transposase: implications for control of the transposition pathway. Genes
Dev 11, 1561-1572.
Li, J., and Sha, B. (2003). Crystal structure of the E. coli Hsp100 ClpB N-terminal domain.
Structure (Camb) 11, 323-328.
Liang, C., Weinreich, M., and Stillman, B. (1995). ORC and Cdc6p interact and determine the
frequency of initiation of DNA replication in the genome. Cell 81, 667-676.
Lindsey-Boltz, L.A., Bermudez, V.P., Hurwitz, J., and Sancar, A. (2001). Purification and
characterization of human DNA damage checkpoint Rad complexes. Proc Natl Acad Sci
USA 98, 11236-11241.
Liu, Z., Tek, V., Akoev, V., and Zolkiewski, M. (2002). Conserved amino acid residues within
the amino-terminal domain of ClpB are essential for the chaperone activity. J Mol Biol
321, 111-120.
Lo, J.H., Baker, T.A., and Sauer, R.T. (2001). Characterization of the N-terminal repeat domain
of Escherichia coli ClpA-A class I Clp/HSP100 ATPase. Protein Sci 10, 551-559.
Lupas, A.N., and Martin, J. (2002). AAA proteins. Curr Opin Struct Biol 12, 746-753.
Magasanik, B. (1989). Regulation of transcription of the glnALG operon of Escherichia coli by
protein phosphorylation. Biochimie 71, 1005-1012.
Makovets, S., Doronina, V.A., and Murray, N.E. (1999). Regulation of endonuclease activity by
proteolysis prevents breakage of unmodified bacterial chromosomes by type I restriction
enzymes. Proc Natl Acad Sci USA 96, 9757-9762.
48
Makovets, S., Titheradge, A.J., and Murray, N.E. (1998). ClpX and ClpP are essential for the
efficient acquisition of genes specifying type IA and IB restriction systems. Mol
Microbiol 28, 25-35.
Matouschek, A. (2003). Protein unfolding--an important process in vivo? Curr Opin Struct Biol
13, 98-109.
Maurizi, M.R., Clark, W.P., Katayama, Y., Rudikoff, S., Pumphrey, J., Bowers, B., and
Gottesman, S. (1990a). Sequence and structure of ClpP, the proteolytic component of the
ATP-dependent Clp protease of Escherichia coli. J Biol Chem 265, 12536-12545.
Maurizi, M.R., Clark, W.P., Kim, S.H., and Gottesman, S. (1990b). ClpP represents a unique
family of serine proteases. J Biol Chem 265, 12546-12552.
Maurizi, M.R., Trisler, P., and Gottesman, S. (1985). Insertional mutagenesis of the lon gene in
Escherichia coli: lon is dispensable. J Bacteriol 164, 1124-1135.
May, A.P., Whiteheart, S.W., and Weis, W.I. (2001). Unraveling the mechanism of the vesicle
transport ATPase NSF, the N-ethylmaleimide-sensitive factor. J Biol Chem 276, 2199121994.
McCarty, J.S., Buchberger, A., Reinstein, J., and Bukau, B. (1995). The role of ATP in the
functional cycle of the DnaK chaperone system. J Mol Biol 249, 126-137.
Mei, J.M., Nourbakhsh, F., Ford, C.W., and Holden, D.W. (1997). Identification of
Staphylococcus aureus virulence genes in a murine model of bacteraemia using
signature-tagged mutagenesis. Mol Microbiol 26, 399-407.
Mhammedi-Alaoui, A., Pato, M., Gama, M.J., and Toussaint, A. (1994). A new component of
bacteriophage Mu replicative transposition machinery: the Escherichia coli ClpX protein.
Mol Microbiol 11, 1109-1116.
49
Misra, N., Habib, S., Ranjan, A., Hasnain, S.E., and Nath, I. (1996). Expression and functional
characterisation of the clpC gene of Mycobacterium leprae: ClpC protein elicits human
antibody response. Gene 172, 99-104.
Missiakas, D., Schwager, F., Betton, J.M., Georgopoulos, C., and Raina, S. (1996). Identification
and characterization of HsIV HsIU (ClpQ ClpY) proteins involved in overall proteolysis
of misfolded proteins in Escherichia coli. EMBO J 15, 6899-6909.
Mogk, A., Deuerling, E., Vorderwulbecke, S., Vierling, E., and Bukau, B. (2003a). Small heat
shock proteins, ClpB and the DnaK system form a functional triade in reversing protein
aggregation. Mol Microbiol 50, 585-595.
Mogk, A., Schlieker, C., Strub, C., Rist, W., Weibezahn, J., and Bukau, B. (2003b). Roles of
individual domains and conserved motifs of the AAA+ chaperone ClpB in
oligomerization, ATP hydrolysis, and chaperone activity. J Biol Chem 278, 17615-17624.
Msadek, T., Dartois, V., Kunst, F., Herbaud, M.L., Denizot, F., and Rapoport, G. (1998). ClpP of
Bacillus subtilis is required for competence development, motility, degradative enzyme
synthesis, growth at high temperature and sporulation. Mol Microbiol 27, 899-914.
Nair, S., Frehel, C., Nguyen, L., Escuyer, V., and Berche, P. (1999). ClpE, a novel member of
the HSP100 family, is involved in cell division and virulence of Listeria monocytogenes.
Mol Microbiol 31, 185-196.
Nakano, M.M., Hajarizadeh, F., Zhu, Y., and Zuber, P. (2001). Loss-of-function mutations in
yjbD result in ClpX- and ClpP-independent competence development of Bacillus subtilis.
Mol Microbiol 42, 383-394.
50
Nakano, S., Nakano, M.M., Zhang, Y., Leelakriangsak, M., and Zuber, P. (2003). A regulatory
protein that interferes with activator-stimulated transcription in bacteria. Proc Natl Acad
Sci USA 100, 4233-4238.
Nanamiya, H., Takahashi, K., Fujita, M., and Kawamura, F. (2000). Deficiency of the initiation
events of sporulation in Bacillus subtilis clpP mutant can be suppressed by a lack of the
Spo0E protein phosphatase. Biochem Biophys Res Commun 279, 229-233.
Neher, S.B., Flynn, J.M., Sauer, R.T., and Baker, T.A. (2003a). Latent ClpX-recognition signals
ensure LexA destruction after DNA damage. Genes Dev 17, 1084-1089.
Neher, S.B., Sauer, R.T., and Baker, T.A. (2003b). Distinct peptide signals in the UmuD and
UmuD´ subunits of UmuD/D´ mediate tethering and substrate processing by the ClpXP
protease. Proc Natl Acad Sci USA 100, 13219-13224.
Neuwald, A.F., Aravind, L., Spouge, J.L., and Koonin, E.V. (1999). AAA+: A class of
chaperone-like ATPases associated with the assembly, operation, and disassembly of
protein complexes. Genome Res 9, 27-43.
O'Connell, M.J., Walworth, N.C., and Carr, A.M. (2000). The G2-phase DNA-damage
checkpoint. Trends Cell Biol 10, 296-303.
O'Toole, G.A., and Kolter, R. (1998). Initiation of biofilm formation in Pseudomonas
fluorescens WCS365 proceeds via multiple, convergent signalling pathways: a genetic
analysis. Mol Microbiol 28, 449-461.
Ogura, T., and Wilkinson, A.J. (2001). AAA+ superfamily ATPases: common structure--diverse
function. Genes Cells 6, 575-597.
Ortega, J., Lee, H.S., Maurizi, M.R., and Steven, A.C. (2002). Alternating translocation of
protein substrates from both ends of ClpXP protease. EMBO J 21, 4938-4949.
51
Ortega, J., Singh, S.K., Ishikawa, T., Maurizi, M.R., and Steven, A.C. (2000). Visualization of
substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 6,
1515-1521.
Pak, M., Hoskins, J.R., Singh, S.K., Maurizi, M.R., and Wickner, S. (1999). Concurrent
chaperone and protease activities of ClpAP and the requirement for the N-terminal ClpA
ATP binding site for chaperone activity. J Biol Chem 274, 19316-19322.
Palleros, D.R., Reid, K.L., Shi, L., Welch, W.J., and Fink, A.L. (1993). ATP-induced proteinHsp70 complex dissociation requires K+ but not ATP hydrolysis. Nature 365, 664-666.
Palleros, D.R., Shi, L., Reid, K.L., and Fink, A.L. (1994). Hsp70-protein complexes. Complex
stability and conformation of bound substrate protein. J Biol Chem 269, 13107-13114.
Park, S.K., Kim, K.I., Woo, K.M., Seol, J.H., Tanaka, K., Ichihara, A., Ha, D.B., and Chung,
C.H. (1993). Site-directed mutagenesis of the dual translational initiation sites of the clpB
gene of Escherichia coli and characterization of its gene products. J Biol Chem 268,
20170-20174.
Patel, S., and Latterich, M. (1998). The AAA team: related ATPases with diverse functions.
Trends Cell Biol 8, 65-71.
Pederson, K.J., Carlson, S., and Pierson, D.E. (1997). The ClpP protein, a subunit of the Clp
protease, modulates ail gene expression in Yersinia enterocolitica. Mol Microbiol 26, 99107.
Petit, M.A., Bedale, W., Osipiuk, J., Lu, C., Rajagopalan, M., McInerney, P., Goodman, M.F.,
and Echols, H. (1994). Sequential folding of UmuC by the Hsp70 and Hsp60 chaperone
complexes of Escherichia coli. J Biol Chem 269, 23824-23829.
52
Pierpaoli, E.V., Sandmeier, E., Baici, A., Schonfeld, H.J., Gisler, S., and Christen, P. (1997). The
power stroke of the DnaK/DnaJ/GrpE molecular chaperone system. J Mol Biol 269, 757768.
Polissi, A., Pontiggia, A., Feger, G., Altieri, M., Mottl, H., Ferrari, L., and Simon, D. (1998).
Large-scale identification of virulence genes from Streptococcus pneumoniae. Infect
Immun 66, 5620-5629.
Porankiewicz, J., Schelin, J., and Clarke, A.K. (1998). The ATP-dependent Clp protease is
essential for acclimation to UV-B and low temperature in the cyanobacterium
Synechococcus. Mol Microbiol 29, 275-283.
Porter, S.C., North, A.K., Wedel, A.B., and Kustu, S. (1993). Oligomerization of NTRC at the
glnA enhancer is required for transcriptional activation. Genes Dev 7, 2258-2273.
Pratt, L.A., and Silhavy, T.J. (1996). The response regulator SprE controls the stability of RpoS.
Proc Natl Acad Sci USA 93, 2488-2492.
Pritchard, A.E., Dallmann, H.G., Glover, B.P., and McHenry, C.S. (2000). A novel assembly
mechanism for the DNA polymerase III holoenzyme DnaX complex: association of ´
with DnaX4 forms DnaX3´. EMBO J 19, 6536-6545.
Rabinovich, E., Kerem, A., Frohlich, K.U., Diamant, N., and Bar-Nun, S. (2002). AAA-ATPase
p97/Cdc48p, a cytosolic chaperone required for endoplasmic reticulum-associated protein
degradation. Mol Cell Biol 22, 626-634.
Rabouille, C., Misteli, T., Watson, R., and Warren, G. (1995). Reassembly of Golgi stacks from
mitotic Golgi fragments in a cell-free system. J Cell Biol 129, 605-618.
53
Rajagopal, S., Sudarsan, N., and Nickerson, K.W. (2002). Sodium dodecyl sulfate
hypersensitivity of clpP and clpB mutants of Escherichia coli. Appl Environ Microbiol
68, 4117-4121.
Ramachandran, R., Hartmann, C., Song, H.K., Huber, R., and Bochtler, M. (2002). Functional
interactions of HslV (ClpQ) with the ATPase HslU (ClpY). Proc Natl Acad Sci USA 99,
7396-7401.
Ranson, N.A., Dunster, N.J., Burston, S.G., and Clarke, A.R. (1995). Chaperonins can catalyse
the reversal of early aggregation steps when a protein misfolds. J Mol Biol 250, 581-586.
Reid, B.G., Fenton, W.A., Horwich, A.L., and Weber-Ban, E.U. (2001). ClpA mediates
directional translocation of substrate proteins into the ClpP protease. Proc Natl Acad Sci
USA 98, 3768-3772.
Rice, D.W., Rafferty, J.B., Artymiuk, P.J., and Lloyd, R.G. (1997). Insights into the mechanisms
of homologous recombination from the structure of RuvA. Curr Opin Struct Biol 7, 798803.
Rohrwild, M., Coux, O., Huang, H.C., Moerschell, R.P., Yoo, S.J., Seol, J.H., Chung, C.H., and
Goldberg, A.L. (1996). HslV-HslU: A novel ATP-dependent protease complex in
Escherichia coli related to the eukaryotic proteasome. Proc Natl Acad Sci USA 93, 58085813.
Roseman, A.M., Chen, S., White, H., Braig, K., and Saibil, H.R. (1996). The chaperonin ATPase
cycle: mechanism of allosteric switching and movements of substrate-binding domains in
GroEL. Cell 87, 241-251.
54
Rouquette, C., de Chastellier, C., Nair, S., and Berche, P. (1998). The ClpC ATPase of Listeria
monocytogenes is a general stress protein required for virulence and promoting early
bacterial escape from the phagosome of macrophages. Mol Microbiol 27, 1235-1245.
Rouquette, C., Ripio, M.T., Pellegrini, E., Bolla, J.M., Tascon, R.I., Vazquez-Boland, J.A., and
Berche, P. (1996). Identification of a ClpC ATPase required for stress tolerance and in
vivo survival of Listeria monocytogenes. Mol Microbiol 21, 977-987.
Rudiger, S., Germeroth, L., Schneider-Mergener, J., and Bukau, B. (1997). Substrate specificity
of the DnaK chaperone determined by screening cellulose-bound peptide libraries.
EMBO J 16, 1501-1507.
Sanchez, Y., and Lindquist, S.L. (1990). HSP104 required for induced thermotolerance. Science
248, 1112-1115.
Santocanale, C., and Diffley, J.F. (1998). A Mec1- and Rad53-dependent checkpoint controls
late-firing origins of DNA replication. Nature 395, 615-618.
Saraste, M., Sibbald, P.R., and Wittinghofer, A. (1990). The P-loop--a common motif in ATPand GTP-binding proteins. Trends Biochem Sci 15, 430-434.
Schirmer, E.C., Glover, J.R., Singer, M.A., and Lindquist, S. (1996). HSP100/Clp proteins: a
common mechanism explains diverse functions. Trends Biochem Sci 21, 289-296.
Schroder, H., Langer, T., Hartl, F.U., and Bukau, B. (1993). DnaK, DnaJ and GrpE form a
cellular chaperone machinery capable of repairing heat-induced protein damage. EMBO J
12, 4137-4144.
Schweder, T., Lee, K.H., Lomovskaya, O., and Matin, A. (1996). Regulation of Escherichia coli
starvation sigma factor (s) by ClpXP protease. J Bacteriol 178, 470-476.
55
Seol, J.H., Yoo, S.J., Kim, K.I., Kang, M.S., Ha, D.B., and Chung, C.H. (1994). The 65-kDa
protein derived from the internal translational initiation site of the clpA gene inhibits the
ATP-dependent protease Ti in Escherichia coli. J Biol Chem 269, 29468-29473.
Seong, I.S., Kang, M.S., Choi, M.K., Lee, J.W., Koh, O.J., Wang, J., Eom, S.H., and Chung,
C.H. (2002). The C-terminal tails of HslU ATPase act as a molecular switch for
activation of HslV peptidase. J Biol Chem 277, 25976-25982.
Seong, I.S., Oh, J.Y., Lee, J.W., Tanaka, K., and Chung, C.H. (2000). The HslU ATPase acts as
a molecular chaperone in prevention of aggregation of SulA, an inhibitor of cell division
in Escherichia coli. FEBS Lett 477, 224-229.
Seong, I.S., Oh, J.Y., Yoo, S.J., Seol, J.H., and Chung, C.H. (1999). ATP-dependent degradation
of SulA, a cell division inhibitor, by the HslVU protease in Escherichia coli. FEBS Lett
456, 211-214.
Singh, S.K., Grimaud, R., Hoskins, J.R., Wickner, S., and Maurizi, M.R. (2000). Unfolding and
internalization of proteins by the ATP-dependent proteases ClpXP and ClpAP. Proc Natl
Acad Sci USA 97, 8898-8903.
Singh, S.K., and Maurizi, M.R. (1994). Mutational analysis demonstrates different functional
roles for the two ATP-binding sites in ClpAP protease from Escherichia coli. J Biol
Chem 269, 29537-29545.
Singh, S.K., Rozycki, J., Ortega, J., Ishikawa, T., Lo, J., Steven, A.C., and Maurizi, M.R. (2001).
Functional domains of the ClpA and ClpX molecular chaperones identified by limited
proteolysis and deletion analysis. J Biol Chem 276, 29420-29429.
56
Sollner, T., Bennett, M.K., Whiteheart, S.W., Scheller, R.H., and Rothman, J.E. (1993). A
protein assembly-disassembly pathway in vitro that may correspond to sequential steps of
synaptic vesicle docking, activation, and fusion. Cell 75, 409-418.
Song, H.K., Hartmann, C., Ramachandran, R., Bochtler, M., Behrendt, R., Moroder, L., and
Huber, R. (2000). Mutational studies on HslU and its docking mode with HslV. Proc Natl
Acad Sci USA 97, 14103-14108.
Sousa, M.C., Kessler, B.M., Overkleeft, H.S., and McKay, D.B. (2002). Crystal structure of
HslUV complexed with a vinyl sulfone inhibitor: corroboration of a proposed mechanism
of allosteric activation of HslV by HslU. J Mol Biol 318, 779-785.
Sousa, M.C., Trame, C.B., Tsuruta, H., Wilbanks, S.M., Reddy, V.S., and McKay, D.B. (2000).
Crystal and solution structures of an HslUV protease-chaperone complex. Cell 103, 633643.
Squires, C., and Squires, C.L. (1992). The Clp proteins: proteolysis regulators or molecular
chaperones? J Bacteriol 174, 1081-1085.
Squires, C.L., Pedersen, S., Ross, B.M., and Squires, C. (1991). ClpB is the Escherichia coli heat
shock protein F84.1. J Bacteriol 173, 4254-4262.
Stephani, K., Weichart, D., and Hengge, R. (2003). Dynamic control of Dps protein levels by
ClpXP and ClpAP proteases in Escherichia coli. Mol Microbiol 49, 1605-1614.
Sternberg, N. (1973). Properties of a mutant of Escherichia coli defective in bacteriophage
lambda head formation (groE). I. Initial characterization. J Mol Biol 76, 1-23.
Stewart, J., Hingorani, M.M., Kelman, Z., and O'Donnell, M. (2001). Mechanism of  clamp
opening by the  subunit of Escherichia coli DNA polymerase III holoenzyme. J Biol
Chem 276, 19182-19189.
57
Story, R.M., and Steitz, T.A. (1992). Structure of the recA protein-ADP complex. Nature 355,
374-376.
Studwell, P.S., and O'Donnell, M. (1990). Processive replication is contingent on the
exonuclease subunit of DNA polymerase III holoenzyme. J Biol Chem 265, 1171-1178.
Swaffield, J.C., Melcher, K., and Johnston, S.A. (1995). A highly conserved ATPase protein as a
mediator between acidic activation domains and the TATA-binding protein. Nature 374,
88-91.
Swamy, K.H., and Goldberg, A.L. (1981). E. coli contains eight soluble proteolytic activities,
one being ATP dependent. Nature 292, 652-654.
Szabo, A., Langer, T., Schroder, H., Flanagan, J., Bukau, B., and Hartl, F.U. (1994). The ATP
hydrolysis-dependent reaction cycle of the Escherichia coli Hsp70 system DnaK, DnaJ,
and GrpE. Proc Natl Acad Sci USA 91, 10345-10349.
Tanaka, T., Knapp, D., and Nasmyth, K. (1997). Loading of an MCM protein onto DNA
replication origins is regulated by Cdc6p and CDKs. Cell 90, 649-660.
ter Huurne, A.A., Muir, S., van Houten, M., van der Zeijst, B.A., Gaastra, W., and Kusters, J.G.
(1994). Characterization of three putative Serpulina hyodysenteriae hemolysins. Microb
Pathog 16, 269-282.
Theyssen, H., Schuster, H.P., Packschies, L., Bukau, B., and Reinstein, J. (1996). The second
step of ATP binding to DnaK induces peptide release. J Mol Biol 263, 657-670.
Thomas, J.G., and Baneyx, F. (1998). Roles of the Escherichia coli small heat shock proteins
IbpA and IbpB in thermal stress management: comparison with ClpA, ClpB, and HtpG in
vivo. J Bacteriol 180, 5165-5172.
58
Thomas, J.G., and Baneyx, F. (2000). ClpB and HtpG facilitate de novo protein folding in
stressed Escherichia coli cells. Mol Microbiol 36, 1360-1370.
Thompson, M.W., and Maurizi, M.R. (1994). Activity and specificity of Escherichia coli ClpAP
protease in cleaving model peptide substrates. J Biol Chem 269, 18201-18208.
Thompson, M.W., Singh, S.K., and Maurizi, M.R. (1994). Processive degradation of proteins by
the ATP-dependent Clp protease from Escherichia coli. Requirement for the multiple
array of active sites in ClpP but not ATP hydrolysis. J Biol Chem 269, 18209-18215.
Todd, M.J., Viitanen, P.V., and Lorimer, G.H. (1994). Dynamics of the chaperonin ATPase
cycle: implications for facilitated protein folding. Science 265, 659-666.
Turgay, K., Hahn, J., Burghoorn, J., and Dubnau, D. (1998). Competence in Bacillus subtilis is
controlled by regulated proteolysis of a transcription factor. EMBO J 17, 6730-6738.
VanBogelen, R.A., Kelley, P.M., and Neidhardt, F.C. (1987). Differential induction of heat
shock, SOS, and oxidation stress regulons and accumulation of nucleotides in
Escherichia coli. J Bacteriol 169, 26-32.
Veinger, L., Diamant, S., Buchner, J., and Goloubinoff, P. (1998). The small heat-shock protein
IbpB from Escherichia coli stabilizes stress-denatured proteins for subsequent refolding
by a multichaperone network. J Biol Chem 273, 11032-11037.
Venclovas, C., and Thelen, M.P. (2000). Structure-based predictions of Rad1, Rad9, Hus1 and
Rad17 participation in sliding clamp and clamp-loading complexes. Nucleic Acids Res
28, 2481-2493.
Viitanen, P.V., Gatenby, A.A., and Lorimer, G.H. (1992). Purified chaperonin 60 (GroEL)
interacts with the nonnative states of a multitude of Escherichia coli proteins. Protein Sci
1, 363-369.
59
Volker, U., Engelmann, S., Maul, B., Riethdorf, S., Volker, A., Schmid, R., Mach, H., and
Hecker, M. (1994). Analysis of the induction of general stress proteins of Bacillus
subtilis. Microbiology 140 ( Pt 4), 741-752.
Walker, J.E., Saraste, M., Runswick, M.J., and Gay, N.J. (1982). Distantly related sequences in
the - and -subunits of ATP synthase, myosin, kinases and other ATP-requiring
enzymes and a common nucleotide binding fold. EMBO J 1, 945-951.
Walter, S., Lorimer, G.H., and Schmid, F.X. (1996). A thermodynamic coupling mechanism for
GroEL-mediated unfolding. Proc Natl Acad Sci USA 93, 9425-9430.
Wang, J., Hartling, J.A., and Flanagan, J.M. (1997). The structure of ClpP at 2.3 Å resolution
suggests a model for ATP-dependent proteolysis. Cell 91, 447-456.
Wang, J., Song, J.J., Franklin, M.C., Kamtekar, S., Im, Y.J., Rho, S.H., Seong, I.S., Lee, C.S.,
Chung, C.H., and Eom, S.H. (2001). Crystal structures of the HslVU peptidase-ATPase
complex reveal an ATP-dependent proteolysis mechanism. Structure (Camb) 9, 177-184.
Wang, L., Elliott, M., and Elliott, T. (1999). Conditional stability of the HemA protein
(glutamyl-tRNA reductase) regulates heme biosynthesis in Salmonella typhimurium. J
Bacteriol 181, 1211-1219.
Wawrzynow, A., Wojtkowiak, D., Marszalek, J., Banecki, B., Jonsen, M., Graves, B.,
Georgopoulos, C., and Zylicz, M. (1995). The ClpX heat-shock protein of Escherichia
coli, the ATP-dependent substrate specificity component of the ClpP-ClpX protease, is a
novel molecular chaperone. EMBO J 14, 1867-1877.
Weber, T., Zemelman, B.V., McNew, J.A., Westermann, B., Gmachl, M., Parlati, F., Sollner,
T.H., and Rothman, J.E. (1998). SNAREpins: minimal machinery for membrane fusion.
Cell 92, 759-772.
60
Weber-Ban, E.U., Reid, B.G., Miranker, A.D., and Horwich, A.L. (1999). Global unfolding of a
substrate protein by the Hsp100 chaperone ClpA. Nature 401, 90-93.
Wedel, A., and Kustu, S. (1995). The bacterial enhancer-binding protein NTRC is a molecular
machine: ATP hydrolysis is coupled to transcriptional activation. Genes Dev 9, 20422052.
Weibezahn, J., Schlieker, C., Bukau, B., and Mogk, A. (2003). Characterization of a trap mutant
of the AAA+ chaperone ClpB. J Biol Chem 278, 32608-32617.
Weichart, D., Querfurth, N., Dreger, M., and Hengge-Aronis, R. (2003). Global role for ClpPcontaining proteases in stationary-phase adaptation of Escherichia coli. J Bacteriol 185,
115-125.
Welty, D.J., Jones, J.M., and Nakai, H. (1997). Communication of ClpXP protease
hypersensitivity to bacteriophage Mu repressor isoforms. J Mol Biol 272, 31-41.
West, S.C. (1997). Processing of recombination intermediates by the RuvABC proteins. Annu
Rev Genet 31, 213-244.
Wickner, S., Gottesman, S., Skowyra, D., Hoskins, J., McKenney, K., and Maurizi, M.R. (1994).
A molecular chaperone, ClpA, functions like DnaK and DnaJ. Proc Natl Acad Sci USA
91, 12218-12222.
Wickner, S., Maurizi, M.R., and Gottesman, S. (1999). Posttranslational quality control: folding,
refolding, and degrading proteins. Science 286, 1888-1893.
Wild, J., Altman, E., Yura, T., and Gross, C.A. (1992). DnaK and DnaJ heat shock proteins
participate in protein export in Escherichia coli. Genes Dev 6, 1165-1172.
61
Wild, J., Rossmeissl, P., Walter, W.A., and Gross, C.A. (1996). Involvement of the DnaK-DnaJGrpE chaperone team in protein secretion in Escherichia coli. J Bacteriol 178, 36083613.
Wojtkowiak, D., Georgopoulos, C., and Zylicz, M. (1993). Isolation and characterization of
ClpX, a new ATP-dependent specificity component of the Clp protease of Escherichia
coli. J Biol Chem 268, 22609-22617.
Wojtyra, U.A., Thibault, G., Tuite, A., and Houry, W.A. (2003). The N-terminal zinc binding
domain of ClpX is a dimerization domain that modulates the chaperone function. J Biol
Chem 278, 48981-48990.
Woo, K.M., Kim, K.I., Goldberg, A.L., Ha, D.B., and Chung, C.H. (1992). The heat-shock
protein ClpB in Escherichia coli is a protein-activated ATPase. J Biol Chem 267, 2042920434.
Wu, W.F., Zhou, Y., and Gottesman, S. (1999). Redundant in vivo proteolytic activities of
Escherichia coli Lon and the ClpYQ (HslUV) protease. J Bacteriol 181, 3681-3687.
Wyman, C., Rombel, I., North, A.K., Bustamante, C., and Kustu, S. (1997). Unusual
oligomerization required for activity of NtrC, a bacterial enhancer-binding protein.
Science 275, 1658-1661.
Xiao, H., Naktinis, V., and O'Donnell, M. (1995). Assembly of a chromosomal replication
machine: two DNA polymerases, a clamp loader, and sliding clamps in one holoenzyme
particle. IV. ATP-binding site mutants identify the clamp loader. J Biol Chem 270:
13378-13383.
Xu, Z., Horwich, A.L., and Sigler, P.B. (1997). The crystal structure of the asymmetric GroELGroES-(ADP)7 chaperonin complex. Nature 388, 741-750.
62
Yamamori, T., and Yura, T. (1980). Temperature-induced synthesis of specific proteins in
Escherichia coli: evidence for transcriptional control. J Bacteriol 142, 843-851.
Yu, R.C., Hanson, P.I., Jahn, R., and Brunger, A.T. (1998). Structure of the ATP-dependent
oligomerization domain of N-ethylmaleimide sensitive factor complexed with ATP. Nat
Struct Biol 5, 803-811.
Yu, X., West, S.C., and Egelman, E.H. (1997). Structure and subunit composition of the RuvABHolliday junction complex. J Mol Biol 266, 217-222.
Zahn, R., Perrett, S., Stenberg, G., and Fersht, A.R. (1996). Catalysis of amide proton exchange
by the molecular chaperones GroEL and SecB. Science 271, 642-645.
Zeth, K., Ravelli, R.B., Paal, K., Cusack, S., Bukau, B., and Dougan, D.A. (2002). Structural
analysis of the adaptor protein ClpS in complex with the N-terminal domain of ClpA. Nat
Struct Biol 9, 906-911.
Zhou, Y., Gottesman, S., Hoskins, J.R., Maurizi, M.R., and Wickner, S. (2001). The RssB
response regulator directly targets s for degradation by ClpXP. Genes Dev 15, 627-637.
Zolkiewski, M. (1999). ClpB cooperates with DnaK, DnaJ, and GrpE in suppressing protein
aggregation. A novel multi-chaperone system from Escherichia coli. J Biol Chem 274,
28083-28086.
63
CHAPTER TWO
C-terminal domain mutations in ClpX uncouple substrate binding
from an engagement step required for unfolding
Published: Shilpa A. Joshi, Tania A. Baker, Robert T. Sauer, 2003.
Molecular Microbiology, 48, pp. 67-76.
64
Summary
ClpX mediates ATP-dependent denaturation of specific target proteins and disassembly of
protein complexes. Like other AAA+ family members, ClpX contains an -ATPase domain
and an -helical C-terminal domain. ClpX proteins with mutations in the C-terminal domain
were constructed and screened for disassembly activity in vivo. Seven mutant enzymes with
defective phenotypes were purified and characterized. Three of these proteins (L381K, D382K
& Y385A) had low activity in disassembly or unfolding assays in vitro. In contrast to wild-type
ClpX, substrate binding to these mutants inhibited ATP hydrolysis instead of increasing it.
These mutants appear to be defective in a reaction step that engages bound substrate proteins and
is required both for enhancement of ATP hydrolysis and for unfolding/disassembly. Some of
these side chains form part of the interface between the C-terminal domain of one ClpX subunit
and the ATPase domain of an adjacent subunit in the hexamer and appear to be required for
communication between adjacent nucleotide binding sites.
65
Introduction
ClpX and related members of the Clp/Hsp100 protein family are AAA+ ATPases (Neuwald et
al., 1999) which serve as specialized energy-dependent molecular chaperones to unfold specific
target proteins, disassemble multimeric complexes, and solubilize protein aggregates (Kim et al.,
2000; Konieczny and Liberek, 2002; Levchenko et al., 1995; Seonga et al., 2000; Wawrzynow et
al., 1995; Weber-Ban et al., 1999; Wickner et al., 1994; Zolkiewski, 1999). The minimal AAA+
unit contains an ATPase domain with a mixed -structure and a C-terminal domain that is
largely -helical, but most family members also contain additional structural domains (Bochtler
et al., 2000; Maurizi and Li, 2001; Neuwald et al., 1999; Schirmer et al., 1996). ClpX subfamily
members, for example, contain a short N-terminal Cys-cluster domain, an AAA+ ATPase
domain, and a C-terminal AAA+ domain (Singh et al., 2001). One of the current challenges in
understanding the biological activities of Clp/Hsp100 family members such as ClpX is to
determine how the structural domains and interactions between these domains contribute to
overall function.
ClpX forms a ring hexamer that unfolds target proteins and dismantles specific protein
complexes in an ATP-dependent fashion; the ClpX hexamer can also serve as the substrate
recognition and unfolding module of the ClpXP protease (Gottesman et al., 1997b). Different
Clp/Hsp100 family members generally have distinct substrate recognition preferences
(Gottesman et al., 1997a; Gottesman et al., 1997b; Suzuki et al., 1997). Well-characterized
substrates of ClpX and ClpXP include the bacteriophage Mu transposase and proteins with ssrA
degradation tags, both of which are recognized via C-terminal peptide sequences (GonciarzSwiatek et al., 1999; Gottesman et al., 1998; Kim et al., 2000; Levchenko et al., 1995;
66
Wawrzynow et al., 1995). Structural information is not available for ClpX, but several structures
of the related HslU (ClpY) hexamer have been determined (Bochtler et al., 2000; Song et al.,
2000; Sousa et al., 2000; Wang, 2001; Wang et al., 2001). The amino-acid sequences of the
AAA+ domains from Escherichia coli ClpX and HslU are 38% identical over 303 common
positions.
Potential roles for the -helical domain in AAA+ family members include structural stabilization
of hexameric or multimeric structures (Bochtler et al., 2000; Singh et al., 2001; Sousa et al.,
2000), providing an arginine that serves as an active-site residue for ATP hydrolysis (Guenther et
al., 1997; Jeruzalmi et al., 2001; Karata et al., 2001; Neuwald et al., 1999), and mediating
recognition of specific protein and peptide substrates (Cashikar et al., 2002; Levchenko et al.,
1997a; Smith et al., 1999). Some of these functions, however, are controversial (Feng and
Gierasch, 1998; Ishikawa et al., 2001; Ortega et al., 2000; Singh et al., 2001; Wickner and
Maurizi, 1999) and additional undiscovered roles for the -helical AAA+ domain may also exist.
In this paper, we present a genetic and biochemical analysis of the functional roles of residues in
the C-terminal domain of E. coli ClpX. By screening sequence substitutions in ClpX, we
identified mutations in the C-terminal domain that caused reduced disassembly activity in vivo.
Three of these proteins had low disassembly activity for one substrate and essentially no
unfolding activity for another substrate in vitro. These mutant enzymes bound substrate proteins
but displayed reduced ATP hydrolysis in response to binding, instead of increased hydrolysis.
We propose that these ClpX mutants are defective in a reaction step that engages bound substrate
proteins. This engagement step is required both for enhancement of ATP hydrolysis and for
initiation of the unfolding and/or disassembly process.
67
Results
Mutant construction, screening, and purification. Using multiple sequence alignments and
the structure of the homologous HslU protein (Bochtler et al., 2000; Song et al., 2000; Sousa et
al., 2000; Wang, 2001; Wang et al., 2001) as guides, mutations were introduced into the Cterminal domain of E. coli ClpX by site-directed mutagenesis of a plasmid-borne gene encoding
a variant of ClpX. We initially mutated residues that appeared to be solvent exposed based on
the HslU structure and that differed between ClpX and other Clp/Hsp100 family members. After
screening this initial set of mutations for activity as described below, we also mutated residues
that neighbored positions found to be defective. In all, 50 mutations representing 44 different
residue positions in ClpX were constructed.
Activity assays in vivo were based on the fact that ClpX is required for lytic growth of
bacteriophage Mu because ClpX must disassemble DNA-bound tetramers of Mu transposase
(MuA) before DNA replication and subsequent phage assembly and cell lysis can occur
(Mhammedi-Alaoui et al., 1994; Nakai and Kruklitis, 1995). In this assay, ClpX-defective host
cells containing a Mu lysogen with a temperature-sensitive repressor did not lyse and continued
to grow normally after a shift to the restrictive temperature (Fig. 1A). When wild-type ClpX was
provided on a plasmid, however, the host cells lysed roughly 30-40 minutes after the temperature
shift (Fig. 1A). Most of the ClpX mutants (37 of 50) showed lysis kinetics indistinguishable
from wild-type ClpX in this assay (Figs. 1A & 2A; data not shown). However, seven variants
showed moderate delays in lysis (Figs. 1B & 2A) and an additional six ClpX mutants showed
significant delays in lysis kinetics (Figs. 1C & 2A).
68
Figure 1. ClpX mutants support cell lysis by bacteriophage Mu to different
degrees. (A) Kinetics of wild-type ClpX and three mutants (A364D, Q406E, &
I412Q) with no lysis delay. (B) Kinetics of ClpX mutants displaying moderate
lysis delays. (C) Kinetics of mutants with long lysis delays.
A. No delay
2
1.5
No ClpX
A364D
Q406E
I412Q
Wild type
1
0.5
Absorbance (540 nm)
0
0
20
40
60
80
B. Moderate delay
2
No ClpX
V402D
V396A
K335E
M390D
S407L
V397T
A357E
Wild type
1.5
1
0.5
0
0
20
40
60
80
C. Long delay
2
No ClpX
Y385A
V393G
L381K
D382K
D399R
I403A
Wild type
1.5
1
0.5
0
0
20
40
60
Time (min)
80
70
Figure 2.
Positions
of mutations
in the
C-terminal
domain
of ClpX.
A
325
326
348
327
349
328
350
329
351
330
314
352
331
315
353
332
316
354
333
317
355
334
318
356
335
319
357
336
320
358
337
321
359
338
322
360
339
323
361
340
324
362
341
363
342
364
343
365
344
366
345
His
Lys
Lys Arg
Ala Arg
Leu Asn Glu Leu Ser Glu Glu Ala Leu Ile Gln
347
390
391
392
411
393
412
394
413
395
414
396
415
397
416
398
417
399
418
400
419
401
420
402
421
403
422
404
423
405
424
406
407
408
409
————————— Cluster II ——————————
|
|
|
|
|
Asp
Gly
Ala Thr
Arg
Asp Asp Ala
Leu
| Lys
| Arg
|
|
| Arg Ala |
| Arg Asn Glu | Glu
Met Glu Asp Val Glu Lys Val Val Ile Asp Glu Ser Val Ile Asp Gly Gln Ser Lys Pro
—— Cluster I ——
|
|
|
Asp
Lys Lys
Ala
Arg
|
| Ile
| Asn
Ala Arg Gly Leu Arg Ser Ile Val Glu Ala Ala Leu Leu Asp Thr Met Tyr Asp Leu Pro
369 370 371 372 373 374 375 376
Sensor-II helix 381 382 383 384 385 386 387 388
367
Glu
Glu
Ser
Glu
Lys
Val Lys Lys
Ala
Tyr Asp
Asp Leu Glu Phe Arg Asp Glu Ala Leu Asp Ala Ile Ala Lys Lys Ala Met Ala Arg Lys Thr
Glu
Glu
Ser
|
Lys
Ile Leu Lys Glu Pro Lys Asn Ala Leu Thr Lys Gln Tyr Gln Ala Leu Phe Asn Leu Glu Gly
(A) Mutations
causing neutral Val
phenotypes in 346
the Mu lysis
assay are shown
in green above
the wild-type
Gly
sequence.
368
Moderately
defective
and severely
defective
mutations are Leu
Ser
shown in
389
yellow
or red boxes,
respectively.
410
Gln
Glu
Lys Tyr
Leu Leu Ile Tyr Gly Lys Pro Glu Ala Gln Gln Ala Ser Gly Glu
Figure 2. (B) Mutations were mapped onto the homologous E. coli HslU
structure10, based on sequence alignments. The ATPase domains of subunits in
the ClpX hexamer are colored dark or light gray. Residues in the C-terminal
domain are colored dark blue (not mutated), green (neutral), yellow (moderately
defective), or red (severely defective) based on their Mu lysis phenotypes.
B
Cluster I
Cluster II
Most of the defective mutations fell into one of two clusters on a homology model of ClpX (Fig.
2B). The first group, cluster I, included the L381K, D382K, and Y385A mutations and lies near
the interface between the C-terminal domain of one subunit and the ATPase domain of the
adjacent subunit (Figs. 2A & 2B). The second group, cluster II, included the M390D, V396A,
V397T, D399R, and V402D mutations (Figs. 2A & 2B). The mutants that comprise clusters I
and II were subjected to a purification protocol that included Ni++-NTA chromatography and
Mono Q anion-exchange chromatography in the presence of ATP. The D399R mutant could not
be purified, presumably because of altered stability or solubility properties. The remaining seven
mutant proteins purified in a manner similar to wild-type ClpX, except for slightly earlier or later
elution from the ion-exchange column for mutants with additional positive/negative charges
(data not shown).
Disassembly of MuA-DNA complexes in vitro. Based upon their phenotypes in vivo, we
expected that the purified mutants would be defective in disassembling DNA complexes of the
MuA tetramer in vitro. Indeed, the cluster-I mutants had substantial defects in disassembly in
vitro, whereas the cluster-II mutants had disassembly activities that were usually intermediate
between those of wild-type ClpX and the cluster-I mutants (Figs. 3A & 4B). There was general
agreement between the phage Mu lysis assay and the MuA-DNA disassembly assay.
In
particular, the cluster-I mutants (L381K, D382K & Y385A) were among the most defective both
in vivo and in vitro (Figs. 4A & 4B). Most cluster-II mutants, by contrast, showed intermediate
levels of activity in both assays.
73
Unfolding of GFP-ssrA. The purified ClpX mutants were tested for their ability to denature
green-fluorescent protein with a C-terminal ssrA tag (GFP-ssrA). The ssrA tag provides a
recognition site for ClpX (Gottesman et al., 1998; Kim et al., 2000) and for SspB (Levchenko et
al., 2000), a specificity factor that enhances recognition of ssrA-tagged substrates by ClpX. As
assayed by loss of native GFP-ssrA fluorescence, the cluster-I mutants had no detectable
unfolding activity either in the presence of SspB (Figs. 3B & 4E; data not shown) or in its
absence (data not shown). By contrast, the cluster-II mutants showed robust GFP-ssrA unfolding
activities (Figs. 3B & 4E; data not shown). In fact, the M390D, V397T, and V402D mutants,
which showed reduced disassembly of MuA-DNA complexes, were slightly more active than
wild-type ClpX in denaturing GFP-ssrA.
ATP hydrolysis and substrate-binding effects. In ATPase assays, each of the seven purified
ClpX mutants showed substantial rates of basal hydrolysis, although usually less than the wildtype rate (Fig. 5). L381K and D382K had the lowest ATPase activities, which were roughly half
of the wild-type value. ATP hydrolysis by wild-type ClpX is stimulated in the presence of
protein substrates (Wawrzynow et al., 1995).
The ATPase activities of the mutants were
therefore assayed in the presence of different concentrations of GFP-ssrA and Arc-MuA10. ArcMuA10 (Arc repressor fused to the 10 C-terminal residues of MuA) was used for these studies
because intact MuA was not sufficiently soluble at the concentrations required for these assays,
and the C-terminal residues of MuA are sufficient for mediating interactions with ClpX
(Levchenko et al., 1997b). The ATPase activity of wild-type ClpX was not stimulated by
untagged Arc repressor (data not shown).
74
Figure 3. Disassembly and unfolding assays.
(A) Disassembly of MuA-DNA complexes in the presence of wild-type ClpX
(with and without ATP) or the Y385A, V396A, and M390D mutants.
(B) Unfolding kinetics of GFP-ssrA, assayed by the loss of native fluorescence
at 511 nm, catalyzed by wild-type or mutant ClpX in the presence of SspB.
MuA-DNA disassembly
A
Wild
type
}
% disassembled in
20 minutes
100
+ATP
M390D
50
V396A
-ATP
0
Y385A
GFP-ssrA unfolding
B
Y385A
100
% folded
GFP-ssrA
L381K
50
V396A
Wild type
M390D
0
0
5
10
Time (min)
15
76
int
-log (K
)
6
4
2
0
Arc-MuA10/ClpX
interaction
20
-ATP
0
F
6
4
2
0
-40
Wild
type
80
+ATP
V396A
V397T
V402D
-20
M390D
GFP-ssrA/ClpX
interaction
V402D
M390D
V396A
V397T
Wild
type
40
M390D
V396A
V397T
V402D
E
}
% change in rate
0
L381K
D382K
Y385A
Wild
type
M390D
V396A
V397T
V402D
20
L381K
D382K
Y385A
MuA-DNA disassembly
% unfolded in 20 min.
V402D
V397T
0
60
Wild
type
0
M390D
+ATP
D
L381K
D382K
Y385A
C
Mu cell lysis
-log (Kint )
20
V396A
60
M390D
V396A
V397T
V402D
Wild type
}
-1
.03
Wild
type
40
Y385A
B
L381K
D382K
Y385A
.01
L381K
D382K
Y385A
[50% lysis time (min)]
.02
L381K
D382K
% disassembled in 20 min.
A
Substrate effect
on ATPase rate
+Arc-MuA10
+GFP-ssrA
GFP-ssrA unfolding
60
40
-ATP
Figure 4. Summary of characterized ClpX mutant activities.
(A) Mu lysis activity in vivo.
(B) MuA-DNA disassembly activity in vitro.
(C) Kint values for Arc-MuA10.
(D) Effect of substrate (Arc-MuA10 or GFP-ssrA) at saturating
substrate concentration on ATP hydrolysis activity.
(E) GFP-ssrA unfolding activity in the presence of SspB.
(F) Kint values for GFP-ssrA.
0
200
Cluster
I
100
Cluster
II
V402D
V397T
V396A
M390D
Y385A
Wild
type
300
D382K
L381K
ATP turnover
(ATP min-1 [ClpX6]-1)
Figure 5. Basal ATPase activity.
80
Figure 6. Substrate-dependent changes in ATP hydrolysis. The panels
on the left side show changes in ATP hydrolysis in the absence of SspB
as a function of increasing concentrations of the substrate, GFP-ssrA, for
wild-type ClpX, a cluster-I mutant (L381K), and a cluster-II mutant (M390D).
The right-side panels show parallel experiments with a different substrate,
Arc-MuA10.
Wild type
Wild type
320
280
280
240
240
ATP turnover (ATP min-1 [ClpX6]-1)
320
0
10
20
L381K
120
100
80
80
10
20
M390D
240
240
200
200
160
160
0
10
[GFP-ssrA] (mM)
20
20
0
20
120
100
0
0
0
20
40
L381K
40
M390D
40
[Arc-MuA10] (mM)
82
The ATPase rates of each of the cluster-I and cluster-II mutants changed when GFP-ssrA or ArcMuA10 was present (Figs. 4D & 6).
The four cluster-II mutants, like wild type, showed
increased ATP hydrolysis activity in the presence of both protein substrates.
Strikingly,
however, the ATPase activity of each cluster-I mutant decreased when either GFP-ssrA or ArcMuA10 was added. Thus, the cluster-I mutants bind protein substrates, but respond to this
binding in a manner opposite from the wild-type enzyme.
The changes in ClpX ATPase rates with protein-substrate addition were fit to obtain interaction
constants (Kint) for GFP-ssrA and Arc-MuA10. Kint for ClpX interaction with GFP-ssrA was 2.0
± 0.59 M, a value indistinguishable from the published Km for ClpXP degradation of GFP-ssrA
(1.9 ± 0.3 M; Kim et al., 2000). Six of the seven purified ClpX mutants, including all of the
cluster-I mutants, had Kint values for GFP-ssrA within 50% of the wild-type value (Fig. 4F).
Hence, the inability of the cluster-I mutants to unfold GFP-ssrA is not caused by an inability to
bind GFP-ssrA. One cluster-I mutant (D382K; Kint = 0.99 ± 0.58 M) and one cluster-II mutant
(V397T; Kint = 0.34 ± 0.12 M) interacted with GFP-ssrA slightly more tightly than the wildtype enzyme (Fig. 4F).
Kint for ClpX interaction with Arc-MuA10 was 12.5 ± 4 M, a value similar to a previously
reported interaction constant for Arc-MuA10 with ClpXP (14 ± 3 M; Kim et al., 2000). The
cluster-I mutants had Kint values for Arc-MuA10 within 50% of the wild-type value (Fig. 4C).
Once again, the poor MuA disassembly activities of these mutants cannot be explained by their
inability to bind to substrate. The cluster-II mutants showed a wider range of interaction
constants for Arc-MuA10. For example, relative to wild type, Kint for Arc-MuA10 was increased
83
about 1.5-fold for the M390D mutant and decreased about 3-fold for the V396A and V397T
mutants (Fig. 4C).
Discussion
We constructed and studied the properties of E. coli ClpX mutants with site-directed amino-acid
substitutions in the C-terminal domain. In the set of 50 mutants, 37 supported the replication of
phage Mu and subsequent cell lysis in a manner indistinguishable from wild type. These
positions are marked in green in the homology model of the ClpX hexamer shown in Fig. 2B. In
13 cases, however, we identified mutations that caused delays in Mu lysis. The majority of the
defective mutations that caused modest lysis delays (colored yellow in Fig. 2B) or severe delays
(colored red in Fig. 2B) formed two clusters on the surface of the C-terminal domain.
Cluster II includes the M390D, V396A, V397T, D399R, and V402D mutations. Most cluster-II
mutants had relatively subtle lysis defects in vivo and the mutants that were purified from this
group generally showed small changes in disassembly or unfolding activities, ATP hydrolysis
rates, or substrate interaction constants. Compared to wild-type ClpX, most cluster-II mutants
showed reduced activity in MuA-DNA disassembly but had slightly increased activity in GFPssrA unfolding. This distinctive substrate-specific behavior of the cluster-II mutants suggests
that ClpX interacts with MuA and GFP-ssrA in somewhat different ways during protein
unfolding. These differences are not, however, correlated with changes in binding to GFP-ssrA
or to an Arc variant (Arc-MuA10) bearing the C-terminal ClpX-recognition residues of MuA.
For example, the V396A ClpX mutant showed the poorest MuA-DNA disassembly activity
among the cluster-II mutants but bound Arc-MuA10 about 3-fold more strongly than wild type.
84
The adjacent V397T mutant also bound Arc-MuA10 about 3-fold better than wild type but had
the best MuA-DNA disassembly activity among the cluster-II mutants.
Because most of the cluster-II mutants caused relatively small changes in activity and substrate
binding, it seems clear that these mutations do not define a substrate interaction site or a site
required for ClpX unfolding activity.
The cluster-II mutations probably exert their effects
indirectly, possibly by destabilizing one enzyme conformation, thereby increasing the population
of an alternative conformation.
The purified cluster-I mutants were quite defective in disassembling MuA-DNA complexes in
vitro and displayed essentially no ability to catalyze denaturation of another ClpX substrate,
GFP-ssrA. These activity deficiencies do not appear to result from major folding or stability
defects, as the L381K, D382K, and Y385A mutants had purification and solubility properties
similar to wild-type ClpX, and had basal ATPase activities within two-fold of wild type.
Moreover, the most severely deficient mutant (Y385A) formed oligomeric complexes
indistinguishable from wild-type ClpX in formaldehyde crosslinking experiments in the presence
of ATP (data not shown). The dramatic reduction in GFP-ssrA denaturation does not result from
an inability of the cluster-I mutants to bind substrates. Each of these mutants have Kint values for
GFP-ssrA within 50% of the wild-type value. Similarly, all three mutants have Kint values for
Arc-MuA10 within 2-fold of the wild-type value. One of the most interesting characteristics of
the cluster-I mutants is that they display reduced ATPase activities in response to GFP-ssrA or
Arc-MuA10 binding. Wild-type ClpX and all of the cluster-II mutants, by contrast, hydrolyze
ATP more rapidly in response to substrate binding.
85
In the structure of the HslU hexamer, each C-terminal domain packs tightly against the ATPase
domain of an adjacent subunit, providing a potential avenue of communication between
ATP/ADP binding sites in adjacent subunits. The sensor-II helix in the C-terminal AAA+
domain (residues 369-387 in ClpX) plays a central role in this interaction. At the N-terminal end
of this helix, the side chain of an invariant Arg residue (Arg370 in E. coli ClpX) contacts the
nucleotide bound to its own ATPase domain. At the C-terminal end of the sensor-II helix, the
side chain of an aromatic residue (Tyr385 in E. coli ClpX) intercalates between side chains from
-helix I and II of the adjacent subunit's ATPase domain, and side chains in the box-II loop
connecting helices I and II contact the nucleotide in this domain. This arrangement is illustrated
in Fig. 7. Hence, the Y385A ClpX mutation could disrupt subunit-subunit interactions which, in
turn, regulate coordinated ATP hydrolysis.
One model that could explain the properties of the cluster-I mutants is as follows. In an initial
uncommitted binding step, protein substrates bind to ClpX in the ATP state. This uncommitted
complex has decreased ATPase activity relative to the basal rate. Normally, the enzyme then
engages the bound protein in a second reaction step that requires communication between
adjacent subunits of the ClpX hexamer and some type of conformational change. The ATPase
activity of the protein-bound ClpX in this engaged state is higher than the basal rate. The
engaged-state could be one in which the native protein is subjected to mechanical forces by the
enzyme which increase the probability of substrate denaturation. By this model, progression
from the uncommitted binding state to the engaged state would occur significantly more slowly
for the cluster-I mutants. We assume that the rate of ATP hydrolysis sets the cycle time for
86
Figure 7. Y385A could disrupt communication between
nucleotide-binding sites in adjacent subunits.
The position of Tyr385 in the sensor-II helix of ClpX is shown in a
homology model based on amino-acid sequence alignments between
E. coli ClpX and H. influenzae HslU19. A conserved arginine (Arg370
in E. coli ClpX; Arg394 in H. influenzae HslU) at the N-terminal end
of the sensor-II helix contacts the nucleotide in its own subunit. At the
C-terminal end of the sensor-II helix, a conserved aromatic side chain
(Tyr385 in E. coli ClpX; Phe409 in H. influenzae HslU) intercalates
between side-chains from a-helices I and II in the ATPase domain of
the adjacent subunit. In turn, the box-II loop connecting a-helices I and
II contacts the bound nucleotide in this subunit.
Arg370
Sensor-II
helix
One subunit
Tyr385
88
ClpX. Because the overall rate of hydrolysis is similar for the free enzyme, the uncommitted
complex, and the engaged complex, mutants that spend substantially more time in uncommitted
complex will spend less time in the engaged complex.
For substrates like GFP-ssrA,
denaturation might not occur in the cluster-I mutants for two reasons. The GFP-ssrA could
dissociate from the uncommitted complex before engagement occurs, or the GFP-ssrA could
reach the engaged complex slowly but spend too little time in this state to have a significant
probability of being denatured.
The model presented above makes several predictions. First, if communication between domains
mediated by the sensor-II helix is important, then mutations at the N-terminus of this helix and/or
in the parts of the ATPase domain that contact this helix should cause phenotypes similar to the
cluster-I mutations. Second, complexes of the cluster-I ClpX mutants with substrates like GFPssrA should have kinetic and/or physical properties that differ markedly from complexes of wildtype ClpX with GFP-ssrA. Experimental tests of these predictions are in progress.
Note added in proof: The P67H mutation, which alters an ATPase-domain residue that contacts
Y385, causes severe defects in Mu lysis and in GFP-ssrA unfolding. This mutant, like the
cluster-I mutants, binds GFP-ssrA like wild type and is a substrate-inhibited ATPase, supporting
the model that contacts between Y385 and the ATPase domain regulate the coordination of ATP
hydrolysis.
89
Experimental procedures
Strains and Plasmids. To construct E. coli strain SJ9, a bacteriophage Mu-cts62 lysogen of
strain W3110 (kindly provided by Maurice Fox, MIT) was infected with a P1 lysate prepared
from E. coli strain SG22177 (Gottesman et al., 1998) and clpX::kan transductants were selected.
The wild-type ClpX protein with an N-terminal His6-tag was encoded in a pET-14b-ClpX vector
(Levchenko et al., 1997b). Overlap extension PCR mutagenesis (Ho et al., 1989) was used to
introduce each point mutation and a new restriction site into a portion of the clpX gene encoding
the C-terminal domain, and the resulting DNA fragments were cloned into pET-14b-ClpX
containing the rest of the clpX gene. Following transformation into E. coli strain DH5, plasmid
DNA was isolated from ampicillin-resistant candidates and screened for the new restriction site
by digestion with an appropriate restriction enzyme. The plasmid region that was PCR amplified
during mutagenesis was sequenced to confirm the presence of the expected mutations.
Solutions. NB buffer contained 50 mM HEPES (pH 7.6), 500 mM NaCl, 40 mM imidazole,
15% glycerol, and 5 mM -mercaptoethanol. NE buffer was NB buffer with 500 mM imidazole
instead of 40 mM imidazole. QB buffer contained 50 mM HEPES (pH 7.6), 15% glycerol, 3
mM dithiothreitol (DTT), 100 M ZnSO4, 100 M ATP, 1 mM MgCl2, and 0.01% Nonidet P-40
(NP-40). ML buffer contained 43 mM Tris•Cl (pH 8.0), 6% sucrose, 66 mM NaCl, 3.3 mM
EDTA, 0.66 mM DTT, 13.3 mM spermidine-HCl, and 19.4 mg mL-1 ammonium sulfate. H
buffer contained 25 mM HEPES (pH 7.6), 0.1 mM EDTA, 1 mM DTT, and 10% glycerol. AH
buffer contained 42 mM HEPES (pH 7.6), 6.2 mM Tris•Cl (pH 8.0), 133 mM KCl, 5.3 mM
MgCl2, 312 M EDTA, 1 mM DTT, 34 M ZnSO4, 34 M ATP, 0.035% NP-40, and 15%
90
glycerol. ATP mix I contained 2.5 mM ATP, 1 mM NADH, 7.5 mM phosphoenolpyruvate,
18.75 U mL-1 pyruvate kinase, and 21.45 U mL-1 lactate dehydrogenase.
Proteins. His6-tagged ClpX proteins were purified from E. coli strain BL21(DE3)/pET-14bClpX cells grown at 30 °C in 1.33% tryptone, 2.66% yeast extract, 0.44% glycerol, and 100 g
mL-1 ampicillin. Protein expression was induced with 0.6 mM IPTG, and cells were harvested
three hours after induction. Cell pellets were resuspended at 4 °C in 3 mL of NB buffer per gram
of cells, and protease-inhibitor cocktail set II (Calbiochem) was added as recommended by the
manufacturer. Cells were lysed using a French press, and the lysate was centrifuged for one hour
at 15,000 rpm in a Sorvall SA-600 rotor. The supernatant was filtered through a 0.2 m filter
and mixed by rotation with Ni-NTA resin (equilibrated in NB buffer) for 45 minutes at 4 C. A
column was packed by gravity flow at 4 C and then washed with 100 column volumes of NB
buffer. Bound protein was eluted with five column volumes of NE buffer and buffer exchanged
into QB buffer plus 150 mM KCl using a PD-10 desalting column (Amersham Pharmacia
Biotech).
This material was applied to a Mono Q HR 5/5 column (Amersham Pharmacia
Biotech), which was developed with a 10-volume linear gradient in QB buffer from 150 to 500
mM KCl. Peak absorbance fractions were analyzed by SDS-polyacrylamide gel electrophoresis,
and those containing ClpX protein were pooled, divided into aliquots, and stored at –80 C.
MuA protein, expressed in E. coli strain BL21(DE3)/pMK591 (Baker et al., 1993), was purified
using a protocol based on a previously published procedure (Williams et al., 1999). Cells were
resuspended in ML buffer and lysed by French press. After precipitation with ammonium
sulfate, pellets were collected by centrifugation, resuspended in H buffer plus 100 mM NaCl,
91
filtered through 0.2 m filters, and buffer exchanged into H buffer plus 300 mM NaCl using a
PD-10 column.
This material was applied to a SP Sepharose Fast Flow HR 5/5 column
(Amersham Pharmacia Biotech), and a five-volume linear gradient in H buffer from 300 to 1000
mM NaCl was run. Fractions containing MuA were pooled, buffer exchanged into H buffer plus
300 mM NaCl, and loaded onto a Mono S HR 5/5 column (Amersham Pharmacia Biotech),
which was developed with a five-volume linear gradient in H buffer from 300 to 1000 mM NaCl.
Peak fractions were pooled, divided into aliquots, and stored at –80 C.
GFP-ssrA was purified using a published protocol (Yakhnin et al., 1998). Purified SspB protein
was a gift from David Wah (MIT). Arc-MuA10 (Levchenko et al., 1997b) was purified using a
protocol developed for other Arc variants (Burton et al., 2001b). UV absorbance or dye binding
(Bradford, 1976) were used to determine final protein concentrations.
Mu lysis screen. Overnight cultures of SJ9 host cells containing wild-type or mutant pET-14bClpX plasmids were grown in Luria-Bertani broth at 30 C with 100 g mL-1 ampicillin. These
cultures were diluted 1:100 and grown until the absorbance at 540 nm was between 0.40 and
0.45. The cultures were shifted to 42 C for 30 minutes and then shifted to 37 °C for 50 minutes.
After the initial temperature shift, absorbance readings at 540 nm were taken every 10 minutes.
Biochemical assays. MuA-DNA disassembly gel-shift assays were performed as described
(Burton et al., 2001a), except 0.1 M ClpX6 was used in each reaction. GFP-ssrA unfolding
reactions (30 °C) were performed as described (Kim et al., 2000) using 0.26 M ClpX6 with or
without 0.46 M SspB protein. ATPase assays (30 °C) were performed with 0.3 M ClpX6 in
92
AH buffer containing ATP mix I but otherwise followed the published protocol (Kim et al.,
2001).
93
Acknowledgements
We thank Scott Boyd, Briana Burton, Randy Burton, Robert Grant, Igor Levchenko, Hillel
Schwartz, David Wah, Robert Horvitz, and Jonathan King for technical help and discussions,
materials, and use of equipment.
T.A.B. is an employee of the Howard Hughes Medical
Institute. Supported by NIH grant AI-16892.
94
References
Baker, T.A., Mizuuchi, M., Savilahti, H., and Mizuuchi, K. (1993) Division of labor among
monomers within the Mu transposase tetramer. Cell 74: 723-733.
Bochtler, M., Hartmann, C., Song, H.K., Bourenkov, G.P., Bartunik, H.D., and Huber, R. (2000)
The structures of HsIU and the ATP-dependent protease HsIU-HsIV. Nature 403: 800805.
Bradford, M.M. (1976) A rapid and sensitive method for the quantitation of microgram
quantities of protein utilizing the principle of protein-dye binding. Anal Biochem 72: 248254.
Burton, B.M., Williams, T.L., and Baker, T.A. (2001a) ClpX-mediated remodeling of Mu
transpososomes: selective unfolding of subunits destabilizes the entire complex. Mol Cell
8: 449-454.
Burton, R.E., Siddiqui, S.M., Kim, Y.I., Baker, T.A., and Sauer, R.T. (2001b) Effects of protein
stability and structure on substrate processing by the ClpXP unfolding and degradation
machine. EMBO J 20: 3092-3100.
Cashikar, A.G., Schirmer, E.C., Hattendorf, D.A., Glover, J.R., Ramakrishnan, M.S., Ware,
D.M., and Lindquist, S.L. (2002) Defining a pathway of communication from the Cterminal peptide binding domain to the N-terminal ATPase domain in a AAA protein.
Mol Cell 9: 751-760.
Feng, H.P., and Gierasch, L.M. (1998) Molecular chaperones: clamps for the Clps? Curr Biol 8:
R464-467.
Gonciarz-Swiatek, M., Wawrzynow, A., Um, S.J., Learn, B.A., McMacken, R., Kelley, W.L.,
Georgopoulos, C., Sliekers, O., and Zylicz, M. (1999) Recognition, targeting, and
95
hydrolysis of the O replication protein by the ClpP/ClpX protease. J Biol Chem 274:
13999-14005.
Gottesman, S., Maurizi, M.R., and Wickner, S. (1997a) Regulatory subunits of energy-dependent
proteases. Cell 91: 435-438.
Gottesman, S., Wickner, S., and Maurizi, M.R. (1997b) Protein quality control: triage by
chaperones and proteases. Genes Dev 11: 815-823.
Gottesman, S., Roche, E., Zhou, Y., and Sauer, R.T. (1998) The ClpXP and ClpAP proteases
degrade proteins with carboxy-terminal peptide tails added by the SsrA-tagging system.
Genes Dev 12: 1338-1347.
Guenther, B., Onrust, R., Sali, A., O'Donnell, M., and Kuriyan, J. (1997) Crystal structure of the
´ subunit of the clamp-loader complex of E. coli DNA polymerase III. Cell 91: 335-345.
Ho, S.N., Hunt, H.D., Horton, R.M., Pullen, J.K., and Pease, L.R. (1989) Site-directed
mutagenesis by overlap extension using the polymerase chain reaction. Gene 77: 51-59.
Ishikawa, T., Beuron, F., Kessel, M., Wickner, S., Maurizi, M.R., and Steven, A.C. (2001)
Translocation pathway of protein substrates in ClpAP protease. Proc Natl Acad Sci USA
98: 4328-4333.
Jeruzalmi, D., O'Donnell, M., and Kuriyan, J. (2001) Crystal structure of the processivity clamp
loader gamma () complex of E. coli DNA polymerase III. Cell 106: 429-441.
Karata, K., Verma, C.S., Wilkinson, A.J., and Ogura, T. (2001) Probing the mechanism of ATP
hydrolysis and substrate translocation in the AAA protease FtsH by modelling and
mutagenesis. Mol Microbiol 39: 890-903.
96
Kim, Y.I., Burton, R.E., Burton, B.M., Sauer, R.T., and Baker, T.A. (2000) Dynamics of
substrate denaturation and translocation by the ClpXP degradation machine. Mol Cell 5:
639-648.
Kim, Y.I., Levchenko, I., Fraczkowska, K., Woodruff, R.V., Sauer, R.T., and Baker, T.A. (2001)
Molecular determinants of complex formation between Clp/Hsp100 ATPases and the
ClpP peptidase. Nat Struct Biol 8: 230-233.
Konieczny, I., and Liberek, K. (2002) Cooperative action of Escherichia coli ClpB protein and
DnaK chaperone in the activation of a replication initiation protein. J Biol Chem 11: 11.
Levchenko, I., Luo, L., and Baker, T.A. (1995) Disassembly of the Mu transposase tetramer by
the ClpX chaperone. Genes Dev 9: 2399-2408.
Levchenko, I., Smith, C.K., Walsh, N.P., Sauer, R.T., and Baker, T.A. (1997a) PDZ-like
domains mediate binding specificity in the Clp/Hsp100 family of chaperones and
protease regulatory subunits. Cell 91: 939-947.
Levchenko, I., Yamauchi, M., and Baker, T.A. (1997b) ClpX and MuB interact with overlapping
regions of Mu transposase: implications for control of the transposition pathway. Genes
Dev 11: 1561-1572.
Levchenko, I., Seidel, M., Sauer, R.T., and Baker, T.A. (2000) A specificity-enhancing factor for
the ClpXP degradation machine. Science 289: 2354-2356.
Maurizi, M.R., and Li, C.C. (2001) AAA proteins: in search of a common molecular basis.
International Meeting on Cellular Functions of AAA Proteins. EMBO Rep 2: 980-985.
Mhammedi-Alaoui, A., Pato, M., Gama, M.J., and Toussaint, A. (1994) A new component of
bacteriophage Mu replicative transposition machinery: the Escherichia coli ClpX protein.
Mol Microbiol 11: 1109-1116.
97
Nakai, H., and Kruklitis, R. (1995) Disassembly of the bacteriophage Mu transposase for the
initiation of Mu DNA replication. J Biol Chem 270: 19591-19598.
Neuwald, A.F., Aravind, L., Spouge, J.L., and Koonin, E.V. (1999) AAA+: A class of
chaperone-like ATPases associated with the assembly, operation, and disassembly of
protein complexes. Genome Res 9: 27-43.
Ortega, J., Singh, S.K., Ishikawa, T., Maurizi, M.R., and Steven, A.C. (2000) Visualization of
substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 6:
1515-1521.
Schirmer, E.C., Glover, J.R., Singer, M.A., and Lindquist, S. (1996) HSP100/Clp proteins: a
common mechanism explains diverse functions. Trends Biochem Sci 21: 289-296.
Seonga, I.S., Oha, J.Y., Leeab, J.W., Tanakab, K., and Chunga, C.H. (2000) The HslU ATPase
acts as a molecular chaperone in prevention of aggregation of SulA, an inhibitor of cell
division in Escherichia coli. FEBS Lett 477: 224-229.
Singh, S.K., Rozycki, J., Ortega, J., Ishikawa, T., Lo, J., Steven, A.C., and Maurizi, M.R. (2001)
Functional domains of the ClpA and ClpX molecular chaperones identified by limited
proteolysis and deletion analysis. J Biol Chem 276: 29420-29429.
Smith, C.K., Baker, T.A., and Sauer, R.T. (1999) Lon and Clp family proteases and chaperones
share homologous substrate-recognition domains. Proc Natl Acad Sci USA 96: 66786682.
Song, H.K., Hartmann, C., Ramachandran, R., Bochtler, M., Behrendt, R., Moroder, L., and
Huber, R. (2000) Mutational studies on HslU and its docking mode with HslV. Proc Natl
Acad Sci USA 97: 14103-14108.
98
Sousa, M.C., Trame, C.B., Tsuruta, H., Wilbanks, S.M., Reddy, V.S., and McKay, D.B. (2000)
Crystal and solution structures of an HslUV protease-chaperone complex. Cell 103: 633643.
Suzuki, C.K., Rep, M., van Dijl, J.M., Suda, K., Grivell, L.A., and Schatz, G. (1997) ATPdependent proteases that also chaperone protein biogenesis. Trends Biochem Sci 22: 118123.
Wang, J. (2001) A corrected quaternary arrangement of the peptidase HslV and ATPase HslU in
a cocrystal structure. J Struct Biol 134: 15-24.
Wang, J., Song, J.J., Franklin, M.C., Kamtekar, S., Im, Y.J., Rho, S.H., Seong, I.S., Lee, C.S.,
Chung, C.H., and Eom, S.H. (2001) Crystal structures of the HslVU peptidase-ATPase
complex reveal an ATP-dependent proteolysis mechanism. Structure (Camb) 9: 177-184.
Wawrzynow, A., Wojtkowiak, D., Marszalek, J., Banecki, B., Jonsen, M., Graves, B.,
Georgopoulos, C., and Zylicz, M. (1995) The ClpX heat-shock protein of Escherichia
coli, the ATP-dependent substrate specificity component of the ClpP-ClpX protease, is a
novel molecular chaperone. EMBO J 14: 1867-1877.
Weber-Ban, E.U., Reid, B.G., Miranker, A.D., and Horwich, A.L. (1999) Global unfolding of a
substrate protein by the Hsp100 chaperone ClpA. Nature 401: 90-93.
Wickner, S., Gottesman, S., Skowyra, D., Hoskins, J., McKenney, K., and Maurizi, M.R. (1994)
A molecular chaperone, ClpA, functions like DnaK and DnaJ. Proc Natl Acad Sci USA
91: 12218-12222.
Wickner, S., and Maurizi, M.R. (1999) Here's the hook: similar substrate binding sites in the
chaperone domains of Clp and Lon. Proc Natl Acad Sci USA 96: 8318-8320.
99
Williams, T.L., Jackson, E.L., Carritte, A., and Baker, T.A. (1999) Organization and dynamics of
the Mu transpososome: recombination by communication between two active sites.
Genes Dev 13: 2725-2737.
Yakhnin, A.V., Vinokurov, L.M., Surin, A.K., and Alakhov, Y.B. (1998) Green fluorescent
protein purification by organic extraction. Protein Expr Purif 14: 382-386.
Zolkiewski, M. (1999) ClpB cooperates with DnaK, DnaJ, and GrpE in suppressing protein
aggregation. A novel multi-chaperone system from Escherichia coli. J Biol Chem 274:
28083-28086.
100
CHAPTER THREE
Communication between ClpX and ClpP during
substrate processing and degradation
Submitted: Shilpa A. Joshi, Greg L. Hersch, Tania A. Baker,
Robert T. Sauer, 2004. Nature Structural and Molecular Biology.
Figure 1: work of G.L.H.
Figures 2-6: work of S.A.J.
101
Summary
ClpXP is a compartmental protease in which ring hexamers of the AAA+ ClpX ATPase bind,
denature, and then translocate protein substrates into the degradation chamber of the double-ring
ClpP14 peptidase. A key question is the extent to which functional communication between
ClpX and ClpP occurs and is regulated during substrate processing. Here, we show that ClpXClpP affinity varies with the protein-processing task in which ClpX is engaged and with the
catalytic engagement of the ClpP active sites.
Functional communication between the
symmetry-mismatched ClpX and ClpP rings depends on the ATPase activity of ClpX and
appears to be transmitted through changes in the conformation, flexibility, or accessibility of its
IGF loops which contact ClpP.
The protein unfolding activity of ClpX mutants with
substitutions at an interface between ATPase modules can be rescued by ClpP in a manner that
suggests that energy-dependent denaturation of protein substrates generates tension and
quaternary distortions at these ClpX interfaces.
Communication between ClpX and ClpP
appears to stabilize ClpP binding to substrate-engaged ClpX molecules and is probably required
for coordination of substrate processing and degradation.
102
Introduction
AAA+ ATPases function as essential components of energy-dependent compartmental proteases
in all biological kingdoms1. For example, the 19S portion of the eukaryotic proteasome consists
predominantly of AAA ATPases, which help recognize and translocate substrates to an
associated 20S core protease2. In bacteria, the ClpXP and ClpAP proteases consist of either the
ClpX or ClpA ATPase and the ClpP peptidase, and the HslUV protease consists of the HslU
ATPase and the HslV peptidase1,3,4. In each of these energy-dependent proteases, the active sites
used for polypeptide cleavage are sequestered in a degradation chamber formed by a multisubunit complex with a barrel-like shape5-14. Entry portals, which are too small to admit native
proteins, provide access to this chamber. The ATPases of the compartmentalized proteases from
bacteria form ring hexamers that bind appropriate protein substrates and, in reactions dependent
upon ATP hydrolysis, unfold these molecules and translocate them through a central proteinprocessing pore and into the peptidase chamber, where degradation to small peptides occurs1,3,4.
One key question for all energy-dependent proteases is how interactions between the ATPase
and the peptidase allow coordination of substrate processing and degradation. For HslUV, the
peptidase and ATPase are both six-fold symmetric and structures of the complex in different
nucleotide-bound states have been solved9,11,13,15,16.
These structures show that nucleotide
binding modulates the size of both the protein-processing pore and the entry portal and also
alters the contacts between HslU and HslV, propagating structural changes to the peptidase
active sites. These structural changes undoubtedly regulate the functional communication which
has been observed between the hexameric HslU and HslV rings11,15-20. For ClpXP and ClpAP,
docking of the ATPases and peptidase involves a symmetry mismatch between a hexameric
ATPase ring and a heptameric ClpP ring7,8,21-23. Although high-resolution structures of ClpXP or
ClpAP have not been solved, flexible surface loops in both ClpX and ClpA, which contain a IGF
or IGL motif, have been implicated in ClpP binding24-27.
For both ClpXP and ClpAP,
coordination between the activities of the 6-fold symmetric ATPase and 7-fold symmetric
103
peptidase must occur, as binding of ClpA activates ClpP peptidase activity28, proteins trapped in
the degradation chamber of inactive ClpP can be released in a reaction that depends upon ATP
hydrolysis by ClpX29, and the ATPase activity of ClpX is depressed upon binding to ClpP24.
To probe functional communication between the ClpX and ClpP enzymes of Escherichia coli,
we have used changes in ATPase activity to monitor and quantify the strength of the ClpX-ClpP
interaction. We find that ClpP binds most tightly to ClpX when the ATPase is denaturing
protein substrates, less tightly during translocation of substrates, and least tightly in the absence
of substrates.
ClpX is also able to detect the catalytic status of the ClpP active sites, as
evidenced by a significant increase in affinity when the active-site serines of ClpP are modified.
ClpP binding suppresses the protein-unfolding defects of ClpX variants with mutations at an
intersubunit interface, but rescue occurs at the expense of binding affinity and reverses the
response to substrate processing. This result suggests that subunit-subunit interactions in wildtype ClpX play an important role in the unfolding of protein substrates by preventing quaternary
distortions that prevent substrate denaturation and weaken ClpP binding.
Finally, we
demonstrate linkage between the nucleotide state of ClpX and functional interactions with ClpP.
Results
Substrate design.
Protein substrates had C-terminal ssrA tags to target them to ClpX30.
Unfolding of GFP-ssrA by ClpX or degradation by ClpXP results in loss of fluorescence29,31.
Unlabelled and
35
S-labelled variants of the human titin-I27-ssrA protein were degraded by
ClpXP either as native proteins or as denatured, carboxymethylated (CM) molecules32.
ClpP interaction requires more than two IGF loops. The IGF loop (residues 264-278) of E.
coli ClpX mediates binding to ClpP24,25. We constructed and purified ClpXloopless, a variant in
which the IGF loop was replaced with a short linker. As expected, ClpXloopless did not support
degradation of an ssrA-tagged protein substrate in the presence of ClpP (Fig. 1A), and did not
104
bind His6-tagged ClpP in Ni++-NTA pull-down assays (data not shown).
ClpXloopless was,
however, as active as wild-type ClpX in unfolding GFP-ssrA as monitored by loss of native GFP
fluorescence (Fig. 1B). Moreover, ClpXloopless formed stable ternary complexes with GFP-ssrA
and the delivery protein SspB as assayed by gel filtration in the presence of ATPS (Fig. 1C).
These mutant complexes chromatographed at the same position as wild-type ternary complexes,
suggesting that ClpXloopless, like ClpX, is hexameric under these conditions33. We conclude that
ClpXloopless fails to interact with ClpP but otherwise assembles normally and is fully active in
binding and denaturing ssrA-tagged substrates.
It is not known how many IGF loops are required for functional interactions between ClpP and a
ClpX hexamer, which assembles as a trimer of stable dimers34. An IGF-loop peptide bound
ClpP very weakly (Kd>200 M; data not shown), suggesting that stable ClpP binding requires
interaction with several IGF loops and/or requires the loop to be held in a specific conformation
by ClpX. To test the activity of hexamers containing mixtures of wild-type and loopless dimers,
we added ClpXloopless to wild-type ClpX and assayed degradation of an ssrA-tagged substrate in
the presence of excess ClpP. As shown in Fig. 1D, a modest excess of ClpXloopless inhibited
degradation, with the data fitting well to a model in which hexamers with only one wild-type
dimer have less than 1% activity, those with two wild-type dimers are roughly 60% active, and
mixing of wild-type and mutant dimers is relatively unbiased. We confirmed that wild-type and
loopless dimers do in fact form mixed multimers by showing that untagged ClpXloopless subunits
bound Ni++-NTA only when mixed with His6-tagged wild-type ClpX (data not shown). Thus, we
conclude that functional interactions between ClpX and ClpP require IGF loops in at least two of
the dimers that comprise the ClpX hexamer.
An ATPase assay for ClpX-ClpP affinity.
To determine ClpX-ClpP affinity, we took
advantage of the observation that ClpP binding decreases the rate of ATP hydrolysis by ClpX24.
ATP turnover by 50 nM ClpX6 was assayed in the presence of ClpP14 concentrations ranging
105
from 0 to 2.5 M. The change in observed rates fit well to a simple binding isotherm, whereas
no significant changes were observed for the ClpXloopless control (Fig. 2A). This experiment
confirms the importance of the IGF loop in binding ClpP and indicates that the changes in ATP
hydrolysis observed when ClpP binds to wild-type ClpX are mediated by the same set of
interactions that permit collaboration in protein degradation. Assays performed with 200 nM
ClpX6 gave similar results, with the combined data obtained for the two ClpX concentrations
fitting a model in which ClpX and ClpP interact with an apparent equilibrium dissociation
constant (Kapp) of 126 ± 15 nM (Fig. 2B). It is important to note that this apparent affinity
constant reflects a population-weighted average of the actual Kd values of ClpP for ClpX in each
of its different enzymatic and conformational states as it passes through its cycles of ATP
hydrolysis and coupled conformational changes.
Substrate processing strengthens the ClpX-ClpP interaction. Recent studies using titin-ssrA
substrates with a range of thermodynamic stabilities have shown that ClpX denatures these
molecules at different rates but translocates the denatured proteins to ClpP at the same rate32.
Hence, we reasoned that by assaying ClpX-ClpP affinity during the degradation of these titin
substrates, it should be possible to determine whether ClpX engaged in substrate degradation
binds ClpP in a different manner than in the absence of substrate and, if so, to deconvolute the
contributions of protein denaturation and translocation to this difference. Changes in the ATPase
rate of ClpX were used to monitor ClpP affinity in the presence of titin-ssrA substrates that were
roughly 10-fold above the Km for denaturation to ensure approximately 90% saturation of the
enzyme by substrate. When ClpP was titrated against ClpX in the presence of the most stable
native substrate, wild-type titin-ssrA, Kapp was 38 ± 4 nM, a decrease of roughly three-fold
compared to the absence of substrate (Figs. 2A & 3A). In the presence of less stable native
variants, Kapp values were 54 ± 6 nM (V15P) and 84 ± 14 nM (V13P). When a denatured titin
substrate (CM-V13P) was present, Kapp was 105 ± 8 nM. With other denatured substrates (CMtitin; CM-V15P), Kapp values were within error of the value determined for CM-V13P (data not
106
Figure 1. Properties of ClpXloopless. (A) ClpX6loopless (100 nM) did not support
degradation of CM-titin-V13P-ssrA (1 mM) in the presence of ClpP14 (300 nM), whereas
wild-type ClpX mediated efficient degradation under these conditions. (B) GFP-ssrA
(1 mM) in the presence of SspB2 (1 mM) was unfolded at essentially the same rate by
ClpX6 or ClpX6loopless (300 nM) as assayed by loss of native GFP-ssrA fluorescence.
(C) ClpXloopless forms stable ternary complexes. ClpX6 or ClpX6loopless (6 mM),
SspB2 (3 mM), GFP-ssrA (6 mM), and ATPgS (5 mM) were chromatographed on a
Superdex 200 column. The elution position of GFP-ssrA was monitored by A500.
Positions of the ternary complex (X6S2G2), binary complex of SspB and GFP-ssrA
(S2G2), and free GFP-ssrA (G) are shown. (D) Degradation of titin-V4A-ssrA (5 mM)
by wild-type ClpXP (100 nM ClpX6; 2 mM ClpP14) was inhibited by addition of
ClpXloopless. The solid line is a fit to eq. 2 (see Methods) with a bias factor of 0.99, an
A21 value of 0.63, and an A12 value of 0.003.
native GFP-ssrA (mM)
B
ClpXloopless/ClpP
0.8
0.6
ClpXP
0.4
0.2
0.0
0
5
1.0
10
15
time (min)
0.9
ClpX
ClpXloopless
0.7
0
50
100
time (sec)
S2G2
G
20
1
D
unfolding
0.8
ClpX
ClpXloopless
absorbance (500 nm)
1.0
X6S2G2
C
degradation
150
fraction initial activity
substrate remaining (mM)
A
1.2
1.4
volume (mL)
1.6
1.0
0.8
0.6
0.4
0.2
0.0
0
*Work of Greg L. Hersch
4
8
12
loopless
[ClpX
]/[ClpX]
108
Figure 2. An assay for ClpX-ClpP interaction in solution. (A) Changes in the ATP
hydrolysis rate of 50 nM ClpX6 or ClpX6loopless as a function of ClpP concentration.
(B) Percent inhibition of ATP turnover as a function of ClpP concentration. The filled
symbols are for experiments performed with 200 nM ClpX6 and the open symbols for
50 nM ClpX6. The fitted line is that expected for a one-to-one binding reaction with
an apparent affinity of 126 ± 15 nM.
A
300
ATP turnover
(min-1[ClpX6]-1)
ClpXloopless
250
200
ClpX
150
0.01
B
0.1
1
[ClpP14] (mM)
% inhibition
ATP turnover
40
20
0
0
0.5
1.0
[ClpP14] (mM)
1.5
110
Figure 3. ClpX-ClpP affinity changes in an ATPase-dependent fashion during
substrate denaturation and translocation. (A) Binding of ClpP to 50 nM ClpX6
in the presence of 15 mM concentrations of four variants of titin-ssrA. The fits
represent Kapp values of 38 ± 4 nM for wild-type titin-I27-ssrA, 54 ± 6 nM for the
native V15P mutant, 84 ± 14 nM for the native V13P variant, and 105 ± 8 nM for the
denatured CM-V13P protein. (B) Kapp values for the ClpX-ClpP interaction in the
presence of titin-ssrA substrates vary linearly with the rate constant for ClpXP degradation
of these substrates32. The line is a fit to eq. 1 (see Methods), with affinities of 38 and
99 nM, respectively, when ClpXP is denaturing or translocating titin-ssrA substrates.
(C) Kapp for the ClpX-ClpP interaction correlates linearly with the ClpXP ATPase rate
(R = 0.99) and the ClpX ATPase rate (R = 0.97) during the processing of different titin-ssrA
substrates.
A
B 120
500
400
100
Kapp (nM)
ATP turnover
(min-1[ClpX6]-1)
CM-V13P
V13P
V15P
300
80
V13P
60
V15P
titin-ssrA
200
0.01
40
0.1
[ClpP14] (mM)
1
120
C
CM-V13P
ClpXP
100
Kapp (nM)
CM-V13P
V13P
80
ClpX
60
V15P
40
titin-ssrA
200
300
400
500
ATP turnover
(min-1[ClpX6]-1)
titin-ssrA
1
2
3
4
kdeg (min-1)
5
112
shown).
These results show that the apparent affinity for ClpP is stronger when ClpX is
processing protein substrates, with the strongest binding observed for native substrates that are
denatured most slowly.
Kapp for ClpX-ClpP binding in the presence of different titin-ssrA substrates showed a linear
correlation (R = 0.99) with the rate constant for degradation of these proteins (Fig. 3B). This
relationship is expected if ClpX has one affinity for ClpP while denaturing titin substrates and a
different affinity while translocating these substrates (see Methods). Indeed, the linear fit to the
data in Fig. 3B represents a model in which Kapp is 38 nM when ClpX is denaturing a titin-ssrA
substrate and 99 nM when ClpX is translocating these substrates. These results show that the
interaction between ClpX and ClpP is stronger during substrate denaturation than during
translocation and demonstrate the existence of a molecular mechanism by which ClpP is able to
"sense" the protein-processing task in which ClpX is engaged.
The apparent ClpX-ClpP
affinities were also well correlated with the overall rates of ATP turnover by ClpXP and ClpX in
the presence of these titin-ssrA substrates (Fig. 3C). Based on this observation, we suggest that
changes in the ATPase rate of ClpX during substrate processing may be an important factor in
determining the apparent affinity for ClpP.
Active-site communication between ClpP and ClpX. If ClpP can sense whether ClpX is
processing protein substrates, then ClpX may be able to detect whether ClpP is degrading
substrates. To address this possibility, we used ClpPDFP, a variant in which the active-site serines
(S97) were covalently modified by reaction with di-isopropyl-fluorophosphate29.
Although
ClpPDFP is inactive in protein degradation, the chemical modification mimics the acyl-enzyme
intermediate in the normal peptide-bond hydrolysis reaction. Moreover, an oxygen of DFP binds
in the oxyanion hole of the ClpP active site, resembling the carbonyl oxygen of a bound peptide
substrate7. ClpPDFP bound ClpX so strongly that the bound and total concentrations of ClpX
were essentially the same (Fig. 4). Under these “stoichiometric binding” conditions it was not
113
possible to determine an affinity constant but this value must be less than 5 nM. Hence, ClpX
appears to be capable of sensing whether the ClpP active sites are engaged with substrate. To
ensure that the tighter ClpX binding observed for ClpPDFP was caused by modification of the
active-site S97, we also assayed binding of a ClpPS97A mutant after treatment with DFP35. DFPtreated ClpPS97A bound ClpX with roughly the same affinity as wild-type ClpP and much more
weakly than ClpPDFP (Fig. 4). We conclude that the strong ClpX binding observed for ClpPDFP
results directly from acylation of the active-site serines and/or from concomitant substrate-like
interactions of the covalent modification.
ClpP rescues the unfolding defects of ClpX mutants. In a previous study, we showed that
mutations in the C-terminal portion of the ClpX sensor-II helix, which forms part of the interface
between the C-domain of one subunit and the ATPase domain of an adjacent subunit, caused
defects in substrate unfolding but not in substrate binding36. Because ClpP binds more tightly
when ClpX is denaturing substrates, we reasoned that ClpP binding might suppress the unfolding
defects of the sensor-II mutants. This result was observed. By themselves, the L381K and
D382K mutants of ClpX had undetectable activities in unfolding GFP-ssrA (Fig. 5A). In the
presence of ClpPDFP, however, ClpXL381K and ClpXD382K catalyzed efficient unfolding of GFPssrA (Fig. 5B). Unmodified ClpP also suppressed the unfolding defects of the sensor-II mutants,
albeit less efficiently than ClpPDFP (data not shown).
Because ClpPDFP suppression of the
unfolding defect of the D382K mutant was most efficient, we selected this ClpX variant for more
detailed studies.
In the absence of protein substrate, ClpP bound ClpXD382K with an apparent affinity roughly twofold weaker (230 ± 60 nM; Fig. 5C) than for wild-type ClpX. In the presence of protein
substrates, however, the observed affinity of ClpP for ClpXD382K was reduced even more. For
example, Kapp was almost 10-fold weaker (2.4 ± 0.5 M) in the presence of the V13P variant of
titin-ssrA and about 5-fold weaker (1.2 ± 0.4 M) in the presence of denatured CM-titin-ssrA
114
Figure 4. Modification of the ClpP active sites strengthens ClpX binding.
ClpPDFP bound tightly to 50 nM ClpX6 as assayed by changes in ATP hydrolysis.
ClpP14 can bind two ClpX hexamers and, at half-maximal binding, the total
concentration of ClpP14DFP was roughly 15 nM and the concentration of bound
ClpX6 was 25 nM. Although an accurate Kapp cannot be determined from these
data, the upper limit for this constant is ª 5 nM. Kapp for the interaction of ClpX
with DFP-treated ClpPS97A (130 nM) was similar to that for wild-type ClpP,
confirming that the tight binding of ClpPDFP results from modification of the
active-site S97 side chain.
% max. inhibition
100
ClpX-ClpPDFP
75
50
ClpX-ClpPS97A-DFP
25
0
0
100
200
[total ClpP14] (nM)
300
116
Figure 5. ClpP rescues the unfolding defects of ClpX mutants. The sensor-II
helix mutants, ClpXD382K and ClpXL381K, fail to unfold GFP-ssrA (310 nM)
by themselves (panel A) but unfold this substrate efficiently in the presence of
800 nM ClpP14DFP (panel B). In these experiments, the concentrations of mutant
or wild-type ClpX6 were 260 nM, and 460 nM SspB was present. ClpP binds
200 nM ClpX6D382K with an apparent affinity of 230 ± 60 nM in the absence of
substrate (panel C) but binds more weakly in the presence of a denatured substrate
(13.1 mM CM-titin-ssrA; Kapp = 1.2 ± 0.3 mM) or a native substrate (15 mM
titin-V13P-ssrA; Kapp = 2.4 ± 0.5 mM) (panel D).
A
C
100
ClpXL381K
ClpXD382K
75
% folded GFP-ssrA
ClpX
25
B
0
5
100
10
15
+ClpPDFP
ATP turnover (min-1[ClpX6]-1)
100
50
0
ClpXD382K
110
90
D
No substrate
0
0.5
50
ClpXL381K
2.5
ClpXD382K
CM-titin
200
75
1.5
V13P
160
ClpXD382K
25
ClpX
0
0
5
10
time (min)
15
120
0
2.5
7.5
[ClpP14] (mM)
118
(Fig. 5D).
Thus, whereas the affinity of ClpP for wild-type ClpX strengthens during the
processing of protein substrates, the opposite is true for the D382K mutant.
When ClpXD382K engages a protein substrate, it appears to assume a conformation poorly suited
both for binding ClpP and substrate unfolding. Although ClpP can stabilize ClpXD382K in a
conformation active in substrate processing, binding energy must be used to drive the
accompanying conformational change and thus the observed overall affinity for ClpP is
weakened significantly. Interestingly, addition of ClpP to ClpXD382K increased rates of ATP
hydrolysis in the presence of protein substrates, whereas ATP turnover decreased when ClpP was
added in the absence of protein substrates (Figs. 5C and 5D). This pattern is different from that
observed for the wild-type interaction, where ClpP binding always reduced ClpX ATPase rates
with or without substrate (Figs. 2A & 3A).
Nucleotide-state of ClpX controls ClpP binding. The experiments presented above suggest
that the ATPase activity of ClpX is linked, in some fashion, to interactions with ClpP. In pulldown assays, untagged ClpP bound to Ni++-NTA in the presence of His6-tagged ClpX and
ATPS but did not bind well when ATP or ADP were present (Fig. 6A). ATP is hydrolyzed
rapidly by ClpX, raising the possibility that ADP•ClpX may predominate even with excess ATP
because ADP release is slow. By contrast, ATPS is hydrolyzed slowly by ClpX37, and thus
ATPS•ClpX may be a better mimic of the ATP state.
In the crystal structure of H. pylori ClpX, Arg396 in the N-terminal portion of the sensor-II helix
is positioned to contact bound nucleotide27. This residue is conserved in the ClpX and HslU
families, suggesting an important functional role. We constructed and purified a mutant of E.
coli ClpX in which the corresponding residue, Arg370, was mutated to lysine. ClpXR370K failed
to hydrolyze ATP (Fig. 6B) but bound fluorescent nucleotide analogs with Kd values of 39 M
for mant-ATP and 18 M for mant-ADP (Fig. 6C). These nucleotide affinities were similar to
119
those for wild-type ClpX37 (52 M and 8 M, respectively; data not shown). Hence, the major
defect of ClpXR370K appears to be in ATP hydrolysis rather than ATP binding. Irrespective of the
nucleotide present, addition of His6-tagged ClpXR370K did not result in detectable binding of
untagged ClpP to Ni++-NTA (Fig. 6A). This result further emphasizes that some aspect of the
ATP binding/hydrolysis cycle or linked conformational changes plays a role in modulating the
binding of ClpX to ClpP.
ClpXR370K did not unfold GFP-ssrA or degrade this substrate when ClpP was present (data not
shown). To determine whether hexamers with a mixture of wild-type and R370K dimers were
active, we titrated increasing quantities of the ClpXR370K mutant against a fixed quantity of wildtype ClpX and excess ClpP and assayed degradation of GFP-ssrA. As shown in Fig. 6D,
addition of the mutant in 12-fold excess caused nearly complete inhibition. We also observed
inhibition by ClpXR370K of ClpXP degradation of denatured CM-titin-ssrA and of ClpX
unfolding of GFP-ssrA (data not shown). Hence, mixed hexamers with one wild-type dimer and
two ClpXR370K dimers must be defective in denaturing native substrates and be unable to engage
or translocate denatured substrates to ClpP. The latter defect could reflect an inability to bind
ClpP. The best fit of the Fig. 6D inhibition data was obtained from a model in which the 2:1 and
1:2 mixed hexamers were both inactive, and wild-type dimers had a 3-fold preference for
assembling with themselves rather than with mutant dimers. Untagged ClpP bound Ni++-NTA
resin following incubation with His6-ClpXR370K, untagged wild-type ClpX, and ATPS (Fig. 6D
inset). This result demonstrates that ClpXR370K and wild-type subunits co-assemble and show
that at least one species of mixed hexamer can bind ClpP.
120
Figure 6. ATP binding or hydrolysis is required for productive ClpX-ClpP interactions.
(A) Untagged ClpP bound Ni++-NTA in the presence of His6-ClpX and ATPgS; weak
binding was observed with ATP or ADP in some experiments but is not visible here. No
binding of ClpP to His6-ClpXR370K was detected with any nucleotide. (B) ClpX6R370K
(400 nM) hydrolyzed ATP at less than 1% of the rate of wild-type ClpX6 (400 nM) as
determined in a coupled spectrophotometric assay41. (C) Binding of ClpXR370K to 1 mM
mant-ADP or mant-ATP assayed by changes in fluorescence in the absence of magnesium.
The fitted lines are for Kd’s of 39 mM (mant-ATP) and 18 mM (mant-ADP). (D) Degradation
of GFP-ssrA (7 mM) by wild-type ClpXP (100 nM ClpX6; 2.7 mM ClpP14) was inhibited by
addition of the ClpXR370K mutant. The solid line is a fit to eq. 3 (see Methods) with a bias
factor of 0.33. Wild-type and mutant ClpX were preincubated for 5 min at 30 ∞C, ClpP and
ATP mix I were added, and GFP-ssrA was added two min later to start the reaction. Inset—
untagged ClpP binds Ni++-NTA in the presence of a mixture of His6-ClpXR370K and untagged
wild-type ClpX, but much less in the presence of either single species. All pull-down reactions
contained ATPgS.
B
A
His6-ClpXR370K
His6-ClpX
ATPgS ATP ADP ATPgS ATP ADP
ClpX
ClpXR370K
ATP remaining (nmol)
]
]
60
40
ClpP
ClpX
20
3.0
F445 (x105)
mant-ADP
2.5
fraction initial activity
D
C
1
0.8
mant-ATP
His6ClpXR370K +
+
-
+
-
+
ClpX
0.6
2.0
1.5
100 time (sec) 300
ClpP
0.4
0.2
0
20
40
60
[ClpXR370K] (mM monomer)
0.0
0
4
8
12
[ClpXR370K]/[ClpX]
122
Discussion
The studies presented here provide strong evidence for functional communication between ClpX
and ClpP during the processing and degradation of protein substrates. For example, ClpP
affinity changed as a function of substrate processing by ClpX. In the absence of protein
substrates, Kapp for the ClpX-ClpP interaction was 126 nM. This value dropped to roughly 100
nM during translocation of denatured substrates and to approximately 40 nM when ClpX was
denaturing native substrates. These affinity changes could potentially affect ClpXP function in
two ways. First, the processivity of degradation would almost certainly be reduced if complexes
dissociated in the midst of degradation of a single substrate molecule. We do not believe that
this is a significant problem, however, as the peptide products of degradation were found to be
identical for degradation of a titin-ssrA substrate by ClpXP or ClpXD382K/ClpP (data not shown),
despite the fact that Kapp was more than 20-fold higher for the mutant complex. Second, the total
ClpX6 concentration in E. coli is estimated to be roughly 100 hexamers, which would correspond
to an intracellular concentration of 0.3 M. This is only about twice Kapp for the ClpX-ClpP
interaction in the absence of substrate, and thus substrate binding to free ClpX hexamers might
help drive assembly of ClpXP complexes by increasing the affinity of this interaction. Because
ClpP in the cell can associate with either ClpX or ClpA, this could provide a mechanism to
distribute the peptidase based on whether substrates for one ATPase or the other were most
prevalent.
Irrespective of the functional consequences, the observed changes in ClpX-ClpP affinity
demonstrate the existence of structural changes in ClpX-ClpP interactions that depend upon the
physical state of the engaged substrate. Like other molecular machines, the conformation of
123
ClpX must change during the ATPase cycle. Hence, alterations in the amount of time spent in
different stages of this cycle as a function of substrate processing could modulate ClpX-ClpP
affinity. In support of this model, apparent ClpX-ClpP affinities were well correlated with the
overall rates of ATP turnover by ClpXP and ClpX in the presence of substrates (Fig. 3C). It is
possible, of course, that ClpX assumes distinct conformations when it is denaturing or
translocating protein substrates and/or that the substrates themselves also contribute in some
direct fashion to changes in affinity. During translocation, for example, the denatured protein
substrate must be in loose contact with the protein-processing pore of ClpX and the entry portal
of ClpP.
Indeed, all of these factors–ATPase rate, distinct conformations, and substrate
interactions–may contribute to the apparent ClpX-ClpP affinity to varying degrees.
The IGF loop of E. coli ClpX appears to be the major determinant of ClpP binding24,25 and
homologous peptide motifs are found in all of the AAA+ ATPases that collaborate with ClpP
family members in protein degradation. In the crystal structure of H. pylori ClpX, a homologous
LGF tripeptide sits at the tip of a straight but flexible surface loop that extends away from the
protease-proximal surface of ClpX27. Alignment of the symmetry axes of the mismatched ClpX
and ClpP rings positions these tripeptides from the ClpX hexamer near hydrophobic clefts on the
surface of a heptameric ClpP ring8,24,27. In each ClpX subunit, the IGF loop is preceded by a
short -helix that connects directly to the sensor-I portion of the ATP/ADP binding site. In
ClpA, this loop is disordered26 but is also connected to the sensor-I portion of an active site for
ATP hydrolysis. It is possible that the length and/or flexibility of the IGF loop changes as a
function of nucleotide binding or hydrolysis, thereby mediating communication with ClpP.
Alternatively, changes in ClpX hexamer conformation might alter the number of loops that
contact ClpP. In this regard, it is interesting that both ClpX and ClpA crystallize not as ring
hexamers but in "lock washer" conformations in which subunits are related by a screw axis26,27.
124
In a lock-washer conformation, fewer IGF loops would be capable of contacting the relatively
flat surface of a ClpP ring because of axial displacements.
The sensor-II helix in AAA+ ATPases provides a communication link between the nucleotide
binding sites of adjacent subunits. In E. coli ClpX, Arg370 is near the N-terminus of the sensorII helix, where it should be close to bound nucleotide27. Our results show that this side chain
plays an essential role in ATP hydrolysis and also reveal that ClpX hexamers with two R370K
dimers and one wild-type dimer are inactive in unfolding GFP-ssrA and in mediating ClpP
degradation of unfolded titin-ssrA substrates. The unfolding defect of these mixed hexamers
suggests that the ATPase activities of different ClpX subunits must be coordinated in some
fashion to drive protein unfolding. Wild-type ClpX is also defective in unfolding GFP-ssrA
when ATPS is substituted for ATP37. Because ATPS is hydrolyzed slowly by ClpX, some
type of coordination of NTP hydrolysis rates by different ClpX subunits appears to be required
for productive unfolding of native substrates.
Mutations at the C-terminal end of the sensor-II helix disrupt subunit-subunit packing in the
hexamer and result in ClpX enzymes that bind but fail to unfold ssrA-tagged substrates36. ClpP
binding suppresses the substrate processing defects of several of these sensor-II mutants,
including ClpXD382K. ClpP binding to ClpXD382K is about 30-fold weaker than to wild type in the
presence of native ssrA-tagged substrates, but only 2-fold weaker in the absence of substrate.
These results suggest that when ClpXD382K binds to and attempts to unfold a native ssrA-tagged
substrate, it becomes trapped in a conformation, perhaps similar to the lock-washer state, that is
inactive for protein unfolding and binds ClpP poorly. Hence, maintenance of proper subunitsubunit contacts within ClpX appear to be essential for substrate unfolding and for strong ClpP
interactions.
We propose that subunit-subunit contacts mediated by D382 and surrounding
residues in wild-type E. coli ClpX resist tension that is generated when the enzyme applies an
unfolding force to a native substrate. Such tension would be a natural consequence of the
125
unfolding force creating an equal and opposite force.
If this tension results in quaternary
distortions of the D382K mutant, then binding of ClpP could stabilize the active ATPase
conformation, allowing it to resist distortion and thus to use the energy of ATP hydrolysis for
productive conformational changes that drive unfolding.
ClpX-ClpP affinity becomes at least 10-fold stronger when the active-site serines of ClpP are
acylated by reaction with DFP, revealing the existence of mechanisms that allow the peptidase
active sites to communicate with ClpX. Because the DFP-modified residues are located within
the degradation chamber of ClpP7, they cannot affect the ClpX-ClpP interaction directly.
Instead, modification of the sequestered ClpP active sites must result in a conformational change
that is propagated to the external surface of ClpP, where the number or quality of contacts with
ClpX could be influenced. Despite the fact that the ClpP and HslV peptidases have unrelated
structures and completely different active-site architectures, it is notable that ClpXP and HslUV
both have mechanisms that allow communication between the ATPase and the peptidase active
sites.
This functional conservation in the absence of structural conservation supports an
important role for communication in the degradation of protein substrates by these energydependent proteases.
Other than the affinity changes discussed above, what role could be served by communication
between ClpX and ClpP or HslU and HslV? One possibility is that translocation of denatured
substrates from the ATPase to the peptidase requires coordinated changes in the diameters of the
ATPase processing pore and the peptidase entry portal. This could allow efficient transfer
during the power stroke associated with each cycle of ATP hydrolysis and prevent slippage or
dissociation during the recovery phase. It is also possible that coordination is required to allow
efficient release of cleaved peptides from the degradation chamber at the same time that
uncleaved polypeptide chains are entering the chamber. Finally, when ClpX hexamers are
docked with both peptidase rings of ClpP, translocation appears to occur exclusively from one
126
ClpX hexamer rather than simultaneously from both hexamers38. Communication between ClpX
and ClpP would obviously be critical for regulating substrate traffic under these circumstances.
Methods
Solutions. Buffer A contains 43 mM Hepes-KOH (pH 7.6), 8.5 mM Tris-HCl (pH 8.0), 142 mM
KCl, 15% glycerol, 1.1 mM DTT, 5.4 mM MgCl2, 420 M EDTA, 36 M ZnSO4, 36 M ATP,
0.032% NP-40, and 0.004% Triton X-100. Buffer B contains 35 mM Hepes-KOH (pH 7.6), 4.4
mM Tris-HCl (pH 7.6), 1.5 mM Tris-HCl (pH 8.0), 95 mM KCl, 14% glycerol, 660 M DTT,
7.4 mM MgCl2, 85 M EDTA, 19 M ZnSO4, 19 M ATP, 0.16% NP-40, and 0.002% Triton
X-100. Buffer L contains 50 mM Tris-HCl (pH 7.6), 10% glycerol, 1 mM DTT, and 0.5 mM
EDTA. Buffer M contains 47 mM Tris-HCl (pH 7.6), 284 mM KCl, 9.5% glycerol, 4.7 mM
DTT, 95 M MgCl2, and 10 mM EDTA. Buffer N contains 50 mM sodium phosphate (pH 8.0),
300 mM NaCl, and 250 mM imidazole. Buffer S contains 50 mM Tris-HCl (pH 8.0), 300 mM
KCl, 10% glycerol, 3 mM DTT, and 10 mM MgCl2. ATP mix I contains 5 mM ATP, 16 mM
creatine phosphate, and 0.32 mg ml-1 creatine phosphokinase. ATP mix III contains 2.5 mM
ATP, 1 mM NADH, 7.5 mM phosphoenolpyruvate, 0.05 mg ml-1 pyruvate kinase, and 0.025 mg
mL-1 lactate dehydrogenase.
Strains, Plasmids, and Proteins. E. coli strain CF150 is an X90 derivative, in which the
adjacent clpP, clpX, and lon genes are replaced by a gene for chloramphenicol acetyltransferase,
and was a gift from Chris Farrell (MIT). A plasmid expressing His6-tagged R370K (pET-14bClpXR370K) was produced using overlap extension mutagenesis as described36.
A plasmid
expressing ClpXloopless (pGH003) was constructed from pET-3a-ClpX39 by polymerase chain
reaction. In ClpXloopless, a GSGSG sequence replaces wild-type residues 264-278. The sequence
of
the
synthetic
peptide
containing
the
KKGRYTGSGIGFGATVKAK-CONH2.
127
ClpX
IGF
loop
was
fluorescein-NH-
ClpXloopless and wild-type ClpX were purified as described37. GFP-ssrA and His6-tagged variants
of ClpP, ClpX, and titin-I27-ssrA were purified as described29,32,36.
35
S-titin-ssrA variants were
gifts from Jon Kenniston (MIT), and SspB was a gift from David Wah (MIT). His6-tagged
ClpPS97A was purified from E. coli strain CF150 containing pYK16235 using a published
protocol29 with modifications. After Mono Q chromatography, fractions containing ClpPS97A
were applied to a HiPrep 16/60 Sephacryl S-300HR column (Amersham Biosciences)
equilibrated in buffer S. Fractions containing purified protein were pooled, divided into small
aliquots, and stored at -80 C. Carboxymethylation of titin-ssrA variants and acylation of ClpP
variants with DFP were performed as described29,32.
Chemically modified proteins were
dialyzed extensively prior to use.
E. coli ClpP was purified based on a published protocol40 with modifications. Cells were
resuspended in 3 mL of buffer L plus 150 mM KCl for each gram of cells, lysed by French press,
and centrifuged at 15,000 rpm in a SA-600 rotor for 60 min. The supernatant was passed
through a 0.2 m filter, ammonium sulfate was added to 30% saturation, and the supernatant
containing ClpP was retained after centrifugation.
Ammonium sulfate was added to this
supernatant to 60% saturation, and the pellet containing ClpP was recovered by centrifugation.
The pellet was resuspended, desalted into buffer L plus 150 mM KCl using a PD-10 column
(Amersham Biosciences), and loaded onto a HiLoad 16/10 Q Sepharose HP column (Amersham
Biosciences) equilibrated in buffer L with 150 mM KCl. The column was developed with a 200
mL linear gradient from 150 mM to 400 mM KCl in buffer L. Fractions containing ClpP were
concentrated by ammonium sulfate precipitation (60% saturation), and the pellet was
resuspended, desalted into buffer L plus 100 mM KCl, and loaded onto a HiPrep 16/60
Sephacryl S-300HR column equilibrated in this buffer. Fractions containing purified ClpP were
pooled, concentrated by chromatography on a HiLoad 16/10 Q Sepharose HP column, and stored
in small aliquots at -80 C.
128
Nucleotide hydrolysis and binding assays. Hydrolysis of ATP by ClpX in buffer A was
measured at 30 C using a coupled assay24. ClpX and ClpP were incubated for two min before
addition of substrate and/or ATP mix III. Except where noted, 50 nM ClpX6 was used for all
assays. Binding of mant-ADP or mant-ATP to ClpXR370K was assayed at 4 C in buffer M37.
Unfolding and degradation assays. GFP-ssrA unfolding or degradation29 was performed in
buffer B plus ATP mix I at 30 C. Degradation of
35
S-titin-ssrA substrates was assayed by
release of TCA-soluble peptides as described30,32. For ClpXP degradation of titin-ssrA substrates
at saturating concentrations, the slow steps are denaturation (kden) and translocation (ktrans), with
1/kdeg = 1/kden + 1/ktrans and deg = den + trans32. Kapp for the ClpX-ClpP interaction during
substrate processing can be expressed as (den/deg)•Kappden + (trans/deg)•Kapptrans. The value of
ktrans for different titin-ssrA substrates is essentially constant (4.3 min-1). Substitution of deg –
trans for den and rearrangement of terms, yields the linear equation Kapp = trans•(Kapptrans –
Kappden)•kdeg + Kappden (eq. 1). The general equation for inhibition by mixed hexamer formation
between active and inactive dimers of ClpX is A = (1 + 3•A21•B•R + 3•A12•B2•R2)/(1 + B•R)2
(eq. 2) where A is the fractional activity relative to fully active hexamers, A21 is the fractional
activity of a hexamer with two active and one inactive dimer, A12 is the activity of a hexamer
with one active dimer and two inactive dimers, R is the ratio of total inactive to total active
subunits, and B is the mixing bias. B=1 indicates unbiased mixing of active and inactive dimers;
B<1 indicates a preference of active dimers to associate with other active dimers rather than
inactive dimers. If three active dimers are required for activity, eq. 2 simplifies to A = (1 + B•R)2
(eq. 3).
Pull-down and ternary complex assays.
ClpXP pull-down assays were performed by a
modification of a published protocol24. Each reaction (30 L) contained 300 nM His6-tagged
ClpX6 (wild-type or R370K) and 300 nM untagged ClpP14. Some reactions also contained 300
129
nM untagged wild-type ClpX6. Nucleotide (ATPS, ATP, or ADP) at a concentration of 3 mM
was present during initial complex formation, and in washes after binding to Ni++-NTA agarose.
Protein was eluted in 30 L buffer N and subjected to SDS-PAGE. Gels were stained with
Sypro Orange (Molecular Probes) and visualized using a FluorImager 595 (Molecular
Dynamics). Assays for ternary complexes of ClpX, SspB, and GFP-ssrA were performed by gelfiltration chromatography on a Superdex 200 column (Amersham Biosciences) as described33.
130
Acknowledgements
We thank Scott Boyd, Randy Burton, Elizabeth Courtenay, Chris Farrell, Julia Flynn, Jon
Kenniston, Igor Levchenko, Samia Siddiqui, and David Wah for discussion and materials, and
Bob Horvitz and Jon King for use of equipment. This work was supported by grants from the
NIH and the Howard Hughes Medical Institute. T.A.B. is an employee of HHMI.
131
References
1.
Ogura, T. & Wilkinson, A.J. AAA+ superfamily ATPases: common structure--diverse
function. Genes Cells 6, 575-97 (2001).
2.
Glickman, M.H. et al. A subcomplex of the proteasome regulatory particle required for
ubiquitin-conjugate degradation and related to the COP9-signalosome and eIF3. Cell 94,
615-23 (1998).
3.
Gottesman, S., Wickner, S. & Maurizi, M.R. Protein quality control: triage by chaperones
and proteases. Genes Dev 11, 815-23 (1997).
4.
Gottesman, S., Maurizi, M.R. & Wickner, S. Regulatory subunits of energy-dependent
proteases. Cell 91, 435-8 (1997).
5.
Bochtler, M., Ditzel, L., Groll, M. & Huber, R. Crystal structure of heat shock locus V
(HslV) from Escherichia coli. Proc Natl Acad Sci USA 94, 6070-4 (1997).
6.
Groll, M. et al. Structure of 20S proteasome from yeast at 2.4 Å resolution. Nature 386,
463-71 (1997).
7.
Wang, J., Hartling, J.A. & Flanagan, J.M. The structure of ClpP at 2.3 Å resolution
suggests a model for ATP-dependent proteolysis. Cell 91, 447-56 (1997).
8.
Wang, J., Hartling, J.A. & Flanagan, J.M. Crystal structure determination of Escherichia
coli ClpP starting from an EM-derived mask. J Struct Biol 124, 151-63 (1998).
9.
Bochtler, M. et al. The structures of HsIU and the ATP-dependent protease HsIU-HsIV.
Nature 403, 800-5 (2000).
10.
Groll, M. et al. A gated channel into the proteasome core particle. Nat Struct Biol 7,
1062-7 (2000).
132
11.
Sousa, M.C. et al. Crystal and solution structures of an HslUV protease-chaperone
complex. Cell 103, 633-43 (2000).
12.
Whitby, F.G. et al. Structural basis for the activation of 20S proteasomes by 11S
regulators. Nature 408, 115-20 (2000).
13.
Wang, J. et al. Crystal structures of the HslVU peptidase-ATPase complex reveal an
ATP-dependent proteolysis mechanism. Structure (Camb) 9, 177-84 (2001).
14.
Sousa, M.C. & McKay, D.B. Structure of Haemophilus influenzae HslV protein at 1.9 Å
resolution, revealing a cation-binding site near the catalytic site. Acta Crystallogr D Biol
Crystallogr 57, 1950-4 (2001).
15.
Wang, J. et al. Nucleotide-dependent conformational changes in a protease-associated
ATPase HsIU. Structure (Camb) 9, 1107-16 (2001).
16.
Sousa, M.C., Kessler, B.M., Overkleeft, H.S. & McKay, D.B. Crystal structure of HslUV
complexed with a vinyl sulfone inhibitor: corroboration of a proposed mechanism of
allosteric activation of HslV by HslU. J Mol Biol 318, 779-85 (2002).
17.
Yoo, S.J. et al. Purification and characterization of the heat shock proteins HslV and
HslU that form a new ATP-dependent protease in Escherichia coli. J Biol Chem 271,
14035-40 (1996).
18.
Seol, J.H. et al. The heat-shock protein HslVU from Escherichia coli is a proteinactivated ATPase as well as an ATP-dependent proteinase. Eur J Biochem 247, 1143-50
(1997).
19.
Ramachandran, R., Hartmann, C., Song, H.K., Huber, R. & Bochtler, M. Functional
interactions of HslV (ClpQ) with the ATPase HslU (ClpY). Proc Natl Acad Sci USA 99,
7396-401 (2002).
133
20.
Seong, I.S. et al. The C-terminal tails of HslU ATPase act as a molecular switch for
activation of HslV peptidase. J Biol Chem 277, 25976-82 (2002).
21.
Grimaud, R., Kessel, M., Beuron, F., Steven, A.C. & Maurizi, M.R. Enzymatic and
structural similarities between the Escherichia coli ATP-dependent proteases, ClpXP and
ClpAP. J Biol Chem 273, 12476-81 (1998).
22.
Beuron, F. et al. At sixes and sevens: characterization of the symmetry mismatch of the
ClpAP chaperone-assisted protease. J Struct Biol 123, 248-59 (1998).
23.
Ortega, J., Singh, S.K., Ishikawa, T., Maurizi, M.R. & Steven, A.C. Visualization of
substrate binding and translocation by the ATP-dependent protease, ClpXP. Mol Cell 6,
1515-21 (2000).
24.
Kim, Y.I. et al. Molecular determinants of complex formation between Clp/Hsp100
ATPases and the ClpP peptidase. Nat Struct Biol 8, 230-3 (2001).
25.
Singh, S.K. et al. Functional domains of the ClpA and ClpX molecular chaperones
identified by limited proteolysis and deletion analysis. J Biol Chem 276, 29420-9 (2001).
26.
Guo, F., Maurizi, M.R., Esser, L. & Xia, D. Crystal structure of ClpA, an Hsp100
chaperone and regulator of ClpAP protease. J Biol Chem 277, 46743-52 (2002).
27.
Kim, D.Y. & Kim, K.K. Crystal structure of ClpX molecular chaperone from
Helicobacter pylori. J Biol Chem 278, 50664-70 (2003).
28.
Thompson, M.W., Singh, S.K. & Maurizi, M.R. Processive degradation of proteins by the
ATP-dependent Clp protease from Escherichia coli. Requirement for the multiple array
of active sites in ClpP but not ATP hydrolysis. J Biol Chem 269, 18209-15 (1994).
134
29.
Kim, Y.I., Burton, R.E., Burton, B.M., Sauer, R.T. & Baker, T.A. Dynamics of substrate
denaturation and translocation by the ClpXP degradation machine. Mol Cell 5, 639-48
(2000).
30.
Gottesman, S., Roche, E., Zhou, Y. & Sauer, R.T. The ClpXP and ClpAP proteases
degrade proteins with carboxy-terminal peptide tails added by the SsrA-tagging system.
Genes Dev 12, 1338-47 (1998).
31.
Singh, S.K., Grimaud, R., Hoskins, J.R., Wickner, S. & Maurizi, M.R. Unfolding and
internalization of proteins by the ATP-dependent proteases ClpXP and ClpAP. Proc Natl
Acad Sci USA 97, 8898-903 (2000).
32.
Kenniston, J.A., Baker, T.A., Fernandez, J.M. & Sauer, R.T. Linkage between ATP
consumption and mechanical unfolding during the protein processing reactions of an
AAA+ degradation machine. Cell 114, 511-20 (2003).
33.
Wah, D.A., Levchenko, I., Baker, T.A. & Sauer, R.T. Characterization of a Specificity
Factor for an AAA+ ATPase. Assembly of SspB Dimers with ssrA-Tagged Proteins and
the ClpX Hexamer. Chem Biol 9, 1237-45 (2002).
34.
Wojtyra, U.A., Thibault, G., Tuite, A. & Houry, W.A. The N-terminal zinc binding
domain of ClpX is a dimerization domain that modulates the chaperone function. J Biol
Chem 278, 48981-90 (2003).
35.
Flynn, J.M., Neher, S.B., Kim, Y.I., Sauer, R.T. & Baker, T.A. Proteomic discovery of
cellular substrates of the ClpXP protease reveals five classes of ClpX-recognition signals.
Mol Cell 11, 671-83 (2003).
135
36.
Joshi, S.A., Baker, T.A. & Sauer, R.T. C-terminal domain mutations in ClpX uncouple
substrate binding from an engagement step required for unfolding. Mol Microbiol 48, 6776 (2003).
37.
Burton, R.E., Baker, T.A. & Sauer, R.T. Energy-dependent degradation: Linkage
between ClpX-catalyzed nucleotide hydrolysis and protein-substrate processing. Protein
Sci 12, 893-902 (2003).
38.
Ortega, J., Lee, H.S., Maurizi, M.R. & Steven, A.C. Alternating translocation of protein
substrates from both ends of ClpXP protease. EMBO J 21, 4938-49 (2002).
39.
Levchenko, I., Luo, L. & Baker, T.A. Disassembly of the Mu transposase tetramer by the
ClpX chaperone. Genes Dev 9, 2399-408 (1995).
40.
Levchenko, I., Yamauchi, M. & Baker, T.A. ClpX and MuB interact with overlapping
regions of Mu transposase: implications for control of the transposition pathway. Genes
Dev 11, 1561-72 (1997).
41.
Karon, B.S., Nissen, E.R., Voss, J. & Thomas, D.D. A continuous spectrophotometric
assay for simultaneous measurement of calcium uptake and ATP hydrolysis in
sarcoplasmic reticulum. Anal Biochem 227, 328-33 (1995).
136
Biographical note
Shilpa Joshi grew up in Kennewick, Washington, the daughter of Vandana and Arun and the
younger sister of Sampada and Suraj. In Maharashtra, India, her maternal grandfather, Gopal
Ranade, was the founder of the G.A. Ranade school for uneducated laborers and disadvantaged
children in Mumbai, and her paternal grandfather, Nilkanth Joshi, was a physicist and inventor.
Before immigrating to the U.S. in 1973, her mother worked as a teacher after being the gold
medallist from Elphinstone College, University of Mumbai, in 1963. Her father worked as a
nuclear engineer and continued this work after immigrating.
After graduating second in her high school class and being named a National Merit Commended
Student, Shilpa went to college at the University of California at Berkeley as a Robert C. Byrd
Scholar from Washington state in August 1993. Her first work in a laboratory was with Michael
A. Kennedy at Pacific Northwest National Laboratory as an Associated Western UniversitiesNorthwest Fellow in the summer of 1995. During that summer and the summer of 1997, she
helped optimize the purification of a protein useful in bioremediation efforts. Back at college,
she worked in Bing K. Jap's laboratory at Lawrence Berkeley National Laboratory and identified
the sequence for an aquaporin from bovine bone marrow. She also volunteered at the Berkeley
Adult School and tutored homeless adults in reading and mathematics during her last two years
of college. After graduating in May 1997 with an A.B. in Molecular and Cell Biology with a
minor in American Literature, Shilpa began graduate school at MIT in September 1997. While
there, she taught an XL seminar for inorganic chemistry through the Office of Minority
Education, and was a teaching assistant for undergraduate biochemistry and experimental
microbial genetics courses. Shilpa will now head to Eric Brown's laboratory at the University of
California at San Francisco, where she has accepted a postdoctoral position to study
Mycobacterium marinum.
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