Final Thesis

Mechanisms of Substrate Recognition by the AAA+ Protease HslUV

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

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

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 C- terminus, 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

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. :)

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 9 Concentration-dependent mechanisms 10 Inhibition and activation mechanisms 12 Cleavage-site recognition 14 Compartmental proteases: The 26S proteasome 15

ATP-dependent Proteolysis in Bacteria 18 FtsH 20 Lon 22 ClpXP 24 ClpAP 28

The HslUV Protease 29 Cellular roles of HslUV 30 Structural and biochemical insights 32 Research Approach 34

CHAPTER TWO: Multiple Sequence Signals Direct Recognition and Degradation of Protein Substrates by the AAA+ Protease HslUV 36

Abstract 37 Introduction 38 Results 40 Effects of terminal λcIN sequences on HslUV degradation 40 Effects of terminal Arc sequences on degradation 42 Terminal sequences do not alter substrate stability 43 Protein binding to the C-terminus of substrates slows HslUV degradation 43 ClpS binding to the N-terminus of substrates inhibits HslUV degradation 44 Synergistic effects of N- and C-terminal blocking 44 Single-turnover inhibition 45

Discussion 45 Materials & Methods 49

Acknowledgements 52 Figures & Tables 53

CHAPTER THREE: The Intermediate Domain of the AAA+ HslUV Protease Coordinates Substrate Binding and Stimulation of ATP Hydrolysis 60

Abstract 61 Introduction 62 Results 65 Effects of HslU pore-loop mutations on substrate binding and degradation 65 Degradation of an Arc fusion protein starts at the N-terminus 67 The intermediate domain of HslU plays a major role in determining ATP-hydrolysis rates 68 ∆175-209linker HslUV degrades Arc-st11-ssrA with a lower Vmax and weaker KM 70 Roles of the GYVG and 175-209 loops in substrate stimulation of ATP hydrolysis 71

Discussion 72 Materials & Methods 76 Acknowledgements 79 Figures & Tables 80

CHAPTER FOUR: Perspectives for Future Research 88

Significance of HslUV Function in Cellular Physiology 89 Trapping experiments to identify biologically relevant substrates 89 Dissection of substrate recognition rules 90 Investigating protease function in other organisms 91

Detailed Mechanisms of HslUV Machine Function 92 Amino-acid level characterization of substrate binding by the I domain 92 Understanding how the I domain regulates ATPase function 94 Role of the I domain in intra-enzyme communication 95

Potential Lessons for the Study of Other AAA-Motors 95

APPENDIX A: Structures of HslU and HslUV 97

REFERENCES 103

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

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

Chapter One:

Introduction to Regulation and Substrate Specificity in Proteolysis

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 & Jacobs-

Wagner, 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

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

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.

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,

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 self- cleavage, 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 pro- segment 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

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 immuno- proteasome, 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

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 Cell Biology 309 |

fully understand how the RP subunits are arranged in the subcomplexes and how the RP stimulates the opening of the CP gate and facilitates substrate unfolding and translocation. The recent pub- lication of several crystal structures of the essential domains of the proteasomal ATPases from archaea (named proteasome- activating nucleotidase or PAN) and actinobacteria (named ATPase forming ring-shaped complex or ARC) ﬁnally provides some important clues to understand the action of the 19S RP (Djuranovic et al., 2009; Zhang et al., 2009a). PAN and ARC are orthologous to the eukaryotic Rpt subunits, sharing more than 40% sequence identity (Smith et al., 2006). The N-terminal region of PAN contains a coiled-coil (CC) segment followed by an oligonucleotide/oligosaccharide-binding (OB) domain. The Figure 1 Schematic diagram of the 26S proteasome. The 26S protea- OB domains from six PAN N-terminal fragments form a hexameric Figure 1.1 – Schematicsome consists diagram of the of20 S the core particle 26S proteasome (CP) and the 19 S (Xie, regulatory 2010).ring Components with a pseudo 6 -foldand symmetry. The axial pore of the OB ring particle (RP). The CP is formed by four stacked rings: two outside assemblage of the 20S CP and 19S RP is shown. Importantly, the is hexameric13 A˚. The CC Rpt segments ring of the six PAN fragments display only a-rings and two inner b-rings. Each ring is formed by seven different ! 3-fold symmetry. Thus, the N-terminal fragment of PAN is (comprising Rpt1-6) issubunits. the ATP The- RPdependent is divided intomotor two subcomplexes:and the proteins the lid andin the lid are responsible for

assembled into a crown-shaped subcomplex: three CC segments Downloaded from ubiquitin recognition. base. Certain The subunits subunits forming of the the lid CP and theβ-rings base are conta shown.in Rpn the10 proteolytic active sites, sprouting from the hexameric OB ring (Djuranovic et al., 2009; which face into the chamberstabilizes and the associationaway from between solution. the lid and the base. Zhang et al., 2009a). The N-terminal region of ARC carries a CC segment and two OB domains. The OB domains of ARC form favorable for antigen processing (Boes et al., 1994; Gaczynska double hexameric rings (Djuranovic et al., 2009). The ATPase et al., 1994; Cardozo and Kohanski, 1998). b5i is replaced by domains of PAN and ARC are located at their C-termini. The another proteolytic active subunit b5t in cortical thymic epithelial ATPase domain of PAN was also crystallized (Zhang et al., jmcb.oxfordjournals.org cells, forming the so-called thymoproteasome (Murata et al., 2009a). It appears to form the canonical AAA ATPase ring + 2007). Different from eukaryotic CP, there are only one type of with a narrow channel.15 Together, these crystal structures a subunit and one type of b subunit in prokaryotic proteasomes, suggest a model of double-ring structure for the PAN hexamer. with the exception of the proteasome isolated from bacterium This model is consistent with a three-dimensional model of PAN Rhodococcus, which contains two types of a and b subunits generated from tomographic reconstruction of negatively (Voges et al., 1999). stained particles (Medalia et al., 2009). Zhang et al. (2009b) Crystal structures of the CP from archaeon, yeast and mammals further demonstrated that the PAN ATPase subcomplex associ- by guest on March 28, 2011 have been solved (Lo¨we et al., 1995; Groll et al., 1997; Unno ates with the CP and drives ATP-dependent substrate unfolding, et al., 2002). There is a very narrow pore or gate at the center whereas the CC–OB subcomplex serves as the entry port of the of the a subunit ring where protein substrates enter the CP substrate translocation channel. This PAN–CP model explains chamber. The gate is closed in a free CP by interactions among why an unfolded segment is an essential determinant of a protea- the N-termini of the a subunits, mainly a2, a3 and a4, blocking somal substrate (Schrader et al., 2009). According to this model, substrate entry into the proteolytic chamber. In the 26S protea- only the unfolded segment can enter the narrow port of the CC– some, the RP acts as an activator to facilitate the opening of OB channel. The leading segment is then captured by the ATPases the gate by interacting with the a subunits (Smith et al., 2007; and drawn deeper into the channel by the force generated by ATP Rabl et al., 2008). Besides the RP, the PA200/Blm10 protein hydrolysis. At the same time, the pulling of the leading segment and the PA28/11S complex can also bind the a subunits and par- results in unfolding of the substrate. tially or fully open the gate of the CP, thereby facilitating sub- How are the six different Rpt ATPase subunits arranged in the strate access to the proteolytic chamber (Whitby et al., 2000; eukaryotic proteasome? Taking advantage of the observation Sadre-Bazzaz et al., 2010). that the residues in the interfaces between the PAN OB Unlike the CP, no crystal structures of the 19S RP have been domains can be aligned with the sequences of the eukaryotic solved, perhaps due to its unstable conformation and/or variable ATPases in these regions, Tomko et al. (2010) used a clever dis- composition. Our current understanding of the 19S RP structure ulfide engineering approach to demonstrate that the six different mainly relies on the information obtained from cryo-electron ATPases of eukaryotic RP form a heterohexameric ring with the microscopy and biochemical analyses (da Fonseca and Morris, arrangement of Rpt1–Rpt2–Rpt6–Rpt3–Rpt4–Rpt5. This novel 2008; Nickell et al., 2009). The 19S RP is a multi-subunit study also reveals that the central port of the hexameric ATPase complex, which can be divided into two subcomplexes called ring is very narrow and unlikely to accommodate the large Rpn1 the base and the lid (Figure 1). The base consists of six AAA and Rpn2 subunits. This finding challenges a current model, + ATPases (Rpt1-6) and three non-ATPase subunits (Rpn1, Rpn2 which suggests that Rpn1 and Rpn2 stack on one another to and Rpn13), whereas the lid includes at least nine non-ATPase form a toroid surrounded by the ATPases (Rosenzweig et al., subunits (Rpn3, Rpn5-9, Rpn11, Rpn12 and Rpn15/Sem1). The 2008; Effantin et al., 2009). Indeed, some other EM structures connection between the lid and the base is stabilized by the of the 26S proteasome have already pointed out that the Rpn1 Rpn10 subunit. The lack of structural data makes it difficult to and Rpn2 subunits may be more peripherally connected to the

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

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

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 ATP- hydrolysis 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.

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.

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 N- terminus 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

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 non- polar C-terminal sequences target many substrates to FtsH, which can degrade unfolded

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

onnurineus (Park et al., 2006; Cha et al., 2010). In addition to the central AAA-module and C- terminal 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 substrate- delivery 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.

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

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.

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-

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 stationary- phase transcription factor, during exponential bacterial growth (Bougdour et al., 2008). The

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).

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

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

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.

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

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.,

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 N- and 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).

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 γ- proteobacteria, 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.

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.

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 peptide- binding 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.

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 translocation- mediated unfolding (Siddiqui et al., 2004; Hinnerwisch et al., 2005a; Martin et al., 2008a). This

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 (Gonciarz-

Swiatek 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 N- terminal 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 C- terminal 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

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

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 35S- 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 S1LλcIN-RSYYF and

S1L N λcI -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 N- termini 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

N dimers of λcI 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.

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

DD sequence was replaced by DD (Arc-st11-ssrA ) (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

-1 -1 Arc-ssrA and Arc-st11-ssrA with Vmax values of 4.9 and 12 min enz , respectively. Thus, inserting the st11 tag before the ssrA tag increased the maximum degradation rate more than 2- fold. 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 N- terminal 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.

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 S1LλcIN-RSYYF to ~10% of the uninhibited rate but inhibited degradation of the S1LλcIN-

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 M1LArc-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.

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 S1LλcIN-RSEYE and somewhat weaker inhibition of S1LλcIN-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 M1LArc-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 S1LλcIN-RSYYF was more severe in the presence of ClpS and the DegS PDZ domain than with either single protein (Fig. 2.5A, bottom).

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 35S-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 35S-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

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 multi- degron 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

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

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,

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 multi- step 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

S1L N designated λcI were initially cloned, expressed, and purified as fusions to an N-terminal H6- tagged 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

DD sequence (H6KNQHD), ssrA tag (AANDENYALAA), ssrA tag (AANDENYALDD), or ssrAN3A tag (AAADENYALAA). The wild-type N-terminus of Arc is Met1 (Vershon et al.,

M1L 1985). Arc 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 Arc- st11-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

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. 35S-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 35S-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

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

35 with S-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 35S-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.

Figures & Tables

Table 2.1 – Properties of λcIN and Arc substrates.

epitopes for binding to substrates Vmax KM Vmax / KM Tm SspB DegS ClpS (min-1) (µM) min-1µM-1 (°C) PDZ

ext1-λcIN-RSEYE 4.3 ± 0.1 120 ± 19 0.04 56.7 no no no N ext1-λcI -ISVTL 9.9 ± 1.1 89 ± 12 0.11 56.1 no no no ext1-λcIN(I84S)-ISVTL 9.9 79 0.13 50.5 no no no S1LλcIN-RSEYE 3.5 ± 0.3 120 ± 24 0.03 56.5 no no yes S1LλcIN-RSYYF 7.1 ± 1.0 40 ± 5.0 0.18 55.5 no yes yes λcIN-RSYYF 5.7 ± 0.6 54 ± 6.7 0.11 54.5 no yes no

Arc-ssrA 4.9 ± 0.1 24 ± 1.4 0.20 58.6 yes no no Arc-st11-ssrA 12 ± 1.9 31 ± 0.9 0.38 56.5 yes no no DD Arc-st11-ssrA 7.0 ± 0.3 51 ± 4.8 0.14 59.3 yes no no ext2-Arc-ssrA 3.0 ± 0.5 55 ± 1.3 0.06 58.2 yes no yes M1LArc-ssrA 4.7 ± 1.5 21 ± 4.0 0.22 56.9 yes no yes M1LArc-ssrAN3A 5.7 ± 0.5 23 ± 7.7 0.25 56.3 no no 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.

€

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 35 N concentrations of S-λcI variants by 100 nM HslU6, 300 nM HslV12 were determined by assaying acid-soluble radioactivity. Lines represent non-linear-least-squares fits to the Michaelis- Menten equation: rate = Vmax•[S]/([S]+KM). Kinetic parameters are listed in Table 2.1.

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.

Figure 2.3 – Occluding the C-terminus of λcIN and Arc substrates inhibits HslUV 35 N degradation. (A) Degradation of S-labeled λcI 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 35 rate with no PDZ domain. (B) Degradation of S-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.

Figure 2.4 – ClpS binding to the N-terminus of Arc or λcIN substrates slows HslUV 35 degradation. (A) Proteolysis of S-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 N HslUV (0.5 µM HslU6, 1.5 µM HslV12) degradation of λcI 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).

Figure 2.5 – Combinatorial and single-turnover inhibition. (A) (top) A combination of SspB 35 M1L and ClpS inhibited degradation of S- Arc-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 35 S1L N and ClpS also inhibited degradation of S- λcI -RSYYF (50 µM) by 0.5 µM HslU6, 1.5 µM 35 HslV12 more than either single protein alone. (B) Degradation of different S-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.

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.

Chapter Three:

The Intermediate Domain of the AAA+ HslUV Protease Coordinates Substrate Binding and Stimulation of ATP Hydrolysis

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 substrate- mediated 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.

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 double- ring 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

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

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 C- terminal 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 N- terminal 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.

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 90-

93) 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

35S-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

-1 -1 kinetic parameters was a ~20% reduction in Vmax, with values of 11 ± 0.7 min enz for the

-1 -1 wild-type enzyme and 8.7 ± 0.1 min enz 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

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.

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

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-GFP- st11-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

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 I- domain 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

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

GG ∆175-209 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 wild- type enzyme to hydrolyze ATP rapidly.

linker ∆175-209 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-st11-

-1 -1 ssrA with a KM of ~130 µM and a Vmax of ~3.5 min enz . Thus, in comparison with wild-type

HslUV, the ∆175-209linker mutation slowed the maximal rate of Arc-st11-ssrA degradation ~4- fold 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

hydrolysis decreases. However, slower degradation, by itself, would be expected to decrease

linker rather than increase KM. Thus, the increased KM for Arc-st11-ssrA degradation by ∆175-209

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 ATPase- stimulation 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

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-st11- ssrA 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

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

linker ∆175-209 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 175-

209 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

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

linker hydrolyze ATP rapidly. Strikingly, the ~4-fold decrease in Vmax for ∆175-209 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 wild- type 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

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 ~2- fold 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 Arc- st11-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-st11- ssrA. 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).

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 membrane- binding 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, 35S-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

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

-1 -1 extinction coefficient of 70,000 M cm 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

-1 -1 were calculated as ∆A340/(∆time*path_length*6.23 mM cm ), 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.

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-Arc- st11-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-st11- ssrA (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 HRP- conjugated 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 Arc- st11-ssrA, 3 mM ATPγS, varying concentrations of HslU, and if appropriate, 4.8 µM HslV12.

Readings were corrected for scattering, and G-factors of 1.392 (gt1 peptide) or 1.627 (Arc-st11- ssrA) 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.

Figures & Tables

Table 3.1 – Functional parameters of HslU enzyme variants.

** Degradation Parameters Basal ATPase HslV12 Activation KATP HslU enzyme -1 -1 -1 -1 HslUV (min HslU6 ) (peptides•min HslU6 ) (µM) Vmax KM -1 -1 (min HslU6 ) (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.

Figure 3.1 – Functional properties of HslU pore-loop mutants. (A) Steady-state kinetics of -1 Arc-st11-ssrA degradation by wild-type HslUV (circles; KM = 20 µM; Vmax = 11 min ), Y91F -1 HslUV (squares; KM = 26 µM; Vmax = 8.7 min ), 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.

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 N- terminal residues 1-31 of this substrate were not recovered. (C) Western blot analysis of the Arc- GFP-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.

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 linker main plot shows representative experiments for 300 nM wild-type HslU6 (●) or ∆175-209 GG HslU6 (❍). The inset shows representative experiments for 300 nM ∆175-209 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 linker concentration of wild-type HslU6 (●) or ∆175-209 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 linker GG wild-type HslU6, ∆175-209 HslU6, and ∆175-209 HslU6. Gray bars show stimulation by addition of 800 nM HslV12. (D) Rates of ATP hydrolysis for 300 nM wild-type HslU6 (●), linker GG ∆175-209 HslU6 (❍), or ∆175-209 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).

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) linker or ∆175-209 HslU6 (right panel) assayed by SDS-PAGE. The main plot shows changes in the linker Arc-st11-ssrA degradation rate by 300nM ∆175-209 HslU6 and 800 nM HslV12 as a function of substrate concentration. The solid line is a fit to the Michaelis-Menten equation (KM = 130 ± -1 -1 23 µM; Vmax = 3.5 ± 2.1 min enz ). For comparison, KM for wild-type HslUV degradation was -1 -1 20 µM and Vmax was 11 min enz (Fig. 3.1A).

Figure 3.5 – ∆175-209linker HslU binding to substrate and peptide. Increasing concentrations linker of ∆175-209 HslU6 were titrated against 200 nM FL-gt1 peptide or 400 nM FL-Arc-st11- ssrA 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.

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 y- intercept 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).

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.

Chapter Four:

Perspectives for Future Research

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 condition- specific 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

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

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.

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 α- proteobacteria (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.

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),

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.

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

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.

Appendix A:

Structures of HslU and HslUV

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.

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.

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.

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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.

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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 3- 3- (PO4 ). LEGEND: AAA+ domains = green; GYVG loop = pink; ADP and PO4 = red.

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