Mechanisms of Substrate Recognition by the
AAA+ Protease HslUV
by
Shankar Sundar
B.A., Molecular and Cellular Biology (BMB)
University of California, Berkeley – 2006
SUBMITTED TO THE DEPARTMENT OF BIOLOGY IN PARTIAL FULFILLMENT OF
THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
AT THE
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
MAY 2011
 2011 Shankar Sundar. All rights reserved.
The author hereby grants to MIT permission to reproduce and to distribute publicly paper and
electronic copies of this thesis document in whole or in part in any medium now known or
hereafter created.
Signature of Author______________________________________________________________
Department of Biology
May 19, 2011
Certified by____________________________________________________________________
Robert T. Sauer
Salavador E. Luria Professor of Biology
Thesis Supervisor
Accepted by___________________________________________________________________
Alan D. Grossman
Praecis Professor of Biology
Chairman, Graduate Committee
Mechanisms of Substrate Recognition by the AAA+ Protease HslUV
by
Shankar Sundar
Submitted to the Department of Biology on May 19, 2011
in partial fulfillment of the requirements for the degree of Doctor of Philosophy at the
Massachusetts Institute of Technology
ABSTRACT
Protein degradation is a central component of all biological processes. The proteome must
constantly change in response to environmental stimuli. As a result, protein synthesis and
regulated proteolysis are vital to cell survival. In Escherichia coli, the protease HslUV is one of
five ATP-dependent proteases that shoulder the major burden of intracellular protein
degradation. Although ample data exist for describing the structural architecture of the HslUV
protease, very little is known about its mechanisms of function. I took a two-pronged approach to
understand the functional principles that govern this protease.
My first goal was to understand the rules of substrate recognition. To do so, I performed a
variety of experiments on two model proteins: Arc repressor and λcIN repressor. I found that
both substrates had common requirements for HslUV degradation, suggesting a conserved mode
of recognition by this protease. Mutagenesis of either substrate terminus affected binding and
degradation kinetics. While degron mutations generally affect only enzyme-substrate binding
properties in other bacterial proteases, the changes described here often affected the maximal rate
of HslUV degradation. Moreover, specific occlusion of either the N-terminus or C-terminus of
these substrates resulted in a substantial defect in degradation. A synergistic inhibitory effect was
observed for the simultaneous masking of both termini. These results suggested a mechanism of
tethering prior to engagement for degradation of HslUV substrates.
I then sought to define the regions of HslU that were important for recognition and found that
two segments, the GYVG pore loop and the intermediate (I) domain, played crucial roles.
Investigation of mutants altered at these sites supported a mechanism of tethering of the substrate
C-terminus to the I domain and engagement of the substrate N-terminus in the pore. I showed
that degradation of an Arc substrate proceeds processively from the N-terminus towards the Cterminus, lending further support to this idea. Interestingly, I also discovered that the I domain
plays a very important role in ATP hydrolysis by HslU and coordinates substrate recognition and
stimulation of ATP turnover. This trait appears to be unique for HslU and is not a property of the
accessory domains of other AAA+ protein unfolding machines.
Thesis Supervisor: Robert T. Sauer
2 ACKNOWLEDGEMENTS
This work would not have been possible without the help and support of several people…
First and foremost, I would like to thank my advisor, Bob Sauer, for continually challenging me
scientifically, for our spirited discussions, for unwavering support in the face of challenging
circumstances, for excellent guidance in experimental design and for being genuinely vested in
my ambitions both within science and outside of it. From Bob, I have learned how to think
creatively and critically as a scientist, how to successfully communicate my work, and how to be
an effective leader. I aspire to his ideal in all my future endeavors.
Tania Baker’s creative input, enthusiasm and positive energy have been instrumental in keeping
me on track and motivated during my graduate career. Without her, I am certain I would have
chased several wild geese and been very frustrated. For these reasons and many others, I have
enjoyed having her as my co-advisor.
The members of my thesis committee – Michael Laub and Uttam RajBhandary – have always
provided me a fresh perspective on my work and offered excellent experimental suggestions and
criticisms. My work would be severely lacking if not for their input. A special thanks goes to
Thomas Schwartz and Andres Leschziner for quickly getting up to speed on my work, making
useful suggestions on my thesis, and attending my thesis defense seminar.
Occupants of the 5th floor and the Schwartz Lab (on the 4th floor) deserve a world of thanks for
making coming into work every day a lot of fun and for providing a wonderful intellectual
environment.
The MIT Biology department has been an excellent place to pursue science. I want to
particularly highlight the contributions of Betsey Walsh, Frank Solomon, and the BioREFS
program members (past and present) in helping to make the department a nurturing place to do
research.
My new friends and classmates have made New England winters infinitely more bearable.
Special thanks in this respect go to Dave Christie, Mary Lee and Sofia Sasse. Also, the timely
transplantation of the Cupertino connection and the hope that we may all one day settle again in
California allowed me to both have a life and do science while pursuing life sciences research
and gave me so much to look forward to.
My parents and my brother have been my foundation for my whole life and graduate school has
been no different. Their love and compassion (even absent any understanding of my work) has
gotten me through so many tough times. I love them dearly for who they are and for who they
have allowed me to become.
Of course, the most hectic year of graduate school would have never seemed so easy or been so
much fun, if not for beebs. Thank you so much. :)
3 TABLE OF CONTENTS
ABSTRACT
2
ACKNOWLEDGEMENTS
3
TABLE OF CONTENTS
4
LIST OF FIGURES & TABLES
6
CHAPTER ONE: Introduction to Regulation and Substrate Specificity in Proteolysis
8
Mechanisms for Regulating Proteolysis in Biological Systems
Concentration-dependent mechanisms
Inhibition and activation mechanisms
Cleavage-site recognition
Compartmental proteases: The 26S proteasome
9
10
12
14
15
ATP-dependent Proteolysis in Bacteria
FtsH
Lon
ClpXP
ClpAP
18
20
22
24
28
The HslUV Protease
Cellular roles of HslUV
Structural and biochemical insights
Research Approach
29
30
32
34
CHAPTER TWO: Multiple Sequence Signals Direct Recognition and Degradation
of Protein Substrates by the AAA+ Protease HslUV
36
Abstract
Introduction
Results
Effects of terminal λcIN sequences on HslUV degradation
Effects of terminal Arc sequences on degradation
Terminal sequences do not alter substrate stability
Protein binding to the C-terminus of substrates slows HslUV degradation
ClpS binding to the N-terminus of substrates inhibits HslUV degradation
Synergistic effects of N- and C-terminal blocking
Single-turnover inhibition
37
38
40
40
42
43
43
44
44
45
Discussion
Materials & Methods
45
49
4 Acknowledgements
Figures & Tables
CHAPTER THREE: The Intermediate Domain of the AAA+ HslUV Protease
Coordinates Substrate Binding and Stimulation of ATP Hydrolysis
52
53
60
Abstract
Introduction
Results
Effects of HslU pore-loop mutations on substrate binding and degradation
Degradation of an Arc fusion protein starts at the N-terminus
The intermediate domain of HslU plays a major role in
determining ATP-hydrolysis rates
∆175-209linker HslUV degrades Arc-st11-ssrA with a lower
Vmax and weaker KM
Roles of the GYVG and 175-209 loops in substrate stimulation
of ATP hydrolysis
61
62
65
65
67
Discussion
Materials & Methods
Acknowledgements
Figures & Tables
72
76
79
80
CHAPTER FOUR: Perspectives for Future Research
68
70
71
88
Significance of HslUV Function in Cellular Physiology
Trapping experiments to identify biologically relevant substrates
Dissection of substrate recognition rules
Investigating protease function in other organisms
89
89
90
91
Detailed Mechanisms of HslUV Machine Function
Amino-acid level characterization of substrate binding by the I domain
Understanding how the I domain regulates ATPase function
Role of the I domain in intra-enzyme communication
92
92
94
95
Potential Lessons for the Study of Other AAA-Motors
95
APPENDIX A: Structures of HslU and HslUV
97
REFERENCES
103
5 LIST OF FIGURES & TABLES
Figure 1.1 – Schematic diagram of the 26S proteasome
15
Figure 1.2 – Cartoon depiction of ATP-dependent degradation
by bacterial proteases
19
Figure 1.3 – Domain organization of bacterial AAA+ enzymes
20
Figure 1.4 – Alignment of residues 159-210 from the HslU I domain
(E. coli numbering) from different organisms
35
Table 2.1 – Properties of λcIN and Arc substrates
53
Figure 2.1 – C-terminal λcIN sequences alter HslUV degradation
54
Figure 2.2 – C- and N-terminal sequences of Arc substrates
affect HslUV degradation
55
Figure 2.3 – Occluding the C-terminus of λcIN and Arc substrates
inhibits HslUV degradation
56
Figure 2.4 – ClpS binding to the N-terminus of Arc or λcIN substrates
slows HslUV degradation
57
Figure 2.5 – Combinatorial and single-turnover inhibition
58
Figure 2.6 – Model of HslUV recognition of a substrate (S)
with degrons at each terminus
59
Table 3.1 – Functional parameters of HslU enzyme variants
80
Figure 3.1 – Functional properties of HslU pore-loop mutants
81
Figure 3.2 – Degradation of Arc-GFP-st11-ssrA proceeds from the N-terminus
82
Figure 3.3 – The intermediate domain of HslUV is important
for robust ATP hydrolysis
83
Figure 3.4 – Degradation of Arc-st11-ssrA by ∆175-209linker HslUV
84
Figure 3.5 – ∆175-209linker HslU binding to substrate and peptide
85
Figure 3.6 – Stimulation of ATP hydrolysis by gt1-peptide or
Arc-st11-ssrA substrate
86
6 Figure 3.7 – Cutaway cartoon of substrate recognition by HslUV
87
Figure 4.1 – Sequence logos of residues 159-210 of the HslU I domain
from proteobacteria
93
Figure A.1 – Structure of HslU from Bochtler et al. (2000)
98
Figure A.2 – Structure of HslUV from Sousa et al. (2000)
99
Figure A.3 – Structure of HslU from Wang et al. (2001)
100
Figure A.4 – Structure of HslUV from Sousa et al. (2002)
101
Figure A.5 – Structure of HslU from Kwon et al. (2003)
102 7 Chapter One:
Introduction to Regulation and
Substrate Specificity in Proteolysis
8 Mechanisms for Regulating Proteolysis in Biological Systems
Life, at its very essence, is driven by chemical reactions. Enzymes allow these reactions to
proceed at biologically relevant rates and are required for diverse processes, ranging from the
transport of molecules within the cell to the locomotion of whole organisms (Cabeen & JacobsWagner, 2010). Enzymatic function, however, must be controlled. Biology has evolved many
mechanisms to regulate activity and thus ensure that chemical reactions only occur at the
appropriate time and place. These mechanisms are ubiquitous across all kingdoms of life and
classes of enzymes. Unsurprisingly, dysregulation of enzymatic activity is the primary cause of
many diseases, including numerous cancers and neurological disorders (Vande Walle et al.,
2008; Bedford et al., 2011; Wolfe et al., 2010).
Synthesis of new proteins by the ribosome represents one part of the cell’s effort to respond to
ever-changing environmental or developmental conditions. Equally important, proteolysis of
unneeded proteins and polypeptides is also required for proper maintenance of the cellular
proteome. Accordingly, proteases play key biological roles and their activities are tightly
regulated to prevent superfluous degradation.
Pepsin, a well-characterized digestive enzyme, offers an example of three distinct classes of
proteolytic regulation. First, when substrates are present in the stomach, the chief cells of the
stomach lining secrete pepsinogen, the inactive zymogen or precursor, into the digestive lumen
(Richter et al., 1998; Khan & James, 1998). This secretion is only brought about as a result of
changes in pH, osmotic pressure, and peptide binding to cell-surface receptors that occur in the
presence of food (Khropycheva et al., 2009). Next, the acidic environment of the stomach causes
9 the 44 amino-acid pro-segment (PS) to lose inhibitory contacts with the enzyme active site and to
reveal a specific cleavage site. Finally, the liberated active site recognizes and cleaves this PS
sequence. The cleaved PS fragment dissociates from the enzyme, now called pepsin, and the
fully active protease is free to degrade its substrates (Richter et al., 1998; Dee et al., 2009). Prior
to secretion, pepsinogen is inactive both because of autoinhibition by the pro-segment and
because the pH optimum for the mature protease is ~1.6, which is far below the intracellular pH.
Below, I discuss similar types of proteolytic regulation that operate in other biological programs.
Concentration-dependent mechanisms
Bimolecular and higher-order processes, such as substrate degradation, are dependent on the
concentrations of the reactants. Therefore, controlling enzyme/substrate concentration or
localization is a method of regulating proteolysis. The eukaryotic signal-peptidase complex
(SPC) and the mitochondrial stress-sensor HtrA2 provide examples of regulation by localization
and control of enzyme abundance.
Enzyme and substrate localization is used to control the activity of the signal-peptidase complex.
In yeast, the SPC is a bundle of five polypeptides embedded in the membrane of the endoplasmic
reticulum (Paetzel et al., 2002). The active sites for cleavage are positioned inwards toward the
ER lumen. When a secretory protein is synthesized on the ribosome, it is bound and targeted to
the translocon complex in the ER membrane by the signal-recognition particle (Paetzel et al.,
2002; Janda et al., 2010). The signal sequence required for targeting is encoded at the N-terminus
of the nascent protein and also contains a specific sequence motif for cleavage. During synthesis
and transfer to the ER, the signal sequence is preserved, as cytoplasmic proteases cannot access
10 and cleave this peptide. However, once the newly synthesized protein moves into the ER lumen,
the concentration of the membrane-bound SPC relative to the signal sequence, which is still
associated with the translocon, is very high, leading to cleavage. Thus, co-localization of the
protease and its substrate controls proteolytic activity in this instance.
Modulating protease expression is another method of regulation, as observed for the mammalian
HtrA2 protease, which is located primarily in mitochondria (Vande Walle et al., 2008). Upon
heat shock of neuroblastoma cells at 45 °C, Gray and colleagues (2000) found that HtrA2 protein
levels were elevated ten-fold. Several lines of evidence suggest that the concentration of HtrA2
is increased under heat-stress conditions to cope with the increased load of unfolded and
misfolded proteins at this temperature. Firstly, in mice, point mutants of the catalytic serine in
HtrA2 exhibit a phenotype similar to homozygous null alleles (Jones et al., 2003; Martins et al.,
2004). This result implicates the protease function of HtrA2 as being crucial. Furthermore, yeast
and bacterial homologs of HtrA2 act as proteases at high temperatures to clear the proteome of
damaged proteins (Padmanabhan et al., 2009; Vande Walle et al., 2008). When cells encounter
stress conditions, they increase the concentration of HtrA2 and related proteases to degrade
unfolded and misfolded substrates.
Secretion, co-localization, and altered expression are three methods by which protease activity
can be controlled. These types of regulation are common, not only for proteases, but also for
enzymes in general.
11 Inhibition and activation mechanisms
The second type of regulation seen in the pepsin example is the control of accessibility and
function of the active site. Electrostatic contacts between the pro-segment and the core protease
are lost in the acidic environment of the stomach and subsequent auto-cleavage of the PS fully
activates the protease (Richter et al., 1998). Thus, precisely when pepsin activity is needed, its
active site is fully formed and available to cleave substrate proteins. Similarly, pathways for
caspase activation utilize inhibitor removal, zymogen cleavage, and multimerization to control
protease activity in programmed cell death (Pop & Salvesen, 2009).
The irreversible nature of programmed cell death (also known as apoptosis) means that this
process must be very tightly regulated. A key point of regulation involves caspase-9, the protease
that initiates the cell-death program (Wang, 2001). To ensure that cell death does not happen
erroneously, inhibitor of apoptosis (IAP) family proteins bind procaspase-9 and prevent
dimerization and autocatalytic processing, which are required for formation of the active
protease (Shiozaki et al., 2003). Part of the IAP-binding site involves the N-terminus (ATPF) of
caspase-9 (Deveraux & Reed, 1999; Li et al., 2004; Pop & Salveson, 2009). Mitochondrial
destruction, a major signal in the progression of apoptosis, releases another protein Smac into the
cytosol. The N-terminus (AVPI) of Smac resembles that of procaspase-9 and so, is able to
compete away IAP binding (Srinivasula et al., 2001; Wang, 2001), allowing procaspase-9 to
participate in downstream steps required for activation and initiation of the apoptotic program
(see below). Thus, inhibition of an inhibitor plays a key role in caspase-9 activation. A similar
method of protease control is seen in HtrA family members from bacteria to humans, which
contain PDZ-domains that stabilize an inactive conformation of the protease. During cell stress,
12 binding of specific peptide sequences to the PDZ-domain relieves this inhibition and activates
the protease (Walsh et al., 2003; Murwantoko et al., 2004). Unlike IAP competition by Smac,
however, peptide binding to the PDZ domain results in allosteric rearrangements of the HtrA
active site, making it competent for proteolysis. Whether directly or indirectly, protease activity
is frequently regulated by controlling changes in tertiary and/or quaternary structure that are
necessary to form the functional enzyme.
The production of active caspase-9 and subsequent cell-death program require two additional
steps in which procaspase-9 forms a dimer and then cleaves itself to yield the active protease.
Dimerization is mediated by the recruitment of two procaspase-9 molecules to the heptameric
apoptosome scaffold, where “induced proximity” promotes formation of a procaspase-9 dimer
(Rodriguez & Lazebnik, 1999; Wang, 2001; Pop & Salveson, 2009). Dimerization of
procaspase-9, in turn, stimulates several auto-cleavage events, which produce active caspase-9
(Pop et al., 2006). The active enzyme then initiates a proteolytic cascade by cleaving the
executioner procaspase-3 molecule, activating caspase-3 to cleave other substrates that induce
cell death (Pop & Salveson, 2009). All initiator caspases are activated by intra-molecular selfcleavage, whereas downstream executioner caspases can be activated by cleavage in trans by the
initiator enzymes.
As we have seen, proteases can be regulated by cleavage and multimerization. Cleavage of a prosegment in a zymogen, either in cis- or in trans-, can remove inhibitory contacts at the active site
or cause a rearrangement of residues required for catalysis. Other processes, such as blood
clotting and initiation of the immunological complement pathway, also take advantage of
13 zymogen cascades for regulation (Davie & Kulman, 2006; Gal et al., 2009). It is equally
apparent that many protease systems, including HtrA family proteins and the immunoproteasome, require multimerization for activity (Sawa et al., 2010). Although the mechanisms
described here for proteolytic inhibition and activation, exemplified by pepsin, caspase-9, and
others, differ greatly, particularly in their reversibility, all are frequently used to modulate the
function of proteases across biology.
Cleavage-site recognition
Once a protease has been activated in a given intracellular or extracellular compartment, how are
specific substrates selected for cleavage? In general, the substrate sequence to be cleaved must
be complementary to the protease active site, affording some selectivity at the amino-acid
sequence level. For example, pepsin cleaves acid-denatured proteins in the stomach after almost
any Phe or Leu residue (Inouye et al., 1966; Fruton, 2002). Similarly, the digestive proteases
chymotrypsin and trypsin, cleave polypeptides in the intestines after aromatic/hydrophobic
residues and basic residues, respectively (Pozsgay et al., 1981; Di Cera, 2009). These very broad
specificities ensure that virtually any unfolded polypeptide in these compartments gets cleaved
multiple times. Some proteases are more selective. For example, the TEV protease cleaves after
the glutamine (Q) in sequences that match the pattern E-[xx]-Y-[x]-Q-[GS] (Dougherty et al.,
1989; Phan et al., 2002). However, the complexity of the intracellular proteome generally
precludes the use of cleavage-site specificity as a general method of substrate selection by
proteases for several reasons. First, most intracellular proteins are folded and thus most potential
sites of proteolytic cleavage are inaccessible. Second, with the exception of irreversible
14 processes such as apoptosis, intracellular degradation must be highly selective, allowing only
damaged or unneeded proteins to be proteolytically destroyed.
Compartmental proteases: The 26S proteasome
Many intracellular proteases sequester their active sites into a barrel-like chamber and then use
accessory proteins for substrate recognition. For example, the 26S proteasome consists of the
20S core particle (CP) and the 19S regulatory particle (RP) (Fig. 1.1). Studies of the assembly of
the CP revealed that the α-rings serve as nucleating scaffolds for β-ring formation. Once the
entire 20S particle is assembled, it forms an enclosed chamber, gated by the narrow axial pore at
the center of the α-ring. This architecture ensures that the proteolytic active sites (found on the
β1, β2, and β5 subunits) within the chamber will not encounter substrates by random diffusion
(Xie, 2010).
Structure, assembly and homeostatic regulation of the 26S proteasome
Journal of Molecular C
fully understand how the RP subunits are arran
plexes and how the RP stimulates the opening
facilitates substrate unfolding and translocatio
lication of several crystal structures of the es
the proteasomal ATPases from archaea (na
activating nucleotidase or PAN) and actin
ATPase forming ring-shaped complex or AR
some important clues to understand the act
(Djuranovic et al., 2009; Zhang et al., 2009a)
orthologous to the eukaryotic Rpt subunits,
40% sequence identity (Smith et al., 2006
region of PAN contains a coiled-coil (CC) se
an oligonucleotide/oligosaccharide-binding
Figure 1 Schematic diagram of the 26S proteasome. The 26S proteaOB domains from six PAN N-terminal fragment
some consists
of the of
20S the
core particle
(CP) and the 19S (Xie,
regulatory
Figure 1.1 – Schematic
diagram
26S proteasome
2010).
andsymmetry. The axial
ring Components
with a pseudo 6-fold
particle (RP). The CP is formed by four stacked rings: two outside
assemblage of the 20S CP and 19S RP is shown. Importantly, the ishexameric
Rpt
ring of the six PAN frag
!13 Å. The CC
segments
a-rings and two inner b-rings. Each ring is formed by seven different
3-fold
symmetry.
Thus,
(comprising Rpt1-6) issubunits.
the ATP-dependent
motor
and
the
proteins
in
the
lid
are
responsible
forthe N-terminal fra
The RP is divided into two subcomplexes: the lid and the
assembled
into
a
crown-shaped
subcomplex:
ubiquitin recognition. base.
Certain
subunits
ofthethelid CP
β-rings
contain
the proteolytic active sites,
The subunits
forming
and the
base are
shown. Rpn10
sprouting from the hexameric OB ring (Djura
stabilizesand
the association
between
the lid and the base.
which face into the chamber
away from
solution.
Zhang et al., 2009a). The N-terminal region o
segment and two OB domains. The OB dom
favorable for antigen processing (Boes et al., 1994; Gaczynska
double hexameric rings (Djuranovic et al., 2
et al., 1994; Cardozo and Kohanski, 1998). b5i is replaced by
domains of PAN and ARC are located at th
another proteolytic active subunit b5t in cortical thymic epithelial
ATPase domain of PAN was also crystalliz
cells, forming the so-called thymoproteasome (Murata et al.,
2009a). It appears to form the canonical A
2007). Different from eukaryotic CP, there are only one type of
15 Together, these
with a narrow channel.
a subunit and one type of b subunit in prokaryotic proteasomes,
suggest a model of double-ring structure for
with the exception of the proteasome isolated from bacterium
This model is consistent with a three-dimensi
Rhodococcus, which contains two types of a and b subunits
generated from tomographic reconstructi
Cleavage of substrates by the proteasome is only possible when the RP binds substrates, unfolds
them, and translocates the resulting polypeptide into the chamber of the CP (Xie, 2010). As a
result of this compartmentalization, the activity of the proteasome can be tightly controlled
depending on the specificity and behavior of the 19S regulatory particle.
The ubiquitin-proteasome pathway (UPP) is responsible for the majority of protein degradation
in the eukaryotic cell. It is also responsible for conferring specificity to proteasomal degradation.
In the UPP, the 76-residue protein Ubiquitin (Ub) is activated by an activating-enzyme (E1) and
transferred onto a specific Ub-carrier protein (E2). Ub-ligase enzymes (E3) recruit the E2 and the
protein marked for degradation to form a ternary complex. The Ub moiety is then transferred
onto the substrate protein. Multiple rounds of this process (or extension of the Ub-chain by E4
elongation factors) yield a substrate protein that has been poly-ubiquitinated. The lid of the 19S
particle recognizes this motif and uses its isopeptidase activity to disassemble the ubiquitin
chain. Subsequently, the target protein is mechanically unfolded by the Rpt1-6 ATPases in an
energy-dependent process and translocated into the 20S chamber, where it is cleaved into
peptides (Liu et al., 2007; Gallastegui & Groll, 2010). Specific regulation of substrate
degradation is attributable to the E2 and E3 accessory proteins. Mammalian cells contain tens of
E2 carriers and hundreds of E3 ligases. The diversity of the E3 Ub-ligases in structure,
recognition specificity, and post-translational modification confers differences in rates of
ubiquitination of various substrates. In turn, this leads to differential degradation of these
substrates by the 26S proteasome (Liu et al., 2007). In many processes, Ub-ligases also function
to bind the proteasome directly and enhance degradation in a ligase-independent manner
(Kulikov et al., 2010; Um et al., 2010). They achieve enhancement by stabilizing the 19S particle
16 or facilitating the substrate-proteasome interaction. These latter techniques of regulation are
highly analogous to those used by bacterial adaptor proteins, which will be discussed later.
Regardless of the mechanism, it is clear that accessory proteins of the UPP are the primary
regulators of proteasomal activity.
In the much simpler prokaryotic proteasome from Mycobacterium tuberculosis, a coiled-coil
region resides above the ATPase domains and is required for recognition of a ubiquitin-like
protein (Wang et al., 2010). Additionally, a second domain that sits above the ATPase ring (the
interdomain) seems to be important for initial binding and translocation of unfolded polypeptides
(Wang et al., 2009). Although the exact role of each subunit in the 19S regulatory particle is not
fully understood, it is possible that some mammalian RP subunits may also contribute to
substrate choice in this way.
The specificity of the 26S proteasome depends upon the sequestration of its active sites, the
ability of E2 and E3 enzymes in the UPP to tag appropriate substrates, and the ability of the RP
to recognize these tagged substrates, unfold them in an ATP-dependent manner, and then spool
them into the CP for degradation. These reactions allow the 26S proteasome to destroy large,
stably folded proteins with great speed and specificity. Unlike pepsin, whose specificity stems
entirely from the shape of its active-site binding pocket, the complex mechanisms used by the
proteasome allow it to be a key player in many important cellular processes. Defects in the
function and regulation of the 26S proteasome are implicated in numerous human diseases,
including but not limited to cancer, cardiovascular disease, viral pathogenesis, and neurological
17 disorders (Bedford et al., 2011). For this reason, significant effort has been invested in
understanding the function of this complex enzyme and related protein-degrading machines.
ATP-dependent Proteolysis in Bacteria
Some of the mechanisms that regulate 26S-proteasome activity are also observed in bacterial
AAA+ proteases, which are simpler and thus more amenable to detailed experimental
investigation. For example, Escherichia coli contains five ATP-dependent proteases (FtsH, Lon,
ClpXP, ClpAP, and HslUV) that sequester their active sites in internal chambers and utilize
various modes of direct or adaptor-assisted substrate recognition to ensure regulated proteolysis.
These proteases possess distinct, yet overlapping, roles in maintaining the cellular proteome and
in regulating biochemical reactions (Sauer et al., 2004; Baker & Sauer, 2006; Schmidt et al.,
2009).
All five ATP-dependent proteases in E. coli share a common architecture, which resembles that
of the 26S proteasome. Each protease has a hexameric “regulatory particle” that couples ATPhydrolysis to substrate recognition and unfolding, although the intrinsic strength of unfolding
may vary significantly depending upon the substrate and enzyme (Koodathingal et al., 2009).
The unfolded polypeptide is then translocated through a central pore into the peptidase chamber,
where it is cleaved by otherwise inaccessible active sites (Fig. 1.2). For the Lon and FtsH
proteases, the unfoldase and peptidase units are encoded on a single polypeptide.
18 Figure 1.2 – Cartoon depiction of ATP-dependent degradation by bacterial proteases
(Baker & Sauer, 2006). The unfoldase recognizes and binds a substrate degradation tag. It then
applies a mechanical force to unfold and translocate the substrate protein into the peptidase
chamber. The peptidase active sites cleave the unfolded polypeptide and the resulting fragments
are free to leave by diffusion.
The hexameric AAA ring used for substrate unfolding and translocation is functionally similar to
the ring formed by ATPases of the 19S base (Rpt1-6), whereas the peptidase resembles the 20S
core particle. Each bacterial unfoldase also has accessory domains that play roles in substrate
recognition and degradation (Dougan et al., 2002a; Xia et al., 2004; Rotanova et al., 2006;
Abdelhakim et al., 2008; Erbse et al., 2008; Chowdhury et al., 2010). These domains often reside
above the unfolding ring and probably serve some functions analogous to those of proteins in the
19S lid of the eukaryotic proteasome (Wang et al., 2001; Botchler et al., 2000; Kim & Kim,
2003). The domain architectures of different classes of bacterial AAA+ unfoldases and related
enzymes are shown in Figure 1.3.
19 Figure 1.3 – Domain organization of bacterial AAA+ enzymes (adapted from Dougan et al.,
2002a). Similarities and differences in domain architecture between the five ATP-dependent
proteases in E. coli are shown here. Lon and FtsH contain both the unfoldase and peptidase
components on a single polypeptide. Interestingly, while four of the enzymes have an N-terminal
accessory domain, HslU instead has an intermediate (I) domain.
Significant differences between the 26S proteasome and these ATP-dependent bacterial
proteases include the overall complexity of the machine in terms of the number of distinct
subunits and the intricacy of the mechanisms used to ensure substrate specificity.
FtsH
Of the five AAA+ proteases in E. coli, FtsH is the only one that is membrane anchored (Ogura et
al., 1991; Tomoyasu et al., 1993a). FtsH contains two transmembrane segments near its Nterminus that tether it to the inner face of the cytoplasmic membrane. This proteolytic machine is
also the only AAA+ protease that is essential for E. coli growth (Ogura et al., 1991; Tomoyasu et
al., 1993b). FtsH degrades LpxC, which is required for the biosynthesis of lipopolysaccharide
(LPS) and thus for maintenance of outer-membrane integrity (Ogura et al., 1999); this is the FtsH
20 activity required for cell viability (Jayasekera et al., 2000; Ito & Akiyama, 2005). Both depletion
and overexpression of LpxC are lethal, and degradation by FtsH is thought to maintain a level of
this enzyme that results in the proper ratio of LPS to other phospholipids (Führer et al., 2007).
Biochemical and genetic experiments suggest that other physiological functions of FtsH pertain
to protein quality control and stress sensing at the membrane (Akiyama & Ito, 2000; Shimohata
et al., 2002; Ito & Akiyama, 2005).
Like the other ATP-dependent unfoldases in bacteria, FtsH is active as a homo-hexamer
(Akiyama & Ito, 2000; Suno et al., 2006). For FtsH, however, the periplasmic region, the AAA+
domains, the protease domain, and perhaps even the C-terminal coiled-coil segment all appear to
contribute to hexamerization (Akiyama et al., 1998; Akiyama & Ito, 2000; Shotland et al., 2000;
Suno et al., 2006).
Substrate recognition by FtsH is mediated by three distinct regions: the N-terminal periplasmic
domain, the conserved pore-1 loop of the AAA-module, and the C-terminal coiled-coiled domain
(Akiyama et al., 1998; Shotland et al., 2000; Yamada-Inagawa et al., 2003). A periplasmic
complex, HflKC, interacts with the N-terminal domain of FtsH and modulates degradation of
specific membrane proteins (Akiyama et al., 1998). In this fashion, HflKC acts as an adaptor
molecule, enhancing or inhibiting degradation of membrane-bound substrates. Of the other
recognition elements, the pore-1 loop is required for processive translocation of all substrates and
the coiled-coiled region helps bind cytosolic proteins. Biochemical studies have shown that nonpolar C-terminal sequences target many substrates to FtsH, which can degrade unfolded
21 polypeptides and metastable proteins but has a very limited ability to unfold stable protein
domains (Asahara et al., 2000; Herman et al., 2003).
Lon
Degradation by the Lon protease is important for several physiological processes in E. coli (for
review, see Van Melderen & Aertsen, 2009). Deletion of the lon gene is not lethal, but these
mutants are highly mucoid, because the RcsA protein is not degraded efficiently, and are hyper
sensitive to DNA damage, because the SulA protein is not degraded completely (Mizusawa &
Gottesman, 1983; Stout et al., 1991). Interestingly, the E. coli HslUV protease (see below) also
degrades RcsA and SulA (Wu et al., 1999). Overlapping specificity of this type is a relatively
common theme, suggesting that proper degradation of certain proteins is an important cellular
priority. Transcription of the sulA gene is normally repressed by the LexA stress-response
regulator. Upon DNA damage, LexA undergoes an autocleavage reaction that reduces its affinity
for operator DNA, resulting in increased expression of many SOS-response proteins, including
SulA, which functions as a cell-division inhibitor (Huisman et al., 1984). Once DNA repair is
complete, enhanced expression of SulA ceases and degradation by Lon and/or HslUV is required
to relieve cell-division arrest and to resume normal growth. If SulA is not degraded, cells form
filaments and eventually die (Gottesman et al., 1981), illustrating the importance of degradation
in regulating cell growth and responses to stress.
From a structural perspective, Lon is similar to FtsH in having the ATPase and protease domains
on a single polypeptide (Fig. 1.3). A ring-hexamer structure for Lon was established by electron
microscopy of the E. coli enzyme and crystallography of an ortholog from Thermococcus
22 onnurineus (Park et al., 2006; Cha et al., 2010). In addition to the central AAA-module and Cterminal protease domain, Lon also possesses an N-terminal accessory domain that consists of
two sub-domains, one primarily β-sheet and the other consisting of an α-helical coiled coil (Li et
al., 2010). Interestingly, the N-terminal domain of Lon shares significant structural similarity
with parts of RssB (Levchenko et al., personal communication; PDB: 3eq2, 3f7a), a substratedelivery adaptor for ClpXP (see below). Functionally, this domain of Lon appears to be involved
in specific recognition of some substrates. For example, mutation of Glu240 in the coiled-coil
portion of the N domain eliminates intracellular degradation of RcsA but not of SulA by Lon
(Ebel et al., 1999; Li et al., 2010). Removal of the N domain by limited proteolysis also results in
an enzyme with low basal ATPase activity, which cannot be stimulated by substrates, and has
severely reduced degradation activity (Rotanova et al., 2006; Melnikov et al., 2008).
Destabilization of hexamers, however, could be responsible for some of these effects.
In E. coli, Lon is responsible for approximately half of the degradation of proteins caused by
premature translational termination or the incorporation of amino-acid analogs (Kowit &
Goldberg, 1977), suggesting that it must recognize most proteins that cannot fold properly.
Indeed, one class of peptide signals in substrates recognized by Lon consists of hydrophobic
residues that would normally be buried in folded protein structures (Gonzalez et al., 1998; Gur &
Sauer, 2008a). It has been suggested that in addition to degrading protein substrates, Lon may
have chaperone activity, regulated allosterically by substrate sequence elements (Gur & Sauer,
2009). Although this proposal remains to be tested rigorously, Lon clearly functions to degrade
specific proteins, such as RcsA and SulA, and also serves as a general stress-response protease
maintaining protein quality in the cell.
23 ClpXP
ClpXP is arguably the best characterized of all of the AAA+ proteases. Unlike FtsH and Lon, the
ClpXP protease is composed of two distinct polypeptides. The ClpP peptidase is formed by two
stacked heptameric rings, which enclose the degradation chamber (Wang et al., 1997; Bewley et
al., 2006; Kim & Kim, 2008). ClpX assembles as a ring hexamer (Grimaud et al., 1998; Glynn et
al., 2009) and associates non-covalently with ClpP14. ClpX recognizes substrates and, using the
energy of ATP binding and hydrolysis, unfolds and translocates them into ClpP for degradation
(Sauer et al., 2004).
In E. coli, one cellular role of ClpXP is degradation of ssrA-tagged proteins (Gottesman et al.,
1998; Moore & Sauer, 2007). When translation of an mRNA stalls because of an unoccupied A
site, the tmRNA molecule binds the stalled ribosome, adds a charged alanine to the nascent
chain, and then directs addition of ten additional amino acids before normal termination of
translation (Keiler et al., 1996). The resulting C-terminal peptide sequence (AANDENYALAA)
is called the ssrA tag. Approximately, 1 in 200 translational events terminates in ssrA tagging
(Moore & Sauer, 2005), and ClpXP is the primary cytosolic protease responsible for recognition
and degradation of these ssrA-tagged proteins (Gottesman et al., 1998; Farrell et al., 2005; Lies
& Maurizi, 2008).
In addition to its role in protein quality control, ClpXP plays roles in regulating the cell cycle and
cell division (Jenal & Fuchs, 1998; Camberg et al., 2009; Camberg et al., 2011). In Caulobacter
crescentus, for example, ClpXP degrades CtrA, the master cell-cycle transcription factor (Jenal
& Fuchs, 1998). Precisely regulated CtrA degradation is such a vital part of the developmental
24 program in this bacterium that cells lacking the clpX gene are inviable. In E. coli, ClpXP
modulates the function of FtsZ, a protein that forms a large, multimeric ring at the mid-cell
during cell division. The ClpX unfoldase can inhibit FtsZ multimerization simply by binding this
protein (Sugimoto et al., 2010), thus modulating ring formation. The complete ClpXP protease,
however, degrades FtsZ monomers and polymers at a modest rate in wild-type cells (Flynn et al.,
2003; Camberg et al., 2009). Overproduction of the protease leads to increased degradation and a
filamentous phenotype caused by the disruption of ring formation (Camberg et al., 2009). Thus,
ClpXP appears to balance formation and destruction of the FtsZ ring during cell division. ClpXP
also degrades many other cellular proteins (Flynn et al., 2003; Neher et al., 2006), including
enzymes induced during the SOS-response to DNA damage and transcription factors that
mediate environmental stress-response systems. As observed for Lon and FtsH, peptide signals,
including the ssrA tag and other sequence motifs target substrates to ClpXP (Flynn et al., 2003;
Neher et al., 2006).
Some peptide sequences that target substrates to ClpX bind in the axial pore of the hexamer. For
example, mutations in the pore-1, pore-2, and RKH loops that form the pore substantially
increase KM for degradation of ssrA-tagged substrates (Siddiqui et al., 2004; Martin et al., 2007;
2008a; Farrell et al., 2007). Furthermore, cysteines introduced into the pore-1 and pore-2 loops
can be crosslinked to cysteines introduced into the ssrA tag (Martin et al., 2008b). The pore-1
loop of ClpX has a highly conserved GYVG sequence motif. Importantly, mutations in this loop
have also been shown to alter the intrinsic ability of ClpX to unfold and translocate saturating
concentration of substrates (Martin et al., 2008a), suggesting that these loops help grip and pull
on the substrate during the conformational changes responsible for these mechanical activities.
25 Indeed, Lon, FtsH, ClpA, and HslU all have similar pore-1 loop sequences that also appear to be
important for substrate recognition, unfolding, and translocation (Yamada-Inagawa et al., 2003;
Park et al., 2005; Hinnerwisch et al., 2005a).
Deletion of the family-specific N-terminal domain of ClpX does not affect binding to ClpP or the
rate of ATP-dependent degradation of native ssrA-tagged proteins, demonstrating that the
hexameric AAA-module carries out the mechanical functions of ClpX (Singh et al., 2001;
Wojtyra et al., 2003; Martin et al., 2005; 2007). N-domain dimers appear to be flexibly tethered
to the AAA+ ring of ClpX and they stabilize hexamer formation at low protein concentrations by
maintaining higher concentrations of the tethered AAA+ subunits. These domains are also
needed for recognition of some protein substrates, including FtsZ, MuA transposase, phage λO
replication protein, and the Bacillus subtilus Spx stress-response regulator (Singh et al., 2001;
Wojtyra et al., 2003; Zhang & Zuber, 2007; Abdelhakim et al., 2008; Sugimoto et al., 2010).
Modeling places each N-domain dimer above the AAA+ ring in a position that would potentially
allow a peptide tag on a tethered substrate to interact concurrently with the axial pore (Kim &
Kim, 2003; Glynn et al., 2009).
Proteomic studies, in which endogenous E. coli substrates were trapped in the chambers of
inactive ClpXP proteases and then identified, confirmed the importance of peptide targeting and
revealed multiple types of degradation signals (Flynn et al., 2003; Neher et al., 2006).
Bioinformatic analysis of the trapped sequences, peptide-blotting experiments, tag-transfer
experiments, and mutational studies suggested a minimum of five classes of ClpXP degradation
tags. Two classes of C-terminal targeting signals shared homology with the ssrA-tag and MuA-
26 tag sequences, respectively. The substrate-identification experiments also identified several types
of N-terminal recognition signals, including one similar to a sequence that targets λΟ to ClpXP
(Gonciarz-Swiatek et al., 1999). Just a few residues define most of these targeting motifs and are
necessary for robust degradation by ClpXP (Flynn et al., 2001; Flynn et al., 2003).
Controlling the accessibility of recognition sequences can determine if and how quickly specific
substrates are degraded by ClpXP. For example, MuA tetramers interact with ClpXP differently
than monomers because formation of the multimer causes a conformational change that reveals
previously hidden degradation signals (Abdelhakim et al., 2008). Endoproteolytic cleavage of the
LexA and RseA proteins also produces fragments that conclude with an “ssrA-like” AA-COOH
sequence, which targets these proteins for ClpXP degradation (Neher et al., 2003; Flynn et al.,
2004).
ClpXP recognition of RseA and ssrA-tagged substrates can be enhanced by the adaptor protein
SspB, which binds both to these substrates and to the N-domain of ClpX (Levchenko et al.,
2000; 2003; 2005; Flynn et al., 2004; Park et al., 2007). In the ternary substrate-delivery
complex, the local concentration of the substrate relative to the axial pore is increased, resulting
in lower KM’s and an increased Vmax for degradation (Levchenko et al., 2000; Hersch et al.,
2004; Flynn et al., 2004; McGinness et al., 2007; Davis et al., 2009). Another ClpXP adaptor,
RssB, functions using similar principles (Zhou & Gottesman, 1998; Becker et al., 1999; Zhou et
al., 2001; Studemann et al., 2003). RssB is required for degradation of RpoS, the stationaryphase transcription factor, during exponential bacterial growth (Bougdour et al., 2008). The
27 combined use of latent degradation tags, multivalent degrons, and adaptor proteins allows
intricate levels of proteolytic control (Baker and Sauer, 2006).
ClpAP
ClpAP also utilizes the compartmental ClpP14 peptidase but in combination with hexamers of the
ClpA unfoldase, which consists of two stacked AAA+ rings (D1 and D2), both of which are
required for optimal protease activity (Kress et al., 2009). ClpA alone can unfold native proteins,
function as a disassembly chaperone, and act independently of ClpP in regulating RpoS during
stationary phase (Wickner et al., 1994; Pak et al., 1999; Weber-Ban et al., 1999; Hoskins et al.,
2000; Hoskins et al., 2002; Weichart et al., 2003). ClpAP degrades ssrA-tagged proteins
(although ClpXP is responsible for most degradation in the cell), other substrates during
stationary phase, and N-end rule substrates with the aid of the ClpS adaptor (Gottesman et al.,
1998; Farrell et al., 2005; Lies & Maurizi, 2008; Chenoweth & Wickner, 2008; Dougan et al.,
2010).
Loops in the axial pore of ClpA are important for translocation, unfolding, and recognition of
some substrates (Hinnerwisch et al., 2005a). The ClpA N-terminal domain is important for
recognition of RepA, recognition of the ClpS adaptor, and appears to be needed for ClpAP
degradation of unfolded proteins (Pak et al., 1999; Hinnerwisch et al., 2005b; Xia et al., 2004;
Ishikawa et al., 2004; Hoskins & Wickner, 2006). Flexible tethering of the monomeric N
domains to the D1 ring of ClpA allows them to be highly dynamic and to assume many different
positions relative to the axial pore (Cranz-Mileva et al., 2008).
28 The ClpS adaptor serves negative and positive functions in substrate recognition by ClpA. For
example, unlike ClpAP, ClpAPS complexes do not bind and degrade ssrA-tagged proteins
(Dougan et al., 2002b). However, ClpS binds substrates with N-terminal Tyr, Phe, Trp, or Leu
residues and delivers them for ClpAP degradation (Erbse et al., 2006; Hou et al., 2008; Wang et
al., 2008; Roman-Hernandez et al., 2009). Interestingly, simple tethering of these N-end rule
substrates to ClpA is not sufficient to promote degradation (Hou et al., 2008), suggesting that
active handoff of the substrate from the adaptor to the protease is required for proteolysis. ClpS
beautifully illustrates the role of adaptor-mediated substrate delivery in regulating substrate
choice and thus protease activity. For example, an increase in the concentration of ClpAP
relative to ClpS during stationary phase in E. coli allows increased degradation of ssrA-tagged
proteins and presumably other substrates (Farrell et al., 2005). During exponential growth,
however, proteolytic activity is probably restricted to substrates that can be degraded by ClpAPS.
The HslUV protease
HslUV (sometimes called HslVU or ClpYQ) was discovered in a screen for proteins induced by
heat shock (Chuang & Blattner, 1993). Indeed, the expression of HslU (a hexameric AAA+
unfoldase) and HslV (a double-ring dodecameric compartmental peptidase) is driven by a
promoter controlled by σ32, the heat-shock transcription factor (Chuang et al., 1993; Lien et al.,
2009a). Since its initial discovery and characterization, HslUV has been found to play roles in
other cellular stress responses as well. Interestingly, significant sequence and structural
homology exists between the HslV peptidase and the catalytic β-subunits of the 20S proteasome
core (Rohrwild et al., 1996; Missiakas et al., 1996; Bochtler et al., 1997). Multiple crystal
structures have been determined for HslU, HslV, and the HslUV complex (Appendix A; for
29 review, see Groll et al., 2005), making HslUV the best characterized system from a structural
perspective.
Although nearly 20 years have passed since the discovery of HslUV, relatively little is known
about the detailed biochemical mechanisms of machine function, especially with respect to
substrate specificity. Below, I summarize what is known about this AAA+ protease and
introduce my dissertation work, which aims to address some of the unanswered questions.
Cellular roles of HslUV
In addition to heat-shock roles, HslUV has been found to participate in the SOS response to
DNA damage, the acid-stress response, the unfolded-protein response, and the suppression of
mucoidy (Missiakas et al., 1996; Kanemori et al., 1997; Khattar, 1997; Kuo et al., 2004; Kannan
et al., 2008). In E. coli, HslUV is not essential for survival, although deletion of the hslU gene
causes temperature sensitivity above 43 °C (Missiakas et al., 1996). Synthesis of these studies
strongly suggests that HslUV is a “backup” protease that plays an increased role at higher
temperatures. Two lines of evidence support this idea: (i) each of the well-characterized HslUV
substrates is also degraded by a second AAA+ protease; and (ii) many of its attributed cellular
functions are a result of protease-overexpression studies.
HslUV degrades the cell-division inhibitor SulA and the heat-shock transcription factor σ32,
providing an example of feedback regulation (Kanemori et al., 1997; 1999a; 1999b; Seong et al.,
1999; Wu et al., 1999). In these experiments, the effects of hslU deletion on the half-lives of
SulA and σ32 were probed by western blots of cell extracts at various times after inhibition of
30 protein synthesis, convincingly showing that maximal rates of protein destruction require HslU.
However, for both substrates, a second protease seemed to play the primary role in degradation.
For example, Lon is the main protease that degrades SulA, and FtsH is the principal enzyme that
degrades σ32 (Kanemori et al., 1997; 1999a; 1999b; Seong et al., 1999; Wu et al., 1999). The
contribution of HslUV to the overall intracellular rates of degradation of SulA and σ32 was
increased considerably at temperatures above 41.5 °C. This result makes sense for two reasons.
First, because the HslUV operon is under σ32 control, the levels of this protease are increased
significantly at higher temperatures. Second, HslUV is more proteolytically active at higher
temperatures (Burton et al., 2005).
Degradation of the capsule-synthesis activator RcsA (again, primarily a Lon substrate) and of
proteins that unfold during heat-shock conditions also varies depending on HslUV concentration
(Missiakas et al., 1996; Kanemori et al., 1997; Khattar, 1997; Kuo et al., 2004). For all of these
substrates, increased cellular degradation occurred upon HslUV overexpression. These
investigations of the physiological role of HslUV highlight a common theme. Under normal
growth conditions, HslUV seems to function in a secondary role to other cellular proteases, such
as Lon and FtsH. It is only upon increased expression and/or activity (both of which are achieved
at higher temperatures) that it takes on a more prominent role in proteolysis. Because HslUV
expression is controlled by σ32, future work probing the cellular role of this protease should
focus on conditions which result in increased amounts of σ32 and thus increased amounts of
HslUV.
31 Structural and biochemical insights
Our understanding of the HslUV machine has come mainly from x-ray crystallographic studies
in combination with some biochemical experiments. These studies have identified how HslU
binds nucleotides, how it interacts with HslV, and have begun to probe how HslU may recognize
some substrates.
Multiple high-resolution structures of HslU, alone or in complex with HslV, have been solved
(Appendix A; Botchler et al., 2000; Sousa et al., 2000; Wang et al., 2001; Sousa et al., 2002;
Kwon et al., 2003; Groll et al., 2005). As expected, these structures show that ATP/ADP binding
involves the Walker A, Walker B, sensor I, sensor II, and “arginine finger” motifs that are
characteristic of all AAA+ enzymes. Strikingly, however, some crystal structures showed
symmetric binding of six nucleotides to each HslU hexamer, whereas others show asymmetric
binding to only three or four subunits of the hexamer. Solution studies revealed that a maximum
of three or four nucleotides bind the hexamer (Yakamavich et al., 2008), suggesting that the
asymmetric structures are more physiologically relevant. Such asymmetry would also be
consistent with structural and biochemical results for ClpX (Hersch et al., 2005; Glynn et al.,
2009), whose AAA+ module shares ~50% sequence homology with that of HslU.
Structures of HslV alone and in complex with HslU revealed that the proteolytic active sites are
only well formed in the complete HslUV protease (Sousa et al., 2002). A malformed active site
is also seen in the crystal structure of CodW, an HslV-like peptidase found in Bacillus subtilis
(Rho et al., 2008). Indeed, free HslV has almost no ability to hydrolyze small peptide substrates.
Addition of HslU and the non-hydrolyzable ATP analog, ATPγS, however, allosterically
32 activates the peptidase activity of HslV (Rohrwild et al., 1996; Yoo et al., 1996; Sousa et al.,
2002; Park et al., 2008). Multiple biochemical studies have established that the highly conserved
C-terminal tails of HslU are responsible for communication with the peptidase (Seong et al.,
2002; Park et al., 2008). In fact, a peptide corresponding to the final 10 residues of HslU is
sufficient to elicit HslV cleavage of model peptide substrates. Interestingly, activation of the 20S
proteasome core particle by C-terminal tails of subunits in the AAA+ ring is thought to occur in
a similar fashion both in archaea (by PAN) and in eukaryotes (by 19S RP) (Smith et al., 2007;
Yu et al., 2010). Although some crystal structures corroborate the importance of the HslU tails in
contacting and activating HslV (Sousa et al., 2000; Sousa et al., 2002; Kwon et al., 2003), others
show no interactions between these tails and the peptidase in the complex (Wang et al., 2001). It
remains to be determined whether HslU can interact with HslV in fundamentally different ways
and if different complexes have different activities.
Although basic HslUV functions, such as nucleotide binding and peptidase activation, are
understood in some detail, a key feature of the enzyme that permits regulated function – substrate
choice – remains poorly studied. For example, very little is known about the determinants that
result in recognition and degradation of physiological substrates. In one report, it was shown that
appending the C-terminal 11-residues of SulA to Arc repressor was sufficient for HslUV
degradation (Kwon et al., 2004). However, Arc repressor alone is a good substrate for HslUV
degradation, and the amino-terminal parts of this substrate seemed most important for HslUV
recognition (Burton et al., 2005). Other experiments suggested that the GYVG pore loop and the
intermediate (I) domain of HslU (see Fig. 1.3) were important for degradation of folded
substrates (Song et al., 2000; Kwon et al., 2003; Lee et al., 2003; Park et al., 2005; Lien et al.,
33 2009b). Because the pore loops and accessory domains of other AAA+ proteases are also
involved in substrate recognition, these results were not surprising. However, none of the
published studies provided any detailed characterization or convincing evidence that linked the
pore loops or I domain to specific roles in substrate recognition, unfolding, or translocation.
Research Approach
In this chapter, I have summarized mechanisms used by all organisms to modulate proteolytic
activity, with an emphasis on the function of ATP-dependent proteases in bacteria. The ability of
this class of enzymes to participate in the destruction of cellular substrates depends entirely on
the capacity of the AAA+ unfoldase to recognize specific proteins. Because the molecular
mechanisms of substrate recognition by HslUV were poorly characterized, I have carried out
detailed biochemical studies of its interactions with several model substrates.
In chapter two, I use mutagenesis and sequence-occlusion experiments to show that both the Nand C-terminal sequences of substrates act as degrons for HslUV recognition. These results
resolve the apparent conflict in the literature regarding the location of the degradation signal in
Arc-repressor variants (Kwon et al., 2004; Burton et al., 2005). In chapter three, I undertake a
more thorough investigation into the role of the GYVG loop and the I domain in HslUV
function. Although the GYVG loop functions similarly in HslU and other AAA+ unfoldases, the
I domain is unlike other accessory domains. Crystal structures that contain a significant portion
of the I domain reveal that the apical region of this domain is highly disordered (Bochtler et al.,
2000; Wang et al., 2001). However, closer examination of this unstructured region reveals strong
sequence conservation across all species that contain this protease (Fig. 1.4).
34 Figure 1.4 – Alignment of residues 159-210 from the HslU I domain (E. coli numbering)
from different organisms. These residues correspond to the apical tip of the HslU I domain; the
residue 175-209 region is disordered in all crystal structures. The first four sequences are from γproteobacteria, O. formigenes is a β-proteobacterium and C. crescentus is an α-protobacterium.
T. maritima is a more distantly related bacterial species. Finally, the pea aphid and leishmania
are eukaryotes. There are several highly conserved basic and acidic residues in this stretch as
well as I/V/L residues at positions 177, 175 and 173. Interestingly, Pro190 is also quite invariant
across species. This figure was generated using Geneious (Drummond et al.,
http://www.geneioius.com).
Why would a “floppy” region have such ubiquitous conservation? I find that this region of the I
domain participates in substrate recognition but is also required for coordinating ATP-hydrolysis
and substrate degradation. My results are consistent with a model in which the GYVG loops are
important for binding the N-terminal residues of Arc variants, whereas the I domain is important
for recognition of C-terminal residues. In chapter four, I suggest future lines of investigation. It is
my hope that continuing study of specificity and regulation in HslUV will lead to a better overall
understanding of AAA+ enzyme function in biological systems.
35 Chapter Two:
Multiple Sequence Signals Direct Recognition and Degradation of
Protein Substrates by the AAA+ Protease HslUV
This chapter is published:
Sundar, S., McGinness, K.E., Baker, T.A., Sauer, R.T. (2010).
Journal of Molecular Biology. 403, 420-429.
Kathleen McGinness initiated some of the studies reported here, but I performed all of the
experiments in this chapter.
36 Abstract
Proteolysis is important for protein quality control and for the proper regulation of many
intracellular processes in prokaryotes and eukaryotes. Discerning substrates from other cellular
proteins is a key aspect of proteolytic function. The E. coli HslUV protease is a member of a
major family of ATP-dependent AAA+ degradation machines. HslU hexamers recognize and
unfold native protein substrates and then translocate the polypeptide into the degradation
chamber of the HslV peptidase. Although a wealth of structural information is available for this
system, relatively little is known about mechanisms of substrate recognition. Here, we
demonstrate that mutations in the unstructured N-terminal and C-terminal sequences of two
model substrates alter HslUV recognition and degradation kinetics, including changes in Vmax.
By introducing N- or C-terminal sequences that serve as recognition sites for specific peptidebinding proteins, we also show that blocking either terminus of the substrate interferes with
HslUV degradation, with synergistic effects when both termini are obstructed. These results
support a model in which one terminus of the substrate is tethered to the protease and the other
terminus is engaged by the translocation/unfolding machinery in the HslU pore. Thus,
degradation appears to consist of discrete steps, which involve the interaction of different
terminal sequence signals in the substrate with different receptor sites in the HslUV protease.
37 Introduction
AAA+ proteases are responsible for quality-control surveillance of the proteome and for
regulating key intracellular processes in all kingdoms of life (Gottesman, 2003; Striebel et al.,
2009). These ATP-fueled molecular machines recognize specific substrates, forcibly denature
these proteins, and then spool the denatured polypeptide in a processive fashion through an axial
channel and into a sequestered chamber for degradation (Inobe & Matouschek, 2008). In the
HslUV protease, for example, ring hexamers of HslU perform the recognition, unfolding, and
translocation functions, whereas the double-ring HslV dodecamer degrades polypeptides that are
translocated into its proteolytic chamber (Bochtler et al., 2000; Song et al., 2000; Sousa et al.,
2000; Wang et al., 2001). Recent studies have shown that HslUV is an extremely powerful
protein unfoldase when degrading proteins in the N-terminal to C-terminal direction but has
modest denaturation activity when degradation initiates at the C-terminus of a substrate
(Koodathingal et al., 2009).
For all proteases, substrate recognition is critical in ensuring that the correct molecules are
degraded. Exposed peptide sequences in specific substrates, called degrons or degradation tags,
typically serve as recognition signals for bacterial AAA+ proteases (Baker & Sauer, 2006; Inobe
& Matouschek, 2008). Peptide sequences can also target proteins for HslUV degradation (Kwon
et al., 2004; Burton et al., 2005; Koodathingal et al., 2009), but relatively little is known about
the underlying recognition events. Based on studies with related AAA+ proteases, it is clear that
some degrons are bound by pore loops in the central translocation channel of the hexameric
unfoldase, allowing these enzymes to grasp the attached substrate and to begin translocationmediated unfolding (Siddiqui et al., 2004; Hinnerwisch et al., 2005a; Martin et al., 2008a). This
38 mechanism is also likely to be important for HslUV degradation, because mutations in the highly
conserved GYVG pore loop of HslU diminish or block degradation of specific substrates (Park et
al., 2005). Other degradation signals tether substrates to other parts of the protease, raising the
effective concentration of the substrate relative to the enzyme and facilitating recognition of a
weak degron elsewhere in the substrate. Indeed, there is growing evidence in both natural and
engineered substrates that the existence of multiple degradation signals can play important roles
in determining how well a protein is recognized and degraded by AAA+ proteases (GonciarzSwiatek et al., 1999; Neher et al., 2003; Mettert & Kiley, 2005; Hoskins & Wickner, 2006;
Abdelhakim et al., 2008; Davis et al., 2009; Schrader et al., 2009).
Although a handful of substrates for HslUV have been identified (Seong et al., 1999; Nishii &
Takahashi, 2003; Kwon et al., 2004; Burton et al., 2005; Park et al., 2005; Koodathingal et al.,
2009), relatively little is known about the ways in which sequence signals in these proteins
influence degradation. To explore this issue, we employ two model substrates, the Arc repressor
of phage P22 and the N-terminal domain of phage λ cI repressor. In solution, both proteins
unfold in seconds but refold in milliseconds and thus maintain metastable structures (Milla &
Sauer, 1994; Huang & Oas, 1995). As a consequence, unfolding of these substrates is not a
serious impediment to HslUV degradation. We find that mutations in the unstructured Nterminal or C-terminal sequences of both proteins alter the steady-state kinetics of HslUV
proteolysis, including changes in the maximal rate of degradation. By introducing N- or Cterminal sequences that serve as recognition sites for specific peptide-binding proteins, we also
demonstrate that blocking either substrate terminus by protein binding interferes with HslUV
degradation. Our results support a model in which one terminus of the substrate is tethered to the
39 protease and the other terminus is engaged by the translocation/unfolding machinery in the HslU
pore. This model explains how mutations in either N- or C-terminal sequence can affect the
maximal degradation rate and suggests that substrate recognition by HslUV consists of discrete
steps, with the efficiency of degradation ultimately depending on the interplay of multiple
sequence signals in the substrate with appropriate receptor sites in the protease.
Results
Effects of terminal λcIN sequences on HslUV degradation
By screening a library in which five unstructured C-terminal residues of the N-terminal domain
of λ cI repressor (residues 1-102; λcIN) had been randomized, Parsell et al. (1990) identified
variants that were degraded rapidly in vivo. To determine if some of these proteins were
substrates for Escherichia coli HslUV, we recloned the parental domain and several variants to
add an N-terminal MGS2H6S3H tag (called ext1), purified these proteins, and assayed HslUV
degradation by SDS-PAGE. We identified one variant in which the C-terminal sequence was
changed from RSEYE to ISVTL (ext1-λcIN-ISVTL) that HslUV degraded faster than the
parental protein (ext1-λcIN-RSEYE; Fig. 2.1A), suggesting that the altered C-terminal residues
of this mutant make it a better substrate.
Although substrate degradation by HslUV is a multi-step reaction, simple Michaelis-Menten
kinetics are observed (Kwon et al., 2004; Burton et al., 2005), indicating that the overall reaction
can be modeled by substrate binding and dissociation steps, and by a rate-limiting enzymatic step
(kdeg), which could correspond to substrate engagement, unfolding, or translocation. Because
degradation signals usually affect protease binding, we expected that the ISVTL sequence would
40 reduce KM for HslUV degradation when compared to degradation of the wild-type RSEYE
sequence. Indeed, when we determined steady-state kinetic parameters for degradation of
35
S-
labeled proteins, a slight reduction in KM was observed (Fig. 2.1B; Table 2.1). Surprisingly,
however, the major effect on degradation of the ISVTL substrate was a ~2-fold increase in Vmax
(Fig. 2.1B; Table 2.1).
For inhibition experiments presented below, we changed the N-terminal residue of mature λcIN
from serine to leucine and/or changed the C-terminal sequence from RSEYE to RSYYF.
Determination of the kinetic parameters for HslUV degradation of the
S1L
S1L
λcIN-RSYYF and
λcIN-RSEYE variants revealed a ~2-fold difference in Vmax and ~3-fold difference in KM (Fig.
2.1C; Table 2.1). Because this pair of substrates differs only at the final and antepenultimate
amino acids, these C-terminal residues apparently affect both HslUV binding and the maximum
velocity of degradation. λcIN substrates with the same C-terminal sequences but different Ntermini sequences were degraded by HslUV with similar kinetic parameters (Table 2.1).
We used another λcIN variant to test if dimerization affected HslUV degradation. Monomers and
dimers of λcIN equilibrate rapidly in solution (KD ~300 µM), but the I84S mutation reduces
dimerization to undetectable levels (Weiss et al., 1987). We introduced the I84S mutation into
ext1-λcIN-ISVTL and determined kinetic parameters for HslUV degradation, which were within
error of the parental protein (ext1-λcIN-ISVTL; Table 2.1). We conclude that λcIN dimerization
is not a prerequisite for HslUV degradation.
41 Effects of terminal Arc sequences on degradation
Arc repressor and variants, including those with C-terminal sequences of H6KNQHD (st11 tag)
and/or AANDENYALAA (ssrA tag), are substrates for the E. coli and Haemophilus influenzae
HslUV enzymes (Kwon et al., 2004; Burton et al., 2005). Using E. coli HslUV, we determined
steady-state degradation parameters for Arc variants with a C-terminal ssrA tag (Arc-ssrA), the
st11 and ssrA tags (Arc-st11-ssrA), or st11 and a mutant ssrA tag in which the terminal AA
sequence was replaced by DD (Arc-st11-ssrADD) (Fig. 2.2; Table 2.1). KM for degradation of
these substrates varied over a ~2-fold range (Table 2.1; Fig. 2.2). Importantly, HslUV degraded
Arc-ssrA and Arc-st11-ssrA with Vmax values of 4.9 and 12 min-1 enz-1, respectively. Thus,
inserting the st11 tag before the ssrA tag increased the maximum degradation rate more than 2fold. Changing the C-terminal residues of Arc-st11-ssrA from AA to DD reduced Vmax from 12
to 7 min-1 enz-1. Hence, the C-terminal sequences of both λcIN and Arc substrates influence
binding as well as the maximal rate of HslUV degradation.
Previous studies demonstrated that N-terminal residues of Arc were important for HslUV
degradation (Burton et al., 2005). Consistently, we found that addition of an 18-residue Nterminal extension (ext2; LA3SGAG2SEG2TSGAT) to Arc-ssrA reduced Vmax and increased KM
(Fig. 2.2; Table 2.1). Interestingly, extending the C-terminal region of Arc-ssrA by insertion of
the st11 sequence increased Vmax, whereas extending the N-terminal region decreased Vmax.
Thus, degradation efficiency is not a simple function of the total number of unstructured residues
at the ends of a substrate.
42 Terminal sequences do not alter substrate stability
Sequences at the termini of λcIN and Arc substrates might influence the kinetics of HslUV
degradation by altering protein thermodynamic stability. However, no substantial differences in
the melting temperatures of different λcIN or Arc substrates were observed when thermal
denaturation was monitored by changes in circular-dichroism ellipticity (Table 2.1). These
results support a model in which interactions between the terminal sequences of these substrates
and HslUV are responsible for the observed changes in degradation kinetics.
Protein binding to the C-terminus of substrates slows HslUV degradation
As noted above, λcIN variants were engineered to contain a C-terminal YYF sequence, which can
be bound with sub-µM affinity by the PDZ domain of E. coli DegS (Walsh et al., 2003; Sohn &
Sauer, 2009). Saturating concentrations of the DegS PDZ domain inhibited HslUV degradation
of
S1L
λcIN-RSYYF to ~10% of the uninhibited rate but inhibited degradation of the
S1L
λcIN-
RSEYE control protein only marginally (Fig. 2.3A). Thus, inhibition by the PDZ domain is
specific and depends on the C-terminal sequence of the substrate.
The E. coli SspB protein binds to the AANDENY segment of the ssrA tag with sub-µM affinity
(Levchenko et al., 2000; Flynn et al., 2001). At saturating concentrations, SspB slowed HslUV
degradation of
M1L
Arc-ssrA to ~15% of the uninhibited value (Fig. 2.3B). SspB addition had a
much smaller effect on degradation of an otherwise identical protein with a mutant ssrAN3A tag
that binds SspB poorly (Levchenko et al., 2000). Again, these results show that SspB inhibits
HslUV degradation in a substrate-specific fashion. Thus, blockade of the C-terminal sequences
of λcIN and Arc substrates by protein binding prevents efficient HslUV degradation.
43 ClpS binding to the N-terminus of substrates inhibits HslUV degradation
E. coli ClpS binds with µM affinity to proteins with an N-terminal leucine, phenylalanine,
tyrosine, or tryptophan but binds substantially more weakly to proteins with N-terminal
methionine or other amino acids (Erbse et al., 2006; Wang et al., 2007; Roman-Hernandez et al.,
2009). To test if ClpS blockade of the N-terminus of λcIN or Arc influenced HslUV degradation,
we engineered variants of these substrates containing N-terminal leucines. Addition of ClpS to
an Arc variant with an N-terminal leucine (M1LArc-ssrA) slowed HslUV degradation to ~20% of
the uninhibited rate at saturation but did not inhibit degradation of an otherwise identical
substrate with an N-terminal methionine (Fig. 2.4A). Thus, ClpS binding to the N-terminal
leucine of M1LArc-ssrA is responsible for slowing HslUV degradation.
We also observed strong ClpS inhibition of HslUV degradation of
somewhat weaker inhibition of
S1L
S1L
λcIN-RSEYE and
λcIN-RSYYF (Fig. 2.4B). This difference may reflect the
lower KM for HslUV degradation of the latter substrate (Table 2.1).
Synergistic effects of N- and C-terminal blocking
If protein binding to the N- and C-terminus of substrates inhibits HslUV degradation by
independent mechanisms, then blocking both termini should produce greater inhibition than
blocking just one terminus. Indeed, inhibition of HslUV degradation of
M1L
Arc-ssrA by the
combination of SspB and ClpS was greater than inhibition by either protein alone (Fig. 2.5A,
top). Similarly, inhibition of HslUV degradation of
S1L
λcIN-RSYYF was more severe in the
presence of ClpS and the DegS PDZ domain than with either single protein (Fig. 2.5A, bottom).
44 Single-turnover inhibition
In principle, protein binding to the N- or C-terminal sequences of substrates could block
substrate sequences required for HslUV binding or engagement. Alternatively, the protein-bound
substrate could be recognized, engaged, unfolded, and translocated like the free substrate, but
steady-state degradation could be slowed because of the inability of the enzyme to initiate the
next cycle of degradation. The latter possibility could occur, for example, if the protein that was
bound at the terminus (ClpS, SspB, or the DegS PDZ domain) had to dissociate or be
enzymatically stripped in order to complete the last steps of degradation. By this model, the
initial release of
35
S-peptides from a substrate would occur faster than subsequent steps, and
inhibition by bound protein might not be evident if enzyme were present in excess of substrate.
Thus, we assayed single-turnover degradation of low concentrations of
35
S-labeled substrate in
the presence of excess enzyme and inhibitor. Under these conditions, specific inhibition by ClpS,
SspB, and the DegS PDZ domain was still observed (Fig. 2.5B), suggesting that inhibition
affects an early step in degradation.
Discussion
The simplest mechanism of targeting substrates for degradation by specific AAA+ proteases is
exemplified by the ssrA-tagging system, in which a short peptide signal targets a wide variety of
proteins for ClpXP or ClpAP degradation (Keiler et al., 1996; Gottesman et al., 1998). It is
becoming evident, however, that substrate recognition mediated by a single tag or degron is
generally the exception rather than the rule. Indeed, one substrate signal often mediates binding,
whereas another signal ensures engagement by the translocation machinery of the AAA+
protease (Inobe & Matouschek, 2008). The results presented here show that sequences at both
45 the N- and C-terminal ends of Arc and λcIN substrates influence the rate of HslUV degradation
and thus support a multi-degron model of recognition.
Unfolded monomers and native dimers of Arc equilibrate rapidly in solution, but the strong
dimerization constant (~10 nM) ensures that most molecules are dimers at the concentrations
used for degradation studies (Bowie & Sauer, 1989; Milla & Sauer, 1994). Thus, each Arc
substrate has two N-terminal sequences and two C-terminal sequences that potentially could
interact with HslU. By contrast, the dimerization constant of λcIN is substantially weaker (~300
µM) (Weiss et al., 1987), and a mixture of monomers and dimers are present in our degradation
assays. Importantly, however, a mutation (I84S) that weakens λcIN dimerization >30-fold did not
cause significant changes in the steady-state kinetics of λcIN degradation (Table 2.1), suggesting
that HslUV recognizes monomers and dimers of λcIN comparably.
HslUV degradation of Arc variants was initially suggested to depend exclusively on C-terminal
sequences and was subsequently proposed to depend completely on N-terminal sequences
(Kwon et al., 2004; Burton et al., 2005). Our present studies resolve this apparent conflict, as
mutational and inhibitor experiments demonstrate that both termini of Arc substrates need to be
recognized for efficient HslUV degradation. Similar results with λcIN variants suggest that multidegron recognition may be common for HslUV substrates. The terminal sequences of both Arc
and λcIN variants are unstructured in the free proteins and could mediate binding/engagement by
the translocation machinery of the HslU pore or tethering to an independent receptor site
elsewhere in the enzyme. At present, we do not know which termini serve which function or
46 even if one specific terminus always binds the pore while the other always serves a tethering
role, but the results presented here should allow future experiments to address this question.
Fig. 2.6 shows a model with three types of HslUV•substrate complexes and provides a
framework for understanding how sequence changes at either terminus of a substrate can alter
KM and Vmax for degradation. In complex P, one terminus of the substrate binds to the enzyme
pore. In complex T, the other terminus of the substrate binds to a distinct tethering site. In
complex TP, both termini of the substrate bind to their respective sites in HslUV. Only
complexes P and TP are proteolytically active, because binding to the pore is required for
degradation. The distribution of these species depends on the equilibrium constants for binding
the tethering site in a bimolecular reaction (K1), binding the pore in a bimolecular reaction (K2),
binding the pore in a unimolecular reaction (K3), and binding the tethering site in a unimolecular
reaction (K4). Because K1K3 = K2K4, specifying three of these constants determines the fourth
constant. Thus, the apparent KM and the fraction of active enzymes at substrate saturation (fact)
can be expressed as a function of just K1, K2, and K3 (Fig. 2.6). In principle, changing the
sequence at either terminus could alter KM and the maximal rate of degradation (fact•kdeg). In
practice, however, substantial sequence-dependent changes in either kinetic parameter would
only be observed if the T complex is present in excess of the P and TP complexes, which
requires stronger bimolecular binding to the tethering site than to the pore site (K1 < K2) and a K3
value > 1. Under these conditions, mutations at either substrate terminus could alter both KM and
Vmax by affecting binding to one of the enzyme sites (altering K1 or K2) and might also change
these parameters by altering the length or flexibility of the terminal sequences (affecting K3).
Sequence changes at the substrate terminus that binds the pore of HslUV could also directly
47 affect the efficiency of engagement by the translocation machinery and thus alter Vmax by
changing kdeg as well as fact. Studies of adaptor-mediated delivery of substrates to the ClpXP
protease provide general precedents for changing the steady-state kinetics of degradation via
alterations in tethering sequences, pore-contact sequences, and the flexibility and geometry of
intervening domains (McGinness et al., 2007; Davis et al., 2009).
From a biological perspective, multi-degron degradation permits combinatorial control of
proteolysis through masking of one or both degrons in a substrate, as demonstrated by our
observation that protein binding to the N- and C-terminal sequences of Arc and λcIN substrates
inhibits HslUV degradation. Such effects can be readily rationalized in terms of the model of
Fig. 2.6, as substrates with proteins bound to their terminal sequences would be expected to have
different K1, K2, and/or K3 values. Many proteins are multimeric, and it is straightforward to
envision regulatory strategies in which two degrons are properly displayed only in a monomer or
only in a multimer, allowing just one species to be targeted for degradation (Baker & Sauer,
2006). Degrons could also be masked by a protein’s native structure and only be revealed when
the protein unfolds at high temperatures. In this regard, we note that HslUV is a heat-shock
protease and that degradation signals in the interior of protein sequences have been shown to act
as recognition signals for other AAA+ proteases (Chuang & Blattner, 1993; Hoskins & Wickner,
2006; Gur & Sauer, 2008a; Schrader et al., 2009).
AAA+ proteases are allosteric machines that undergo continual cycles of ATP-powered
conformational changes, even in the absence of substrate. As a consequence, it may be more
difficult for the translocation machinery to bind and engage a degron in a single concerted step,
48 but easier if the substrate is first tethered to the protease and engagement occurs subsequently.
The latter situation could be viewed analogously to mid-air refueling, in which the speeds and
general orientations of two airplanes are matched in one step and the fuel-transfer machinery is
engaged in a subsequent step. We propose that HslUV carries out degradation in a similar multistep manner.
Materials & Methods
Protein expression and purification
Genes encoding protein substrates were constructed by standard methods of molecular biology
using the polymerase-chain reaction and/or site-directed mutagenesis. λcIN substrates contained
varied N-terminal sequences, a common core corresponding to residues 2-97 of λ cI repressor,
and C-terminal sequences of RSEYE (residues 98-102 of λ cI repressor), RSYYF, or ISVTL.
The initiator methionine of wild-type λ repressor is removed post-translationally (Sauer &
Anderegg, 1978), leaving Ser1 as the N-terminal residue. Variants designated ext1-λcIN had an
additional N-terminal extension (MGS2H6S3H; glycine at N-terminus after processing of initiator
methionine), which was derived from the pACYCDuet-1 expression vector (Novagen). Variants
designated S1LλcIN were initially cloned, expressed, and purified as fusions to an N-terminal H6tagged SUMO domain in a pET23b vector (Novagen), and the fusion protein was then purified
and cleaved with Ulp1 protease to generate the mature substrate with Leu1 at the N-terminus
(Malakhov et al., 2004). Arc substrates contained variable N-terminal sequences, a common core
consisting of residues 2-53 of P22 Arc repressor, and C-terminal sequences including the st11
sequence (H6KNQHD), ssrA tag (AANDENYALAA), ssrADD tag (AANDENYALDD), or
ssrAN3A tag (AAADENYALAA). The wild-type N-terminus of Arc is Met1 (Vershon et al.,
49 1985). M1LArc variants were expressed as H6-SUMO fusions from pET23b, purified, and cleaved
to generate Leu1 at the N-terminus. Substrates designated ext2-Arc contained the N-terminal
extension LA3SGAG2SEG2TSGAT and were also purified as SUMO fusions and processed by
Ulp1 cleavage.
H6-tagged variants of E. coli HslU6 and HslV12 were purified essentially as described (Burton et
al., 2005), except buffer B (pH 8.0) contained 50 mM NaHPO4, 300 mM NaCl, and 250 mM
imidazole, and buffer C (pH 7.5) contained 20 mM Tris-HCl, 100 mM NaCl, 1 mM EDTA, and
10% (v/v) glycerol. Pooled HslU6 or HslV12 fractions after Ni++-NTA chromatography were
chromatographed on a Sephacryl-300HR26/60 gel filtration column (GE Healthcare) and then on
an HR16/10 Q-Sepharose column (GE Healthcare), developed with a gradient from 0.1 M to 1 M
NaCl in buffer C. Appropriate fractions were pooled, concentrated, and exchanged into buffer B
using Amicon Ultra-15 centrifugation filters.
Arc and λcIN proteins were expressed in E. coli strain X90 (λDE3). Cells were grown at 37 °C to
an OD600 of ~0.8, and shifted to room temperature. Expression was induced by the addition of 1
mM IPTG, and cells were harvested 4 h later, lysed by sonication, and cleared lysates were
applied to a Ni++-NTA column as described for the HslU6 and HslV12 purifications. The Arcst11-ssrA, Arc-st11-ssrADD, ext1-λcIN-ISVTL and ext1-λcIN-RSEYE proteins were subjected to
additional purification on a HiLoad 16/60 Superdex-100 gel-filtration column (GE Healthcare)
equilibrated in buffer D (20 mM Tris (pH 7.5), 200 mM NaCl, 1 mM EDTA). Appropriate
fractions were pooled and concentrated using Amicon Ultra-15 centrifugation filters. Following
the Ni++-NTA step, substrates expressed as H6-SUMO fusions were exchanged into 50 mM
50 NaHPO4 (pH 8.0), 100 mM NaCl, and 20 mM imidazole and cleaved with Ulp1 protease (a gift
from J. Davis and J. Kaplan, MIT) at 37 °C overnight. Cleaved samples were re-applied to a
Ni++-NTA column, and the flow-through fraction, which contained the desired product, was
pooled and chromatographed on a HiLoad 16/60 Superdex-100 gel-filtration column (GE
Healthcare) equilibrated in buffer D. Appropriate fractions were pooled and concentrated using
Amicon Ultra-15 centrifugation filters.
35
S-substrates were expressed and purified as described
(Burton et al., 2005), with additional steps of Ulp1 cleavage, Ni++-NTA re-purification, and S100
purification for substrates expressed as H6-SUMO fusions.
An E. coli SspB expression clone was a gift from Igor Levchenko (MIT) and an E. coli ClpS
expression clone (pET23b-H6-SUMO-ClpS) was a gift from G. Román-Hernández (MIT). SspB
was purified as described (Levchenko et al., 2000). ClpS was purified by the protocol used for all
SUMO-fusion proteins. The purified E. coli DegS PDZ domain was a gift from Jungsan Sohn
(MIT). Protein concentrations were determined by absorbance at 280 nm using extinction
coefficients calculated from the amino-acid sequence.
Degradation assays
For degradation assays monitored by SDS-PAGE, substrates (10 µM) were incubated at 37 °C
for different times with 300 nM E. coli HslU6, 800 nM E. coli HslV12, and an ATP regeneration
system (5 mM ATP, 16 mM creatine phosphate, 10 µg/mL creatine kinase). Steady-state HslUV
degradation of
35
S-labeled proteins at 37 °C was carried out as described (Burton et al., 2005)
using 100 nM HslU6, 300 nM HslV12, and the ATP regeneration system. Aliquots were quenched
at appropriate time points by adding ice-cold trichloroacetic acid (TCA) to a final concentration
51 of 10% (w/v) and were separated into pellet and supernatant fractions by centrifugation.
Radioactivity in the acid-soluble fraction was used as a measure of degradation. For experiments
probing the inhibitory effects of SspB, ClpS, or the DegS PDZ domain, each protein was mixed
with 35S-labeled substrate for 5 min at 37 °C before addition of 500 nM HslU6, 1.5 nM HslV12,
and the ATP regeneration mix. At different times, degradation was assayed by acid-soluble
radioactivity as described above. For degradation of
35
S-labeled proteins under single turnover
conditions, each reaction contained 2 µM substrate, 5 µM HslU6, 6 µM HslV12, and the ATP
regeneration system, and degradation was assayed as described above for steady-state kinetic
experiments.
Thermal denaturation
Denaturation experiments were monitored by changes in circular-dichroism ellipticity at 222 nm
using an AVIV Model 60DS spectrometer. Proteins (2 µM) in 10 mM KPO4 (pH 6.8), 200 mM
NaCl were heated from 25 to 95 °C in 2 °C steps, the sample was allowed to equilibrate for 1
min at each temperature, and the ellipticity was averaged for 30 s and recorded. Melting data
were fit to a two-state model in which the intercepts and slopes of the native and denatured
baselines, TM, and ΔHU were determined by a non-linear least squares fitting protocol.
Acknowledgements. We thank J. Davis, J. Kaplan, I. Levchenko, G. Román-Hernández, and J.
Sohn for materials and helpful discussion. T.A.B. is an employee of the Howard Hughes Medical
Institute. Supported by NIH grant AI-16892.
52 Figures & Tables
Table 2.1 – Properties of λcIN and Arc substrates.
epitopes for binding to
SspB
DegS
ClpS
PDZ
substrates
Vmax
(min-1)
KM
(µM)
Vmax / KM
min-1µM-1
Tm
(°C)
ext1-λcIN-RSEYE
ext1-λcIN-ISVTL
ext1-λcIN(I84S)-ISVTL
S1L
λcIN-RSEYE
S1L
λcIN-RSYYF
λcIN-RSYYF
4.3 ± 0.1
9.9 ± 1.1
9.9
3.5 ± 0.3
7.1 ± 1.0
5.7 ± 0.6
120 ± 19
89 ± 12
79
120 ± 24
40 ± 5.0
54 ± 6.7
0.04
0.11
0.13
0.03
0.18
0.11
56.7
56.1
50.5
56.5
55.5
54.5
no
no
no
no
no
no
no
no
no
no
yes
yes
no
no
no
yes
yes
no
Arc-ssrA
Arc-st11-ssrA
Arc-st11-ssrADD
ext2-Arc-ssrA
M1L
Arc-ssrA
M1L
Arc-ssrAN3A
4.9 ± 0.1
12 ± 1.9
7.0 ± 0.3
3.0 ± 0.5
4.7 ± 1.5
5.7 ± 0.5
24 ± 1.4
31 ± 0.9
51 ± 4.8
55 ± 1.3
21 ± 4.0
23 ± 7.7
0.20
0.38
0.14
0.06
0.22
0.25
58.6
56.5
59.3
58.2
56.9
56.3
yes
yes
yes
yes
yes
no
no
no
no
no
no
no
no
no
no
yes
yes
yes
Vmax values are expressed as turnover numbers (the degradation rate at substrate saturation
divided by the total enzyme concentration). Vmax and KM values are generally averages of two
2
determinations (error estimated as
∑ (value − mean)
2
). Values with no error are from a single
1
experiment.
€
53 Figure 2.1 – C-terminal λcIN sequences alter HslUV degradation. (A) Degradation of λcIN
variants (10 µM) bearing RSEYE or ISVTL C-terminal sequences by 300 nM HslU6, 800 nM
HslV12 was analyzed by SDS-PAGE. (B, C) Rates of steady-state degradation of different
concentrations of 35S-λcIN variants by 100 nM HslU6, 300 nM HslV12 were determined by
assaying acid-soluble radioactivity. Lines represent non-linear-least-squares fits to the MichaelisMenten equation: rate = Vmax•[S]/([S]+KM). Kinetic parameters are listed in Table 2.1.
54 Figure 2.2 – C- and N-terminal sequences of Arc substrates affect HslUV degradation.
Steady-state rates of HslUV degradation of 35S-Arc variants were determined as described in the
Fig. 2.1 legend. Lines are fits to the Michaelis-Menten equation; kinetic parameters are listed in
Table 2.1.
55 Figure 2.3 – Occluding the C-terminus of λcIN and Arc substrates inhibits HslUV
degradation. (A) Degradation of 35S-labeled λcIN substrates (50 µM) by 0.5 µM HslU6, 1.5 µM
HslV12 was measured at different concentrations of the DegS PDZ domain, which binds strongly
to the C-terminal tripeptide YYF but weakly to EYE. Rates are expressed as a percentage of the
rate with no PDZ domain. (B) Degradation of 35S-labeled Arc variants (4 µM) by 0.5 µM HslU6,
1.5 µM HslV12 was measured at different concentrations of SspB, which binds strongly to the
wild-type ssrA tag but weakly to the ssrAN3A mutant. Rates are expressed as a percentage of the
rate with no SspB.
56 Figure 2.4 – ClpS binding to the N-terminus of Arc or λcIN substrates slows HslUV
degradation. (A) Proteolysis of 35S-labeled Arc variants (4 µM) by 0.5 µM HslU6, 1.5 µM
HslV12 was measured at different concentrations of ClpS, which binds strongly to the N-terminal
leucine of M1LArc-ssrA but weakly to the N-terminal methionine of Arc-ssrA. (B) ClpS inhibits
HslUV (0.5 µM HslU6, 1.5 µM HslV12) degradation of λcIN substrates (50 µM) with N-terminal
leucines (S1LλcIN-RSYYF – diamonds; S1LλcIN-RSEYE – circles) more efficiently than variants
with an N-terminal serine (λcIN-RSYYF – triangles) or glycine (ext1-λcIN-RSEYE – squares).
57 Figure 2.5 – Combinatorial and single-turnover inhibition. (A) (top) A combination of SspB
and ClpS inhibited degradation of 35S-M1LArc-ssrA (15 µM) by 0.5 µM HslU6, 1.5 µM HslV12 to
a greater extent than either individual protein. (bottom) A combination of the DegS PDZ domain
and ClpS also inhibited degradation of 35S-S1LλcIN-RSYYF (50 µM) by 0.5 µM HslU6, 1.5 µM
HslV12 more than either single protein alone. (B) Degradation of different 35S-labeled substrates
(2 µM) by 5 µM HslU6, 6 µM HslV12 was assayed in the presence the indicated concentrations of
SspB, ClpS, or the DegS PDZ domain.
58 Figure 2.6 – Model of HslUV recognition of a substrate (S) with degrons at each terminus.
One degron (shown as a triangle) binds to a tethering site on the HslUV enzyme (E). The other
degron (shown as a circle) binds to a site in the translocation pore of HslU. Because binding to
the pore is a prerequisite for degradation, the enzyme•substrate complexes marked P and TP are
proteolytically active but the T complex is inactive. K1 ([E][S]/[T]), K2 ([E][S]/[P]), K3
([T]/[TP]), and K4 ([P]/[TP]) are equilibrium dissociation constants. Note that K1•K3 = K2•K4. If
the rate of substrate dissociation is fast compared to kdeg, then the apparent KM ([S] at half
maximal velocity) is 1/(1/K1+1/K2+1/(K1•K3)). At substrate saturation, the fraction of active
enzymes (fact) = ([P]+[TP])/([T]+[P]+[TP]) = (1/K2+1/(K1•K3)/(1/K1+1/K2+1/(K1•K3)).
Vmax/[Etotal] equals fact•kdeg. The K3-kdeg pathway of degradation corresponds to single-degron
recognition.
59 Chapter Three:
The Intermediate Domain of the AAA+ HslUV Protease
Coordinates Substrate Binding and Stimulation of ATP Hydrolysis
60 Abstract
AAA+ proteases carry out regulated protein degradation in all organisms. In the HslUV protease,
HslU hexamers bind substrates, engage a segment of the protein in an axial pore, unfold the
native structure, and then translocate the polypeptide through the pore and into a chamber of
HslV for degradation. Most of these activities require ATP binding and/or hydrolysis by HslU.
Here, we characterize HslU interactions with an Arc protein substrate that contains degrons at
both termini. Mutational studies suggest that the N-terminal degron binds in the axial pore of
HslU, with important contacts mediated by a conserved tyrosine in the highly conserved GYVG
loop. As expected from this model, HslUV degradation of an Arc-GFP fusion protein begins at
the N-terminus. The 175-209 loop in the intermediate (I) domain of HslU is also important for
substrate degradation. Deletion of this disordered loop weakens substrate affinity, suggesting a
role in recognition, but also decreases the maximal degradation rate. The latter defect results
from a lower maximal rate of ATP hydrolysis. Indeed, deletion of the entire I domain almost
eliminates ATP hydrolysis by HslU, and an intact 175-209 loop is essential for substratemediated stimulation of ATP hydrolysis. Thus, the I domain and 175-209 loop play roles in
tethering substrates to HslU and in allowing rapid rates of ATP hydrolysis.
61 Introduction
Energy-dependent proteolysis is a key process in sculpting the proteomes of cells from all
kingdoms of life. Degradation can clear damaged or misfolded proteins, remove superfluous or
unneeded proteins following a shift in growth conditions or developmental programs, and play
important roles in regulatory circuits that drive the cell cycle or mediate transcriptional responses
to environmental stress (Gottesman, 2003; Baker & Sauer, 2006). The ATP-fueled proteases that
execute these processes must be highly specific to avoid degradation of essential proteins. They
also provide paradigms for a wide range of molecular machines that perform mechanical tasks in
intracellular settings.
Escherichia coli contains five ATP-dependent proteases: HslUV, ClpXP, ClpAP, Lon, and FtsH
(Baker & Sauer, 2006). Related proteases, including the 26S proteasome, are found in most
eubacteria and archaebacteria, in mitochondria and chloroplasts, and in the cytoplasm of
eukaryotic cells (Striebel et al., 2009). Each of these multi-subunit proteases contains a
hexameric AAA+ enzyme, which functions to recognize, unfold, and translocate specific target
proteins into the degradation chamber of a compartmental peptidase. In HslUV, for example, the
HslU hexamer serves as the AAA+ protein unfoldase/translocase, whereas HslV forms a doublering dodecamer that encloses the proteolytic compartment. Like its AAA+ relatives, HslU
contains large and small AAA+ domains that couple ATP binding and hydrolysis to the
conformational changes that drive substrate unfolding and translocation. These mechanical
processes are thought to occur via ATP-powered movements of loops that project into the axial
pore of the hexamer. In HslU, ClpX, and ClpA, these loops contain a highly conserved GYVG
sequence, which appears to contact some substrates and to play roles in translocation and
62 unfolding (Siddiqui et al., 2004; Park et al., 2005; Hinnerwisch et al., 2005a; Martin et al.,
2008a; 2008b).
Substrates are typically targeted to specific AAA+ proteases by peptide sequences (Baker &
Sauer, 2006). For example, the ssrA-tag sequence binds in the axial pores of ClpX and ClpA,
resulting in degradation of ssrA-tagged substrates by ClpXP or ClpAP (Gottesman et al., 1998;
Siddiqui et al., 2004; Hinnerwisch et al., 2005a; Martin et al., 2008a; 2008b). Other peptide
sequences tether substrates or adaptor proteins to family specific auxiliary domains in the AAA+
hexamer. For example, the N domain of ClpX binds to a tethering sequence in the UmuD/D´
protein, helping to mediate ClpXP degradation of this error-prone polymerase (Neher et al.,
2003b). Our understanding of substrate recognition by HslUV is rudimentary. Studies of a
handful of natural or model HslUV substrates show that peptide sequences are important
determinants of targeting for HslUV degradation (Seong et al., 1999; Kwon et al., 2003; 2004;
Burton et al, 2005; Park et al., 2005; Koodathingal et al., 2009; Sundar et al., 2010).
Nevertheless, how peptide signals in substrates interact with HslU to control degradation is
poorly understood.
Numerous crystal structures of HslU and HslUV have been solved, including the first views of a
AAA+ ring unfoldase in complex with its compartmental peptidase (Appendix A; Bochtler et al.,
2000; Sousa et al., 2000; Wang et al., 2001; Sousa et al., 2002; Kwon et al., 2003). Packing
between the highly conserved large and small AAA+ domains of neighboring subunits stabilizes
the HslU hexameric ring. In addition, an intermediate (I) domain, which is only found in the
HslU family, is inserted between two neighboring helices of the large AAA+ domain. One of
63 these helices follows the GYVG loop and the other precedes the Walker-B motif, which plays
important roles in ATP hydrolysis. The I domains project upward from the top surface of the
HslU ring, forming a bowl or funnel shaped cavity above the axial pore.
In chapter two, we characterized a model substrate, Arc-st11-ssrA, in which N-terminal and Cterminal degrons appeared to interact with different sites in HslU (Sundar et al., 2010). Arc is a
dimeric phage repressor (Breg et al., 1990; Raumann et al., 1994). The N-terminal residues of
Arc are disordered in solution and are known to be important for degradation by HslUV; the
unstructured C-terminal st11-ssrA sequence also enhances proteolysis, in part by increasing the
maximal rate of degradation (Burton et al., 2005; Sundar et al., 2010). Here, we probe the
interaction of Arc-st11-ssrA with HslU variants bearing mutations in the GYVG pore loop or the
I domain. Our results support a model in which the GYVG loop in the axial pore binds the Nterminal degron of the substrate, whereas the disordered 175-209 loop in the I domain tethers
another part of the substrate, possibly the C-terminal degron. Surprisingly, we discovered that
deleting most of the I domain reduced the basal level of ATP hydrolysis at least 40-fold, a level
too slow to support normal HslUV function. When we replaced the 175-209 loop in the I domain
with a flexible linker, the resulting mutant was defective in degradation of Arc-st11-ssrA because
of reduced binding to the substrate, an inability of substrates to stimulate ATP hydrolysis, and
therefore reduced levels of ATP hydrolysis. Thus, the I domain plays an active role in substrate
binding as well as in coordinating ATP hydrolysis and substrate processing.
64 Results
Effects of HslU pore-loop mutations on substrate binding and degradation
ATP-dependent unfoldases contain highly conserved axial-pore loops, which appear to function
in substrate binding, unfolding, and translocation (Baker & Sauer, 2006; Martin et al., 2008a;
2008b). Park et al. (2005) showed that mutations in the GYVG pore loop of HslU (residues 9093) affected HslUV degradation of MBP-SulA. For example, the Y91F mutant (GFVG) slowed
degradation as assayed by SDS-PAGE, whereas the Y91A mutant (GAVG) was essentially
inactive, although both mutants hydrolyzed ATP and stimulated the peptidase activity of HslV as
well as wild-type HslU.
To characterize the effects of the Y91F and Y91A substitutions more quantitatively, we purified
35
S-labeled Arc-st11-ssrA (Sundar et al., 2010) and monitored degradation of different
concentrations of this substrate by wild-type HslUV or by enzymes bearing the pore-loop
mutations (Fig. 3.1A; Table 3.1). At each substrate concentration, degradation by wild-type
HslUV was slightly faster than degradation by Y91F HslUV. The main difference in steady-state
kinetic parameters was a ~20% reduction in Vmax, with values of 11 ± 0.7 min-1 enz-1 for the
wild-type enzyme and 8.7 ± 0.1 min-1 enz-1 for Y91F; the KM values for the wild-type and mutant
enzymes were 20 µM and 26 µM, respectively. We observed no significant degradation of the
radiolabeled substrate by Y91A HslUV, emphasizing the importance of an aromatic side chain at
the second residue of the GYVG loop. As expected from the work of Park et al. (2005), we
found that purified Y91A HslU hydrolyzed ATP at a rate slightly faster than wild-type HslU and
stimulated HslV to comparable maximal levels of peptidase activity (Table 3.1). These results
65 show that the inactivity of Y91A HslU in degrading Arc-st11-ssrA is not a consequence of a
failure to form hexamers or defects in ATP hydrolysis or HslV interactions.
Is the very poor degradation observed for Y91A HslUV caused by defects in substrate binding,
engagement, unfolding, or translocation? To address this question, we labeled amino groups in
Arc-st11-ssrA with a fluorescent dye and used changes in fluorescence anisotropy to measure
binding of the modified protein to HslU or the pore-loop variants in the presence of ATPγS, an
ATP analog that HslU does not hydrolyze. As shown in Fig. 3.1B, wild-type HslU bound this
protein with an affinity of ~1.3 µM, the Y91F mutant bound with an affinity of ~20 µM, and the
Y91A variant showed little if any binding. KD for HslU binding to fluorescent Arc-st11-ssrA,
measured in the presence of ATPγS, was much lower than KM for ATP-dependent degradation of
unmodified Arc-st11-ssrA. The fluorescent modifications do not appear to be responsible for this
difference, as a competition experiment also demonstrated a low KD for binding of unmodified
Arc-st11-ssrA to HslU in the presence of ATPγS (data not shown). Another potential cause for
the large difference in KD and KM could be that HslUV binds substrate more weakly than HslU.
However, binding was actually enhanced in the presence of HslV (Fig. 3.1C). Wild-type HslUV
bound the fluorescent substrate with an affinity of 110 nM, and Y91F HslUV bound with an
affinity of 680 nM. These results suggest that HslV binding stabilizes a higher-affinity
conformation of the HslU hexamer, although it is also possible that there are some contacts
between the substrate and HslV. In any event, the Y91 mutations clearly reduce the affinity of
binding of HslU and HslUV to the Arc-st11-ssrA substrate. Other explanations for the observed
differences between KD and KM are considered in the Discussion.
66 We also measured binding of HslU and the pore-loop mutants to a fluorescein-labeled variant of
the gt1 peptide (MRYFFKKKLRFY), which was designed to mimic interactions normally made
by N-terminal residues of Arc substrates (Burton et al., 2005). The Y91A variant did not bind
gt1, the Y91F enzyme bound with ~3.8 µM affinity, and wild-type HslU bound with ~760 nM
affinity (Fig. 3.1D). In chapter two, we showed that both N-terminal and C-terminal sequences of
Arc-st11-ssrA were important for recognition and degradation by HslUV (Sundar et al., 2010).
The failure of Y91A HslU to show significant binding to either Arc-st11-ssrA or gt1 peptide
suggests that N-terminal residues of Arc substrates bind in the axial pore of HslU. In addition,
the reduced Y91F affinity for Arc-st11-ssrA and gt1 indicates that the side-chain hydroxyl group
of Tyr91 contributes to substrate recognition, whereas the dramatic defects of Y91A demonstrate
that the aromatic ring of Tyr91 plays an even more important role in substrate binding.
Degradation of an Arc fusion protein starts at the N-terminus
If the N terminal residues of Arc substrates initially bind in the axial pore of HslU, then
unfolding and translocation should begin at the N-terminus and proceed towards the C-terminus.
To test this model, we constructed a substrate with an N-terminal Arc domain, followed by GFP,
followed by the st11-ssrA sequence (Arc-GFP-st11-ssrA). As assayed by SDS-PAGE, HslUV
degradation of this substrate resulted in accumulation of a truncated product (Fig. 3.2A). We
excised the truncated product from the gel, digested it with trypsin, and characterized the
resulting peptides by liquid-chromatography and mass-spectrometry (LC-MS). Importantly, we
identified a tryptic peptide (NQHDAANDENYALAA) corresponding to the 15 C-terminal
residues of the st11-ssrA tag (Fig. 3.2B). This result demonstrates that the truncated product is a
C-terminal fragment of Arc-GFP-st11-ssrA. A western blot (Fig. 3.2C) and a Ni++ pull-down
67 experiment (data not shown) confirmed this result, as both the intact substrate and the truncated
fragment cross-reacted with an antibody against the His6-tag (found in the st11 sequence) and
both proteins bound Ni++-NTA resin. Finally, we observed no change in GFP fluorescence over
the time that the truncated degradation product was formed (data not shown). These results show
that HslUV degradation of Arc-GFP-st11-ssrA begins at the N-terminus but stalls when it
encounters GFP.
Previous studies showed that binding of the SspB protein to the ssrA tag of Arc substrates
slowed HslUV degradation (Sundar et al., 2010). The same result was observed for Arc-GFPst11-ssrA. Specifically, the presence of equimolar SspB slowed production of the truncated
product when substrate was in excess (Fig. 3.2D) and under single-turnover conditions (data not
shown), as expected if the C-terminal sequence of Arc-GFP-st11-ssrA participates in binding to
the protease. Overall, these results support a model in which N-terminal residues of Arc
substrates bind in the axial pore of HslU, whereas C-terminal residues interact with a tethering
site on the enzyme.
The intermediate domain of HslU plays a major role in determining ATP-hydrolysis rates
The I domain of HslU (residues 108-243) is required for degradation of some native substrates
(Song et al., 2000; Kwon et al., 2003). To probe I-domain function in greater detail, we
constructed and purified HslU variants in which residues 108-243 were replaced with a GG
dipeptide (∆108-243GG), residues 121-231 were replaced with a SGAGGTSGEGGS linker
(∆121-231linker), residues 175-209 were replaced with the same linker (∆175-209linker), or
residues 175-209 were replaced with GG (∆175-209GG; an enzyme originally characterized by
68 Song et al., 2000). The ∆175-209linker and ∆175-209GG proteins lack a loop that is disordered in
all HslU crystal structures. The ∆108-243GG and ∆121-231linker enzymes are missing most of the I
domain, but the latter variant retains the sequences that form parts of two α helices that extend
from the large AAA+ domain of HslU to form the I-domain junctions (Bochtler et al., 2000;
Song et al., 2000; Sousa et al., 2000).
As anticipated from prior studies (Song et al., 2000; Kwon et al., 2003), each of our I-domain
variants purified as a hexamer, and the mutants tested stimulated HslV peptidase activity
normally (Table 3.1). In comparison to wild-type HslU, however, each I-domain mutant had
reduced ATPase activity. For example, the ∆108-243GG and ∆121-231linker enzymes displayed
nearly 40-fold lower rates of ATP hydrolysis, the ∆175-209GG enzyme had ~9-fold reduced
ATP-hydrolysis activity, and the ∆175-209linker variant had ~2-fold lower ATPase activity (Fig.
3.3A; Table 3.1). These results were surprising because Song et al. (2000) reported that E. coli
HslU variants missing the entire I domain or the 175-209 loop had 80-100% of wild-type
ATPase activity. Unfortunately, rates, assay conditions, enzyme concentrations, or ATP
concentrations were not specified for their studies. To understand possible reasons for the
observed discrepancies, we tested the HslU dependence of ATP-hydrolysis rates for our Idomain variants, which scaled linearly with enzyme (Fig. 3.3B). Thus, our results cannot be
explained by effects of the mutations on dissociation of HslU hexamers. We also found that
HslV enhanced ATP turnover of the I-domain variants ∆175-209linker and ∆175-209GG, but only
in proportion to the lower activities observed with HslU alone (Fig. 3.3C). Thus, the differences
between our results and those of Song and colleagues are not caused by the presence/absence of
HslV. Finally, we considered the possibility that the low ATPase activities observed in our
69 assays might be caused by using an ATP concentration (2.5 mM) substantially below KM for
hydrolysis. However, when we assayed the ATP dependence of turnover for ∆175-209linker or
∆175-209GG HslU, the KM values were similar to (or tighter than) the wild-type value but Vmax
was reduced 2- to 9-fold (Fig. 3.3D; Table 3.1). Thus, defective nucleotide binding cannot
account for the low ATP-hydrolysis rates in our experiments. The ∆175-209GG mutant studied
here and by Song et al. (2000) have exactly the same amino-acid sequence, and thus the striking
differences between the two sets of results cannot be explained by differences in the I-domain
variants utilized for these studies. In combination, our results strongly suggest that the I domain
of HslU and the 175-209 loop of the I domain play very important roles in allowing the wildtype enzyme to hydrolyze ATP rapidly.
∆175-209linker HslUV degrades Arc-st11-ssrA with a lower Vmax and weaker KM
We chose ∆175-209linker HslU for studies of degradation, because this I-domain mutant had the
highest rate of ATP hydrolysis. In assays monitored by SDS-PAGE, ∆175-209linker HslUV
degraded Arc-st11-ssrA substantially more slowly than wild-type HslUV (Fig. 3.4, inset). To
determine the basis for slower proteolysis, we measured the steady-state kinetics of degradation
of different concentrations of Arc-st11-ssrA (Fig. 3.4). ∆175-209linker HslUV degraded Arc-st11ssrA with a KM of ~130 µM and a Vmax of ~3.5 min-1 enz-1. Thus, in comparison with wild-type
HslUV, the ∆175-209linker mutation slowed the maximal rate of Arc-st11-ssrA degradation ~4fold and increased KM ~7-fold (Table 3.1).
In principle, the observed decrease in Vmax could largely be explained by slow ATP hydrolysis,
as the rates of both substrate unfolding and translocation will decrease as the rate of ATP
70 hydrolysis decreases. However, slower degradation, by itself, would be expected to decrease
rather than increase KM. Thus, the increased KM for Arc-st11-ssrA degradation by ∆175-209linker
HslUV suggests that the 175-209 loop aids in the binding of this protein substrate by wild-type
HslU. We confirmed this possibility by direct-binding experiments. ∆175-209linker HslU bound
fluorescent Arc-st11-ssrA with a KD of ~4.6 µM (Fig. 3.5), ~3-fold less tightly than wild-type
HslU (Fig. 3.1B). The same trend was observed for substrate binding to the fully assembled
proteolytic enzyme; ∆175-209linker HslUV bound the fluorescent substrate ~4-fold more weakly
than wild-type HslUV (Fig. 3.1C; data not shown). Importantly, ∆175-209linker HslU bound the
gt1 peptide (KD ~570 nM; Fig. 3.5) as well as or better than wild-type HslU (KD ~760 nM; Fig.
3.1D), showing that the binding defect is specific. These results show that the 175-209 loop plays
a role in binding some part of the Arc-st11-ssrA substrate.
Roles of the GYVG and 175-209 loops in substrate stimulation of ATP hydrolysis
We assayed hydrolysis of 2.5 mM ATP by the pore-loop and ∆175-209linker mutants in the
presence of increasing concentrations of gt1 peptide (Fig. 3.6A) or Arc-st11-ssrA (Fig. 3.6B).
Neither gt1-peptide nor Arc-st11-ssrA significantly altered ATP hydrolysis by Y91A HslU,
consistent with the observed binding defects of this mutant. Half-maximal stimulation
concentrations (Kstim) for the gt1 peptide were ~300 µM for Y91F HslU and ~7.6 µM wild-type
HslU. Kstim values for Arc-st11-ssrA were ~260 µM for Y91F HslU and ~34 µM for wild-type
HslU. Thus, Y91F HslU shows a defect in binding gt1 and Arc-st11-ssrA both in the ATPasestimulation assay and in a direct-binding assay. Surprisingly, neither gt1 nor Arc-st11-ssrA
stimulated ATP hydrolysis by ∆175-209linker HslU (Fig. 3.6A & 3.6B), even though both the
peptide and protein substrate bound this enzyme (Fig. 3.5). Thus, these results demonstrate that
71 the 175-209 loop in the I domain of wild-type HslU plays a critical role in allowing bound
substrates to regulate the rate of ATP hydrolysis.
Discussion
Our results support a degradation model in which the N-terminal residues of Arc substrates
initially bind in the axial pore of the HslU hexamer. Specifically, we find that the Y91F
mutation, which changes the highly conserved GYVG motif in the axial pore of HslU to GFVG,
increases KD for binding Arc-st11-ssrA and the gt1 peptide (which mimics interactions made by
the N-terminus of Arc), and increases the concentrations of Arc-st11-ssrA and gt1 needed for
half-maximal stimulation of ATP hydrolysis. Thus, the side-chain hydroxyl group of Tyr91 in
wild-type HslU plays a role in recognition of Arc substrates. The aromatic ring of Tyr91 is even
more important, as the alanine-substitution mutation abolishes detectable binding of Arc-st11ssrA and gt1. Previous studies showed that the Y91G and Y91A mutations prevented HslUV
degradation of SulA and MBP-SulA but did not distinguish between defects caused by
diminished binding versus poor unfolding/translocation (Song et al., 2000; Park et al., 2005).
ClpX and ClpA also have GYVG pore loops but have different substrate specificities than HslU
(Flynn et al. 2001; Kwon et al., 2004; Burton et al., 2005). Thus, the GYVG loop must contribute
to binding but not determine specificity. Indeed, the GYVG loop of ClpX collaborates with two
additional pore loops to bind the ssrA tag specifically (Farrell et al., 2007; Martin et al., 2008b).
If the N-terminal residues of Arc substrates bind in the axial pore, then degradation of these
proteins should begin at the N-terminus. We confirmed this prediction by showing that HslUV
degradation of an Arc-GFP-st11-ssrA fusion protein generates a truncated product in which the
72 GFP and C-terminal portions of the substrates remain intact. HslUV appears to be a more
powerful protein unfoldase when operating in the N- to C-terminal direction (Koodathingal et al.,
2009). Nevertheless, our results suggest that HslUV degradation of Arc-GFP-st11-ssrA stalls,
because the enzyme cannot unfold GFP by pulling on it from the N-terminus. Previous studies
showed that HslUV degraded most of the C-terminal SulA portion of a GFP-SulA fusion protein
but failed to degrade the GFP portion (Kwon et al., 2004). Thus, the β-barrel architecture of GFP
resists HslUV unfolding from both the C-terminal and N-terminal ends.
The intermediate domains of HslU emerge from the top surface of the hexameric ring, like the
petals of a flower that enclose a cavity shaped roughly like a bowl or funnel. The 175-209 loop is
disordered in all HslU and HslUV crystal structures but would project into the bowl and thus be
positioned to interact with substrates as they approach the axial pore (Fig. 3.7). This loop
contains a 21-residue sequence, GVEIMAPPGMEEMTSQLQSMF (residues 183-203 in E. coli
HslU), that is ~90% conserved among diverse species of γ-proteobacteria, suggesting an
important functional role. Deletion of the I domain or 175-209 loop was known to prevent or
greatly diminish degradation of SulA or Arc-SulA, without apparent changes in the structure of
the remaining portions of HslUV (Song et al., 2000; Kwon et al., 2003). Similarly, we find that
∆175-209linker HslUV shows poor degradation of Arc-st11-ssrA, with a ~7-fold increase in KM
and ~4-fold decrease in Vmax when compared with the wild-type enzyme. The increased KM
suggests a defect in substrate recognition, which we confirmed in direct binding assays. The 175209 loop probably interacts with the st11-ssrA portion of our substrate, but this remains to be
established conclusively and it is possible that one N-terminal segment of the dimeric Arc
substrate interacts with the GYVG loop and the other binds the 175-209 loop. A role for the I
73 domain in tethering protein substrates is consistent with results for other AAA+ proteases, in
which family specific auxiliary domains frequently play tethering roles in substrate recognition
(Sauer & Baker, 2011).
We were surprised to find that the I domain of HslU plays an important role in ATP hydrolysis.
For example, we observed that large I-domain deletions reduced the rate of ATP hydrolysis to
nearly zero, whereas replacing the 175-209 loop with different linkers decreased ATPase activity
from 2- to 9-fold. These defects were seen both in the presence and absence of HslV and were
independent of enzyme concentration. By contrast, Song et al. (2000) reported that the same or
very similar deletions caused almost no change in ATP hydrolysis but did not report assay
conditions or show experimental data. Why our results differ from those of Song et al. (2000) is
unclear. It is possible that they used an end-point assay with a single time that was not in the
linear kinetic range and thus did not reflect actual rates. Nevertheless, our results unambiguously
demonstrate that the I domain and the 175-209 loop play important roles in allowing HslU to
hydrolyze ATP rapidly. Strikingly, the ~4-fold decrease in Vmax for ∆175-209linker HslUV
degradation of Arc-st11-ssrA closely matched the observed decrease in ATP-hydrolysis activity,
with part of this decline resulting from reduced basal activity and part from the failure of
substrate to stimulate ATP turnover in the mutant enzyme.
Is substrate binding to the 175-209 loop responsible for stimulating ATP hydrolysis in the wildtype enzyme? This model nicely accounts for the inability of Arc-st11-ssrA to enhance the
ATPase activity of ∆175-209linker HslU. However, the gt1 peptide also failed to stimulate ATP
hydrolysis by this mutant, which bound gt1 as well as the wild-type enzyme. Thus, gt1 would
74 need to make strong pore contacts and weaker contacts with the 175-209 loop if binding to this
loop is required for ATPase stimulation. Alternatively, an intact 175-209 loop may play
independent roles in substrate tethering and in allowing rapid enzymatic ATP turnover. The ~2fold defect in basal ATP hydrolysis by ∆175-209linker HslU arises largely from a decrease in
Vmax, indicating that the missing loop affects the rate-limiting step in ATP turnover. The I
domain is followed by the Walker-B sequence of HslU, which plays roles in coordinating Mg++
binding to the β and γ phosphates of ATP and in activating a water molecule for nucleophilic
attack on the terminal phosphate (Bochtler et al., 2000; Sousa et al., 2000; Zhang & Wigley,
2008). Changes in I-domain conformation could plausibly affect these active-site residues.
Alternatively, I-domain structure could be linked to conformational changes in the hexameric
ring of HslU, which limit the overall rate of ATP turnover.
For wild-type HslUV and variants with the Y91F and ∆175-209linker HslU mutations, we
consistently observed a large difference between KM for ATP-dependent degradation of Arcst11-ssrA and KD for ATPgS-dependent binding to fluorescent Arc-st11-ssrA. For example, KM
for wild-type degradation by HslUV was ~20 µM, whereas KD for binding was ~0.1 µM. These
differences do not reflect the fluorescent modifications of the substrate, as competition
experiments revealed that HslU bound roughly equally well to labeled and unlabeled Arc-st11ssrA. The difference between KM and KD suggests that the rate of substrate dissociation from
ATPγS-saturated HslUV is significantly slower than the combined rates of substrate degradation
and substrate release that occur as a consequence of steps dependent on ATP hydrolysis. For
example, failed enzymatic attempts to unfold the substrate could result in release, as has been
observed for ClpXP (Kenniston et al., 2005).
75 Among AAA+ enzymes, HslU is unique in having an I domain. The Tip48/49 family of snoRNA
remodeling enzymes also have a family-specific insertion in a similar region of the large AAA+
domain, as does the LonB family of archaeal AAA+ proteases (Iyer et al., 2004). In LonB, this
insertion forms a pair of membrane helices that tether each subunit in the hexameric ring to the
inner surface of the cytoplasmic membrane. It will be important to determine if these membranebinding domains, like the I domain of HslU, play roles in substrate binding and regulation of
ATP hydrolysis by LonB proteases.
Materials & Methods
Proteins and peptides
E. coli HslU variants were constructed by PCR in a pET12b-H6-HslU plasmid background. E.
coli HslU, E. coli HslU mutants, E. coli HslV, Arc-st11-ssrA,
35
S-Arc-st11-ssrA, and E. coli
SspB were expressed and purified as described (Sundar et al., 2010). To engineer a gene
encoding Arc-GFP-st11-ssrA, unique NheI (GCTAGC) and SalI (GTCGAC) sites were
introduced by PCR at the Arc/st11 junction in pET21b-Arc-st11-ssrA, unique NheI and SalI sites
were introduced flanking the N- and C-terminal GFP coding sequence in a PCR fragment, these
molecule were cut with NheI and SalI and appropriate fragments were ligated to generate the
expression plasmid. Arc-GFP-st11-ssrA was purified using the Arc-st11-ssrA procedure (Sundar
et al., 2010), except a HiLoad 16/60 Superdex-200 column was used for the final gel-filtration
step.
Labeling of purified Arc-st11-ssrA with FL-BODIPY-CASE and labeling of the synthetic gt1
peptide with fluorescein were performed as described (Burton et al., 2005). Peptides were
76 purified by HPLC, and their molecular weights were confirmed by MALDI mass spectrometry
prior to use. Protein concentrations were determined from A280, using extinction coefficients
calculated from the amino-acid sequence. Peptide concentrations were determined using an
extinction coefficient of 70,000 M-1cm-1 for fluorescein at A492.
Biochemical assays
ATPase assays – HslU or variants were incubated at hexamer concentrations of 150, 300, 600, or
1200 nM in 25 µL PD buffer (25 mM HEPES (pH 7.6), 5 mM KCl, 5 mM MgCl2, 0.032%
(vol/vol) Igepal CA-630 (NP-40), and 10% (vol/vol) glycerol) at 37 °C. We then added 25 µL of
a buffer-matched solution containing 5 mM ATP, 2 mM NADH, 4 mM phosphoenolpyruvate, 6
U/mL lactate dehydrogenase, and 6 U/mL pyruvate kinase and monitored NADH oxidation,
which occurred in a coupled reaction as ADP was generated by ATP hydrolysis, by decreased
absorbance at 340 nm (Nørby, 1988). ATPase activities in units of mM ATP hydrolyzed min-1
were
calculated
as
∆A340/(∆time*path_length*6.23
mM-1cm-1),
plotted
against
HslU
concentration (mM), and specific activities in units of ATPs hydrolyzed HslU-1 min-1 were
determined from a linear fit of this curve. Reactions were carried out in a 96-well plate, with a
path length of 0.17 cm for the 50 µM volume used. To determine the ability of the peptidase to
stimulate ATPase activity, we incubated HslU (300 nM) and HslV (800 nM) together before
beginning the reaction with the addition of ATP. To determine the nucleotide dependence of
hydrolysis activity, we fixed the final HslU concentration (100 nM) and varied the ATP
concentration. To determine if substrates or gt1 peptide altered ATP-hydrolysis rates, we fixed
the final HslU concentration (100 nM) and varied the concentration of peptide/protein which was
included in the 25 µL mix with ATP and the regeneration system.
77 HslV activation assays – Assays probing the ability of HslU variants to bind HslV and activate
cleavage of 20 µM 7-amino-4-methylcoumarin (GGL-AMC) were performed in PD buffer at 37
°C and monitored by increased fluorescence at 440 nm using a PTI QM-2000-4SE
spectrofluorimeter. Reactions contained ATPγS (2 mM), HslV12 (100 nM), and HslU6 (400 nM).
Degradation – Degradation assays were performed at 37 °C in PD buffer supplemented with
ATP (5 mM), 16 mM creatine phosphate, and 10 µg/mL creatine kinase. Proteolysis of 35S-Arcst11-ssrA by 100 nM HslU6, 300 nM HslV12 was assayed by the production of TCA-soluble
radioactive peptides (Sundar et al., 2010). For assays monitored by SDS-PAGE, reactions
contained 10 µM Arc-st11-ssrA (300 nM HslU6, 800 nM HslV12) or 4 µM Arc-GFP-st11-ssrA (1
µM HslU6, 3 µM HslV12, with or without 4 µM SspB) and were quenched at different times by
addition of SDS loading buffer (125mM Bis-Tris pH 6.8, 5% glycerol, 1% SDS, 0.5 mM EDTA,
100mM DTT, .015% Coomassie Blue G-250). For single-turnover degradation of Arc-GFP-st11ssrA (1 µM), reactions contained HslU6 (5 µM), HslV12 (6 µM), SspB2 (10 µM when present)
and kinetics were monitored by SDS-PAGE. Western blots were probed with an HRPconjugated antibody raised against a His6 peptide tag and visualized using ECL substrate (GE
Healthcare).
Binding assays – The binding of HslU or variants to gt1 peptide or Arc-st11-ssrA was assayed at
37 °C in PD buffer by monitoring increases in fluorescence anisotropy (excitation 492 nm;
emission 520 nm) using the SoftMax Pro5 fluorescence plate reader (Molecular Devices).
Samples (20 µL) contained 200 nM fluorescein-labeled gt1 or 400 nM fluorescein-labeled Arcst11-ssrA, 3 mM ATPγS, varying concentrations of HslU, and if appropriate, 4.8 µM HslV12.
78 Readings were corrected for scattering, and G-factors of 1.392 (gt1 peptide) or 1.627 (Arc-st11ssrA) were used for analysis.
LC-MS – Mass spectrometry analysis of tryptic peptides of the truncated Arc-GFP-st11-ssrA
product, produced by HslUV degradation, was performed by Ioannis Papayannopoulos (MIT
Biopolymers & Proteomics Facility).
Acknowledgements
We thank I. Levchenko, I. Papayannopoulos, S. Glynn, D. Nager, S. Kim, R. Mauldin, and A.
Olivares for assistance and helpful discussions. Supported by NIH grant AI-16892.
79 Figures & Tables
Table 3.1 – Functional parameters of HslU enzyme variants.
HslU enzyme
Degradation Parameters
HslUV
KM
Vmax
(min-1 HslU6-1)
(µM)
Basal ATPase
(min-1 HslU6-1)
HslV12 Activation
(peptides•min-1HslU6-1)
KATP**
(µM)
WT
11 ± 0.7
20 ± 2.2
130 ± 23
530
7.3 ± 1.3
Y91F
8.7 ± 0.1
26 ± 0.0
270 ± 25
620
-
Y91A
No fit
No fit
200 ± 22
560
-
∆175-209linker
3.5 ± 2.1
130 ± 23
67 ± 10
550
10.3 ± 2.4
∆175-209GG
-
-
14 ± 3.9
680
3.6 ± 1.1
∆121-231linker
-
-
2.3 ± 1.9
520
-
∆108-243GG
-
-
3.8 ± 2.5
-
-
**KATP represents the concentration of ATP required to achieve half-maximal basal ATPase
activity.
Values for HslUV degradation of Arc-st11-ssrA, basal ATP-hydrolysis rates by HslU, KATP, and
HslU activation of HslV peptidase activity were determined as described in Materials &
Methods. Degradation experiments were done in duplicate, basal ATPase measurements were
done in quadruplicate, and the ATP dependence of hydrolysis was measured in duplicate. For
values that are an average of multiple experiments, errors represent standard deviations.
80 Figure 3.1 – Functional properties of HslU pore-loop mutants. (A) Steady-state kinetics of
Arc-st11-ssrA degradation by wild-type HslUV (circles; KM = 20 µM; Vmax = 11 min-1), Y91F
HslUV (squares; KM = 26 µM; Vmax = 8.7 min-1), and Y91A HslUV (triangles). Reactions
contained 100 nM HslU6 and 300 nM HslV12. Data were fit to the Michaelis-Menten equation.
(B) Wild-type HslU (circles; KD = 1.3 µM), Y91F HslU (squares; KD = 20 µM), and Y91A HslU
(triangles) were titrated against 400 nM FL-Arc-st11-ssrA. (C) Increasing amounts of wild-type,
Y91F, or Y91A HslU6 were titrated against 400 nM FL-Arc-st11-ssrA in the presence of 4.8 µM
HslV12. Except for Y91A (no fit), data were fit to a hyperbolic isotherm (wild-type KD = 110
nM; Y91F KD = 680 nM). (D) Wild-type HslU (circles; KD = 760 nM), Y91F HslU (squares; KD
= 3.8 µM), and Y91A HslU (triangles) were titrated against 200 nM FL-gt1 peptide. The
experiments shown in panels B-D were performed in the presence of 3 mM ATPγS, and (raw
anisotropy measured) / (maximum raw anisotropy for each enzyme) is plotted.
81 Figure 3.2 – Degradation of Arc-GFP-st11-ssrA proceeds from the N-terminus. (A)
Degradation of 4 µM Arc-GFP-st11-ssrA by 1 µM HslU6 and 3 µM HslV12 was monitored by
SDS-PAGE and Coomassie staining. The asterisk marks the major truncated product. (B) The
band corresponding to the truncated product was excised and analyzed by LC-MS. The
fragmentation spectrum shown is consistent with the sequence NQHDAANDENYALAA, which
corresponds to the C-terminal peptide of Arc-GFP-st11-ssrA. Peptides corresponding to Nterminal residues 1-31 of this substrate were not recovered. (C) Western blot analysis of the ArcGFP-st11-ssrA degradation experiment shown in panel A using α-His6 antibody. The truncated
product (*) retains the His6-tag of the st11 sequence. (D) The truncated product (*) accumulates
more slowly using 4 µM SspB in addition to the components present in panel A.
82 Figure 3.3 – The intermediate domain of HslUV is important for robust ATP hydrolysis.
(A) Rates of hydrolysis of 2.5 mM ATP were assayed by coupling ADP production to NADH
oxidation, resulting in decreased absorbance at 340 nm (normalized to the starting value). The
main plot shows representative experiments for 300 nM wild-type HslU6 (●) or ∆175-209linker
HslU6 (❍). The inset shows representative experiments for 300 nM ∆175-209GG HslU6 (❒) and
the ∆121-231linker and ∆108-243linker enzymes (*). The y-axis of the inset ranges from 0.97-1.01.
(B) ATP-hydrolysis rates were calculated for a range of enzyme concentrations from
experiments like those in panel A. The main plot shows linear variation of rates with the
concentration of wild-type HslU6 (●) or ∆175-209linker HslU6 (❍). The inset shows the same
result for the ∆175-209GG (❒), ∆121-231linker (+), and ∆108-243linker (*) enzymes. The y-axis of
the inset ranges from 0-16 ATP min-1. (C) Black bars show ATP-hydrolysis rates by 300 nM
wild-type HslU6, ∆175-209linker HslU6, and ∆175-209GG HslU6. Gray bars show stimulation by
addition of 800 nM HslV12. (D) Rates of ATP hydrolysis for 300 nM wild-type HslU6 (●),
∆175-209linker HslU6 (❍), or ∆175-209GG HslU6 (❒) were determined at different ATP
concentrations and fitted to the Hill form of the Michaelis-Menten equation. Half-maximal rates
were observed at ATP concentrations ranging from 4-10 µM (Table 3.1).
83 Figure 3.4 – Degradation of Arc-st11-ssrA by ∆175-209linker HslUV. The inset shows
degradation of 10 µM Arc-st11-ssrA by 800 nM HslV12 and 300 nM wild-type HslU6 (left panel)
or ∆175-209linker HslU6 (right panel) assayed by SDS-PAGE. The main plot shows changes in the
Arc-st11-ssrA degradation rate by 300nM ∆175-209linker HslU6 and 800 nM HslV12 as a function
of substrate concentration. The solid line is a fit to the Michaelis-Menten equation (KM = 130 ±
23 µM; Vmax = 3.5 ± 2.1 min-1 enz-1). For comparison, KM for wild-type HslUV degradation was
20 µM and Vmax was 11 min-1 enz-1 (Fig. 3.1A).
84 Figure 3.5 – ∆175-209linker HslU binding to substrate and peptide. Increasing concentrations
of ∆175-209linker HslU6 were titrated against 200 nM FL-gt1 peptide or 400 nM FL-Arc-st11ssrA and binding was assayed by changes in fluorescence anisotropy. The solid lines are fits to a
hyperbolic binding isotherm with KD’s of 570 nM for gt1 and 4.6 µM for Arc-st11-ssrA.
85 Figure 3.6 – Stimulation of ATP hydrolysis by gt1-peptide or Arc-st11-ssrA substrate.
Changes in the ATPase activity of wild-type HslU or variants (100 nM hexamer) were measured
as a function of the concentration of gt1 peptide (panel A) or Arc-st11-ssrA (panel B). For each
experiment, ATPase rates were normalized to the rate with no added peptide or substrate. No
significant stimulation was observed for the Y91A enzyme or the ∆175-209linker enzyme. The
solid lines for the wild-type and Y91F enzymes are fits to a hyperbolic equation with a yintercept of 1, with half-maximal stimulation values of 7.6 µM (gt1; wild-type HslU), 300 µM
(gt1; Y91F HslU), 34 µM (Arc-st11-ssrA; wild-type HslU), and 260 µM (Arc-st11-ssrA; Y91F
HslU).
86 Figure 3.7 – Cutaway cartoon of substrate recognition by HslUV. The I domains of HslU6
contain the 175-209 loops (yellow), which project inward towards the axis and play roles in
modulating substrate choice. Unstructured terminal sequences of the substrate could interact with
the 175-209 loops in the “funnel” created by the I domains and with the conserved GYVG pore
loops (Tyr91 shown in red). Engagement by the GYVG loops is required for substrate unfolding,
translocation, and degradation. Stars mark the proteolytic active sites in the HslV chamber.
87 Chapter Four:
Perspectives for Future Research
88 Protein degradation is an important cellular process. In Escherichia coli, compartmental AAA+
proteases have evolved several mechanisms to carry out proteolysis in a regulated and conditionspecific manner. For the HslUV protease, I have shown that degradation specificity is dependent
on recognition of multivalent substrate degrons, assisted by the enzyme intermediate (I) domain.
I have also shown that the I domain, unlike accessory domains of other bacterial unfoldases, is a
vital component of enzyme function. These discoveries are only initial steps in understanding
HslUV function. Below, I outline important experiments to address yet unanswered questions
regarding this protease’s role in cellular physiology and its mechanism of degradation. Gaining a
comprehensive understanding of HslUV function may also provide clues into the role and
regulation of similar energy-dependent enzymes in other organisms.
Significance of HslUV Function in Cellular Physiology
Trapping experiments to identify biologically relevant substrates
Our current knowledge of HslUV relegates it to a back-up role in the specific degradation of
RcsA, SulA, and σ32 (Kanemori et al., 1997; Khattar, 1997; Kuo et al., 2004). The only in vivo
defect seen for a strain lacking this protease is a growth defect at very high temperatures
(Missiakas et al., 1996). I suspect that this is unlikely to be the full extent of HslUV function in
the cell, as other bacterial AAA+ proteases are involved in so many different processes. A
systematic search for natural HslUV substrates could reveal or suggest additional cellular
functions.
For ClpXP, Flynn and colleagues (2003) developed a mass spectrometric approach to identify
putative substrates. Following mutation of the active-site serine of ClpP, polypeptides that were
89 translocated into the peptidase chamber by the ClpX unfoldase were sequestered without being
degraded. These trapped proteins were then co-purified with ClpP, and the whole complex was
subjected to mass spectrometry and protein identification. A similar method could be used for
identifying endogenous substrates of HslUV. An active site mutant of HslV would likely be able
to trap proteins that are specifically recognized by the unfoldase. Identification of the trapped
proteins and analysis of their functions could then present a more detailed picture of the role of
HslUV in the cell.
The efficiency of trapping HslUV substrates would, of course, depend on the cellular
concentrations of these proteins and their availability for recognition by HslU. Protein levels are,
in turn, dependent on the cell environment. Therefore, utilizing a method of condition-specific
trapping could add a more quantitative view of HslUV protease function. Such a method was
used by Neher et al. (2006) to examine the shift in ClpXP substrate choice before and after the
induction of the SOS-response. Because HslUV is involved in both heat stress and acute acid
stress (Kannan et al., 2008), it is conceivable that it is also a player in additional stress responses.
Looking at the extent and diversity of its substrate profile during various types of cellular stress
could provide vital clues on how the function of this enzyme is regulated. Orthogonal techniques,
such as yeast two-hybrid and in vitro binding assays, could be used to confirm the validity of
suggested protease•substrate interactions.
Dissection of substrate recognition rules
The trapping experiments described above should result in a compilation of putative HslUV
substrate sequences in E. coli. Besides clarifying the physiological role of the protease in this
90 bacterium, these sequences would also offer a unique opportunity to understand substrate
recognition on a proteome-wide level. Through my work, we now know that for two unrelated
model proteins, Arc repressor and the λcIN repressor, degrons present at both termini are
important for efficient recognition by HslU (Sundar et al., 2010; Chapter 2). It is likely that
searching the list of trapped protein sequences may yield natural substrates that have a similar
“split-degron” arrangement for HslUV degradation. Other types of recognition signals may also
become apparent during bioinformatic analysis of trapped protein sequences. In combination
with biochemical validation, we can begin to understand the various modes of cellular substrate
recognition by HslU at the molecular level.
Investigating protease function in other organisms
Upon identifying a list of potential HslUV substrates and probing their recognition, it should
become easier to examine how organisms besides E. coli regulate important processes. Basic
methods of genetics, biochemistry, and cell biology can be used to uncover whether HslUV
performs similar roles in E. coli and in other species. For example, the Lon protease plays a key
role in protein quality control during heat stress in bacteria (Bissonnette et al., 2010). Bender and
colleagues (2011) used this knowledge to induce temperature-dependent protein aggregation and
confirmed the same role for Lon in yeast mitochondria. However, proteases can also serve
different functions in different organisms. In Mycoplasma, for example, Lon is responsible for
degradation of ssrA-tagged proteins (Gur & Sauer, 2008b), a novel function that could not have
been predicted given its role in γ-proteobacteria. Therefore, even though complete understanding
of HslUV in related organisms could be challenging, having prior knowledge of its function in E.
coli function would simplify efforts to understand its activities in new contexts.
91 Detailed Mechanisms of HslUV Machine Function
Amino-acid level characterization of substrate binding by the I domain
My most interesting discovery is the integral role of the I domain in HslU proteolysis and
machine function. Currently, we know that deletion of the I-domain loop comprising residues
175-209 results in defects of substrate binding and ATP hydrolysis. Within γ-proteobacteria, this
region is very highly conserved (Fig. 4.1, bottom). Comparing conservation patterns with αproteobacteria (Fig. 4.1, top) and β-proteobacteria (Fig 4.1, middle), it is clear that several
residues are invariant across all species. However, within the loop comprising positions 175-209,
there are several amino acids that are similar within each branch of bacteria, but are different
across branches.
92 Figure 4.1 – Sequence logos starting at amino acid 159 of the HslU I domain from
proteobacteria. Twenty sequences were taken from the indicated taxonomic sub-family of
enterobacteria and aligned in Uniprot. The alignment was then used to create sequence logos
(Crooks et al., 2004; http://weblogo.berkeley.edu). A nearly universally conserved EI motif is
shaded purple, while residues that may potentially confer varying substrate specificities are
shaded yellow.
Why would a potential recognition sequence be conserved in this class-specific manner? It is
quite possible that these specific positions (e.g., the residues after the universally conserved EI
found at position ~186 – shaded purple) are responsible for differential substrate recognition.
HslUV may have a certain mode of recognition (shaded yellow) in gamma species (MAPPG),
93 which is slightly different than its specificity in beta species ((F/M)(A/G)PPG), and significantly
different than in alphas (PGMPG). Directed mutagenesis of these residues may reveal the precise
nature of each amino acid in substrate binding and/or regulation of ATP hydrolysis.
Physiological differences across species could then be probed in greater detail.
Understanding how the I domain regulates ATPase function
Although the entire I domain seems to be crucial for nucleotide hydrolysis, it is not clear how
this is accomplished. The most puzzling aspect is that changes made at positions 175-209, at
least 50 Å away from the ATP-binding site, still affect rates of ATP hydrolysis. Interestingly,
replacement of this 35-amino-acid region with a 12-amino-acid flexible linker has less of an
effect on ATPase activity than replacement with diglycine. Deletion of the entire I domain has
the most severe effect. These data suggest that the I domain undergoes significant
conformational changes during the ATPase cycle, potentially constraining distant regions of the
AAA+ domains that are directly involved in binding and hydrolyzing ATP. The primary
candidates for important structural change are the helices that comprise the bridge between the
large AAA+ domain and the I domain since these helices are near in sequence and space to the
Walker-B motif, which is important for ATP hydrolysis (Bochtler et al., 2000; Sousa et al.,
2000). Dynein, another AAA-machine, contains a coiled coil domain that undergoes precisely
this kind of structural re-arrangement during function (Carter & Vale, 2010). Further
crystallographic study of HslU, with particular focus on the I domain, may help in testing this
model. Alternatively, introducing physical constraints into the I domain by means other than
wholesale deletion (e.g., disulfide-bond formation or chemical crosslinking) could help elucidate
the role of the I domain in ATP hydrolysis by HslU.
94 Role of the I domain in intra-enzyme communication
Even upon dissection of I-domain substrate recognition and the detailed mechanism of control of
ATPase activity, there will be a major gap in understanding HslUV. Specifically, the mechanism
that couples substrate binding to ATP hydrolysis and subsequent protein degradation will not be
known. Tools must be developed to probe distinct kinetic states and the dynamics between these
states. Again, X-ray crystallography to capture different states of protein degradation may be one
option. The drawback, of course, is that the disordered nature of the important I-domain loop
(residues 175-209) will probably make electron-density visualization difficult. In fact, this loop
is disordered and absent in all of the currently solved structures of full-length HslU. This being
the case, single-molecule fluorescence or NMR spectroscopy studies of I-domain dynamics may
provide a more informative approach. Even though identification of appropriate labeling sites in
single-molecule fluorescence experiments is non-trivial, some of the experiments discussed
above may help in selection. Overall, the investigations suggested here should substantially
enhance our knowledge of HslUV I-domain function.
Potential Lessons for the Study of Other AAA-Motors
An atomic-resolution understanding of the I-domain coordinated steps in protein degradation
coupled with physiological functional data provides a strong base for future investigations.
Studying the crosstalk between the I domain and the AAA+ domains of HslU may provide clues
about the dynein motor, whose coiled-coil domains are responsible for both microtubule binding
and for modulation of ATPase activity (Carter & Vale, 2010). To date, no structures exist of the
motor domain plus the coiled-coil domain segment of this 1.5 MDa machine. Archaeal Lon
enzymes may also share mechanistic features with HslUV. The apical insertion domain and
95 membrane anchor regions of Thermococcus onnurineus Lon extrude from the large AAA+
domain in a similar manner to the I domain in HslU (Cha et al., 2010). Could this region be a site
for tethering membrane protein substrates? Can new substrates for Lon be identified in this
archaeon using rules of recognition found for E. coli HslU? These are distinct possibilities.
Study of HslUV can improve our understanding of proteolysis in Escherichia coli. In my thesis
work, I have begun to elucidate the functional mechanisms employed by this protease. Continued
studies of this enzyme may further our knowledge of proteases in other organisms and improve
our general understanding of molecular motors.
96 Appendix A:
Structures of HslU and HslUV
97 Figure A.1 – Structure of HslU from Bochtler et al. (2000). (A) Top and (B) side views of the
hexameric unfoldase HslU from E. coli. The six nucleotide-binding sites are occupied by four
ATP molecules and two sulfate molecules. LEGEND: AAA+ domains = green; I domains
(residues 108-243) = blue; GYVG loops = pink; nucleotides = red; sulfates = orange.
98 Figure A.2 – Structure of HslUV from Sousa et al. (2000). (A) Top and (B) side views of the
HslUV protease from H. influenzae. This is a “double-capped” structure, with a single HslV
flanked on both ends by HslU. The conserved GYVG loop is missing in this structure. All
nucleotide-binding sites contain ATP. LEGEND: AAA+ domains = green; I domains (residues
108-243) = blue; nucleotides = red; HslV molecules = yellow.
99 Figure A.3 – Structure of HslU from Wang et al. (2001). Side view of an HslU unfoldase
monomer from E. coli. ADP is bound in the nucleotide-binding site. LEGEND: AAA+ domains
= green; I domain (residues 108-243) = blue; GYVG loop = pink; ADP = red.
100 Figure A.4 – Structure of HslUV from Sousa et al. (2002). (A) Top and (B) side views of
HslUV from H. influenzae. This is a “double-capped” structure, with a single HslV peptidase
flanked on both ends by HslU. The conserved GYVG loop is missing. All nucleotide-binding
sites contain ATP. The peptidase active sites are bound to the vinyl sulfone inhibitor NLVS.
LEGEND: AAA+ domains = green; I domains (residues 108-243) = blue; nucleotides = red;
HslV molecules = yellow; NLVS = cyan.
101 Figure A.5 – Structure of HslU from Kwon et al. (2003). (A) Top and (B) side views of the
HslU unfoldase from H. influenzae. An HslU construct lacking the I domain (residues 108-243)
was used for crystallography. All nucleotide-binding sites seen here contain ADP and phosphate
(PO43-). LEGEND: AAA+ domains = green; GYVG loop = pink; ADP and PO43- = red.
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