REGULATORY PARTICLES of PROTEASOMES and DETERMINANTS of PROTEIN LEVEL REGULATION in the HALOARCHAEON Haloferax Volcanii

REGULATORY PARTICLES OF PROTEASOMES AND DETERMINANTS OF PROTEIN LEVEL REGULATION IN THE HALOARCHAEON Haloferax volcanii

CHRISTOPHER J. REUTER

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2006

This document is dedicated in loving memory to my grandmother, Gladys Reuter, whom throughout my early schooling would reward me with a dollar for every “A” grade I received. This paper would have certainly been worth at least five.

ACKNOWLEDGMENTS

I would like to thank Dr. Julie Maupin-Furlow for allowing me to be in the position that I am today. By taking her undergraduate prokaryotic physiology class, I immediately gained a fascination for the world of molecular biology, and without her constant support and encouragement none of this would have been possible for me. I would also like to thank the members of my committee, Drs. L.O. Ingram, Madeline

Rasche, Nemat Keyhani, and Anita Wright, for their time and patience. Although I rarely sought their counsel, I always knew their doors were always open. I am especially grateful to Dr. L.O. Ingram, whose guest presentation on microbial ethanol production in my undergraduate general microbiology class really opened up my mind to the power of microbes. I would also like to thank Donna Williams and the late Dr. Henry Aldrich for their assistance in microscopy and photography. Both, although I hardly knew them at the time, were so generous in their efforts involving my research. I also thank Drs. James

Preston and K.T. Shanmugam and members of their labs for allowing me the use of their equipment and time. I would also like to thank Drs. William Gurley, Paul Shirk, Mark

Donnelly, Arthur Horwich, Moshe Mevarech, and Jerry Eichler for gifts of plasmids, cell cultures, or use of instrumentation. I would like to thank former members of Dr.

Maupin’s lab including Mark Ou and Drs. Heather Wilson, Steve Kaczowka, and

LeeAnn Talarico-Blalock for taking the time to teach many laboratory techniques. I would especially like to thank Dr. Steve Kaczowka for many nights of scientific (and some not so scientific) conversations and for collaboration of some of the Pan work. I

iii would also like to thank some of my fellow graduate students, with whom I became good friends, for their helpful discussions along the way, including Franz St. John, P. Aaron

Kirkland and Drs. T. Brice Causey, Celeste Johnson-Causey, and Stuart Underwood.

Other graduate students I would like to thank include Gosia Gil for creating Pan mutants and helpful discussion and Matt Humbard for computer assistance and helpful discussions.

I would like to thank my very good friends Drs. Cassidy Sedacca and Emily Bille for their constant encouragement and partnership outside the lab. I would also like to thank my family members including my sister, Sarah Reuter, my grandfather Robert

Reuter, and my Uncle Joseph Reuter for their encouragement in my pursuance of my

Ph.D. I would especially like to thank my parents, Gregory and Debra Reuter, for without their support and love throughout my life, none of this would be possible. Last and most certainly not least, I would love to thank my wife and best friend, Jennifer

Reuter, for being my biggest fan, and sometimes, hardest critic. She has instilled in me the confidence that has made me what I am today and has made many sacrifices that have allowed me to pursue this degree.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... iii

LIST OF TABLES...... viii

LIST OF FIGURES ...... ix

LIST OF ABBREVIATIONS...... xi

ABSTRACT...... xv

CHAPTER

1 LITERATURE REVIEW ...... 1

Introduction...... 1 ClpP Protease...... 1 HslUV Protease ...... 6 Lon Protease ...... 9 FtsH Protease...... 12 20S Proteasome ...... 14 AAA+ ATPases ...... 20 AAA+ Domain ...... 21 Walker Motifs...... 22 Sensor Regions ...... 22 Second Region of Homology ...... 23 Coiled-coil Domains...... 23 Pore Motifs ...... 24 Mechanistic Action of ATPases ...... 25 Substrate Recognition...... 26 Ubiquitination...... 27 N-end Rule...... 29 SsrA Tagging...... 30 Other Modes of Proteolytic Substrate Recognition...... 31 Green Fluorescent Protein (GFP) as A Model Substrate...... 32 Research Objective ...... 36

2 ANALYSIS OF PROTEASOME-DEPENDENT PROTEOLYSIS IN Haloferax volcanii, USING SHORT-LIVED GREEN FLUORESCENT PROTEINS ...... 39

v Introduction...... 39 Materials and Methods ...... 41 Materials...... 41 Strains, Media and Plasmids...... 41 DNA Purification and Transformation...... 42 Protein Techniques and Immunoblot...... 43 Fluorescence Measurements of GFP Variants in Whole Cells ...... 44 Purification of EGFP and 20S Proteasomes from H. volcanii ...... 45 Peptide Hydrolyzing Activity...... 46 RNA Isolation and Analysis...... 46 Results and Discussion ...... 47 Functional Synthesis of an Archaeal GFP Reporter Protein ...... 47 C-terminal Modifications Reduce the Level of smRS-GFP Protein in vivo ...... 48 C-terminal Modifications Do Not Impact the Level of smRS-GFP-specific mRNA in H. volcanii ...... 51 Clasto-lactacystin β-lactone as an in vivo Inhibitor of H. volcanii 20S Proteasomes ...... 51 Inhibition of 20S Proteasomes Enhances the Level of smRS-GFP-SsrA Protein in H. volcanii ...... 53 Conclusion ...... 54

3 DIFFERENTIAL POST-TRANSLATIONAL REGULATION OF GREEN FLUORESCENT PROTEIN IN THE HALOARCHAEON HALOFERAX VOLCANII IS DETERMINED IN PART BY ITS EXTREME CARBOXYL TERMINAL RESIDUE...... 67

Introduction...... 67 Materials and Methods ...... 70 Materials...... 70 Strains, Media, and Plasmids...... 70 Construction of H. volcanii Expression Plasmids ...... 70 DNA Isolation and Transformation...... 71 Absorbance and Fluorescence of Cell Cultures...... 71 Fluorescence Curves of H. volcanii with Expression Plasmids ...... 72 Summary and Conclusions ...... 73

4 IDENTIFICATION AND PARTIAL CHARACTERIZATION OF PROTEASOME-ACTIVATING NUCLEOTIDASE REGULATTORY PARTICLES OF HALOFERAX VOLCANII ...... 93

Introduction...... 93 Materials and Methods ...... 95 Materials...... 95 Strains, Media, and Plasmids...... 95 DNA isolation, Analysis, and Transformation...... 96 Protein Techniques and Antibody Production...... 97 Protein Expression in E. coli ...... 98

vi Protein Expression in H. volcanii...... 99 Protein Purification...... 99 Nucleotide-hydrolyzing Activity...... 101 DNA and Protein Sequence Analyses ...... 102 Nucleotide Sequence Accession Numbers ...... 102 Results and Discussion ...... 102 H. volcanii Synthesizes Two Pan Proteins...... 102 Protein Sequence Analysis of Pan from H. volcanii ...... 103 Pan Purification from Recombinant H. volcanii ...... 104 Independent Pan Expression from H. volcanii...... 106 Native Pan Forms Oligomers ...... 106 ATP-hydrolyzing Activity of PanA...... 107 SmRS-GFP-SsrA as a Substrate for Pan/20S Proteasomes in vitro...... 109 Summary...... 110

5 SUMMARY AND CONCLUSIONS...... 123

LIST OF REFERENCES...... 129

BIOGRAPHICAL SKETCH ...... 153

vii

LIST OF TABLES

Table page

2-1. Strains and plasmids used for this study ...... 55

2-2. Modification of the C-terminus of smRS-GFP influences whole cell fluorescence and the level of fluorescent protein in recombinant H. volcanii...... 59

3-1. Strains and plasmids used in this study...... 76

3-2. Optimal codon usage of Haloferax volcanii...... 80

4-1. Strains and plasmids used in this study...... 112

viii

LIST OF FIGURES

Figure page

1-1. Predicted structures of proteasomes. Molecular surface of (a) the archaeal, (b) the eukaryotic 20S and (c) the HslV proteasomes...... 38

2-1. Optimization of fluorescent protein expression in recombinant H. volcanii...... 60

2-2. Protein level quantification of smRS-GFP reporter proteins expressed in recombinant H. volcanii by immunoblot...... 61

2-3. Colonies of H. volcanii expressing smRS-GFP exhibit fluorescence...... 62

2-4. H. volcanii expressing smRS-GFP displays uniform cell fluorescence...... 63

2-5. Liquid cultures of H. volcanii expressing smRS-GFP exhibit fluorescence that parallels cell growth ...... 64

2-6. H. volcanii 20S proteasomes are inhibited by clasto-lactacystin β-lactone...... 65

2-7. Inhibition of H. volcanii 20S proteasomes enhances the level of smRS-GFP- ssrA in cell culture...... 66

3-1. Growth and Fluorescence of H. volcanii DS70 with GFP reporter-protein expression plasmids...... 81

3-2. Single C-terminal amino acid additions to smRS-GFP variably affect the fluorescence of the H. volcanii DS70 culture from which they are being expressed...... 92

4-1. Amino acid sequence alignment of H. volcanii PanA and PanB...... 115

4-2. PanA and PanB are produced in H. volcanii...... 116

4-3. Plasmid map of pJAM1012...... 117

4-4. PanA and PanB associate in vivo in H. volcanii...... 118

4-5. Gel filtration chromatographs of PanA-His and PanB-His proteins purified by IMAC and hydroxyapatite ionic exchange chromatography from recombinant H. volcanii...... 119

ix 4-6. Purified Pan complexes from H. volcanii...... 120

4-7. ATP-hydrolyzing activity of PanA-His purified from H. volcanii GG101 (pJAM650)...... 121

4-8. SmRS-GFP-SsrA purification from recombinant E. coli...... 122

LIST OF ABBREVIATIONS

A…………………………………..Absorbance

AAA ………………………………ATPases associated with diverse cellular activities

ADP………………………………. Adenosine 5’-diphosphate

Amc.…………………………….…7-amido-4-methyl-coumarin

AP…………………………………Alkaline phosphatase

Apr.………………………………..Ampicillin resistant

ARC……………………………….AAA ATPase forming ring-shaped complexes

ATCC……………………………...American Type Culture Collection

ATP………………………………..Adenosine 5’-triphosphate

BCIP……………………………….5-bromo-4-chloro-3-indolyl phosphate p-toluidine

BFP………………………………..Blue fluorescent protein

Bp……………………………….…Base pair

ºC………………………………….Celcius

C-………………………………….Carboxy-

CAPSO……………………………3-cyclohexylamino-2-hydroxy-1-propanesulfonic acid

Clβl………………………………..Clasto-lactacystin β-lactone

CP…………………………………Core particle

CSN……………………………….COP9 signalosome

CTP………………………………..Cytidine-5'-triphosphate

xi DMSO……………………………..Dimethyl sulfoxide

Ec………………………………….Escherichia coli

EDTA……………………………...Ethylenediaminetetraacetic acid

EGFP………………………………Enhanced GFP

EM…………………………………Electron microscopy

FDA………………………………..Food and Drug Administration g……………………………………Acceleration due to gravity

GFP………………………………..Green fluorescent protein

GTP………………………………..Guanosine-5'-triphosphate

HRP………………………………..Horseradish peroxidase

Hv………………………………….Haloferax volcanii

I-…………………………………...Intermediate

IMAC………………………………Immobilized metal-affinity chromatography

IPTG……………………………….Isopropyl-β-D-thiogalactopyranoside

ITC………………………………...Isothermal titration calorimetry

Kmr………………………………...Kanamycin resistant

KDa………………………………..Kilodalton

LAN……………………………….Lon N-terminal domain

LB………………………………….Luria broth

LPS………………………………...Lippolysaccharide

MES……………………………….2-morpholinoethanesulfonic acid

Mevr……………………………….Mevinolin resistant

xii Mj………………………………….Methanococcus jannaschii

MOPS……………………………..3-(N-Morpholino)-propanesulfonic acid

N-………………………………….Amino

NBT………………………………..Nitro blue tetrazolium

Ntn…………………………………N-terminal nucleophile

Nvr…………………………………Novobiocin resistant

ORF………………………………..Open reading frame

PAGE………………………………Polyacrylamide gel electrophoresis

PAN………………………………..Proteasome-activating nucleotidase

PCR………………………………...Polymerase chain reaction

Pi…………………………………...Inorganic phosphate

Psi………………………………….Pounds per square inch

PVDF………………………………Polyvinylidene difluoride

RP…………………………………. Regulatory particle rpm…………………………………Revolutions per minute

Rpn…………………………………Regulatory particle non-ATPases

Rpt………………………………….Regulatory particle triple-A type I proteins

RT-PCR…………………………….Reverse transcriptase-PCR

SDS………………………………...Sodium dodecyl sulfate

SRH………………………………...Second region of homology

SmRS-GFP…………………………Soluble modified red shifted-GFP

SsrA………………………………...Small stable RNA A

Ta…………………………………..Thermoplasma acidophilum

xiii TEV…………………………………Tobacco etch virus

TM………………………………….Transmembrane

Tris………………………………….N-tris(hydroxymethyl) aminomethane

TTP…………………………………Thymidine 5’-triphosphate

Ub…………………………………..Ubiquitin

UV…………………………………..Ultraviolet

xiv

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

REGULATORY PARTICLES OF PROTEASOMES AND DETERMINANTS OF PROTEIN LEVEL REGULATION IN THE HALOARCHAEON Haloferax volcanii

Christopher J. Reuter

August 2006

Chair: Julie Maupin-Furlow Major Department: Microbiology and Cell Science

Energy-dependent proteolysis is a vital process in all three domains of life. Protein turnover removes damaged and extraneous proteins, releasing free amino acids that can be recycled or used to generate energy and is responsible for the regulation of many cellular processes. A nano-compartmental, multicatalytic protease complex termed the

20S proteasome is the central component in many of these proteolytic events.

Proteasomes often cooperate with other proteins for the purpose of substrate selection.

Typically these proteins belong to the AAA (ATPases associated with various cellular activities) superfamily and associate with proteasomes for the purpose of substrate differentiation and energy-dependent pre-protein hydrolysis events. The 20S proteasesome is highly conserved from eukaryotes to archaea, making archaeal 20S proteasomes a useful model for understanding the eukaryal 20S proteasomal pathway.

In this study we employed the haloarchaeaon, Haloferax volcanii, to study energy- dependent proteolysis, including the 20S proteasome and its associated regulatory

xv particles, Pan (proteasome-activating nucleotidase). Through this study, two pan genes

have been identified in H. volcanii, which encode proteins (PanA and PanB) that

assemble into at least three distinct high molecular weight homo- and hetero-oligomeric

complexes (between 270 and 650 kDa). We have biochemically characterized the ATP-

hydrolyzing activity of one of these complexes (650 kDa homo-oligomeric PanA) and

determined its optimal activity to be in high concentrations of KCl (2-3.5 M), at 45-50ºC, near neutral pH buffered by MOPS, which relates well to previous reported activities of

20S proteasomes. We have also developed an in vivo system for reporting proteolysis in this organism using 32 unique green fluorescent proteins with variable C-terminal amino acids or peptide sequence fusions. While all additions to smRS-GFP negatively affected its protein level, effects were variable throughout DS70 growth and were least impacted by negatively charged amino acids and most impacted by hydrophobic amino acids. It has been determined through specific inhibition by clasto-lactacystin β-lactone that 20S proteasomes are, at least in part, responsible for the regulation of at least one of these variants, smRS-GFP-SsrA, which ends in a hydrophobic rich, 11-residue peptide sequence. This inhibitor is highly specific toward the proteasome, having a Ki of 40 nM.

xvi CHAPTER 1 LITERATURE REVIEW

Introduction

This literature review is designed to present the most current knowledge of energy- dependent proteolytic systems with emphasis being placed on protease-associated AAA regulatory particles. This review will highlight the mechanisms by which these regulatory particles are believed to operate and some of the properties of their substrate proteins that designate them as so. The final section will discuss the properties of green fluorescent proteins that have made them an excellent tool for the study of energy- dependent proteolysis.

ClpP Protease

ClpA was the first ATPase of the two-component protease systems to be described.

Together with the proteolytic complex ClpP (caseinolytic protease, named for their ability to stimulate the proteolysis of casein) they form the two-component protease,

ClpAP (Hwang et al., 1987; Katayama et al., 1988). ClpP, along with its associated regulatory components such as ClpX and ClpA, is well conserved throughout bacteria and eukaryotes, where they are found in plant cytosol and chloroplasts, and mammalian mitochondria (Gottesman et al., 1990; Corydon et al., 1998; Porankiewicz et al., 1999;

Kang et al., 2002). ClpP homologues are not conserved in archaea, although an open reading frame (ORF) from several methanogen genomes clusters to those of the ClpA

ATPase family. Characterization of proteins respective to these ORFs is needed before relevant functions can be assigned.

1 2

ClpP encompasses a vast number of physiological roles within the cell, and with a variety of similar yet distinct associated ATPases (e.g. ClpA, ClpX) it is proposed to form a number of complexes granting it a diversified substrate selection. Together these

Clp complexes are involved in such events as protein quality control (e.g., degradation of abnormal proteins and incompletely translated proteins tagged with SsrA), survival during stationary phase and other cellular stresses, the disintegration process of bacterial inclusion bodies, and even protein remodeling (Gottesman et al., 1998; Weichart et al.,

2003; Burton and Baker, 2005; Vera et al., 2005). Clp is also involved in sporulation, motility, and development of competence in Bacillus subtilis and in the virulence of bacterial pathogens such as Streptococcus pneumoniae and Salmonella typhimurium, among others (Msadek et al., 1998; Porankiewicz et al., 1999). In cyanobacteria and plants, ClpP seems to be less associated with cell stress and more involved in response to light and regulation of photosynthesis (Porankiewicz et al., 1999). Further study of ClpP and its associated ATPases is destined to yield an even greater understanding of its physiological impact.

The Escherichia coli ClpP is a serine protease whose subunits associate into the characteristic form of a dimer of heptameric rings creating an internal chamber that houses the proteolytic active sites (Kessel et al., 1995; Coux et al., 1996). Alone, ClpP is capable of degrading only short peptides and unfolded proteins and requires a regulatory

ATPase for effective proteolysis of native or aggregated proteins (Thompson and

Maurizi, 1994). Access to the central pore of ClpP is restricted due to the 10 Å diameter of its axial opening, which is just large enough for unfolded proteins and therefore limits the entrance of folded proteins (Wang et al., 1997). Like most AAA proteins, ClpA and

ClpX form homo-hexameric rings. One of these ATPases rings flanks either side of ClpP, where it unfolds substrate proteins and assists in translocation into ClpP (Beuron et al.,

1998). A loop region extending from each Clp ATPase subunit of the hexamer called the

“ClpP loop” is tipped with a conserved hydrophobic motif, [LIV]-G-[FL] (where G, the predominantly conserved residue in the middle of the triad, is typically flanked N- terminally by L, I, or V and C-terminally by F or L), and proposed to interact with the hydrophobic clefts of the ClpP heptamer asymmetrically (Beuron et al., 1998; Guo et al.,

2002; Kim and Kim, 2003). This interaction is believed to be responsible for ClpAP and

ClpXP complex formation. A homologous “ClpP loop” region has been found in Clp

ATPase subfamily members (ClpA/X/C/E) throughout many ClpP-containing organisms.

Furthermore, Clp ATPase subfamily members in ClpP-lacking organisms and non-ClpP associating ATPases (HslU) are devoid of a similar motif (Kim et al., 2001).

Binding of Clp ATPases to ClpP stimulates changes in both the protease and the chaperone, which suggests coordination between the two. ClpA/X stimulation of ClpP peptidase activity occurs simultaneously with a depression in ATPase activity, which is proposed to be due to conformational changes in ClpA/X (Thompson et al., 1994; Hwang et al., 1988; Kim et al., 2001). ATP hydrolysis by ClpX, promoted by additional substrate binding, releases proteins trapped in the degradation chamber of inactive ClpP

(Kim et al., 2000). One model suggests that there is a two-way communication between

ClpX and ClpP based on the nucleotide binding state (ADP vs. ATP) by ClpX or substrate association with ClpP, which affects the binding affinity of ClpX to ClpP (Joshi et al., 2004). In this model, the binding affinities of ClpX to substrate and ClpP are strong in an ATP-bound state and weak in an ADP-bound state. Communication in the

4 other direction is shown by an extreme increase in ClpX to ClpP binding affinity upon acylation of the ClpP active site serine, which mimics the acyl-enzyme intermediate in peptide hydrolysis (Joshi et al., 2004). The location of the acylated serine residues is within the degradation chamber of ClpP and thus cannot directly affect interactions with

ClpX (Wang et al., 1997). Although the mechanism is still not completely understood, this indicates that ClpX may be able to sense whether ClpP active sites are engaged with substrate.

The molecular contacts of ClpA/X to ClpP are located on the C-terminal end of the chaperone, while the outer face of the hexamer, including the distal extending N-terminal domain (N-domain), is proposed to recognize and interact with adaptor proteins and proteolytic substrates (Wickner et al., 1994; Ortega et al., 2000; Dougan et al., 2002a).

Adaptor proteins assist Clp ATPases in recognition and delivery of some substrates and are required in the case of certain substrates (Dougan et al., 2002a). The SspB adaptor forms a homodimer with flexible C-terminal tails that anchor the substrate-adaptor complex to the N-domain of ClpX (Dougan et al., 2003; Levchenko et al., 2005). This association actually lowers the Km for ClpXP degradation of SsrA-tagged substrates.

The master regulator of the general cell stress response in E. coli, σS (RpoS), is a substrate of ClpXP only when bound to the ClpX adaptor protein RssB (Zhou et al.,

2001). The affinity of RssB for σS, and therefore the turnover of σS by ClpXP, is dependent upon phosphorylation of RssB (Becker et al., 1999; Studemann et al., 2003).

Binding of the ClpA adaptor protein, ClpS, switches ClpAP mediated degradation of

SsrA-tagged proteins to aggregated proteins by changing the conformation of ClpA N- domains which prevents further binding of SsrA-tagged substrates and forms a higher

5 affinity binding site for aggregated proteins such as aggregated malate dehydrogenase

(Dougan et al., 2002b; Zeth et al., 2002). It has been determined through cryo-electon microscopy that the N-domains of ClpA are highly mobile and are proposed to permit fluctuations of up to 30 Å (Ishikawa et al., 2004). Ishikawa and colleagues conclude that mobility may allow for several scenarios. The N-domain may act as an “antenna,” initiating a weak binding of substrate before its handoff to another, stronger binging site for unfolding and translocation. Alternatively, it may act as an “entropic brush” by repelling other, non-specific proteins, whose unproductive binding may impede the binding of genuine substrates, therefore reducing the efficiency of ClpAP. Regardless of its purpose, the N-domain is not absolutely required for unfolding/proteolysis by ClpAP or ClpXP (Singh et al., 2001). This was demonstrated by the ability of ClpA∆N to retain similar ATPase activity as ClpA and maintain 50-100% unfoldase activity and ClpP activation (based on substrate used) (Singh et al., 2001). The deletion of ClpX N-domain had a more severe effect; however ClpX∆N still maintained 50% ATPase activity of

ClpX and maintained 30-80% unfoldase activity and ClpP activation (based on substrate used). In contrast to ClpXP there was one test substrate that ClpX∆N-P could not unfold/degrade, demonstrating that the necessity of the N-domain is dependent on the substrate. This raises the possibility that another substrate binding site exists in Clp

ATPases. Indeed a conserved motif [GYVG] in the central pore of ClpX contains residues essential for substrate binding (Siddiqui et al., 2004). ClpA contains three substrate-binding loops (two in D1, and one in D2) which lie in the central channel and also contains residues essential for substrate binding (Hinnerwisch et al., 2005).

Furthermore, Hinnerwisch and colleagues propose that the D2 loop may also participate

in substrate unfolding and/or translocation as mutation of this loop maintains ClpA-like substrate binding and ATP hydrolyzing activity, however is impaired in substrate degradation in the presence of nucleotide and ClpP. Another interesting dilemma involving the Clp ATPase/substrate interaction is determining how many substrate molecules can bind per hexamer. The stoichiometry of substrate to Clp ATPase was shown to be 1:1 when ClpA was tested with the SsrA peptide by isothermal titration calorimetry (ITC) (Piszczek et al., 2005). This is in agreement with the processive nature of the Clp ATPase/ClpP machine, as it would prevent accumulation of excess substrates waiting their turn to be degraded.

Much of the discussion so far has been about ClpA and ClpX ATPases, although it should be pointed out that other well-known Clp ATPases are present in many organisms ranging from bacteria to plants (ClpC, ClpB, ClpE, and ClpY). In Bacillus subtilis, ClpC and ClpE have been shown to associate with ClpP much like ClpA and ClpX (Gerth et al., 2004). The association of ClpY (HslU) with another protease, ClpQ (HslV), will be discussed in the next section. ClpB is not predicted to associate with any protease but, rather to function in the de-aggregation of protein aggregates with the assistance of

DnaK, DnaJ, and GrpE in response to cell stress (e.g., heat shock) (Zolkiewski, 1999).

Even more Clp ATPases have recently been identified (ClpD, ClpL), which are associated with responses to cell stress; but they have yet to be further characterized

(Shen et al., 2003; Suokko et al., 2005).

HslUV Protease

HslUV (heat shock locus UV) was first identified during a screen for bacterial heat

shock genes in E. coli (Chuang and Blattner, 1993). Many of the physiological roles of

HslUV overlap significantly with other ATPase-associated proteases in bacteria,

especially the Lon protease (Kuo et al., 2004; Fredriksson et al., 2005). This is

exemplified by the lack of a phenotype at normal growth conditions in an E . coli hslUV

knockout strain, although growth is impaired at high temperatures (Kanemori et al.,

1997). Arguably, the two most important roles of HslUV found thus far are its regulation

of the heat shock response through degradation of the heat shock factor σ32 and its impact on the SOS response by degradation of the cell-division inhibitor SulA (Kanemori et al.,

1999; Seong et al., 1999).

The sequence of HslU revealed it as a member of the AAA family of ATPases

while HslV was related to β subunits of 20S and 26S proteasomes, sharing approximately

20% sequence similarity (Chuang et al., 1993). The HslV protease is a double ring like

ClpP; however, it differs in subunit composition and type of proteolytic active site (Fig.

1-1). The proteolytic active sites in the HslV rings are threonine, again similar to

proteasomes and unlike the serine active sites of ClpP (Missiakas et al., 1996). Another

difference of the HlsV protease to ClpP, which also contrasts with proteasomes, is its

hexameric ring formation, which causes no symmetry mismatch between the hexameric

HslU and this protease (Kessel et al., 1996; Bochtler et al., 1997). Several structural

analyses have determined that both the N- and C-termini of HslU contact the apical

portion of HslV (Bochtler et al., 2000; Ishikawa et al., 2000). In the crystal structure of

HslUV from Haemophilus influenzae, a pair of HslV apical helices insert into a cleft of

HslU formed from the fold of the N and C domains. Furthermore, the extended C-termini

of the HslU subunits insert into the active-site clefts of HslV nearly reaching the active

sites (Sousa et al., 2000). This region is proposed to allosterically activate HslV due to

conformation changes of HslU during nucleotide binding. It has been shown that the

8 presence of nonhydrolysable ATP is sufficient to stimulate HslUV degradation of substrates that do not require unfolding (Rohrwild et al., 1996; Groll et al., 2005). This indicates that ATP binding, as opposed to ATP hydrolysis, is responsible for protease activation through allosteric effects of ATP-bound HslU on HslV. This should not be confused with the requirement of ATP hydrolysis for unfolding and translocation of substrates by HslU. Further proof of HslV activation is demonstrated by the fact that covalent complex formation between a peptide vinyl sulfone active site inhibitor and

HslV only occurs in the presence of HslU and nucleotide, suggesting that the access to proteolytic active sites are otherwise restricted (Bogyo et al., 1997; Sousa et al., 2002).

Additionally, in a reverse communication, it has been shown that HslV stimulates ATP hydrolysis by HslU (Yoo et al., 1996).

In sync with the Clp ATPases, HslU contains the conserved GYVG pore motif on its inner channel, which is likewise predicted to be involved in substrate association.

Furthermore, mutations in the HslU GYVG motif identify its role in substrate unfolding and potentially substrate translocation (Park et al., 2005). Considering small substrates require neither unfolding nor translocation, these HslU mutations had no effect on the ability of HslUV to hydrolyze the small fluorogenic peptide substrate Z-Gly-Gly-Lue-7- amino-4-methylcoumarin (AMC). In the presence of HslV and ATP, select HslU GYVG motif mutants are severely impaired in their ability to degrade the folded substrate SulA.

In contrast, these mutations inhibit degradation of the natively unfolded protein, casein, to a much lesser, but still significant, extent. Together these results suggest a role for the pore motif in protein unfolding as well as protein translocation.

Another domain of HslU implicated to be involved in substrate binding is the previously mentioned Intermediate (I-) domain. Recent studies have shown that the HslU with a deleted I-domain (HslU∆I) oligomerized and associated with HslV similar to wild type. Although the peptidase activity of HslU∆I -HslV on Z-Gly-Gly-Lue-AMC was half that of HslUV, it maintained absolutely no protease activity of a folded fusion substrate

(bacteriophage P22 Arc repressor fused to the SulA C-terminus) in contrast to the HslUV wild type (Kwon et al., 2003). This is consistent with earlier studies involving three I- domain mutants of E.coli HslU (∆137-150, ∆175-205, ∆111-239), all of which

maintained wild type-like activity toward Z-Gly-Gly-Lue-AMC and casein, while the

latter two were unable to degrade the folded fusion substrate, MBP-SulA (Maltose-

binding protein-SulA) (Song et al., 2000).

Lon Protease

The Lon protease was the first ATP-dependent protease to be studied (Charette et

al., 1981; Chung and Goldberg, 1981). Lon has been found throughout all domains of

life, functioning in the cytosol of eubacteria, and in mitochondria, peroxisomes and plant

plastids of eukaryotes (Chandu and Nandi, 2004). Most archaeal Lon proteases contain

transmembrane (TM) regions and, therefore appear to act as membrane-bound proteases

(Fukui et al., 2002; Besche et al., 2004;Botos et al., 2005). A homologue of the Lon

protease has also been found in infectious bursal disease virus strain P2 (IBDVP2),

although it lacks an ATPase domain (Birghan et al., 2000). Lon is responsible for the

degradation of many abnormal proteins and regulatory proteins involved in important

cellular functions such as radiation resistance, cell division, filamentation, capsular

polysaccharide production, and eukaryal mitochondria viability, among others (Goldberg

et al., 1994; Gottesman, 2003; Levine, 2005; Fredriksson et al., 2005). Interestingly, Lon

has also been found to degrade several ribosomal proteins in the presence of high levels

of inorganic polyphosphate (polyP), such as those found during amino acid starvation and

stationary phase (Kuroda et al., 2001). PolyP has also been proposed to play a role with

Lon in another of its functions, as a DNA-binding protein (Nomura et al., 2004).

The distinguishing feature between Lon and the previously mentioned chaperone-

assisted proteases is that the ATPase domain and protease domain of Lon are encoded on

the same protein. An additional Lon N-terminal (LAN) domain is found in the bacterial

and mitochondrial Lon, which presumably together with the AAA module can interact

with substrate proteins (Ebel et al., 1999; Maupin-Furlow et al., 2005). The presence of a

LAN domain is one factor distinguishing the subfamily LonA from the LonB subfamily.

Other characteristics classifying the subfamilies is the absence of TM regions in LonA

(found only in LonB subfamily members) as well as multiple other intra-family

homologous regions (Rotanova et al., 2004). Being a single component ATP-dependent protease complex, the protease Lon maintains symmetry between the ATPase and protease domain. However, variable accounts of the number of Lon subunits that form a complex have been reported. Archaeal Lon appears to form hexameric rings while the yeast Lon has been shown to form heptameric rings (Stahlberg et al., 1999; Besche and

Zwickl, 2004). The same E. coli Lon which formed hexameric rings in several studies was also reported in another study to form tetramers and octamers (based on the absence or presence of polyP, respectively) (Botos et al., 2004; Nishii et al., 2005). Therefore, it

remains inconclusive at this point whether or not Lon forms several physiologically

important oligomeric states from any one or more organism.

The classical proteolytic active site of Lon proteases studied is a Ser-Lys dyad

(Rotanova et al., 2004). Recently one group has proposed a second type of active site based on a Methanococcus jannaschii Lon, which contains a putative Ser-Lys-Asp triad as the active site (Im et al., 2004). Mutational analysis of two homologous acidic “triad”

residues (Asp508Ala or Glu506Ala) from Archaeoglobus fulgidus Lon displayed little to

no decrease in proteolytic activity, respectively. This result alone does not rule out an

active MjLon triad, as its Asp547 is not universally conserved (Botos et al., 2005).

Further molecular characterization of MjLonB is needed to settle this debate.

Similar to the other proteases to be described, mutational analysis of the

Thermoplasma acidophilum Lon suggests that there is communication between the

ATPase and protease domains via the arginine finger and sensor 2 regions (Besche et al.,

2004). In fashion with other ATPase associated proteases, many Lon proteases have been shown to processively degrade folded proteins in conjunction with ATP hydrolysis and can degrade unfolded proteins in the absence of ATP (Van Melderen et al., 1996;

Fukui et al., 2002; Ondrovicova et al., 2005).

Despite the abundance of Lon substrates known, properties targeting them for destruction by Lon have yet to be fully elucidated. It has been suggested that substrate recognition by Lon is more based on structural properties of the substrate than specific

tag sequences such as SsrA (von Janowsky et al., 2005). This has raised the question of

whether there is a distinct selection between substrates and non-substrates or whether

there is substrate selection based on varying degrees of affinity for Lon (Gottesman,

1996). Further studies on the characterization of Lon will hopefully lead to differentiation

of physiologically relevant substrates.

FtsH Protease

The FtsH protease was initially described from a filament-forming temperature-

sensitive E. coli mutant, which was later found to contain two mutant genes, one

responsible for the filamentation phenotype (ftsI, encoding a penicillin-binding protein)

and the other responsible temperature-sensitivity (ftsH) (Santos and De Almeida, 1975;

Begg et al., 1992; Zellmeier et al., 2003). FtsH is a cytoplasmic Zn++ metalloprotease found anchored in the cytoplasmic membrane of bacteria via two N-terminal transmembrane regions (Tomoyasu et al., 1995; Ito and Akiyama, 2005). It is also found

in plant chloroplasts and has homologs in yeast mitochondria, however, to date, it has not

been found in archaea (Tauer et al., 1994; Bailey et al., 2001). FtsH degrades some

short-lived cytosolic proteins, although appears to be more important in the degradation

of abnormal membrane bound proteins, highlighted by the following examples. FtsH

degrades some soluble substrates encoded by the bacteriophage λ, such as the cII

transcription factor of genes necessary for lysogenization (Shotland et al., 2000). FtsH is

essential for viability in E. coli for degrading LpxC deacetylase, thereby preventing a

lethal overaccumulation of lipopolysaccharide (LPS) and maintaining a balanced

LPS/phospholipid ratio (Ogura et al., 1999). It also appears to be required for sporulation

in Bacillus subtilis (Kotschwar et al., 2004). FtsH is also responsible for the degradation

of unassembled membrane proteins of the Sec translocation pathway (e.g. SecY) and F1F0

ATPase (Akiyama et al., 1996b; Akiyama et al., 1996a). Both SecY and subunit a of the

F1F0 ATPase have half-lives of ~2 minutes when not associated with SecE and F0, respectively. The turnover of photodamaged proteins of the light-harvesting Photosystem

II complex is implicated to be the responsibility of one of the numerous FtsH homologues

found in both cyanobacteria and plants (Garcia-Lorenzo et al., 2005; Komenda et al.,

2005).

Analogous to the Lon protease, the FtsH ATPase region and protease domain are

encoded on the same protein. FtsH is believed to form homo-hexameric rings which

undergo conformational changes upon ATP-binding like many other AAA proteins

(Akiyama and Ito, 2001; Niwa et al., 2002). E. coli FtsH undergoes a self-processing

event which removes seven C-terminal residues, an exception to its normally processive

nature. The physiological significance of this event remains undetermined as both forms

are proteolytically active (Akiyama, 1999). FtsH has a Zn++ metalloprotease proteolytic

active site (HEXXH) located in the C-terminal protease region of FtsH. Both Zn++ and

ATP (hydrolyzed in the AAA ATPase domain) are necessary for degradation of substrates (Tomoyasu et al., 1995; Ito and Akiyama, 2005). Although only enzymatic activity of the FtsH hexamer has been studied, the physiologically relevant form is a FtsH holoenzyme that associates with the proteins HflK and HflC into what is believed to be two hexamers (Kihara et al., 1996). The N-terminal domain of the FtsH homo-hexamer binds the membrane and interacts with HflKC and extends the rest of the FtsH hexamer outward to the cytoplasm. HflKC is composed as a trimer of hetero-dimers, also connected via a transmembrane region near their N-terminus, that associate with the transmembrane regions of FtsH. The rest of the HflKC hexamer extends opposite of

FtsH, into the periplasm (Kihara and Ito, 1998). Interestingly, the HflKC complex has been shown to negatively regulate FtsH activity towards some substrates such as SecY and to a varying degree cII (Kihara et al., 1998; Kihara et al., 2001). FtsH has been shown to degrade a cytosolic SsrA-tagged protein, which was tail specific, as mutations

in the C-terminal Ala residues of the tag (A10D and A11D) blocked its degradation

(Herman et al., 1998). However the degradation of membrane proteins is believed to be

dependent on the presence of ~20 amino acid cytoplasmic tail independent of its

sequence or end location (C- or N-terminal) (Chiba et al., 2000; Chiba et al., 2002).

This degradation does appear to be length dependent as successive truncations in the tail

continue to decrease the ability of FtsH to degrade the substrates. Interestingly, the

unfoldase activity of the FtsH ATPase appears to be less capable than that of some other

ATPases mentioned (e.g. ClpX, ClpA, and Pan) as it is unable to degrade the tightly-

folded secondary structure of GFP-tagged SsrA (Herman et al., 2003). Ito and Akiyama

even speculate that the AAA domain may act primarily as a “dislocase” for membrane-

integrated substrate (Ito and Akiyama, 2005).

20S Proteasome

Proteasomes are large multicatalytic proteases found universally distributed throughout Eukaryal and Archaeal domains and among gram-positive actinomycetes of the Bacterial domain (Lupas et al., 1997b; Maupin-Furlow et al., 2004; Wolf and Hilt,

2004). One main function of proteasomes is to maintain quality control of abnormal, misfolded, and denatured proteins for general protein turnover and in response to various forms of cell stress (Coux et al., 1996). In addition, proteasomes are central players in the regulation of many cellular processes such as cell division, transcription, metabolism,

and DNA repair among others (Dahlmann et al., 1989; Lupas et al., 1997a; De Mot et al.,

1999; Ustrell et al., 2002). In higher eukaryotic organisms the proteasome is also

involved in signal transduction, cell cycle control, cell differentiation, apoptosis,

embryonic development, and generation of antigenic peptides displayed to the immune

system by MHC class I molecules (Voges et al., 1999; Rock et al., 2004; Dhananjayan et

al., 2005; Hershko, 2005). Clinical relevance of the ubiquitin-26S proteasome pathway is demonstrated by its role in neurodegenerative diseases such as Alzheimer's disease and in many forms of cancer (Dimakopoulos, 2005; Mani and Gelmann, 2005). In fact, the

FDA has recently approved a proteasome inhibition drug, for treatment of relapsed multiple myeloma, which results from selective inhibition of malignant cell growth

(Montagut et al., 2005).

The structure of the 20S core particle (CP) of proteasomes, which is a 650-700 kDa cylinder composed of four, stacked heptameric rings, is strikingly conserved throughout the three domains (Volker and Lupas, 2002; Groll and Clausen, 2003) (Fig. 1-1). The two outer rings are composed of α-subunits and the two inner rings are composed of β-

subunits of a related superfamily (Coux et al., 1994). The complexity of subunit

arrangement, however, is quite variable. The CPs of bacterial species are typically

composed of 1 α-type and 1 β-type subunit and those of archaeal species are typically

composed of 1 α-type and 1 to 2 β-type subunits, although exceptions do exist (e.g. the

haloarchaeon Haloferax volcanii contains 2 α-type subunits) (Zuhl et al., 1997; De Mot

et al., 1998; Kaczowka and Maupin-Furlow, 2003). The composition of eukaryotic

proteasomes is much more complex. Yeast encode seven copies of each subunit which

arrange α1-7 and β1-7 in each respective ring and higher eukaryotes form additional

auxiliary 20S proteasomes (e.g. the immunoproteasome of vertebrates and

spermatogenesis-specific proteasome of insects) with even more subunits (e.g. β1i, β2i,

β5i) (Groettrup et al., 2001; Ma et al., 2002; Froment et al., 2005; Groll et al., 2005).

Furthermore, up to 23 different α- and β-type genes have been identified in plants (Fu et

al., 1998). The central chamber created by the two inner β-rings harbor the proteolytic

active site threonines, which are exposed at the N-terminus after an autocatalytic removal

of a propeptide, classifying the proteasome to the amino-terminal (Ntn) hydrolase family

(Brannigan et al., 1995; Lowe et al., 1995). In archaeal 20S proteasomes, this propeptide

processing occurs during the formation of the inner β-rings and requires the presence of

α-rings, presumably as a scaffold (Zwickl et al., 1994; Maupin-Furlow et al., 1998). The

maturation process in eukaryotic proteasomes is likely to be more complex as it has been

shown to occur in the nucleus and require additional “assembly proteins” (Ramos et al.,

1998; Witt et al., 2000; Lehmann et al., 2002). X-ray crystallographic analysis

determined the dimensions of eukaryal 20S CPs to be approximately 15 nm in length and

12 nm in width (Unno et al., 2002; Groll and Huber, 2005). Similar dimensions were

also found for archaeal 20S CPs from T. acidophilum (Ta20S proteasomes), which in

addition contained 1.3 nm axial pores (Lowe et al., 1995). These pores were presumably

revealed due to the absence of a N-terminal structure of the α-rings, caused by the mutant

proteasome used in the study, which incorporated α-subunits truncated of 12 N-terminal

residues (Groll and Huber, 2005). The small openings of the CP restrict folded proteins

from gaining access to the internal chambers. In addition, 20S proteasomes may be gated

at these pores by the N-termini of the α-subunits, which to open, appear to require the aid of regulatory particles (RP) such as Proteasome-activating nucleotidase (Pan) of archaea

and the 19S cap of eukarya (Groll et al., 2000; Kohler et al., 2001b; Benaroudj et al.,

2003). Deletion of the N-termini of these α-subunits creates an “open gate” formation

and reduces the need for the regulatory particles in protein degradation (Kohler et al.,

2001a).

In addition to gate opening, the proteasome associated AAA ATPase RPs also assist in substrate recognition/binding, substrate unfolding, and translocation of the unfolded substrate into the proteolytic chamber (Glickman and Maytal, 2002; Benaroudj et al., 2003; Smith et al., 2005). The regulatory particle of eukaryotic proteasomes,

termed the 19S cap, is composed of at least 17 subunits, forming 2 multi-subunit

substructures. The 19S RP caps either one or both ends of the 20S CP, possibly with the

assistance of a tethering protein, to form 26S proteasomes (Leggett et al., 2002; Kopp

and Kuehn, 2003). The outermost substructure, the “lid”, is composed of nine

Regulatory particle non-ATPases (Rpn) subunits and “base”, the substructure responsible

for ATP-hydrolysis and connection with the CP, is composed of six Regulatory particle triple-A type I proteins (Rpt) and 2 Rpn proteins (Rubin et al., 1997; Finley et al., 1998).

The lid portion of the 19S cap appears to be responsible for substrate recognition as

several of its subunits have been shown to recognize poly-Ubiquitinated (poly-Ub)

proteins and poly-Ub-binding proteins (Fu et al., 1998; Saeki et al., 2002). Furthermore,

removal of the lid from the 26S proteasome inhibits its ability to degrade poly-Ub

proteins (Glickman et al., 1998). Although Ub-lacking substrates can be degraded, the

ubiquitination pathway is predicted to be the most common method for substrate

recognition and degradation by 26S proteasomes, and will be discussed in more detail

later (Hoyt and Coffino, 2004). The 19S cap is not the only regulatory complex that

associates with eukaryotic proteasomes. The COP9 signalosome (CSN) also associates

with eukaryal proteasomes in participation of ubiquitination-dependent proteolysis and

has been proposed to compete with the lid for complex formation with the base of the

19S cap (Huang et al., 2005). In fact there exists numerous proteins that assist eukaryal

20S proteasome-dependent proteolysis acting either upstream or in direct association with

proteasomes. An important representative of these “middlemen” is the AAA protein

Cdc48. Cdc48 assists in the degradation of many proteasomal substrates, especially

Endoplasmic Reticulum membrane proteins (ER) (Romisch, 2005). Cdc48 functions in the binding and delivery of ubiquitinated proteins to the proteasome and may be involved in substrate preprocessing via an unfoldase activity (Elsasser and Finley, 2005; Richly et

al., 2005).

Like their 20S proteasome counterparts, prokaryotic proteasomal RPs consist of a

more simplistic subunit arrangement. A sole eubacterial protein, ARC (AAA ATPase

forming Ring-shaped Complexes) forms double-stacked homo-hexameric rings and is

believed to act as the regulatory complex to 20S CPs of actinomycetes (Wolf et al.,

1998). The regulatory proteins of many archaea, the aforementioned Pan, are homologs

of eukaryal Rpts (~40% similar) which have ATPase activity and unfoldase activity

toward substrates, alone or in concert with substrate degradation by 20S CPs (Zwickl et

al., 1999; Wilson et al., 2000). Despite much evidence that Pan is necessary for the

degradation of folded substrates by archaeal proteasomes, and although it is widely

accepted that Pan and 20S CPs associate during this process, only a few studies exist that

demonstrate this association. The first of these studies demonstrated Methanococcus

jannaschii (Mj)Pan/Ta20S CP association by electron microscopy (EM) (Wilson et al.,

2000). A more recent study demonstrated ATP-dependent association between Pan and

20S proteasomes using immunoprecipitation (IP) and surface plasmon resonance (SPR)

in addition to EM; however, 20S proteasomes of T. acidophilum, one of several archaea

lacking a Pan homologue were used (Smith et al., 2005). Evidence from both studies

19 suggests Pan/20S CP association requires ATP and Mg2+ and appears to be short-lived or require post-translational modifications or accessory proteins, as all proteins used in the study were from recombinant E. coli. The contact is likely occur between the C-terminus of Pan and the outer axial portion of the α-ring of the 20S CP as a truncation of the Pan

C-terminus by one residue or mutation of a highly conserved α-subunit Lys residue inhibits Pan-dependent degradation of substrates (Forster et al., 2005). Similar to other protease-associated AAA proteins described, the N-terminal region of Pan extends outward from the proteasome and is proposed to be involved in substrate recognition/binding (Smith et al., 2005).

Some archaea encode two Pan (PanA and PanB) proteins which can share up to

60% sequence identity and may result in a diversification of substrates targeted for proteolysis by the 20S proteasome (Maupin-Furlow et al., 2004). Another archaeal protein, related to the eukaryal Cdc48 proteins, may further promote variable substrate recognition in association with 20S proteasomes and is proposed to be the primary RP in archaea lacking Pan homologs. This protein, known as VAT (Valosin-Containing Protein

(VCP)-like ATPase), is universally conserved in archaea (Golbik et al., 1999). The T. acidophilum VAT forms homo-hexameric rings and has been shown to degrade folded substrates in an ATP-dependent manner (Rockel et al., 2002). It shares a pore motif

(KFIG, also highly conserved in Pan) with other AAA unfolding proteins (GYVG in

ClpA and ClpX) which is exposed in its pore and is crucial in the binding and unfolding of GFP-SsrA (Gerega et al., 2005). Interestingly, VAT has been shown to unfold non- tagged GFP, a trait not shared by other AAA+ proteins studied, possibly suggesting

substrate recognition in response to lack of structure (as in the unstructured 9-residue C-

terminal tail of GFP) as opposed to a specific sequence (Gerega et al., 2005).

To date, ubiquitin and Rpn homologues have not been discovered in prokaryotes,

suggesting an alternative route of substrate recognition by the RPs that has yet to be

characterized. Several groups have suggested the existence of ubiquitin-like proteins in

prokaryotes based on structural rather than sequence similarity, however further

characterization of these proteins is needed before any conclusions can be drawn to their

physiological relevance (Nercessian et al., 2002; Bienkowska et al., 2003). It is

reasonable to assume that other players exist in prokaryotic proteasome-dependent

proteolysis which are yet undiscovered. Also elusive are the factors that make

proteasomal substrates in archaea susceptible to recognition and subsequent degradation.

AAA+ ATPases

Proteins of the AAA+ superfamily (ATPases associated with various cellular activities) are universally related by the presence of an AAA domain (Neuwald et al.,

1999). Many of these proteins share a similar mechanism and structure, often forming oligomeric rings and interacting with other molecules (proteins and nucleic acids) for the purpose of substrate assembly, disassembly, or rearrangement (Iyer et al., 2004). The importance of AAA+ proteins is demonstrated by their existence in high abundance in all domains of life (Ogura and Wilkinson, 2001; Iyer et al., 2004). The limited requirement for membership to the AAA+ family compiled with the extreme diversity in cellular function undoubtedly led to further classification into subfamilies and numerous clades

(Iyer et al., 2004). The AAA subfamily is distinguished in part by the presence of a

second region of homology (SRH) although moreover by statistical values of

phylogenetic analyses (Frickey and Lupas, 2004). This is just one example of the ever-

evolving, tangled web that is the AAA+ classification system. Focus in this review will

be placed on the features and mechanisms of AAA ATPases associated with energy- dependent proteolysis including ClpX, ClpA, HslU, and Lon and classical AAA subfamily members FtsH and ATPases associated with proteasomes.

AAA+ Domain

The defining AAA+ domain is a conserved region of approximately 200-250 residues consisting of Walker A and B motifs, and sensor 1 and 2 regions (Hanson and

Whiteheart, 2005). AAA+ proteins are classified as type I and type II depending on the presence of one or two AAA+ domains, respectively. In type II AAA+ proteins these domains are often referred to as nucleotide-binding domain 1 and 2 (NBD1 and NBD2) and respectively located at the N- and C-termini. Although the NBD1 and NBD2 domains of one protein are somewhat divergent, they are well conserved with their respective domains in orthologous proteins (Maurizi and Xia, 2004). The AAA+ domain of type I members is more closely related to NBD 2 of type II members. Type I members include ClpX, HslU, Lon, FtsH, Pan, Rpt proteins, whereas type II members include

ClpA, VAT, and Cdc48 (Fu et al., 1999; Maupin-Furlow et al., 2003; Wang et al., 2004;

Weibezahn et al., 2004). The secondary structure of AAA proteins forms an α/β

(Rossman) fold from the middle to N-terminal region while the C-terminal end forms an

α-helical subdomain (Maurizi and Xia, 2004; Hanson and Whiteheart, 2005). The

following sections will highlight each subset of the AAA+ domains, and the SRH unique

to the AAA subfamily.

Walker Motifs

The Walker motifs are primarily responsible for ATP and Mg++ binding and nucleotide hydrolysis (Walker et al., 1982). The Walker A motif forms a phosphate-

binding loop (P-loop) between an α-helix and a β-strand and contains the consensus

sequence GX4GK[T/S] (where X is any amino acid) (Saraste et al., 1990). This motif

coordinates Mg++ and forms hydrogen bonds with the β- and γ-phosphates of nucleotide

triphosphates (NTP). A mutation of a universally conserved Lys residue abolishes

nucleotide binding and, therefore, function of the enzyme (Mogk et al., 2003; Besche et

al., 2004; Gerega et al., 2005). Forming the other side of the nucleotide-binding pocket

is the Walker B motif, which binds nucleotide as well as Mg++. The Walker B consensus sequence is typically hhhhDE (where h is a hydrophobic residue) and mutations in either acidic residue impair ATP hydrolysis (Weibezahn et al., 2003).

Sensor Regions

The sensor regions interact with the γ-phosphate NTP and as the name implies can sense the state of the nucleotide-binding pocket (ADP vs. ATP). Sensor 1 region slightly overlaps the N-terminal region of the SRH, and it contains an essential conserved polar residue (typically Thr or Asn) (Guenther et al., 1997; Hanson and Whiteheart, 2005). A mutation of this polar residue impairs ATP hydrolysis (Hattendorf and Lindquist, 2002b;

Besche et al., 2004). The sensor 2 region is located near the C-terminal end of the AAA+ module in the α-helical region and appears to interact with the γ-phosphate of the NTP and may regulate activity by means of inter-subunit conformational changes associated with nucleotide binding and hydrolysis (Hattendorf and Lindquist, 2002a). Expectedly,

mutations in this region impair ATP hydrolysis and in some cases ATP binding (Joshi et

al., 2003; Ogura et al., 2004).

Second Region of Homology

The SRH domain unique to AAA proteins is positioned immediately C-terminal of the sensor 1 motif where it coordinates NTP hydrolysis coupled with inter-subunit conformational changes (Hanson and Whiteheart, 2005). There is an arginine finger in the consensus sequence (X[T/S][N/S]X5DXAX2RX2RX[D/E]) (where X is any amino

acid) which is crucial for all known activities of the SRH, although altering most of these

residues causes impaired ATP hydrolysis (Wang et al., 2005). Mutations of the Arg

residues affect ATP hydrolysis but not ATP binding. This is demonstrated by an

impaired ATP hydrolyzing activity despite continued conformational changes caused by

ATP binding (Karata et al., 1999). In the case of some proteins, mutations of the Arg

residue also inhibits oligomerization of the ATPase (Ogura et al., 2004; Wang et al.,

2005). Interestingly, this deficiency in oligomerization results from substituting the Arg

with the acidic amino acid Glu and is not observed with a hydrophobic amino acid

replacement (Leu or Ala) (Wang et al., 2005). These mutations, which in two separate

occasions were the result of an Arg replaced by an acidic amino acid (R/E), did not affect

oligomerization in two occasions where the Arg was replaced by a hydrophobic amino

acid (Lue or Ala) (Ogura et al., 2004).

Coiled-coil Domains

Coiled-coil domains are found in an extensive group of proteins and, although not

universal to AAA+ proteins, are found in many of the protease-associated ATPases

described in this review. Coiled-coils are mechanically rigid structures made up of

intertwined α-helices with amphiphilic properties (Lupas, 1996). In protease-associated

ATPases, coiled-coil domains are proposed to bind and interact with substrate proteins,

possibly by recognizing exposed hydrophobic residues of non-native proteins. Most of

these domains are located at the N-terminus, where they extend outward from the

ATPase-protease complex, a structurally relevant position for their proposed function.

This proposed function has also been reinforced by a deletion of the coiled-coil domains

in several of these proteins (Zhang et al., 2004). In HslU, the coiled-coil domains are part

of an Intermediate-domain (I-domain) that is not at the extreme N-terminus (Maurizi and

Xia, 2004). Although there is still a debate as to where the I-domain is spatially located, it has been shown that deletion of this domain is essential for the degradation of folded substrates (Song et al., 2000; Kwon et al., 2003; Groll et al., 2005). It is interesting to speculate that variability in the coiled-coil domains of homologues intra-species protease- associated ATPases creates an ability of diversified substrate recognition. For instance, homologues PanA and PanB of Haloferax volcanii share an overall 60% identical and

78% similar protein sequence; however, they are only 17% identical in the coiled-coil domains (Reuter et al., 2004).

Pore Motifs

The pore motif, in addition to the coiled-coil domain, appears to be responsible for substrate binding and may participate in substrate unfolding and translocation. A ring of pore motifs, which have the conserved motif (hahG) (where “h” is a hydrophobic and “a” is an aromatic amino acid), form short loops and line the interior of the pore of protease- associated AAA+ complexes (Hanson and Whiteheart, 2005). Mutations in this motif, particularly those that alter the aromatic and Gly residues, severely impair substrate binding and processing, often times with little affect on ATP hydrolysis (Yamada-

Inagawa et al., 2003; Siddiqui et al., 2004; Gerega et al., 2005; Park et al., 2005).

Mechanistic Action of ATPases

Protease-associated ATPases act as protein unfolding machines fueled by ATP.

Substrate binding successively follows ATP-binding and likewise stimulates ATP hydrolysis (Hwang et al., 1988; Benaroudj et al., 2003) (Ogura and Wilkinson, 2001;

Martin et al., 2005). Given that the majority of AAA+ ATPases form hexameric rings, most or all of whose subunits are capable of binding/hydrolyzing ATP, there has been much debate as to the order of ATP hydrolysis among subunits. Several models have been proposed for the order of this event. The concerted hydrolysis model predicts a simultaneous ATP binding/hydrolysis by most or all of the ATPase subunits (Gai et al.,

2004). The sequential hydrolysis model suggests hydrolysis in one subunit is sequentially followed by hydrolysis in an adjacent subunit and so on, creating clockwise

ATP hydrolysis (Stitt and Xu, 1998; Singleton et al., 2000). Most recently proposed is the probabilistic hydrolysis model where ATP hydrolysis is randomly occurring among subunits (Martin et al., 2005). While the former two models were predicted using DNA- associating AAA+ ATPases, the later used ClpX as a model. Whether or not there is a difference in ATP hydrolyzing order due to a divergence in function remains to be seen.

Regardless of the order, there is no debate that ATP hydrolysis is necessary for substrate unfolding. The question however, lies in how the unfolding occurs. Mechanisms of translocation have been proposed for ClpA and ClpX that use ATP hydrolysis to actively push or pull the substrate through the pore, simultaneously creating a strain on the folded portion of the protein, therefore denaturing it en route to the proteolytic core of ClpP

(Matouschek, 2003). This scenario, which implicates substrate unfolding and translocation are a linked process, is contradicted by a study with Pan where substrate was unfolded even when translocation was prevented (Navon and Goldberg, 2001).

Recently Smith et al. (2005) demonstrated that ATP binding alone to Pan or the 26S

ATPases can stimulate gate opening and translocation. The authors conclude that

unfolding is the only step that requires ATP hydrolysis and that translocation is therefore

passive. Furthermore they believe that retrograde movement is blocked by a Brownian

ratchet mechanism that that has been documented in other translocation processes. In the

middle of these two possibilities is the model proposed by Kenniston et al. (2003), where substrate denaturation is a combination of unfolding and translocation by ClpX and each require ATP hydrolysis. Unfolding is a consequence of a constant repeated application of an ATP-fueled force to a substrate. Respectfully, this author concludes that despite many similarities between protease-associated ATPases, differences are bound to exist. They go on to include as an example the unfolding of GFP substrate as being “kinetically irrelevant for the degradation by Pan/20S proteasome, an impediment to degradation by

ClpXP and a step that FtsH is unable to perform” (Kim et al., 2000; Benaroudj et al.,

2003; Herman et al., 2003). Overall the changes in protease-associated ATPases that are

responsible for substrate unfolding are not well understood. Based on conformational

changes of protease-associated ATPase upon ATP hydrolysis, it is speculated that a mechanical force is somehow applied to substrate in order to break down the secondary structure (Wang et al., 2001; Kenniston et al., 2003; Wang, 2004; Hanson and

Whiteheart, 2005).

Substrate Recognition

Considering the massive impact of energy-dependent proteolysis upon vast cellular processes, it is no wonder there is a requirement for carefully controlled regulation.

Energy-dependent proteases have the ability to alter protein half-life from years to

seconds. Without a strict set of guidelines, proteases would quickly ravage the proteome

27 by degrading every protein in reach leading to cell death. The majority of proteolytic regulation is based on substrate selection. Numerous degradation signals can be exposed, altered or added to proteins that allow their recognition by protease-associated ATPases.

Many of the known substrate recognition signals will be discussed here, keeping in mind that not all are completely understood and some have yet to be discovered (e.g. substrate recognition in Archaea).

Ubiquitination

The ubiquitin-26S proteasome (UPS) pathway is the major mechanism for protein degradation by eukaryotic organisms (Ciechanover, 1994). Ubiquitin (Ub), a highly conserved 76 amino acid protein, is added to proteasome substrates in a multiple step process. The first three steps in this pathway are mediated by a group of generically designated proteins; E1, E2, and E3. E1, the Ub-activating enzyme, activates Ub in an

ATP hydrolyzing reaction creating a thiolester bond on its C-terminal Gly residue which forms an E1-Ub intermediate. Activated Ub is then transferred to E2, the Ub- conjugating enzyme (UBC), by transesterification. E3, the Ub-protein ligase, then attaches the Ub Gly residue to a free Lys residue (typically Lys48) of a previously bound substrate via an isopeptide bond (Ciechanover, 1998). E3 enzymes are the components of the UPS pathway responsible for substrate recognition. Their importance is demonstrated by the existence of a vast group of corresponding genes (>1200 in plants) which suggests a non-redundant role (Vierstra, 2003). At this point not all proteasomal substrates have a designated individual E3 enzyme, although there have been classes of

E3 enzymes proposed, each of which recognize variable proteolytic substrate determinants (Robinson and Ardley, 2004). After the initial Ub ligation multiple rounds of ubiquitination usually follow, sometimes with the assistance of an additional

28 conjugation factor (E4), ultimately producing a poly-ubiquitinated substrate that is either directly or indirectly presented to the 26S proteasome (Hoppe, 2005). Intrinsic proteasomal subunits can directly bind poly-Ub substrates. Several Rpn proteins and possibly one Rpt protein of the 19S cap have the ability to bind Ub, possibly sequestering via structural and hydrophobic interactions, for unfolding of the associated substrate by other Rpt proteins in the 19S cap (Deveraux et al., 1994; Elsasser et al., 2002; Lam et al.,

2002; Smalle et al., 2003). Ubiquitin, however, appears to escape proteolytic destruction, as several Rpn subunits have an activity that releases poly-Ub chains from the substrate before it is degraded (Verma et al., 2002; Yao and Cohen, 2002). Eukaryotes additionally have a family of debiquitinating enzymes (DUBs) which facilitate ubiquitin removal (Wilkinson, 2000). Indirect transport to the 26S proteasome can be achieved by escorting poly-Ub substrates via Ub receptor proteins. These proteins typically have a

Ub-associated domain (UBA) which binds Ub, and a Ub-like domain (UBL) which is structurally similar to Ub and is recognized by 26S proteasomes (Elsasser and Finley,

2005). The AAA ATPase Cdc48 belongs to a related family of receptor proteins and will be discussed in more detail later. It should be noted that not all ubiquitinated proteins are destined for destruction by the UPS pathway. Factors including the amount of Ub moieties added to a protein and specific Lys residue linkage among others influence the fate of a protein in a myriad of other cellular processes. Alternatively, not all proteins recognized and degraded by 26S proteasomes are ubiquitinated. A subset of proteasomal substrates are degraded without Ub, usually in conjunction with the protein antizyme which can compete with Ub for proteasomal recognition (Hoyt and Coffino, 2004).

N-end Rule

The N-end rule of proteolytic substrate recognition is based on the half-life (due to proteolysis) of a protein depending on the extreme N-terminal residue (N-degron) of that protein (Gonda et al., 1989). The N-end rule has thus far been demonstrated in eukaryotes and eubacteria with some variation, and its presence in archaea has yet to be determined. Primary, secondary and tertiary destabilizing residues and stabilizing residues compose a hierarchy of amino acids that designate the increasing half-life of a protein respectively (eubacteria lack tertiary order). The lone nonconformist residue, Pro, is designated as having an uncertain status as proteins with this N-terminal residue have variable half-lives (Varshavsky, 1997). The primary destabilizing residues of eubacteria

(Lue, Phe, Trp, and Tyr) assign proteins a half-life of 2-3 minutes. Leu/Phe-tRNA- protein transferase (L/F-transferase) mediates the conjugation of a primary destabilizing residue (predominantly Leu) to secondary destabilizing residues (Arg or Lys) subsequently decreasing their protein half-life (Tobias et al., 1991). Proteins with primary destabilizing residues are typically degraded by ClpAP, possibly with the assistance of accessory proteins (Lupas and Koretke, 2003). Proteins with N-termini consisting of stabilizing residues (all others except Pro) grant proteolytic exemption for

>10 hours.

As with many pathways, the N-end rule in eukaryotes is slightly more complex.

The conversion of tertiary destabilizing residues (Gln and Asn) into secondary destabilizing residues (Glu and Asp) (Cys is also a secondary destabilizing residue in mammals) is mediated by an N-terminal amidohydrolase (Nt-amidase) (Levy et al.,

1996). Addition of a primary destabilizing residue is achieved by conjugation of an Arg residue to a secondary destabilizing residue by Arg-tRNA-protein transferase

(Varshavsky, 1997). Primary destabilizing residues are further subdivided into type 1

(basic residues; Arg, Lys, and His) and type 2 (bulky hydrophobic residues; Phe, Leu,

Trp, Tyr, and Ile) based on the specificity of their N-recognin binding site (Varshavsky,

1997). The N-recognin proteins are a class of N-degron recognizing E3 proteins,

discussed previously, that tag N-end rule proteins with Ub for degradation by the 26S

proteasome. An additional N-recognin of mammals has demonstrated binding of another

group of primary destabilizing residues termed type 3 (Ala, Thr, and possibly Ser)

(Hershko, 1991; Levy et al., 1996). All residues not designated as destabilizing (except

Pro) grant proteolytic exemption for >30 hours.

SsrA Tagging

A unique system of eubacteria, SsrA (small stable RNA A) protein tagging employs a transfer-messenger RNA (tmRNA) that encodes an 11-residue tag recognized

by protease-associated ATPases (Komine et al., 1994; Gillet and Felden, 2001). An Ala-

charged SsrA tmRNA is recruited to the A-site of a stalled ribosome due to an incomplete

or untranslatable message. The faulty mRNA is then replaced by the open reading frame

of the tmRNA which encodes a 10-residue SsrA sequence resulting in the addition of an

11-residue SsrA tag to the C-terminus of the protein (AANDENYALAA) (where A

represents the original Ala charged to the tmRNA) (Karzai et al., 2000). The SsrA-

tagged protein is then targeted for degradation by proteases such as ClpXP, ClpAP, FtzH,

and Lon (Gottesman et al., 1998; Herman et al., 1998; Smith et al., 1999). The

recognition of SsrA-tagged proteins by multiple proteases suggests its role in serving as a

relatively generic proteolytic signal. In fact, the Pan/20S proteasome has been shown to

degrade SsrA-tagged proteins, despite the lack of the SsrA system in archaea (Benaroudj

and Goldberg, 2000). The determinants for SsrA signal recognition have been

determined for ClpAP and ClpX and its adaptor protein SspB. ClpA interacts with

residues 1, 2, and 8-10 whereas ClpX interacts with residues 9-11 (Flynn et al., 2001).

Altering only 2 residues (A10D and A11D) greatly reduced degradation by ClpXP

(Gottesman et al., 1998). SspB interacts with residues 1-4, and 7, therefore

complementing SsrA tag binding with ClpX and impairing SsrA tag binding to ClpA

(Flynn et al., 2001). This demonstrates how multiple protein interactions with substrate

can influence proteolytic substrate recognition.

Other Modes of Proteolytic Substrate Recognition

Additional signaling factors of proteolytic substrates exist including post-

translational modification and the presence of certain groups of amino acids. Several

forms of post-translational modification can stimulate or alter the proteolytic fate of

proteins. N-glycosylation of some endoplasmic reticulum (ER) proteins can serve as a degradation signal recognized by a subclass of E3 proteins (Yoshida et al., 2002).

Phosphorylation of proteins has been shown to modulate the poly-ubiquitination capacity

of substrates therefore altering their recognition by the UPS pathway (Harari-Steinberg

and Chamovitz, 2004; Gross et al., 2005). Competition of Ub can be achieved in some

proteins by acetylation of shared Lys residues, so that ubiquitination and therefore

degradation is prevented upon acetylation (Li et al., 2002; Giandomenico et al., 2003).

Oxidative modification of proteins has also been demonstrated to act as a signal of 20S

proteasomes and eubacterial HslUV and Lon proteases (Darwin et al., 2003; Iwai, 2003;

Fredriksson et al., 2005). In eukaryotes, proteolytic recognition of oxidatively modified

proteins appears to be Ub-independent and heavily relies on the exposure of hydrophobic

surface patches due to partial unfolding upon oxidation (Grune et al., 2003). In fact, it

has been known for some time that surface exposure of hydrophobic residues, due to

32 many events, is an important determinant for proteolytic recognition (Bohley, 1996).

Alternatively, hydrophilic patches of amino acids are also a proposed signal for proteolysis such as PEST sequences (regions enriched with the residues Pro(P), Glu(E),

Ser(S), and Thr(T)) (Rechsteiner and Rogers, 1996).

Many proteolytic substrates, and the factors that designate them as so, have been discovered to date. However there are still many more unidentified substrates and substrate recognition factors. Substrate-binding arrays, substrate traps, reporter proteins, and mass spectrometry-coupled 2-D PAGE proteome analysis have and will continue to be valuable tools for the characterization of proteolytic substrates (Flynn et al., 2003;

Weibezahn et al., 2003; Reuter and Maupin-Furlow, 2004; Burton et al., 2005). One of the most widely used reporter proteins for the study of proteolysis is the Green

Fluorescent Protein (GFP). The unique properties of this protein, which make it an ideal reporter of proteolysis, will be discussed in the following section.

Green Fluorescent Protein (GFP) as A Model Substrate

Since the GFP protein was fist isolated from jellyfish (Aequorea victoria) in 1962 by Shimomura and colleagues, its gene has been cloned and optimized for a vast array of molecular biological uses (Shimomura et al., 1962). Hundreds of mutations and fusions have produced a genetic library of GFP genes encoding proteins with characteristics ranging from a variety of light intensities and wavelength excitation and emissions to signals determining its cellular fate (e.g. protein secretion, degradation or localization).

This section will discuss properties of GFP, especially those advantageous for its use as a reporter of energy-dependent proteolysis.

GFP is a 238 amino acid protein with a secondary structure resembling a β-can

(or β-barrel), formed by an eleven strand antiparallel β-sheet, that houses and protects the fluorophore (Ormo et al., 1996). This β-barrel has a diameter of 30 Å, a length of 40 Å, and has “caps” on both ends in the form of short helices and loops. GFP can exist as a monomer or dimer, where dimerization occurs in high protein (5-10 mg per ml) and salt concentrations and is inhibited in the red-shifted mutant (discussed below) (Ward and

Chang, 1982; Tsien, 1998;). The centrally located internal fluorophore is part of an irregular helical structure consisting of a Ser-Tyr-Gly sequence, of which the Gly residue appears to be essential (Heim et al., 1995; Cormack et al., 1996). The mechanism of fluorophore formation begins with a rapid cyclization of the Ser and Gly residues to form an imidazolin-5-one intermediate. This is followed by a rate-limiting oxygenation of the

Tyr side-chain by molecular oxygen ultimately producing the fluorophore as a p- hydroxybenzylidene-imidazolidone (Cody et al., 1993; Heim et al., 1994). This autocatalytic fluorophore generation occurs independently of co-factors or other enzymatic components and is necessary for fluorescence. Although the Ser-Tyr-Gly sequence of residues is found in many other proteins, the lack of a suitable surrounding inhibits fluorophore formation. In addition, an array of other amino acid side chains also contact the fluorophore in various ways, adding to fluorescent properties of GFP.

Many mutations of GFP have produced extremely useful variants exhibiting increased solubility, preferential codon usages, and a multitude fluorogenic light intensities and color emissions (Cubitt et al., 1995; Lippincott-Schwartz and Patterson,

2003). Green fluorescence is ultraviolet (UV)-stimulated in wild type GFP with optimal excitation (ex.) and optimal emission (em.) at 395 nm and 509 nm, respectively.

Fluorophore variants of GFP have been produced that have an altered optimal em., ex., or

both. These variants are ideal for use in dual-labeling and in cases where innate

background emission limits detection at particular excitation wavelengths. These include

the mutations S65T which creates a red-shifted GFP (RS-GFP) (ex. 488, em. 509), Y66H

which creates a blue fluorescent protein (BFP)(ex. 380, em. 440), Y66W which creates a cyan fluorescent protein (CFP)(ex. 433, em. 453), and mutations S65G, V68L, S72A, and

T203Y which create a yellow fluorescent protein (YFP)(ex. 513, em. 527), among others

(Cubitt et al., 1995; Heim and Tsien, 1996; Ormo et al., 1996; Miyawaki et al., 1997).

Early attempts at GFP expression in bacteria, plants, and mammals produced inadequate amounts of protein due to the existence of rare codons and in some cases splicing regions

(Haseloff and Amos, 1995). Since then gfp genes with optimized codon usages have been designed for these organisms, resulting in a much higher protein yield (Zolotukhin et al., 1996; Davis and Vierstra, 1998; Franklin et al., 2002). GFP protein production in some organisms has also been hindered by protein insolubility. Crameri and colleagues created more soluble GFP variants by screening for increased whole-cell fluorescence of

E. coli, expressing gfp genes after PCR-induced mutagenesis (Crameri et al., 1996). The most drastic of these proteins was found to have several amino acid substitutions that decreased hydrophobicity (F99S, M135T, and V163A) resulting in less aggregation and a more intense fluorescence signal.

The native structure and advantageous mutations that allow GFP to maintain stability and active over a wide range of conditions make it an ideal reporter protein.

GFP is active through a wide range of pH (5.5-12), temperature (~4-70 °C), and salt concentrations (smRS-GFP is active to at least 3M NaCl or KCl) (Bokman and Ward,

1981; Tsien, 1998; Reuter and Maupin-Furlow, 2004; Reuter, unpublished). GFP is also

stable in the presence of many chemical reagents such as detergents (e.g. 1% SDS),

chaotropes (e.g. 8M urea or 4M guanidine HCl), fixatives (3% formaldehyde), and

organic solvents (50% ethylene glycol) (however, these usually shift the em./ex.

spectrum) (Chalfie et al., 1994; Kain et al., 1995). GFP is less susceptible to

photobleaching than other fluorescent compounds such as fluorescein (Wang and

Hazelrigg, 1994; Niswender et al., 1995;). GFP also demonstrates resistance to many

common proteases (trypsin, chymotrypsin, papain, thermolysin, elastase, ficin,

proteinase K, chymopapain, subtilisin, pancreatin, bromelain, and pronase) (Kain et al.,

1995). GFP avoids degradation by many energy-dependent proteases in vitro, including

EcClpAP and MjPan/Ta20S proteasomes produced from recombinant E. coli (Navon and

Goldberg, 2001). One of the few protease-associated ATPases that has been shown to recognize and unfold GFP is TaVAT, and this event is carried out to a limited extent

(Gerega et al., 2005).

The same properties of GFP that make it resistant to most proteases ensures specific protease recognition of tags and fusion proteins of GFP and not GFP itself. GFP variants are used as reporters of many proteolytic systems by the addition of one or more amino acids or partial or complete fusion proteins typically to the C-terminus. An unstructured nine residue C-terminal “tail” of GFP suggests that additions to this region have no impact on the overall structure (Ormo et al., 1996). Probably the most widely

used recognition tag added to GFP is the SsrA tag. Energy-dependent proteolytic

degradation of GFP-SsrA has been demonstrated in ClpXP, ClpAP, Lon, Pan/20S

proteasome, and VAT/20S proteasome (Gottesman et al., 1998; Navon and Goldberg,

2001; Gerega et al., 2005). A circular dichroism study of GFP vs. GFP-SsrA concluded

the tag has little if any impact on the overall structure or fluorescence of GFP (Gottesman

et al., 1998). Partial or whole proteins known to be proteolytic substrates of certain

proteases have also been fused the C-terminus of GFP for reporting degradation (e.g.

GFP-RepA and GFP-MuA) (Hoskins and Wickner, 2006). Even entire libraries have

been fused to GFP for the purpose of screening for substrates of 26S proteasomes in

eukaryotic cells (Jiang et al., 2004; Salomons et al., 2005). The use of GFP as a

proteolytic reporter protein will undoubtedly continue to advance the characterization of

proteases and their substrates.

Research Objective

The objective of this study is to investigate proteolytic regulation in archaea,

utilizing Haloferax volcanii as a model organism due to its established genetic system,

ease of culture, and presence of multiple proteasome and Pan isoforms. An in vivo assay

has been designed to measure proteolysis using 32 unique green fluorescent reporter

proteins with variable C-terminal amino acid and peptide fusions. This effort attempts to

elucidate a trend in the properties of exposed C-terminal amino acids that illicit

susceptibility of a protein to proteolysis. As with many proteins, the protease-associated

AAA ATPases of archaea are less characterized than their bacterial and eukaryal

counterparts. Investigation of these archaeal proteins should lead not only to a better

understanding of proteolytic regulation in archaea, but also assist in the understanding

AAA ATPases in general. This study focuses on the presence of multiple Pan isoforms in H. volcanii and attempts to characterize these proteins. In doing so, this study marks

the first of several events: Pan characterization from an organism containing multiple

Pan homologs; utilization of Pan expressed from its native organism in in vitro experiments; characterization of Pan from an archaeon other than a methanogen.

Fig.1-1. Predicted structures of proteasomes. Molecular surface of (a) the archaeal, (b) the eukaryotic 20S and (c) the HslV proteasomes. To mark the position of the active sites, the complexes are shown with the bound inhibitor calpain (yellow). (d) Ribbon plot of the free α-ring from Archaeoglobus fulgidus, focusing on the defined N-termini (red). (e,f) Electron density maps of the yeast CP from the wild type (wt) and α3∆N mutant, respectively. The individual N-terminal tails of the α-subunits are depicted in different colors. Reprinted from Current Opinions in Structural Biology, Vol. 13, Michael Groll and Tim Clausen, “Molecular shredders: how proteasomes fulfill their role”, Pages 665-673, Copyright 2003, with permission from Elsevier.

CHAPTER 2 ANALYSIS OF PROTEASOME-DEPENDENT PROTEOLYSIS IN HALOFERAX VOLCANII, USING SHORT-LIVED GREEN FLUORESCENT PROTEINS

Introduction

The tree of life is divided into three main evolutionary lineages, the Archaea,

Bacteria and Eukarya domains (Woese et al., 1990). Despite the fact that archaea are prokaryotes, their information machinery (DNA replication, transcription, translation) and protein turnover/quality control systems (proteasomes, chaperones) are more closely related to their eukaryal than bacterial counterparts. In fact, archaeal homologs of eukaryal 26S proteasomes have greatly assisted in understanding basic mechanisms of energy-dependent proteolysis (Baumeister et al., 1998; Maupin-Furlow et al., 2001). The elaborate ubiquitin/proteasome system of eukaryotes (Hartmann-Petersen et al., 2003) and tmRNA-ssrA system of bacteria (Withey and Friedman, 2003), however, are not conserved in archaea, and the mechanisms of targeting proteins for destruction remain enigmatic.

Genome sequencing has provided insights into the central role proteasomes are likely to play in archaeal physiology. The proteolytic 20S core is universally distributed in Archaea and postulated to be the major cytosolic protease responsible for protein quality control and regulated proteolysis in this domain (Maupin-Furlow et al., 2004).

Global analysis of Halobacterium sp. NRC-1 (Baliga et al., 2002) reveals little correlation between the levels of mRNA and protein, suggesting post-transcriptional control mechanisms are of central importance. Unfortunately, linking the archaeal

39 40

proteasome to post-transcriptional control and other in vivo functions has been hampered

by the lack of substrate proteins that are readily detected in whole cells.

Green fluorescent protein (GFP) and its variants are often used as reporters for investigating proteolysis. The advantage of using these proteins as substrates is multifaceted and includes ease of detection as well as little, if any, perturbation to overall structure upon addition of N- or C-terminal amino acid residues (Gottesman et al., 1998).

Recently, GFP was used to assay the activity of a reconstituted proteasome system consisting of the Methanocaldococcus jannaschii proteasome-activating nucleotidase

(PAN) and Thermoplasma acidophilum 20S proteasome (Benaroudj and Goldberg, 2000;

Navon and Goldberg, 2001; Benaroudj et al., 2003). Energy-dependent proteolysis of

GFP was dependent on the presence of C-terminal residues identical to the Escherichia coli ssrA tag (AANDENYALAA) (Withey and Friedman, 2003). Although ssrA homologs are not predicted based on archaeal genomics, the in vitro study raised the possibility that the amino acid sequence of SsrA has structural similarity to motifs of proteins with short half-lives in archaeal cells. The absence of a suitable GFP expression system, however, precluded the examination of this in vivo.

Here we describe the generation and characterization of a GFP-based reporter system that enabled the quantification of proteasome-dependent proteolysis in archaeal cells. The haloarchaeon Haloferax volcanii was chosen as the host based on its ease of culture, established genetic exchange system, and synthesis of two 20S proteasome isoforms (Wilson et al., 1999; Kaczowka and Maupin-Furlow, 2003) and PAN proteins

(Reuter et al., 2004a). Although the concentration of inorganic ions (2.5 to 6 M) in the cytosol of haloarchaea can limit the synthesis of non-halophilic proteins in an active form

(Nomura and Harada, 1998; Mevarech et al., 2000), purified GFP is resistant to many denaturants (e.g., 6M urea) (Weber-Ban et al., 1999) suggesting it, or derivatives thereof, may be suitable for expression. A soluble modified derivative of GFP (F99S, M153T,

V163A) (Crameri et al., 1996) with a red-shifted mutation (S65T) (smRS-GFP) (Davis and Vierstra, 1998) was found to be ideal for use as a fluorescent reporter protein to examine the proteolytic activity of 20S proteasomes in H. volcanii.

Materials and Methods

Materials

Organic and inorganic analytical grade chemicals were from Fisher Scientific

(Atlanta, Ga.) and Sigma Chemical Co. (St. Louis, Mo.) unless otherwise indicated.

Clasto-lactacystin β-lactone was from Boston Biochem (Cambridge, Mass.). Restriction endonucleases and DNA-modifying enzymes were from New England BioLabs (Beverly,

Mass.). Desalted oligonucleotides were from Operon Technologies, Inc. (Alameda,

Calif.) and Integrated DNA Technologies (Coralville, Iowa).

Strains, Media and Plasmids

Strains, oligonucleotide primers, template DNA and plasmids are summarized in

Table 2-1. H. volcanii DS70 and E. coli BL21(DE3) were used as hosts for synthesis of recombinant fluorescent protein derivatives. E. coli strains were grown in Luria-Bertani medium (LB medium) and H. volcanii strains were grown in ATCC 974 medium (37˚C,

200 rpm). Media was supplemented with 100 mg ampicillin, 50 mg kanamycin, and/or

0.1 mg novobiocin per liter as needed. The fidelity of all polymerase chain reaction-

(PCR-) amplified products was confirmed by DNA sequencing using the dideoxy termination method (Sanger et al., 1977) with Perkin Elmer/Applied Biosystems and

LICOR automated DNA Sequencers (DNA Sequencing Facilities, Interdisiplinary Center

42 for Biotechnology Research and Department of Microbiology and Cell Science and,

University of Fla.).

PCR was used to introduce restriction enzyme sites for directional cloning of EGFP into plasmid pET28b and synthesis of His6-EGFP in E. coli. A 0.97-kb DNA fragment of this plasmid (pJAM801) was subcloned into shuttle-expression vector pBAP5010 for synthesis of His6-EGFP in H. volcanii. Plasmids expressing soluble modified blue, green and red-shifted fluorescent proteins in H. volcanii were generated by blunt-end ligation of

DNA fragments from plasmids pSMBFP, pSMGFP and pSMRSGFP into the pBAP5010- derived plasmid pJAM202. To introduce C-terminal modifications to smRS-GFP, an internal 356-bp DNA fragment from the smRS-GFP gene was ligated into pGFP-11 using

BalI and MunI. The resulting pET-based plasmid (pJAM1022) encoded smRS-GFP with a C-terminal ssrA tag (AANDENYALAA). To create smRS-GFP-ssrA variants in pET22b, PCR was used to modify the ssrA sequence and the PCR products were ligated with pJAM1022 fragments as indicated in Table 2-1. Genes encoding these smRS-GFP- ssrA variants were cloned into plasmid pJAM202 as indicated in Table 2-1 for expression in H. volcanii. The Halobacterium cutirubrum rRNA P2 promoter and T7 terminator were used for transcription of the cloned-gene in H. volcanii. For all expression plasmids, the presence and orientation of the gene with respect to the promoter and terminator were determined by restriction endonuclease cleavage, PCR and/or DNA sequence.

DNA Purification and Transformation

Plasmids were isolated by Qiagen Miniprep (Qiagen Inc.,Valencia, Calif.). DNA fragments were eluted from 0.8% (wt/vol) SeaKem GTG agarose (FMC Bioproducts,

Rockland, Maine) gels with 1 × TAE buffer (40 mM Tris-acetate, 2 mM EDTA, pH 8.5)

43 by QIAquick gel extraction (Qiagen). H. volcanii DS70 cells were transformed with plasmid DNA isolated from E. coli GM2163 according to Cline et al. (Cline et al.,

1989a).

Protein Techniques and Immunoblot

Protein concentration was determined using Bradford reagent with bovine serum albumin as the standard (BioRad, Hercules, Calif.). For immunoblot, proteins were separated by reducing and denaturing 12% polyacrylamide gel electrophoresis with β- mercaptoethanol and sodium dodecyl sulfate (SDS-PAGE) (Laemmli, 1970). Molecular mass standards included phosphorylase b (97.4 kDa), serum albumin (66.2 kDa), ovalbumin (45 kDa), carbonic anhydrase (31 kDa), trypsin inhibitor (21.5 kDa), and lysozyme (14.4 kDa) (BioRad). After separation, proteins were transferred to Hybond-P membranes (Amersham Biosciences, Piscataway, N.J.) using 10 mM 2-N- morpholino)ethanesulfonic acid (MES) at pH 6.0 with 10% (vol/vol) methanol (20 V,

4ΕC, 16 h). Antigens were detected using Living Colors Aequorea victoria peptide rabbit Ig anti-GFP antibody as the primary antibody, which is suitable for use with all

GFP variants (Clontech Laboratories, Inc., Palo Alto, Calif.). Horseradish peroxidase-

(HRP-) conjugated anti-rabbit Ig (H+L) antibody raised in goat was used as the secondary antibody (Southern Biotechnology Associates, Inc., Birmingham, Ala.). HRP activity was detected using ECL Plus chemiluminescence (Amersham Biosciences) with a VersaDoc 1000 imaging system (BioRad). For preparation of samples with and without clasto-lactacystin β-lactone, 5 ml cultures of H. volcanii (pJAM1023) were grown to an

O.D. 600 nm of 0.23 units. DMSO (0.01%) with or without the proteasome inhibitor (30

µM) was added, and cells were grown an additional 9.5 h to an O.D. 600 nm of 0.37 and

0.80 units, respectively. Thus, cells were exposed to the inhibitor during exponential

growth when synthesis of new 20S proteasome proteins occurs (Reuter et al., 2004a).

Cells were harvested and resuspended in SDS-PAGE loading dye (50 µl). Boiled sample

(20 µl) was used for immunoblot analysis. For all other immunoblots, samples were

prepared from H. volcanii cultures (60 ml) grown in triplicate to an O.D. 600 nm of 0.8 to

1.0 units. Cells were harvested (10,000 × g, 20 min, 4ºC), resuspended in 0.5 ml dH2O, lysed by sonication, and centrifuged (14,000 × g, 20 min, 4ºC). Samples (10 to 50 µg) were analyzed within the linear range of detection as determined using the His-EGFP

(10.7, 21.4, 42.8, 64.2 and 85.6 ng) purified from recombinant E. coli (pJAM801) by

Ni2+-Sepharose chromatography as previously described (Kaczowka and Maupin-Furlow,

2003). Cell lysate (10 µg) of H. volcanii (pJAM1020) expressing smRS-GFP was also included on all blots for comparison to the smRS-GFP variants.

Fluorescence Measurements of GFP Variants in Whole Cells

Colonies of H. volcanii were viewed and photographed on culture plates using a

Leica fluorescence stereo-microscope operated with and without GFP cube band pass

filters (excitation 480/40 nm and emission 510 nm) (Microoptics of Florida, Davie, Fla.)

and Nikon Coolpix 4500 camera. For whole cell fluorescence, cells were viewed at 100

× using DIC (differential interference contrast) microscopy with a Nikon OptiPhot-2

microscope with Nikon B-1E filter combination, and photographed with a Kodak DC290

zoom camera. Fluorescence of liquid cultures was monitored using an Aminco-Bowman

series 2 luminescence spectrometer (Spectronic Instruments, Rochester, N.Y.).

Excitation and emission wavelengths were as follows: 488 nm and 509 nm for EGFP and

smRS-GFP; 395 nm and 509 nm for smGFP; and 380 nm and 440 nm for smBFP. One

unit is defined as the fluorescence intensity of fluorescein at 0.025 nM.

Purification of EGFP and 20S Proteasomes from H. volcanii

One liter cultures of H. volcanii DS70 (pJAM1011) and (pJAM202) were grown in

a 2.8 L Fernbach flasks (42°C, 200 rpm) to late stationary phase (OD 600 nm of 3.5).

Cells (9 g wet wgt) were harvested (10,000 × g, 20 min, 4ºC) and resuspended in 20 ml

of 20 mM Tris buffer at pH 7.2 supplemented with 2 M NaCl (Buffer A). Cells were

passaged through a chilled French Pressure cell at 20,000 lb/in2. Cell debris was removed by centrifugation (16,000 × g, 20 min, 4ºC). Filtrate (45 µm filter) (Nalge Nunc

International, Rochester, N.Y.) was applied to a 5 ml Ni2+-Sepharose column (HiTrap

chelating) (Amersham Biosciences) equilibrated with Buffer A and washed with a step

gradient of 5 mM and 60 mM imidazole in Buffer A. Fractions containing His-tagged

proteins were eluted in Buffer A with 500 mM imidazole. Proteasomes were further

purified by application to a 5 ml Bio-scale hydroxyapatite type I column (BioRad)

equilibrated in 10 mM sodium phosphate buffer at pH 7.2 with 2 M NaCl (Buffer B).

The column was washed with 15 ml of Buffer B and developed with a 30 ml linear

sodium phosphate gradient (10 to 500 mM sodium phosphate at pH 7.2 with 2 M NaCl).

Protein fractions (300 µg per ml) with peptide hydrolyzing activity which eluted at ~350

mM sodium phosphate were pooled. Sample (0.5 ml per run) was applied to a Superose-

6 HR 10/30 gel filtration column (Amersham Bioscience) equilibrated in Buffer A.

Peptide hydrolyzing fractions containing 20S proteasomes of 600 kDa, determined to be

homogeneous by Coomassie blue R-250 stained SDS-PAGE gels, were pooled and used

for kinetic and inhibitor analysis. To determine the percent inhibition of 20S

46 proteasomes by clasto-lactacystin β-lactone in cell culture, H. volcanii DS70 (pJAM202) was grown in 5 ml culture to an O.D. 600 nm of 0.28 units. DMSO (0.01%) with and without proteasome inhibitor (15 µM) was added, and cells were harvested after an additional 9.5 h of cultivation (OD 600 nm of 1.58 and 2.0, respectively). His-tagged

20S proteasomes were purified from these cultures by Ni2+-Sepharose chromatography

(as described above) and assayed for peptide hydrolyzing activity.

Peptide Hydrolyzing Activity

Chymotrypsin-like peptide hydrolyzing activity was assayed in Buffer A with 20 mM N-Suc-Leu-Leu-Val-Tyr-7-amido-4-methylcoumarin (-Amc) unless otherwise indicated. Release of 7-amino-4-methylcoumarin was monitored as an increase in fluorescence over time using excitation and emission wavelengths of 350 and 460 nm with an Aminco Fluoro-colorimeter (Spectronic Instruments). Assays were performed at the approximate temperature (60°C), pH (pH 7.2) and NaCl (2 M) optimum determined previously for H. volcanii 20S proteasomes (Wilson et al., 1999b).

RNA Isolation and Analysis

Cultures (2 ml) of DS70, DS70 (pJAM1020), DS70 (pJAM1023) and DS70

(pJAM1025) strains were grown to an O.D.600 nm of 0.8. Total RNA was isolated using the RNeasy MiniKit for bacteria (Qiagen) with the following modification. Cells were centrifuged for 1 min at 18,000 × g and immediately resuspended in 100 µl RNase-free deionized water. Total RNA was treated with DNaseI (Sigma) and NucAway spin columns. RNA quality was confirmed by ethidium bromide stained 0.8% (w/v) agarose gels. RNA concentration was determined by absorbance at 260 nm. Total RNA (1.5µg) was used as a template for One-Step reverse transcriptase-PCR (RT-PCR) (Qiagen) with

oligonucleotides specific for the coding region of smGFP (5’-

GGAGAAGAACTTTTCACTGGAGT-’3 and 5’-

ATGTTCATCCATGCCATGTGTAATC-’3). cDNA products of 0.7-kb were detected

by agarose gel electrophoresis (as above). Control reactions, incubated on ice (vs. 50°C)

for the RT step, were included to ensure that cDNA products were not due to

contamination of total RNA with recombinant plasmid DNA.

Results and Discussion

Functional Synthesis of an Archaeal GFP Reporter Protein

A variety of GFP variants were explored with the goal of generating of a

fluorescent reporter protein for functional expression in H. volcanii. Initially, EGFP was

examined based on amino acid modifications (Phe64Leu and Ser65Thr) which allow for

enhanced fluorescence compared to GFP. Quantitative immunoblot of a recombinant

strain which expressed the EGFP gene (H. volcanii DS70/pJAM1011) revealed levels of

fluorescent protein that were ~ 0.1% of total cell protein (Fig. 2-1); however, whole cell

fluorescent was not enhanced. An N-terminal polyhistidine tag allowed for rapid

purification of EGFP in high salt (2 M NaCl) buffer from recombinant H. volcanii. The

partially pure EGFP exhibited significant fluorescence (data not shown) suggesting that it

folded in the high ionic strength cytosol of H. volcanii. Thus, the low levels of EGFP

may have limited its detection by fluorescence in whole cells.

To enhance the levels of fluorescent protein, genes encoding soluble modified derivatives of GFP (smGFP, smRS-GFP and smBFP) were expressed in H. volcanii. The soluble modifications (Phe99Ser, M153Thr and Val163Ala) were derived from the UV- optimized Cycle 3 variant which has reduced surface hydrophobicity, increased thermostability, and a lower tendency to aggregate during folding compared to GFP

(Fukuda et al., 2000). The soluble modifications were expected to increase the stability

and/or folding of GFP in the high ionic strength cytosol (2 – 3 M) of H. volcanii. The

genes encoding the fluorescent protein variants also included silent mutations which

removed mRNA processing sites and optimized codons for expression in plants (Davis

and Vierstra, 1998). The red-shift in excitation and emission wavelengths (RS-GFP) was selected to minimize auto-fluorescence of the host cell.

Based on quantitative immunoblot, all three of the soluble modified derivatives of

GFP (smGFP, smBFP and smRS-GFP) were expressed at relatively high levels (> 1% total cell protein) in recombinant H. volcanii (Fig. 2-1). Immunoblot of the 10,000 × g fraction of cell lysate revealed no significant accumulation of the soluble modified GFP derivatives as inclusion bodies (data not shown). Furthermore, recombinant cells which produced the smRS-GFP protein (DS70/pJAM1020) exhibited significant fluorescence over wild type when grown in liquid and plate culture (Figs. 2-3 – 2-5). Fluorescence attributed to smRS-GFP was uniformly distributed throughout the colonies on plates (2-.

3) and in whole cells (Fig. 2-4). The smRS-GFP fluorescence paralleled growth in liquid culture with an exponential increase during log-phase and plateau during stationary-phase

(Fig. 2-5). The persistence of smRS-GFP fluorescence in stationary-phase suggested that this reporter protein was relatively stable in whole cells.

C-terminal Modifications Reduce the Level of smRS-GFP Protein in vivo

C- and N-terminal tags are often fused to GFP proteins to define structural elements or motifs that target proteins for degradation (Chiesa et al., 2001). The apparent stability of smRS-GFP in H. volcanii suggested that a similar approach could be used to determine what residues and/or motifs destabilize proteins in vivo. Thus, a variety of amino acids

49 were fused to the C-terminus of smRS-GFP (Table 2-2). Included in the modifications was the addition of the 11-residue sequence AANDENYALAA or ‘ssrA tag’. Although archaea do not encode SsrA homologs, the 11-residue amino acid sequence had previously been shown to target GFP for degradation by reconstituted archaeal 20S proteasome and PAN complexes purified from recombinant E. coli (Benaroudj and

Goldberg, 2000). Modifications (Table 2-2) also included C-terminal deletions of the ssrA tag and alteration of the C-terminal Ala-10 and Ala-11 residues of ssrA to acidic

(Glu and Asp), basic (Lys), and hydrophobic (Leu) residues. In addition, a random sequence was included which had four contiguous charged residues (Asp-Asp-Lys-Asp or DDKD) in place of the two charged residues of the ssrA tag (Asp-Glu or DE). All of these C-terminal tags were predicted to be on the surface of smRS-GFP and have little if any impact on the overall structure or fluorescence of the intact protein, based on circular dichroism of GFP vs. GFP-ssrA (Gottesman et al., 1998) and the GFP crystal structure

(Ormo et al., 1996).

The majority of amino acid residues which were fused to the C-terminus of smRS-

GFP, including the ssrA-tag, completely abolished whole cell fluorescence (Table 2-2).

In fact, only a few of the tags examined resulted in fluorescent colonies and, of those, the levels of fluorescence were low to moderate compared to the unmodified smRS-GFP. To determine whether the observed reduction in fluorescence reflected changes in the levels of the reporter protein, quantitative immunoblot was performed. Based on this analysis, the levels of most of the C-terminal fusions, including the ssrA-tagged protein, were reduced at least 200-fold compared to the “untagged” smRS-GFP (Table 2-2) (Fig. 2-2).

Exceptions included the addition of AANDDKDLSNN which was of comparable length

to the ssrA tag but had four contiguous charged residues. This modification resulted in

an over 50-fold increase in protein levels compared to smRS-GFP-ssrA. Likewise,

deletion of Ala-2 to Ala-11 of the ssrA tag enhanced the levels of this fusion protein over

16-fold. Replacement of the C-terminal Ala residues (Ala-10 and Ala-11) of ssrA with

acidic residues (Glu or Asp) also enhanced protein levels over 10-fold.

Some general trends in the type of C-terminal modification and level of reporter

protein were apparent. Most prominent was that in all cases addition of a hydrophobic

residue (i.e. Ala or Leu) to the C-terminus dramatically reduced the level of reporter

protein, even when the length of the fusion varied. In fact, addition of a single Ala to

smRS-GFP resulted in an over 10-fold decrease in protein. The type of residue added to

the C-terminus, however, was not the sole determinant of reporter protein levels as

exemplified by fusion proteins with Asn as the terminal residue. These ‘Asn’ fusion

proteins varied from < 0.5% to 30% of protein relative to smRS-GFP. Furthermore, the

length of the tag did not directly correlate with reporter protein levels (e.g., addition of

the 11-residue AANDDKDLSNN tag had less of an impact on smRS-GFP levels than

addition of a single Ala). Interestingly, although the ssrA system that targets proteins for

degradation by the Clp protease in Bacteria is not conserved in Archaea, there are some

similarities in the C-terminal amino acid sequences that reduce/enhance the GFP reporter

protein levels between these two domains. For example, in E. coli, modification of the

Ala-10 and Ala-11 residues of ssrA to acidic residues increases the Km of ClpXP by at least a factor of 100 (i.e., no degradation is observed at saturating protein substrate)

(Flynn et al., 2001). Likewise, in H. volcanii, alteration of the terminal Ala residues to acidic residues enhanced protein levels 10-fold. However, in contrast to E. coli, the

51 levels of these acidic ssrA derivatives were significantly reduced compared to the unmodified smRS-GFP reporter protein in H. volcanii.

C-terminal Modifications Do Not Impact the Level of smRS-GFP-specific mRNA in H. volcanii

Based on the results presented above, a variety of C-terminal fusions were found to reduce the level of smRS-GFP protein in H. volcanii. All of the expression plasmids used for this analysis were designed to carry identical DNA sequence and spacing for the ribosomal binding site, transcriptional promoter and terminator, and the majority of the open reading frame encoding smRS-GFP. Thus, the C-terminal modifications to smRS-

GFP were expected to have little impact on mRNA levels. To confirm this, the levels of smRS-GFP-specific mRNA were analyzed by RT-PCR in cells expressing smRS-GFP, smRS-GFP-ssrA, and the smRS-GFP-ssrA derivative with Ala10Asp and Ala11Asp mutations (i.e. smRS-GFP-ssrDD). Comparable levels of smRS-GFP-specific mRNA were detected in all three recombinant strains (vs. parent strain DS70) (data not shown).

Thus, mRNA levels are not limiting the synthesis of the smRS-GFP variants in these recombinant H. volcanii strains.

Clasto-lactacystin β-lactone as an in vivo Inhibitor of H. volcanii 20S Proteasomes

Clasto-lactacystin β-lactone has been shown to permeate eukaryotic cell membranes and inhibit 20S proteasomes by covalent modification of the active site threonine hydroxyl (Fenteany et al., 1995; Fenteany and Schreiber, 1998). It is considered by many to be the ‘gold standard’ for proteasome inhibitor specificity and is used routinely to investigate proteasome function in eukaryotic cells (Bogyo and Wang,

2002). To determine whether H. volcanii 20S proteasomes are inhibited by this compound, two approaches were taken. First, the peptide hydrolyzing activity of purified

20S proteasomes was assayed in the presence of clasto-lactacystin β-lactone vs. the

general serine protease inhibitor phenylmethanesulfonyl fluoride (PMSF) (Fig. 2-6A).

Compared to PMSF which had little if any influence on the peptide hydrolyzing activity

of the halophilic 20S proteasomes, the lactacystin derivative was highly specific with a Ki

of ~ 40 nM (Fig. 2-6B). Clasto-lactacystin β-lactone increased the Km for N-Suc-Leu-

Leu-Val-Tyr-Amc from 9.85 µM to an apparent Km of 22.2 µM without altering the Vmax

(550 nmoles 7-amino-4-methylcoumarin released per min per mg protein). Based on these results, it appears that the lactacystin derivative competes effectively for the active site of the halophilic 20S proteasomes. Thus, the influence of clasto-lactacystin β- lactone on proteasome activity was examined in cell culture. For this analysis, the H. volcanii strain DS70 (pJAM202) which expresses the β subunit of 20S proteasomes with a His-tag was grown in the presence and absence of clasto-lactacystin β-lactone. The specific activity of the 20S proteasomes was significantly reduced when purified from cells grown in the absence vs. presence of this inhibitor (from 204 to 175 nmol 7-amino-

4-methylcoumarin released per min per mg protein). The reason for the incomplete

inhibition of 20S proteasomes in vivo remains to be determined. Based on our previous

work (Kaczowka and Maupin-Furlow, 2003), expression of the β-His protein from

plasmid pJAM202 enhances the levels of β-protein 2-fold over wild type. Thus, the

levels of 20S proteasomes may be higher in DS70 (pJAM202). However, it is more

likely that clasto-lactacystin β-lactone is not fully soluble at the concentration and

composition of salts required for growth of H. volcanii. This latter possibility is

supported by the 30 to 40% decrease in inhibition of the halophilic 20S proteasome

activity in vitro when clasto-lactacystin β-lactone is preincubated (30 min, 37ºC) in

ATCC medium salts (~ 2.6 M NaCl, 530 mM MgCl2, 30 mM KSO4, and 1.2 mM CaCl2) prior to assay (data not shown). It is also possible that clasto-lactacystin β-lactone does not permeate H. volcanii cells as readily as eukaryotic cells due to differences in cell wall structure. The cell wall of H. volcanii includes a glycoprotein S-layer and cell membrane of isoprenoid ether lipids not found in eukaryotes.

Inhibition of 20S Proteasomes Enhances the Level of smRS-GFP-SsrA Protein in H. volcanii

Of the C-terminal smRS-GFP derivatives, the ssrA-tagged fluorescent protein was chosen as a reporter to investigate whether 20S proteasomes are involved in modulating the levels of this protein in the cell. The rationale for this choice was that an ssrA-tagged derivative of GFP is degraded by 20S proteasomes reconstituted with PAN in vitro

(Benaroudj et al., 2003). Thus, H. volcanii cells expressing smRS-GFP-ssrA were grown in the presence and absence of clasto-lactacystin β-lactone and analyzed for accumulation of the reporter protein. Interestingly, addition of the proteasome inhibitor significantly reduced the growth rate (from a doubling time of ~ 4 h to 6 h) suggesting that 20S proteasomes are needed for normal growth of H. volcanii (Fig. 2-7A).

Immunoanalysis revealed that the reporter protein (smRS-GFP-ssrA) accumulated when cells were grown in the presence of clasto-lactacystin β-lactone (Fig. 2-7B). In contrast, the general serine protease inhibitor PMSF (30 µM) had no influence on cell growth or reporter protein levels (data not shown). Although the ssrA-tagged smRS-GFP protein did not accumulate to levels comparable to smRS-GFP in the presence of clasto- lactacystin β-lactone, H. volcanii 20S proteasomes are not completely inactivated by this inhibitor in cell culture (see previous section).

Conclusion

This study reveals that a soluble modified, red-shifted derivative of GFP (smRS-

GFP) is synthesized and readily detected in recombinant H. volcanii. Fluorescence of smRS-GFP was uniformly distributed in whole cells and paralleled reporter protein levels as determined by immunoblot. Addition of amino acid residues of various sequence and length to the C-terminus of smRS-GFP had a differential effect on the levels of this reporter protein. Proteasomes were found to be responsible, at least in part, for modulating the levels of the ssrA-tagged GFP derivative in vivo.

Recombinant smRS-GFP and C-terminal variant reporter proteins have tremendous

potential in a variety of applications in the haloarchaea. For example, chromosomal mutations that stabilize C-terminal tagged smRS-GFP derivatives, regulated promoters, and chemical libraries of 20S proteasome inhibitors can be rapidly screened in H. volcanii using fluorescent plate assays. The localization of H. volcanii proteins can be

determined by fusion to smRS-GFP and imaging via whole cell fluorescence. Although

the current host strain necessitates the use of immunoblot for the quantitative analysis of

smRS-GFP protein levels, the analysis is bolstered by the rapid and qualitative analysis of

GFP protein levels using whole cell fluorescence. Efforts aimed at reducing the auto-

fluorescence of H. volcanii will allow for greater versatility of this reporter protein in the

future (e.g. sensitive quantification of smRS-GFP protein levels via fluorescence, dual

fluorescent label studies).

Table 2-1. Strains and plasmids used for this study Strain or Phenotype, genotype, description, PCR primers Source plasmid E. coli - - - BL21(DE3) F ompT [lon]hsdSB(rB mB ) (an E. coli B strain) Novagen with DE3, a λ prophage carrying the T7 RNA polymerase gene - - + DH5α F recA1 endA1 hsdR17(rk mk ) supE44 thi-1 gyrA New England relA1 Biolabs GM2163 F- ara-14 leuB6 fhuA31 lacY1 tsx78 glnV44 galK2 New England galT22 mcrA dcm-6 hisG4 rfbD1 rpsL 136 Biolabs dam13::Tn9 xylA5 mtl-1 thi-1 mcrB1 hsdR2 H. volcanii DS70 cured of plasmid pHV2 (Wendoloski et al., 2001) Plasmids pBAP5010 Apr; Nvr; H. volcanii-E. coli shuttle-expression (Jolley et al., vector 1997a;Kaczowka and Maupin- Furlow, 2003) pEGFP Apr; GenBank accession U76561 Clontech pSMGFP Apr; GenBank accession U70495 (Davis and Vierstra, 1998) pSMRSGFP Apr; GenBank accession U70496 (Davis and Vierstra, 1998) pSMBFP Apr; GenBank accession U70497 (Davis and Vierstra, 1998) pGFP-11 Apr; pET22b-derived plasmid expressing GFPuv- A. Horwich SsrA (Weber-Ban et al., 1999) pET22b Apr; E. coli expression vector Novagen pET28b Kmr; E. coli expression vector Novagen pJAM202 Apr; Nvr; H. volcanii-E. coli shuttle expression (Kaczowka and plasmid with psmB-his6 gene; β subunit of 20S CP Maupin-Furlow, expressed with C-terminal His tag in H. volcanii 2003) pJAM801 Kmr; 0.72-kb fragment generated by PCR from This study pEGFP ligated into pET28b using NdeI and blunt- ended BlpI (5’-TTTGATTCATATGGTGAGCA AGGGCG-3’ and 5’TTACTTGTACAGCTCGT CCATGCCGAG -3’; NdeI site in bold) ; EGFP with N-terminal His tag expressed in E. coli pJAM1011 Apr; Nvr; 0.97-kp XbaI-to-BspEI fragment of This study pJAM801 blunt-end ligated with a 9.52-kb KpnI- to-BamHI fragment of pBAP5010; EGFP with N- terminal His tag expressed in H. volcanii

Table 2-1. Continued Strain or Phenotype, genotype, description, PCR primers Source plasmid pJAM657 Apr; Nvr; 0.74-kb BamHI-to-SacI fragment of pSMBFP This study was blunt-end ligated with a 9.94-kb NdeI-to-BlpI fragment of pJAM202; smBFP expressed in H. volcanii pJAM1019 Apr; Nvr; as in pJAM657 except pSMGFP replaced This study pSMBFP; smGFP expressed in H. volcanii pJAM1020 Apr; Nvr; as in pJAM657 except pSMRSGFP replaced This study pSMBFP; smRS-GFP expressed in H. volcanii pJAM1022 Apr; 356-bp BalI-to-MunI fragment of pJAM1020 was This study ligated with a 4-kb BalI-to-MunI fragment of pGFP-11; smRS-GFP-ssrA expressed in E. coli pJAM1023 Apr; Nvr; 910-bp XbaI-to-BlpI fragment of pJAM1022 This study was blunt-end ligated with a 9.94-kb NdeI-to-BlpI fragment of pJAM202; smRS-GFP-ssrA expressed in H. volcanii pJAM1024 Apr; 191-bp fragment generated by PCR from This study pJAM1022 ligated into pJAM1022 using MfeI and BamHI (primer 1: 5’-TATCAACA AAATACTCCAATTGGCGATG-3’ and primer 2: 5’- CCGGATCC TTAGTCGTCTAAAGCGTAGT-3’)(MfeI and BamHI sites in bold); smRS-GFP-ssrDD expressed in E. coli pJAM1025 Apr; Nvr; as in pJAM1023 except pJAM1024 replaced This study pJAM1022; smRS-GFP-ssrDD expressed in H. volcanii pJAM1026 Apr; as in pJAM1024 except 5’- This study GGATCCTTACTCCTCTAAAGCG TAGTT-3’ replaced primer 2 (BamHI site in bold); smRS-GFP-ssrEE expressed in E. coli pJAM1027 Apr; as in pJAM1024 except 5’- This study GGATCCTTAGAGGAGTAAAGCG TAGTT-3’ replaced primer 2 (BamHI site in bold); smRS-GFP-ssrLL expressed in E. coli pJAM1028 Apr; as in pJAM1024 except 5’- This study CGGATCCTTACTTCTTTAAAGC GTAGT-3’ replaced primer 2 (BamHI site in bold); smRS-GFP-ssrKK expressed in E. coli pJAM1029 Apr; as in pJAM1024 except 5’- This study CGGATCCTTAGTCTCATAAAGC GTAGT-3’ replaced primer 2 (BamHI site in bold); smRS-GFP-ssr9 expressed in E. coli pJAM1030 Apr; Nvr; 858-bp XbaI-to-BamHI fragment of pJAM1026 This study was blunt-end ligated with a 9.94-kb NdeI-to-BlpI fragment of pJAM202; smRS-GFP-ssrEE expressed in H. volcanii

Table 2-1. Continued Strain or Phenotype, genotype, description, PCR primers Source plasmid pJAM1031 Apr; Nvr; as in pJAM1030 except pJAM1027 replaced This study pJAM1026; smRS-GFP-ssrLL expressed in H. volcanii pJAM1032 Apr; Nvr; as in pJAM1030 except pJAM1028 replaced This study pJAM1026; smRS-GFP-ssrKK expressed in H. volcanii pJAM1033 Apr; Nvr; as in pJAM1030 except pJAM1029 replaced This study pJAM1026; smRS-GFP-ssr9 expressed in H. volcanii pJAM1035 Apr; 0.85-kb fragment generated by PCR from This study pJAM1022 ligated into pJAM1022 using XbaI and BamHI (primer 3: 5’-GGAGACCACAA CGGTTTCCCTCTAGAAA -3’ and primer 4: 5’- AGCGTAGGAT CCGTCGTTTTAGGCTTT-3’)(XbaI and BamHI sites in bold); smRS-GFP-ssr1 expressed in E. coli pJAM1037 Apr; as in pJAM1035 except 5’- This study AGCGTAGGATCCTTAGTTGGCG GCTTT-3’ replaced primer 4 (BamHI site in bold); smRS-GFP-ssr3 expressed in E. coli pJAM1038 Apr; as in pJAM1035 except 5’- This study TAAGGATCCTTATTCGTCGTTG GCGGC-3’ replaced primer 4 (BamHI site in bold); smRS-GFP-ssr? expressed in E. coli pJAM1039 Apr; 0.85-kb fragment generated by PCR from This study pJAM1022 ligated into pJAM1022 using XbaI and BamHI (primer 5: 5’-CGCGGAGACCAC AACGGTTCCCTCTAGAAA-3’ and primer 6: 5’- AGCGGATCCT TAGTTTTCGTCGTTGGCGGC-3’)(XbaI and BamHI sites in bold); smRS-GFP-ssr6 expressed in E. coli pJAM1040 Apr; as in pJAM1039 except 5’- This study TGCGGATCCTTAGTAGTTTTCGT CGTTGGCGGC-3’ replaced primer 6 (BamHI site in bold); smRS-GFP-ssr7 expressed in E. coli pJAM1041 Apr; as in pJAM1039 except 5’- This study TTAGGATCCTTACGCGTAGTTTT CGTCGTTGGCGGC-3’ replaced primer 6 (BamHI site in bold); smRS-GFP-ssr8 expressed in E. coli pJAM1044 Apr; Nvr; 858-bp XbaI-to-BamHI fragment of pJAM1035 This study was blunt-end ligated with a 9.94-kb NdeI-to-BlpI fragment of pJAM202; smRS-GFP-ssr1 expressed in H. volcanii

Table 2-1. Continued Strain or Phenotype, genotype, description, PCR primers Source plasmid pJAM1046 Apr; Nvr; as in pJAM1044 except pJAM1037 replaced This study pJAM1035; smRS-GFP-ssr3 expressed in H. volcanii pJAM1048 Apr; Nvr; as in pJAM1044 except pJAM1038 replaced This study pJAM1035; smRS-GFP-ssr? expressed in H. volcanii pJAM1049 Apr; Nvr; as in pJAM1044 except pJAM1039 replaced This study pJAM1035; smRS-GFP-ssr6 expressed in H. volcanii pJAM1050 Apr; Nvr; as in pJAM1044 except pJAM1040 replaced This study pJAM1035; smRS-GFP-ssr7 expressed in H. volcanii pJAM1051 Apr; Nvr; as in pJAM1044 except pJAM1041 replaced This study pJAM1035; smRS-GFP-ssr8 expressed in H. volcanii

Table 2-2. Modification of the C-terminus of smRS-GFP influences whole cell fluorescence and the level of fluorescent protein in recombinant H. volcanii. Expression SmRSGFP C- Amino acid tag (abbrev)b Relative Whole cell plasmid terminusa protein fluorescenced (%)c pJAM1025 GITHGMDELYK -AANDENYALAA (ssrAA) UD  pJAM1025 GITHGMDELYK -AANDENYALEE (ssrEE) 6.6± 1.3 + pJAM1025 GITHGMDELYK -AANDENYALDD (ssrDD) 5.5 ± 0.6 + pJAM1031 GITHGMDELYK -AANDENYALLL (ssrLL) <0.5  pJAM1032 GITHGMDELYK -AANDENYALKK (ssrKK) <0.5  pJAM1048 GITHGMDELYK -AANDDKDLSNN (ssr?) 28.9 ± 5.5 ++ pJAM1033 GITHGMDELYK -AANDENYAL (ssr9) <0.5  pJAM1051 GITHGMDELYK -AANDENYA (ssr8) <0.5  pJAM1050 GITHGMDELYK -AANDENY (ssr7) <0.5  pJAM1049 GITHGMDELYK -AANDEN (ssr6) <0.5  pJAM1046 GITHGMDELYK -AAN (ssr3) <0.5  pJAM1044 GITHGMDELYK -A (ssr1) 8.1 ± 1.9 + pJAM1020 GITHGMDELYK 100 ± 4.4 ++++ aC-terminal 11 residues of smRS-GFP are highlighted. bAmino acid residues added to C-terminus of smRS-GFP are separated by a hyphen. Abbreviations used to denote the type of modification are in parentheses. cProtein level based on quantitative immunoblot and expressed as percentage relative to smRS-GFP. Compared to proteins at levels less than 0.5 % of smRS-GFP, the ssrA tagged fusion protein was undetectable (u.d.). dRelative level of fluorescence as observed by fluorescence microscopy of colonies, where (++++) is the highest and () is undetectable.

1 2 3 4 5 6 7 8 9 10

His6-EGFP standard none His-EGFP smBFP smGFP smRS-GFP

Fig. 2-1. Optimization of fluorescent protein expression in recombinant H. volcanii. The levels of fluorescent protein were estimated by quantitative immunoblot as described in methods. His6-EGFP (18, 36, 54, 90 and 180 ng) was purified from recombinant E. coli and used as a standard (lanes 1-5). This was compared to cell lysate (15 – 16 µg) of H. volcanii DS70 (lane 6), DS70/pJAM1011 (lane 7), DS70/pJAM657 (lane 8), DS70/pJAM1019 (lane 9) and DS70/pJAM1020 (lane 10). The recombinant protein expressed in each strain is indicated below the immunoblot.

Fig. 2-2. Protein level quantification of smRS-GFP reporter proteins expressed in recombinant H. volcanii by immunoblot. The levels of fluorescent proteins were estimated by quantitative immunoblot as described in methods. His6- EGFP (10.7, 21.4, 42.8, 64.2, and 85.6 ng) was purified from recombinant E. coli and used as a standard (lanes 1-5). This was compared to cell lysate (15 – 50 µg) of H. volcanii DS70 alone, DS70 expressing EGFP, and DS70 expressing each smRS-GFP reporter protein as indicated below each band. Calculated protein percentages are listed in Table 2-2.

Fig. 2-3. Colonies of H. volcanii expressing smRS-GFP exhibit fluorescence. H. volcanii DS70 and DS70 (pJAM1020) which expresses smRS-GFP were -6 grown to an O.D.600nm of 1 unit and 0.1 ml of 10 dilution was spread onto ATCC 974 solid medium. Cells were incubated for 5 days at 37ºC, and colonies were visualized by bright field (A and C) and fluorescence (B and D) microscopy.

Fig. 2-4. H. volcanii expressing smRS-GFP displays uniform cell fluorescence. Liquid cultures of H. volcanii DS70 and DS70 (pJAM1020) which expresses smRS- GFP were grown to an O.D.600nm of 1 unit. Cultures were transferred (10µl) to glass slides with cover slips, and whole cells were visualized using DIC (differential interference contrast) microscopy with and without fluorescence.

A) 6 5

U 4

n 3

rr 2

F 1

B) 4

m 3

O 1

0 0 20 40 60 80 100 120 140 160

Time (h)

Fig. 2-5. Liquid cultures of H. volcanii expressing smRS-GFP exhibit fluorescence that parallels cell growth. Comparison of the fluorescence (A) and growth (B) of H. volcanii DS70(pJAM1020) expressing smRS-GFP (○) to the parent strain DS70 (□) in liquid culture. Fluorescence was measured at λ ex 488 nm and λ em 510 nm in triplicate cultures. Fluorescence attributed to smRS-GFP expression (●) was calculated as the difference between DS70(pJAM1020) and DS70 cell cultures.

A) 100 90 ) ) 80 (% (% 70 ty ty vi vi 60 i i t t c c 50 A A 40 ve ve 30 ti ti a a l l 20 e e R R 10 0 0 0.2 0.4 0.6 0.8 1.0 30 B) Inhibitor (µM) ) ) 550 –1 –1 500 g g 450 m m

400 y y 350 300 250 Velocit Velocit 200 150 100 (nmol per min • (nmol per min • 50 0 0 1020304050607080 N-Suc-Leu-Leu-Val-Tyr-Amc (µM)

Fig. 2-6. H. volcanii 20S proteasomes are inhibited by clasto-lactacystin β-lactone. A) Purified 20S proteasomes (1 mg per ml; ~ 1.67 nM) were pre-incubated with varying concentrations of clasto-lactacystin β -lactone (●) or PMSF (■) for 30 min at 37ºC in 20 mM Tris-HCl buffer, pH 7.2 with 2 M NaCl and 0.0075% (vol/vol) DMSO or 0.01% (vol/vol) ethanol, respectively. Peptide hydrolysis was assayed as described in methods using 20 mM substrate with an additional 0.004% DMSO. Even at 100 µM PMSF, 92% activity remained. B) Purified proteasomes (0.5 mg per ml; ~ 0.84 nM) were assayed with various concentrations of substrate as indicated, in the absence (●) and presence (■) of 50 nM clasto-lactacystin β -lactone.

A) 4.5 4

3.5

2.5 2 harvest D. 600 nm D. 600 nm . . 1.5 O O 1 inhibitor

0.5

0 0 5 10 15 20 25 30 35 40 45 50 55 60 65 70 Time (h)

B) 1 2 3

30kDa

control —+ proteasome inhibitor

Fig. 2-7. Inhibition of H. volcanii 20S proteasomes enhances the level of smRS-GFP- ssrA in cell culture. A) H. volcanii DS70(pJAM1023) was grown in the absence (●) and presence (■) of clasto-lactacystin β-lactone (30 µM). Arrowhead indicates the time (15 h) inhibitor was added to cell culture. B) Immunoblot with anti-GFP antibody of E. coli purified His6-EGFP (42.8 ng) (lane 1) and log-phase H. volcanii DS70(pJAM1023) expressing smRS-GFP- ssrA without (lane 2) and with clasto-lactacystin β-lactone (lane 3).

CHAPTER 3 DIFFERENTIAL POST-TRANSLATIONAL REGULATION OF GREEN FLUORESCENT PROTEIN IN THE HALOARCHAEON HALOFERAX VOLCANII IS DETERMINED IN PART BY ITS EXTREME CARBOXYL TERMINAL RESIDUE

Introduction

Post-translational protein regulation is critical to the viability of all living cells and is controlled to a large extent by proteolysis. Protein turnover removes damaged and extraneous proteins, resulting in free amino acids, which can be recycled or used to generate energy (Gottesman, 2003; Wolf and Hilt, 2004). Selective removal of these proteins is based on at least one of a variety of factors including exposed hydrophobic groups, post-translation protein modifications (e.g. phosphorylation), specific proteolytic recognition sequences (e.g. SsrA), extreme protein end residue identity (N-end rule), and ubiquitination (in eukaryotes) (Komine et al., 1994; Bohley, 1996; Varshavsky, 1997;

Gross et al., 2005). Energy-dependent proteolysis, carried out by such proteases as ClpP,

FtsH, Lon and 20S proteasomes (HslUV), is responsible for the degradation of the majority of these proteins (Gottesman, 2003). Substrate differentiation based on these proteolytic recognition factors is determined primarily by protease-associated ATPase regulatory particles whose additional responsibilities include substrate unfolding, and in at least some cases, protease gate opening and substrate translocation (Smith et al., 2005).

The ATPase regulatory particles ClpA and ClpX associate with ClpP, and the eukaryotic

19S cap associates with 20S proteasomes for the purpose of protein quality control

(Gottesman, 2003; Schmidt et al., 2005).

67 68

Several proteolytic targeting systems, including those based on the SsrA tagging

system and the N-end rule, have been well characterized. A unique system of eubacteria,

SsrA (small stable RNA A) protein tagging employs a transfer-messenger RNA (tmRNA) that adds an 11-residue sequence (AANDENYALAA) to the end of stalled translation products which conveys a proteolytic targeting signal for ClpXP and ClpAP (Gottesman et al., 1998; Withey and Friedman, 2003). The N-end rule of proteolytic substrate recognition, followed to varying degrees in eubacteria and eukaryotes, is based on the half-life (due to proteolysis) of a protein depending on the extreme N-terminal residue

(N-degron) of that protein (Varshavsky, 1997). In eukaryotes, a vast array of proteolytic substrates, including short-lived N-end rule proteins, are recognized by ubiquitin-protein ligases, which in conjunction with other enzymes of the ubiquitination pathway, target these substrates for degradation by the addition of a poly-ubiquitin chain (Robinson and

Ardley, 2004). Poly-ubiquitination precedes and stimulates proteolytic substrate

targeting of many proteins by 26S proteasomes (composed of the 20S proteasome and

19S cap) in eukaryotes (Vierstra, 2003; Schmidt et al., 2005). Archaea, although

predicted to lack homologs of the ubiquitination pathway, share highly conserved 20S

proteasomes which are also predicted to be the central energy-dependent protease within

the cell (Maupin-Furlow et al., 2004; Lowe et al., 1995). Likewise, a regulatory particle

to 20S proteasomes in many archaea is denoted as the proteasome-activating nucleotidase

(Pan) and is highly similar to Regulatory particle triple-A type I proteins (Rpt) of the base

subcomplex of the 19S cap (Zwickl et al., 1999; Reuter et al., 2004). Despite this high

degree of similarity between 20S proteasomes of archaea and eukaryotes, the lack of

69 archaeal ubiquitination machinery suggests an alternative route of proteolytic substrate recognition by the Pan/20S proteasome, of which little is known.

Recently we demonstrated that a soluble modified, red shifted-green fluorescent protein (smRS-GFP) variant with a C-terminal SsrA-tag fusion (smRS-GFP-SsrA) was degraded by 20S proteasomes in vivo, in the haloarchaeon Haloferax volcanii (Reuter and

Maupin-Furlow, 2004). Although the SsrA system is not predicted in archaea, it had previously been demonstrated that a recombinantly produced Pan and 20S proteasome could degrade GFP-SsrA in vitro, and therefore served as a means by which to study energy-dependent proteolysis in archaea (Navon and Goldberg, 2001). In our study, addition of a single Ala residue to the C-terminus of smRS-GFP resulted in a decrease of intracellular smRS-GFP protein levels by approximately 90%. This led us to question whether the addition of other single C-terminal amino acids would have a similar influence. Here we demonstrate that levels of smRS-GFP variant proteins depend on the addition of a single amino acid to the C-terminus. Proteolysis is suspected to play a role in the fate of these reporter proteins, and current efforts are underway to resolve this issue. In any event, it is important to understand the properties of proteins that mediate their levels within the cell, especially those proteins produced in recombinant archaea that have been derived from other organisms. Archaeal organisms are increasingly being relied upon for use as model systems of eukaryotes and are considered by many to be an attractive host in biotechnological applications due to their ability to survive extreme conditions along with other unique traits and enzymes (Margesin and Schinner, 2001;

Schiraldi and De Rosa, 2002; Kaczowka et al., 2005; Egorova and Antranikian, 2005;

Soppa, 2006). Advancement in these areas of research will excel through the

comprehension of post-translational regulation and optimization of protein expression in

archaea.

Materials and Methods

Materials

Organic and inorganic analytical grade chemicals were from Fisher Scientific

(Atlanta, Ga.) and Sigma Chemical Co. (St. Louis, Mo.) unless otherwise indicated.

Desalted oligonucleotides were from Integrated DNA Technologies (Coralville, Ind.).

Restriction endonucleases and DNA-modifying enzymes were from New England

BioLabs (Beverly, Mass.).

Strains, Media, and Plasmids

Strains, oligonucleotide primers used for cloning, and plasmids are summarized in

Table 3-1. E. coli DH5α was used for routine recombinant DNA experiments. E. coli strains were grown at 37°C (200 rpm) in Luria-Bertani medium. H. volcanii strains were grown at 37oC (200 rpm) in complex medium, ATCC 974. Media was supplemented

with 100 mg ampicillin or 0.1 mg novobiocin per liter as needed. The fidelity of all

polymerase chain reaction- (PCR-) amplified products was confirmed by DNA

sequencing using the dideoxy termination method (Sanger et al., 1992) with an

Amersham Pharmacia Biotech MegaBACE 1000 DNA Sequencer (DNA Sequencing

Facilities, Interdisciplinary Center for Biotechnology Research, University of Fla.).

Construction of H. volcanii Expression Plasmids

Expression plasmids were prepared with oligonucleotides according to Table 3-1.

PCR mutagenesis was used to add and alter optimized codons for H. volcanii

corresponding to each of the twenty essential amino acids at the end of the smrs-gfp gene

directly preceding a stop codon and to introduce restriction enzyme sites for directional

71 cloning into pJAM202, a shuttle plasmid for E. coli and H. volcanii. Plamids contained a

Halobacterium cutirubrum rRNA P2 promoter and T7 terminator for transcription of the cloned genes. A control plasmid was created (pJAM202C) which was identical to these plasmids except for the absence of a gene encoding a smRS-GFP reporter protein.

DNA Isolation and Transformation

Plasmids were isolated by Qiagen Miniprep (Qiagen Inc.,Valencia, Calif.). DNA fragments were eluted from 0.8% (wt/vol) SeaKem GTG agarose (FMC Bioproducts,

Rockland, Maine) gels with 1 × TAE buffer (40 mM Tris-acetate, 2 mM EDTA, pH 8.5) by QIAquick gel extraction (Qiagen). H. volcanii DS70 cells were transformed with plasmid DNA isolated from E. coli GM2163 as previously described (Cline et al., 1989).

Absorbance and Fluorescence of Cell Cultures

Cultures (25 ml) were grown (37ºC, 200 rpm) in 250 ml Erlenmeyer flasks inoculated in triplicate with 50 to 100 µl of a log phase culture of H. volcanii DS70 individually expressing either smRS-GFP or one its variants from the appropriate plasmids (Table 3-1). Optical Density (600 nm) of liquid cultures was measured using a

BioRad Smart Spec 3000 spectrophotometer (BioRad, Hercules, Calif.) at various time points over the course of at least 110 hours (late stationary phase). Fluorescence

(excitation 488 nm, emission 509 nm) of liquid cultures was monitored using an Aminco-

Bowman series 2 luminescence spectrometer (Spectronic Instruments, Rochester, N.Y.).

Innate fluorescence of control cultures, DS70 (pJAM202C), was subtracted from the fluorescence of DS70 cultures grown expressing smRS-GFP variants (data not shown).

One unit is defined as the fluorescence intensity of fluorescein at 0.025 nM as previously described (Reuter and Maupin-Furlow, 2004).

Results and Discussion

Fluorescence Curves of H. volcanii with Expression Plasmids

Previously we demonstrated that the addition of the SsrA motif to the C-terminus of smRS-GFP reduced its cellular protein levels beyond the point of detection by Western blot analysis. Alterations and truncations of the SsrA motif linked to the C-terminus of smRS-GFP had variable effects on the levels of the reporter proteins in H. volcanii in vivo. Surprisingly, a 10 residue truncation of the SsrA motif that left only a single Ala residue fused to the end of smRS-GFP was enough to diminish protein levels to 10% of those of unaltered smRS-GFP.

In order to determine whether other single C-terminal amino acid additions would have a similar effect on smRS-GFP reporter protein levels we constructed 20 smrs-gfp genes, each ending in an H. volcanii optimized codon corresponding to one of each of the twenty essential amino acids. The optical density (O.D. 600 nm) and fluorescence

(excitation 488 nm and emission 509 nm) of H. volcanii DS70 cultures expressing these proteins were measured throughout growth phase and compared to DS70 expressing unaltered smRS-GFP (pJAM1064) and DS70 with a control plasmid (pJAM202C).

Growth curves of H. volcanii containing plasmids expressing smRS-GFP variants were similar to those of the control with plasmid (pJAM202C) (Fig. 3-1). In contrast, the fluorescence of each construct varied substantially dependent on the nature of the amino acid placed at the C-terminus of the reporter protein (Fig. 3-1). Surprisingly, the addition of any single amino acid to the C-terminus of smRS-GFP, regardless of its identity, had a negative effect on the fluorescence of DS70 cultures expressing their respective reporter proteins when compared to DS70 cultures expressing unaltered smRS-GFP (pJAM1064).

DS70 (pJAM1064) displayed a concomitant rise in fluorescence during log phase

followed by a plateau of fluorescence throughout most of stationary phase and ending in

a slight steady increase in fluorescence late in stationary phase (Fig. 3-1). The

fluorescence of all H. volcanii DS70 cultures expressing smRS-GFP variants also began

to rise with log phase (Fig. 3-1). However, their fluorescence profiles then differed by

following a fairly consistent pattern including a peak, a subsequent drop, and a climb in

fluorescence, to varying degrees, throughout the rest of the growth phase. The initial

fluorescence peak ranged from 1 to 4 fluorescent units and occurred between an O.D. of

1 to 2. The variability in the fluorescence drop following the peak was even more

extreme, decreasing as much as 70% in DS70 expressing smRS-GFP-M (pJAM1071) to

being absent altogether in DS70 expressing smRS-GFP-F (pJAM1063). The most

intriguing event in the fluorescent profiles of the DS70 cultures expressing smRS-GFP

variants was the climb in fluorescence that occurred very late in stationary phase. This

also occurred to varying extents.

In an attempt to summarize and compare the multitude of fluorescence profiles, a

bar graph was constructed corresponding to the aforementioned peak (45 h), drop (65 h)

and climb (110 h) in fluorescence of DS70 cultures expressing each smRS-GFP variant

(Fig. 3-2). For simplicity, DS70 cultures expressing smRS-GFP reporter proteins were

arranged from left to right based on the highest to lowest average of these three values.

Summary and Conclusions

Here we have constructed 21 plasmids for the expression of smRS-GFP with variable single amino acids added to the C-terminus. The fluorescence of H. volcanii

DS70 cultures expressing these reporter proteins depends on the identity of the amino acid addition. The addition of a single amino acid to the C-terminus is believed to have little to no impact on the structure and fluorescence of GFP based on the GFP crystal

structure (Ormo et al., 1996) and circular dichroism of GFP vs. GFP-SsrA (Gottesman et

al., 1998), and therefore the fluorescence of these cultures is expected to correlate to

intracellular protein levels.

The addition of any amino acid to the C-terminus of smRS-GFP had a negative

impact on fluorescence of the DS70 culture expressing it compared to DS70 expressing

unaltered smRS-GFP. The reason for this is still unclear, although the unstructured nine-

residue tail of GFP has been implicated as a potential recognition factor of the archaeal

VAT protein, an ATPase regulatory particle of proteasomes (Gerega et al., 2005). In this case, an extension of the unstructured region may potentially increase its vulnerability as a substrate.

Amino acid additions with the least impact on the fluorescence of the DS70 culture expressing them compared to DS70 expressing unaltered smRS-GFP were charged amino acids (Lys, Glu, Asp) (smRS-GFP-K, smRS-GFP-D, and smRS-GFP-E). The addition of an acidic amino acid to the C-terminus would not be predicted to substantially influence the susceptibility of a protein to proteolysis, as they constitute about 20% of amino acids in H. volcanii proteins, many of which are found on the outer surface of the protein.

Likewise smRS-GFP reporter proteins with an addition of the amides of acidic amino acids (Asn and Gln) (smRS-GFP-N and smRS-GFP-Q) had a relatively minor impact on the fluorescence of DS70 cultures expressing them compared to DS70 expressing unaltered smRS-GFP. Although basic amino acids are found in a relatively low abundance in halophilic proteins, it is interesting that the last amino acid of wild type smRS-GFP is a Lys. The fluorescence of cultures expressing smRS-GFP-K is probably not explained by the basic nature of Lys, as additions of other basic amino acids, smRS-

GFP-R and smRS-GFP-H, had a relatively moderate impact upon the fluorescence of

their respective DS70 cultures.

The smRS-GFP reporter proteins whose host cultures displayed the lowest fluorescence profile had an addition of a hydrophobic amino acid to the C-terminus (Ala,

Ile, Tyr, and Val) (smRS-GFP-A, smRS-GFP-I, smRS-GFP-Y, and smRS-GFP-V). One exception is DS70 cultures expressing smRS-GFP-G, whose fluorescence profile was higher than the average. The lower hydrophobic character and single hydrogen side chain of Gly are likely the reason for this. The regulation as determined by the fluorescence profile of smRS-GFP-A is consistent with our earlier study finding of the severely decreased protein half-life of smRS-GFP-A as compared to wild type smRS-GFP. A possible explanation of this could be the exposure of a hydrophobic amino acid, which is of relatively low abundance in halophilic archaea, elicits a post-translational signal for an event such as proteolysis. Proteolytic recognition has been demonstrated in other organisms based on hydrophobic amino acid exposure (Bohley, 1996). These residues are typically found in the interior of a native protein structure and their exposure occurs most often when a protein is damaged and begins to unfold. The appropriate response of the cell would be to dispose of these proteins via proteolysis to prevent their aggregation and maintain protein quality control within the cell. It will be interesting to determine the true fate of these smRS-GFP reporter proteins using experiments such as translation inhibition and select protease inhibition, transcript comparison between cultures, and immunoblot analysis to confirm that protein levels correspond to fluorescence.

Table 3-1. Strains and plasmids used in this study. Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification E. coli - - + DH5α F recA1 endA1 hsdR17(rk mk ) supE44 thi-1 gyrA Life Technologies relA1 GM2163 F- ara-14 leuB6 fhuA31 lacY1 tsx78 glnV44 galK2 New England Biolabs galT22 mcrA dcm-6 hisG4 rfbD1 rpsL 136 dam13::Tn9 xylA5 mtl-1 thi-1 mcrB1 hsdR2 H. volcanii DS70 DS2 cured of pHV2 (Wendoloski et al., 2001) Plasmids: pET22b Apr; expression vector Novagen pBAP5010 Apr; Nvr; H. volcanii-E. coli shuttle-expression (Jolley et al., 1997) vector pJAM202 Apr; Nvr; H. volcanii-E. coli shuttle expression (Kaczowka and plasmid with psmB-his6 gene; β subunit of 20S Maupin-Furlow, proteasomes expressed with C-terminal His tag in H. 2003) volcanii pJAM202C Apr; Nvr; 9.94-kb NdeI-BlpI fragment blunt-end This study ligated together; H. volcanii-E. coli shuttle expression plasmid removed of the psmB-his6 gene for use as a control plasmid. pJAM1022 Apr; pET22b-derived plasmid expressing smRSGFP- (Reuter and Maupin- SsrA in E. coli Furlow, 2004) pJAM1062 Apr; Nvr; 0.73-kb BamHI-BlpI fragment generated by This study PCR (5’-AAGGATCCATGAGTAAAGGAGAAG- ‘3 and 5’-GAGCTCAGCTTTTAGCATTTG TATAG-‘3; BamHI and BlpI in bold) from pJAM1022 ligated into a 9.8-kb BamHI-BlpI fragment from pJAM202; smRS-GFP-C with a C- terminal Cys addition expressed in H. volcanii pJAM1063 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGAATTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-F with a C-terminal Phe addition expressed in H. volcanii pJAM1064 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTATTTGTATAGTTC-‘3 replaced primer 2 (BlpI site in bold); unaltered smRS-GFP expressed in H. volcanii

Table 3-1. Continued Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification pJAM1065 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGGCTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-A with a C-terminal Ala addition expressed in H. volcanii pJAM1066 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGTCTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-D with a C-terminal Asp addition expressed in H. volcanii pJAM1067 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACTCTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-E with a C-terminal Glu addition expressed in H. volcanii pJAM1068 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGAGTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-L with a C-terminal Lue addition expressed in H. volcanii pJAM1069 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGTTTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-N with a C-terminal Asn addition expressed in H. volcanii pJAM1070 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACTTTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-K with a C-terminal Lys addition expressed in H. volcanii pJAM1071 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACATTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-M with a C-terminal Met addition expressed in H. volcanii pJAM1072 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACGGTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-P with a C-terminal Pro addition expressed in H. volcanii

Table 3-1. Continued Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification pJAM1073 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACTGTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-Q with a C-terminal Gln addition expressed in H. volcanii pJAM1074 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGTGTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-H with a C-terminal His addition expressed in H. volcanii pJAM1075 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGATTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-I with a C-terminal Ile addition expressed in H. volcanii pJAM1076 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGCGTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-R with a C-terminal Arg addition expressed in H. volcanii pJAM1077 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACGATTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-S with a C-terminal Ser addition expressed in H. volcanii pJAM1078 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACGTTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-T with a C-terminal Thr addition expressed in H. volcanii pJAM1079 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGTATTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-Y with a C-terminal Tyr addition expressed in H. volcanii pJAM1080 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTACCATTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-W with a C-terminal Trp addition expressed in H. volcanii

Table 3-1. Continued Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification pJAM1081 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGACTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-V with a C-terminal Val addition expressed in H. volcanii pJAM1082 Apr; Nvr; like pJAM1062, except 5’- This study GAGCTCAGCTTTTAGCCTTTGTATAG-‘3 replaced primer 2 (BlpI site in bold); smRS-GFP-G with a C-terminal Gly addition expressed in H. volcanii

Table 3-2. Optimal codon usage of Haloferax volcanii. Downloaded from GenBank Codon Usage Database. (http://www.kazusa.or.jp/codon/cgi- bin/showcodon.cgi?species=Haloferax+volcanii+[gbbct]).

Fig. 3-1. Growth and Fluorescence of H. volcanii DS70 with GFP reporter-protein expression plasmids. The growth of DS70 (pJAM202C) (filled triangles) was compared to the growth of DS70 with GFP reporter-protein expression plasmids (open triangles). The background fluorescence of DS70 cultures with plasmid pJAM202C (data not shown) were subtracted from the fluorescence of DS70 cultures with GFP reporter-protein expression plasmids to generate a net fluorescence curve of each (open circles), as indicated on each figure.

Fig. 3-1. Continued

Fig. 3-2. Single C-terminal amino acid additions to smRS-GFP variably affect the fluorescence of the H. volcanii DS70 culture from which they are being expressed. Bar graph comparison of points corresponding to the “peak” (45 h) (open bars), “drop” (65 h) (shaded bars), and “climb” (110 h) (filled bars) in fluorescence profiles of DS70 expressing smRS-GFP reporter proteins. From left to right indicates highest to lowest fluorescence based on the average of these three values. Letters on the x-axis represent DS70 the amino acid residue added to the C-terminus of smRS-GFP.

CHAPTER 4 IDENTIFICATION AND PARTIAL CHARACTERIZATION OF PROTEASOME- ACTIVATING NUCLEOTIDASE REGULATTORY PARTICLES OF HALOFERAX VOLCANII

Introduction

Energy-dependent proteolysis is a vital process in all three domains of life.

Proteasomes are well-conserved nanocompartmental proteases distributed throughout

Archaea, Eukarya, and eubacterial actinomycetes that are predicted to play a central role in the energy-dependent degradation of proteins (Lupas et al., 1997; Maupin-Furlow et al., 2004). 20S proteasomes are composed of proteins of related α and β superfamiles, which respectively form two outer and two inner heptameric rings that create a barrel- shaped complex (Coux et al., 1996). Access to the inner chamber of 20S proteasomes, or core particles (CPs), which house the catalytically active Thr resides, is through two small pore openings further restricted by axial gates (Groll et al., 2000). Therefore two conditions must be met in order for translocation of folded substrate to occur: protein must be unfolded and the gate must be opened. Both of these events appear to be controlled by the regulatory particles (RP) associated with 20S CPs (Smith et al., 2005).

The eukaryotic 19S cap is a proteasomal regulatory particle composed of 19 subunits further subdivided between a lid and base. The base includes six Rpt (regulatory particle triple-A type I ATPases) subunits belonging to the AAA+ ATPase family, whose responsibilities include substrate unfolding and gate opening (Schmidt et al., 2005).

Their counterparts in archaea, Pan (proteasome-activating nucleotidase) proteins, share

93 94 up to 40 to 50% sequence identity and are involved in similar activities (Zwickl et al.,

1999; Wilson et al., 2000; Reuter et al., 2004). Substrate binding to Pan stimulates ATP hydrolysis which is believed to power mechanistic-like conformational changes of the

Pan complex for the purpose of substrate unfolding (Hanson and Whiteheart, 2005).

Gate opening precedes translocation of the unfolded substrate into the CP and is also controlled by the Pan complex, although this event appears to be energy-independent as it requires only nucleotide binding by Pan (Smith et al., 2005).

Like many eukaryal-related archaeal complexes, the Pan/20S proteasome is composed of a much simpler complexity, allowing for a greater ease of its study.

Eukaryal 26S proteasomes are composed of multitude of proteins and can associate with thousands more for the purpose of regulating substrate selection (Groll and Huber, 2005).

Although not universal to archaea, Pan is typically encoded from one or two homologous genes (Maupin-Furlow et al., 2004). To date, only the single Methanococcus jannaschii

Pan purified from recombinant Esherichia coli has been characterized at the biochemical level (Zwickl et al., 1999; Wilson et al., 2000; Navon and Goldberg, 2001; Benaroudj et al., 2003). Recently our lab cloned and purified two Pan homolgues (PanA and PanB) for antibody production from recombinant E. coli and demonstrated differential regulation of their native proteins in vivo from their host organism, Haloferax volcanii

(Reuter et al., 2004). The presence of these two differentially regulated Pan proteins coupled with the dissimilarity of their N-terminal coiled-coil regions, which are proposed to be responsible for substrate interaction, suggests the existence of multiple functional

Pan isoforms. Multiple Pan complexes could compliment the presence of at least two other known proteasome CP isoforms in this organism (α1α2β and α1β) (Kaczowka and

Maupin-Furlow, 2003). It is possible that the consequence of this slightly increased

complexity results in an alteration of function or modulation of proteolytic substrate

recognition.

Here, for the first time, we have purified and characterized Pan proteins (PanA

and PanB) from an organism that encodes multiple Pan paralogs. This also represents the

first study of Pan from an organism outiside the genus Methanococcus and the first study

utilizing Pan protein expressed from its host organism. We demonstrate that at least three

Pan complexes exist in H. volcanii. Two of these complexes are homo-oligomeric

consisting entirely of either PanA or PanB subunits, while the third forms a hetero-

oligomeric complex consisting of both proteins. This study also presents a biochemical

characterization of these complexes.

Materials and Methods

Materials

Organic and inorganic analytical grade chemicals were from Fisher Scientific

(Atlanta, Ga.) and Sigma Chemical Co. (St. Louis, Mo.) unless otherwise indicated.

Desalted oligonucleotides were from Operon Technologies (Alameda, Calif.) and

Integrated DNA Technologies (Coralville, Ind.). Restriction endonucleases and DNA- modifying enzymes were from New England BioLabs (Beverly, Mass.).

Strains, Media, and Plasmids

Strains, oligonucleotide primers used for cloning, and plasmids are summarized in

Table 4-1. E. coli DH5α was used for routine recombinant DNA experiments. E. coli.

E. coli BL21(DE3) containing the plasmid pSJS1240 (expressing ileX and argU genes

encoding tRNAAUA and tRNAAGA/AGG) was used as a host for optimal expression and

purification of H. volcanii proteins (Kim et al., 1998). E. coli strains were grown at 37°C

(200 rpm) in Luria-Bertani medium. H. volcanii strains were grown at 42oC (200 rpm) in complex medium, ATCC 974. Media was supplemented with 100 mg ampicillin, 50 mg kanamycin, 50 mg spectinomycin, 4 mg mevinolin, and/or 0.1 mg novobiocin per liter as needed. The fidelity of all polymerase chain reaction- (PCR-) amplified products was confirmed by DNA sequencing using the dideoxy termination method (Sanger et al.,

1992) with Perkin Elmer/Applied Biosystems and LICOR automated DNA Sequencers

(DNA Sequencing Facilities, Interdisiplinary Center for Biotechnology Research and

Department of Microbiology and Cell Science and, University of Fla.).

DNA isolation, Analysis, and Transformation

Plasmids were isolated by Qiagen Miniprep (Qiagen Inc.,Valencia, Calif.). DNA

fragments were eluted from 0.8% (wt/vol) SeaKem GTG agarose (FMC Bioproducts,

Rockland, Maine) gels with 1 × TAE buffer (40 mM Tris-acetate, 2 mM EDTA, pH 8.5)

by QIAquick gel extraction (Qiagen). H. volcanii DS70 cells were transformed with

plasmid DNA isolated from E. coli GM2163 according to Cline et al. (Cline et al., 1989).

PCR was used to introduce restriction enzyme sites for directional cloning of panA,

panA∆120 and panB into plasmid pET24b for expression in E. coli (pJAM642,

pJAM643, and pJAM1006)(Table 4-1). Plasmids pJAM642 and pJAM643 were

generously provided by Julie Maupin-Furlow. The start codons of these genes were

positioned 8 bp downstream of the ribosome binding sequence of pET24b using NdeI.

For epitope tagging, the 3’-end of the gene was modified by PCR to remove the stop

codon and provide an in-frame C-terminal addition of poly-histidine residues. PCR was

used to introduce restriction enzyme sites in pMCSG7 and panA and panB for directional

cloning of vectors including upstream His-tags and TEV protease cleavage site encoding regions for plasmids pJAM1060 and pJAM1086. Plasmid pMCSG7 was provided courtesy of Mark Donnelly.

Protein Techniques and Antibody Production

as indicated) with β-mercaptoethanol and sodium dodecyl sulfate (SDS-PAGE). SDS-

PAGE Low Range molecular weight standards (BioRad) include phosphorylase b (97.4 kDa), serum albumin (66.2 kDa), ovalbumin (45 kDa), carbonic anhydrase (31 kDa), trypsin inhibitor (21.5 kDa), and lysozyme (14.4 kDa). After separation, proteins were

transferred to Hybond-P membranes (Amersham Biosciences, Piscataway, N.J.) using 10

mM 2-N-morpholino ethanesulfonic acid (MES) at pH 6.0 with 10% (vol/vol) methanol

(20 V, 4ΕC, 16 h). Prior to antibody preparation, the N-terminal sequences of PanA∆1-40-

His and PanA-His were determined by Edman degradation, and the mass spectra of the tryptic fragments of PanB-His were determined by MALDI-TOF (Applied Biosystems

QSTAR XL Hybrid LC/MS/MS)(Univ. Fla. ICBR Protein Chemistry Core, Fla.).

Preparation of PanA for antibody production was performed by Julie Maupin-Furlow.

PanA∆1-40 –His and PanB-His proteins purified from recombinant E. coli were separated by 7.5% SDS-PAGE, excised from the gel, and used as antigen to raise polyclonal antibodies in rabbit (Cocalico Biologicals, Reamstown, PA)(Reuter et al., 2004).

Horseradish peroxidase- (HRP-) conjugated anti-rabbit Ig (H+L) antibody or alkaline phosphatase- (AP) conjugated anti-rabbit Ig (H+L) antibody raised in goat was used as

the secondary antibody (Southern Biotechnology Associates, Inc., Birmingham, Ala.).

HRP activity was detected using ECL Plus chemiluminescence (Amersham Biosciences)

with a VersaDoc 1000 imaging system (BioRad). AP activity was detected in developing

buffer (100mM Tris pH 9.5, 100mM NaCl, and 5mM MgCl2) with the ordered addition of 100 µl of 50 mg/ml NBT solution (nitroblue tetrazolium in 70% dimethylformamide

(DMF)) and 50 µl 50 mg/ml BCIP solution (5-bromo-4-chloro-3-indolyl-phosphate in

100% DMF).

Protein Expression in E. coli

Haloferax volcanii proteins were expressed in recombinant E. coli BL21(DE3) with pSJS1240 and one of the following; pJAM642 (PanA-His), pJAM1006 (PanB-His), pJAM1086 (His-TEV-PanA), pJAM1060 (His-TEV-PanB), or pJAM643 (PanA∆1-40 -His)

as previously described (Maupin-Furlow et al., 1998;Reuter et al., 2004). Cells were

harvested (6,000 x g, 15 min, 4ºC) lysed in 6 volumes (wet wt per vol) of Buffer A (150

mM NaCl, 20 mM Tris buffer, pH 7.2) by passage through a French pressure cell at

20,000 lb/in2. Cell debris was removed by centrifugation (14,000 × g, 20 min, 4ºC) using

a Sorvall SS-34 rotor and Evolution RC centrifuge (Thermo Electron Corp., Waltham,

Mass.). Cell lysate was dialyzed overnight into Buffer B (2 M NaCl, 20 mM Tris buffer,

pH 7.2) and filtered (0.22 µM) (Nalge Nunc International, Rochester, N.Y.).

SmRS-GFP-SsrA was expressed in E. coli BL21(DE3) by using the bacteriophage

T7 RNA polymerase-promoter system (Tabor and Richardson, 1992). An overnight

culture of cells (1 ml) was inoculated into a 1 L Fernbach flask containing Luria-Bertani

medium (500 ml) supplemented with ampicillin (100 mg/liter) and grown at 37°C and

200 rpm until the cells reached an OD500nm of about 0.7. T7 RNA polymerase-dependent

99 transcription was then induced by incubation with 0.4 mM isopropyl- -D- thiogalactopyranoside (IPTG) for 2 h. Cells were harvested as described above, resuspended in Buffer C (250 mM NaCl, 50 mM sodium phosphate buffer, pH 7.5), lysed, clarified, and filtered as described above.

Protein Expression in H. volcanii

Pan proteins were expressed from H. volcanii from the following combinations of plasmids and strains: DS70 pJAM1012 (PanB-His) and GG102 (pJAM1012) (both expressing PanB-His ) and DS70 (pJAM650) and GG101 (pJAM650) (both expressing

PanA-His). Four one-liter cultures of each were grown separately in 2.8-liter Fernbach flasks to late stationary phase (OD600, 3.5). Cells (~9 g wet wgt) were harvested, resuspended in Buffer B, lysed, clarified, and filtered as described above.

Protein Purification

Cell lysates containing His-tagged Pan were initially separated using IMAC

(immobilized metal-affinity chromatography) with a BioLogic HR workstation (BioRad) as follows. Lysates were applied to a 5 ml HiTrap Sepharose column (4.8 by 0.8 cm) (GE

Healthcare Life Sciences, Piscataway, NJ) charged with 2 ml 0.1 M NiSO4•6H2O, equilibrated in Buffer B supplemented with 5 mM imidazole, and washed with 15 ml step gradients of Buffer B supplemented with 5 and 60 mM imidazole. Fractions containing

His-tagged proteins were eluted in Buffer B with 500 mM imidazole. Fractions of Pan-

His lacking TEV protease cleavage sites were further purified by application to a 5 ml

Bio-scale hydroxyapatite type I column (BioRad) equilibrated in 10 mM sodium phosphate buffer at pH 7.2 with 2 M NaCl (Buffer D). The column was washed with 15 ml of Buffer D and developed with a 30 ml linear sodium phosphate gradient (10 to 500 mM sodium phosphate at pH 7.2 with 2 M NaCl). Protein fractions containing PanA-His

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and PanB-His eluted at 250-300 and 200-250 mM sodium phosphate respectively, while

heter-oligomeric Pan complexes eluted at 230-260 mM sodium phosphate. His-TEV-Pan

fractions were dialyzed post IMAC into Buffer B overnight and the N-terminal His-TEV

region was removed with AcTEV protease according to the manufacturers protocol with

the exception of an additional 2 M NaCl in the reaction mixture (Invitrogen Life

Technologies, Carlsbad, Calif.). All samples (0.5 ml per run) were applied to a Superose-

6 HR 10/30 gel filtration column (Amersham Bioscience) equilibrated in Buffer B.

Native molecular masses of Pan complexes where calculated by a standard curve created

by calibrating the column using the molecular mass standards serum albumin (66 kDa),

alcohol dehydrogenase (150 kDa), β-amylase (200 kDa), apoferritin (443 kDa), and

thyroglobulin (669 kDa) in Buffer A.

The initial step in purification of smRS-GFP-SsrA was performed using IMAC as previously described by Li et al. with a few exceptions (Li et al., 2001;Li and Beitle,

2002). Lysate was applied to a 5 ml HiTrap Sepharose column charged with 2 ml of 0.1

M CuSO4•5H2O, equilibrated in Buffer C supplemented with 5 mM imidazole, washed

with a 10 ml of Buffer C supplemented with 5 mM imidazole, and developed with a 15

ml linear imidazole gradient (5 to 70 mM imidazole in Buffer C). Protein fractions

containing smRS-GFP-SsrA eluted at about 60 mM imidazole were pooled and applied to

a 5 ml Bio-scale hydroxyapatite type I column (BioRad) equilibrated in 150 mM NaCl,

10 mM sodium phosphate buffer, pH 7.5 (Buffer D-2). The column was washed with 15

ml of Buffer D-2 and developed with a 30 ml linear sodium phosphate gradient (10 to

500 mM sodium phosphate at pH 7.2 with 2 M NaCl). Protein fractions containing

smRS-GFP-SsrA eluted at about 450 mM sodium phosphate and were dialyzed into

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Buffer B. Samples were applied to a Superdex 200 10/30 gel filtration column

(Amersham Pharmacia Biosciences Inc., Piscataway, NJ) equilibrated in Buffer B.

SmRS-GFP-SsrA eluting in fractions corresponding to a molecular weight of ~31 kDa

were collected.

Nucleotide-hydrolyzing Activity

Nucleotide-hydrolyzing activity assays were performed under a wide range of temperature, pH, and salt concentrations (KCl and NaCl) as described in Fig 4-8, with a constant concentration of MgCl2 (9.6 mM), ATP (1 mM), and PanA-His purified from

GG102 (pJAM650) (0.0052 mg/ml). Several buffers were used based on their optimal

buffering range and desired pH of the reaction including MES (2-(N-Morpholino)

ethanesulfonic acid hydrate) (pH 6.3), MOPS (3-(N-Morpholino)propanesulfonic acid)

(pH 6.5-7.9), Tris (Tris(hydroxymethyl)aminomethane) (pH 8.4), and CAPSO (3-

(Cyclohexylamino)-2-hydroxy-1-propanesulfonic acid) (pH 8.8-10.3). To avoid

phosphate contamination all glassware was washed with 10% w/v chromic acid, all

plastic ware was new and non-autoclaved, and deionized, double distilled reagent grade

water (Ricca Chemical Co., Arlington, Tx.) was used to prepare all buffers and rinse

glassware. Reactions were carried out for 30 min in 0.5 ml microcentrifuge tubes

(FisherBrand). A portion of the reaction (50 µl) was used to measure inorganic

phosphate (Pi) concentration using solutions E and F as previously described by

Gawronski et al. (Gawronski and Benson, 2004). Samples were analyzed at A655 in clear

polystyrine 96-well F microtiter plates (Nalge Nunc International, Rochester, N.Y.) using

a BioTek Synergy HT microtiter plate reader (Bio-Tek Instruments, Inc., Winooski,

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Vermont). A standard curve was developed in the same manner using concentrations of

Pi from 0-100 nM.

DNA and Protein Sequence Analyses

Clone Manager Professional Suite version 6.0 (Scientific & Educational Software,

Durham NC) and BioEdit Sequence Alignment Editor version 5.0.9 (Hall, 1999) were used for DNA and protein sequence analyses. The probability that a deduced protein sequence will adopt a coiled-coil conformation was predicted using COILS with weighted and unweighted MTK and MTIKL scoring matrices set to scanning windows of

21 and 28 residues (Lupas, 1996).

Nucleotide Sequence Accession Numbers

The nucleotide sequence of the panA and panB genes of H. volcanii DS2 were assigned GenBank accession numbers AY627303 and AY627304.

Results and Discussion

H. volcanii Synthesizes Two Pan Proteins

The synthesis of multiple archaeal Pan proteins has not yet been reported. In fact, most archaea encode only a single Pan homolog, and several do not encode any Pan (e.g.,

Thermoplasma spp., Pyrobaculum). To address whether H. volcanii synthesizes both the

PanA and PanB proteins, an immunoblot approach was employed. Polyclonal antibodies were raised in rabbits against PanA∆1-40-His and PanB-His purified from recombinant E. coli. Antigen fidelity was confirmed by N-terminal sequencing and Q-STAR MALDI-

TOF (Fig. 4-1). Western blot analysis of cell lysate prepared from stationary phase H. volcanii was performed using these antibodies (Fig. 4-2). Both PanA and PanB proteins were detected and based on migration by SDS-PAGE their molecular masses were estimated to be 54 and 65 kDa, respectively. Under similar electrophoretic conditions,

103 the PanA-His and PanB-His purified from recombinant E. coli migrated as slightly larger proteins (55 and 66 kDa) due to the addition of six C-terminal histidine residues. The similar migration of the PanA and PanB proteins purified from recombinant E. coli to those of H. volcanii supports the conclusion that the gene sequences isolated from the H. volcanii chromosome code for the PanA and PanB polypeptides detected in cell lysate. It remains to be determined why the molecular masses calculated from the deduced PanA and PanB protein sequences were at least 8 kDa less than those estimated by SDS-PAGE gels. Although the separation of halophilic proteins by SDS gel electrophoresis usually overestimates molecular weight (Vyazmensky et al., 2000), this alone is unlikely to account for the observed differences. It is possible that the N-terminal region of these

Pan proteins adopts a coiled-coil conformation under these conditions which retards their electrophoretic migration.

Protein Sequence Analysis of Pan from H. volcanii

The deduced PanA and PanB protein sequences are closely related with 60% identical and 78% similar amino acid residues (Fig 4-1). The most significant region of identity was the central P-loop nucleotidase domain, which includes Walker A and B boxes, as well as the second region of homology typical of AAA ATPases (Frickey and Lupas,

2004). N-terminal residues of PanA and PanB (position 17 to 72 and 39 to 73, respectively) are predicted to adopt a coiled-coil conformation. In contrast to the central domain, however, the N-terminal coiled-coil domains were not highly related (17% identity) and differed in length by 21 residues (3 helical turns). Likewise, C-terminal residues 324 to 406 of PanA were only 49% identical to residues 322 to 412 of PanB.

This divergence in the primary sequence of the N- and C-termini, combined with the highly conserved central domain, suggests that PanA and PanB mediate similar yet

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distinct functions in H. volcanii. The N-terminal coiled-coil region of Pan/Rpt has been

proposed to be involved in interactions with other RP subunits and/or substrate proteins

(Gorbea et al., 2000). Furthermore, modification of the C-terminus of Rpt1 (i.e., addition of a FlagHis6 epitope tag in yeast) inhibits RP association with the CP in yeast (Verma et al., 2000). Thus, based on differences in the N- and C-termini, it is possible that the

PanA and PanB proteins of H. volcanii recognize different substrate proteins and/or differ in their affinity for α1β and α1α2β 20S proteasome subtypes.

Pan Purification from Recombinant H. volcanii

To date, in vitro studies on archaeal Pan have exclusively relied upon N-terminal

His-tagged M. jannaschii Pan protein purified from recombinant E. coli (Zwickl et al.,

1999; Navon and Goldberg, 2001; Benaroudj et al., 2003) . The study of Pan from only

one organism vastly limits our understanding of its global function and significance. This

is especially true for organisms that encode paralogous Pan proteins, of which none have

thus far been characterized. It has yet to be determined if multiple Pan proteins

associate in these organisms, and if they do, what affect multiple Pan complexes have

upon substrate differentiation and processing in cooperation with 20S proteasomes.

Equally important to studying Pan from an organism that encodes multiple Pan proteins,

is doing so with Pan protein expressed from its native organism. It is possible that

important post-translational events, accessory proteins, and folding mechanisms, which

occur in protein expression from a native host, are altered or devoid when protein is

recombinantly expressed from another organism. This is especially true of halophilic

protein production from E. coli, whose intracellular environment has a 3-4 fold lower salt

concentration than many halophilic archaea (Engel and Catchpole, 2005). Therefore it is

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likely that recombinant protein production from a native host grants in vitro protein

studies more physiological relevance. For these reasons we initially expressed PanB-His in H. volcanii, one of the few archaea with an established genetic system. This was

accomplished by using H. volcanii strain DS70 carrying plasmid pJAM1012, which

encodes the PanB-His protein (Table 4-1) (Fig. 4-4). Following an IMAC purification

step, fractions were visualized by a Coomassie blue (CB) stained SDS-PAGE gel, which

revealed the presence of three bands. The uppermost band was determined to be a

nonspecific protein termed PitA whose separation by IMAC occurs even from DS70

alone, presumably due to a histidine-rich region determined by mass spectroscopy (Bab-

Dinitz et al., 2006). Furthermore, PitA can be separated from Pan by subsequent ionic

chromatography. The possibility was considered that PanB-His expressed in H. volcanii

could potentially associate with an additional protein in vivo leading to their simultaneous

extraction by IMAC. In an attempt to discern the identity of these proteins, a Western

blot analysis was used with primary anti-PanA antibody, which due the their moderate

protein sequence conservation, interacts with PanA and to a lesser extent PanB. The

middle band corresponded to the mobility of an IMAC purified EcPanB-His (expressed

in E.coli BL21(DE3) from pJAM1006) (65 kDa) (Table 4-1) while the lower band, which

produced a more intense signal despite the apparent equality of loaded protein to the

middle band, corresponded to mobility of EcPanA-His (expressed in E.coli BL21(DE3) from pJAM642) (54 kDa) (Table 4-1) (Fig. 4-5A).

To further investigate a potential PanB and PanA association, a more elaborate

SDS-PAGE and Western blot analysis was performed which included HvPanA-His

(PanA-His expressed in H. volcanii DS70 from pJAM650, HvPanB-His, EcPanA-His,

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and EcPanB-His (Fig. 4-5B). From this result it was concluded that PanA and PanB

associate in H. volcanii in vivo to form a hetero-oligomeric complex, as both

independently expressed His-tagged Pan proteins co-purify with their respective homolog through IMAC, independent of which one is His-tagged. Association of these homologous Pan proteins displays an increased subunit complexity compared to those previously demonstrated.

Independent Pan Expression from H. volcanii

To date there has been only one study that analyzed Pan from its native host, M. jannaschii, although this protein was never purified to homogeneity (Wilson et al., 2000).

It was demonstrated the molecular mass of the native Pan complex was about 550 kDa, however, it was never determined whether or not this complex was associated with other proteins. Since this organism contains only one Pan gene, and the protein self-assembles into a 550 kDa dodecamer in recombinant E. coli, it is assumed that the pool of Pan in this native complex is homogeneous. In order to determine whether PanA and PanB also form oligomers independent of each other, H. volcanii knockout mutants which do not produce PanA (GG102) or PanB (GG101) (Gil and Maupin-Furlow, unpublished) were used to express PanB-His (pJAM1012) and PanA-His (pJAM650), respectively. Proteins were purified by IMAC and each was determined to be independent of its respective homolog by Western blot analysis (data not shown).

Native Pan Forms Oligomers

Previous studies have determined that in the native state Pan associates into oligomeric complexes of between 550 and 650 kDa, which are relative to the size of a dodecamer, possibly consisting of a dimer of oligomeric rings (Zwickl et al., 1999;

Wilson et al., 2000;). Pan belongs to the AAA+ (ATPases associated with various

107 cellular activities) superfamily of proteins of which many form oligomeric rings, often times in the form of hexamers and occasionally heptamers (Hanson and Whiteheart,

2005). In order to distinguish the size of the three Pan isoforms from H. volcanii, fractions of IMAC purified PanA-His (pJAM650 expressed from DS70 and GG101) and

PanB-His (pJAM1012 expressed from DS70 and GG102) were applied to gel filtration chromatography (Fig. 4-5) and analyzed by SDS-PAGE (Fig. 4-6) and Western Blot (data not shown). PanA-His (GG101 pJAM650) predominantly eluted at fractions corresponding to a molecular mass of 600 to 650 kDa, consistent with that of the approximate dodecameric size previously studied Pan complexes. PanB-His (GG102 pJAM1012) also predominantly eluted at fractions corresponding to a molecular mass of

600 to 650 kDa, but also eluted at a peak corresponding to the size of a hexamer (about

270 kDa). PanA-His and PanB-His expressed in DS70 eluted as homo-oligomeric complexes at molecular masses of 600 to 650 kDa, however were only found as apparent hetero-hexameric complexes when eluted in association with their respective homolog.

For reasons yet to be determined, PanA/B hetero-oligomeric complexes appear to be incapable of forming this high molecular mass complex and only associate as a complex about the size of a hexamer.

ATP-hydrolyzing Activity of PanA

PanA-His appears to be specific for ATP as other nucleotides (CTP, GTP, TTP) were not hydrolyzed (data not shown). In order to determine the optimal conditions for

Pan activity, the rate of ATP-hydrolysis was measured using PanA-His purified from H. volcanii GG101 (pJAM650), which displayed the highest ATPase activity of the Pan isoforms. A wide range of conditions were tested and included temperatures from 20 to

75°C, salt concentrations (KCl and NaCl) from 0.5 to 3.5 M, and a pH range of 6.3 to

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10.3 (Fig. 4-7). PanA-His hydrolyzed ATP to an optimal extent in 2.5 M KCl at 45 to

50°C in MOPS, buffered at a pH of 7.5 to 8.5. These data are very consistent with the growth conditions of H. volcanii, which are neutrophilic halophiles whose optimal

growth temperature is around 42 to 45°C. Salt preference by this enzyme, which was as

much as two times more active in KCl than NaCl, also correlates well with the

intracellular environment of many halophilic archaea where K+ is the most prominent ion

(between 1.9 and 5.5 M) and far exceeds that of Na+ (Perez-Fillol and Rodriguez-Valera,

1986). PanA activity was also increased in the reactions buffered with MOPS as opposed

to Tris, although a reason for this is not immediately known. The optimal ATP-

hydrolyzing activity of PanA-His is about 2.5 fold higher than that of M. jannaschii Pan purified from recombinant E. coli (Wilson et al., 2000).

One would expect that the optimal ATP-hydrolyzing conditions of Pan would correlate to the activity of 20S proteasomes. This, in fact, is true for pH and salt concentration as previous biochemical studies of 20S proteasomes of H. volcanii demonstrate its peptide hydrolyzing activity to be optimal at 2 M NaCl (KCl was not tested in this study) and at a pH between 7 and 9 (Wilson et al., 1999). However, the

optimal temperature of 20S proteasome activity was about 65°C, significantly higher than that of PanA-His. At 50°C, 20S proteasomes maintained about 45% of optimal activity.

Several groups have debated between substrate unfolding and substrate translocation as being the rate-limiting step in ATP-dependent proteolysis (Kim et al., 2000; Singh et al.,

2000; Benaroudj et al., 2003). However, all of these groups agree that the rate-limiting step precedes peptide bond cleavage. In this case, energy-dependent proteolysis by the

Pan/20S proteasome complex would proceed efficiently, despite protein cleavage at half

109 of its theoretical capacity. Furthermore and more simply, these organisms grow in regions where temperatures rarely exceed 45°C.

The optimal conditions for ATP-hydrolysis of the other Pan complexes (homo- olgomeric PanA and hetero-oligomeric PanA/B) have yet to be elucidated. Continuous efforts are underway to resolve this issue.

SmRS-GFP-SsrA as a Substrate for Pan/20S Proteasomes in vitro

Several previous studies have demonstrated the requirement of Pan for degradation of folded substrates by 20S proteasomes in vitro (Zwickl et al., 1999; Wilson et al., 2000;

Navon and Goldberg, 2001; Benaroudj et al., 2003). The most common of these substrates used is the reporter protein Green Fluorescent Protein (GFP) with an 11- residue SsrA C-terminal fusion tag (smRS-GFP-SsrA). GFP proteins without this tag are no longer recognized as a proteolytic susbtrate by Pan/20S proteasomes. We have also demonstrated that soluble-modified red-shifted GFP-SsrA is degraded by 20S proteasomes in vivo (Reuter and Maupin-Furlow, 2004) (Chapter 2). In order to elucidate the unfolding and degradation functions of multiple H. volcanii Pan/20S proteasome complexes in vitro, we expressed and purified smRS-GFP-SsrA from recombinant E. coli for use as a substrate. Purified smRS-GFP-SsrA was determined to be homogeneous by

SDS-PAGE (Fig. 4-8A) and displayed fluorescence in a high salt concentration (Fig. 4-

8B). Unfortunately, we have thus far been unable to demonstrate that smRS-GFP-SsrA is degraded by H. volcanii PanA-His/20Sproteasomes in vitro, despite efforts do so under a variety of conditions. PanA-His/20S proteasomes were also unable to degrade several other potential substrates in vitro including: commercially available β-casein;

Halobacterium marismortui malate dehydrogenase (HmMDH) partially purified from

110

recombinant E. coli; and two potential H. volcanii 20S proteasome substrates partially

purified from recombinant E. coli (determined by 2D-PAGE proteome analysis of H.

volcanii cultures incubated with the proteasome specific inhibitor clasto-lactacystin β-

lactone) (Kirkland et al., unpublished). It is likely that the C-terminal His-tag of the Pan

is disrupting association with the 20 CP as the C-terminal region of Pan is proposed to

make this association. In one study, a deletion of a single C-terminal residue of Pan was

shown to render a deficiency in substrate degradation by Pan/20S proteasomes (Forster et al., 2005). Furthermore, the His-tag may also be interfering with Pan-dependent α-ring

gate opening. A prior in vitro study demonstrated dramatically accelerated GFP-SsrA degradation when in the presence of Pan and α-ring gate deleted 20S proteasomes compared to Pan and wild type 20S proteasomes (Benaroudj et al., 2003). Efforts are currently underway to construct a counterpart to these gate-deleted α-subunit mutants in

H. volcanii. E. coli expression plasmids have already been constructed containing genes for panA (pJAM1086) and panB (pJAM1059) with removable N-terminal His-tags via an internal Tobacco etch virus (TEV) protease cleavage region. His-TEV-PanA and His-

TEV-PanB proteins have successfully been expressed and purified from recombinant E. coli and removal of the His-tag has been performed in high salt conditions (data not shown). These genes have also been cloned into H. volcanii expression plasmids pJAM1084 (His-TEV-PanA) and pJAM1085 (His-TEV-PanB) and are currently awaiting expression from recombinant H. volcanii.

Summary

Here we have identified two pan genes (panA and panB) from H. volcanii and

demonstrated that their proteins form homo- and hetero-oligomeric complexes in vivo.

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Consistent with most AAA+ family members, Pan is predicted to form hexameric rings

(Hanson and Whiteheart, 2005). The presence of dodecameric-sized PanA and PanB

homo-oligomers supports studies from M. jannaschii that suggest one state of Pan may be in the form of two, joined hexameric rings (Zwickl et al., 1999; Wilson et al., 2000).

The increased complexity of Pan complexes discovered here may allow for recognition of

a diversified repertoire of proteolytic substrates in H. volcanii. This scenario supports our

previous finding of differential regulation of proteasomal proteins throughout growth

phase in H. volcanii and further implies that distinct Pan/20S proteasome isoforms

perform variable functions within the cell (Kaczowka and Maupin-Furlow, 2003; Reuter et al., 2004). We have also begun characterization of these Pan complexes using the

PanA homo-oligomer, representing the first attempt of Pan characterization utilizing

protein produced from an organism other than M. jannaschii and more importantly from

an organism encoding two paralogous Pan proteins. The optimal ATP-hydrolyzing

activity of PanA is consistent with the peptide hydrolyzing activity of 20S proteasomes

from H. volcanii, which further supports their relation in a cooperative function. Future

studies will attempt to elucidate the in vitro degradation of smRS-GFP-SsrA, as well as

other substrates, by H. volcanii Pan/20S proteasomes, and determine whether a difference

in function exists between the various Pan isoforms.

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Table 4-1. Strains and plasmids used in this study. Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification E. coli - - + DH5α F recA1 endA1 hsdR17(rk mk ) supE44 thi-1 gyrA Life Technologies relA1 GM2163 F- ara-14 leuB6 fhuA31 lacY1 tsx78 glnV44 galK2 New England Biolabs galT22 mcrA dcm-6 hisG4 rfbD1 rpsL 136 dam13::Tn9 xylA5 mtl-1 thi-1 mcrB1 hsdR2 - - - BL21(DE3) F ompT [lon] hsd SB (rB mB ) (an E. coli B strain) Novagen with DE3, a λ prophage carrying the T7 RNA polymerase gene H. volcanii DS70 DS2 cured of pHV2 (Wendoloski et al., 2001) WR480 ∆pyrE2; for use in gene knockouts (Bitan-Banin et al., 2003) GG101 WR480 panB1038 (contains panB deletion) (Gil & Maupin- Furlow, unpublished) GG102 DS70 panA187::hmgA*(MevR) (contains panA (Gil & Maupin- deletion) Furlow, unpublished) Plasmids: pET24b Kmr; expression vector Novagen pSJS1240 Spr; E. coli ileX and argU (Kim et al., 1998) pJAM202 Apr; Nvr; H. volcanii-E. coli shuttle expression (Kaczowka and plasmid with psmB-his6 gene; β subunit of 20S Maupin-Furlow, proteasomes expressed with C-terminal His tag in H. 2003) volcanii pBAP5010 Apr; Nvr; H. volcanii-E. coli shuttle-expression (Jolley et al., 1997) vector pJAM650 Apr; Nvr; 1.2-kb NdeI-HindIII fragment of pJAM642 J. Maupin-Furlow blunt end ligated with a 9.94-kb NdeI-BlpI fragment (Reuter et al., 2004) of pJAM202; H. volcanii-E. coli shuttle expression plasmid with panA-his6 gene; PanA expressed with C-terminal His tag in H. volcanii pJAM642 Kmr; 1.2-kb NdeI-to-HindIII fragment of pJAM638 J. Maupin-Furlow ligated into NdeI and HindIII sites of pET24b; PanA- (Reuter et al., 2004) His expressed in E. coli pJAM643 Kmr; 1.1-kb NdeI-to-HindIII fragment of pJAM639 J. Maupin-Furlow ligated into NdeI and HindIII sites of pET24b; (Reuter et al., 2004) PanA(∆1-40)-His expressed in E. coli pJAM1006 Kmr; 1.24-kb fragment PCR amplified from H. This study volcanii genomic DNA ligated into NdeI and XhoI sites of pET24b; PanB-His expressed in E. coli

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Table 4-1. Continued Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification pJAM1012 Apr; Nvr; 1.24-kb NdeI-XhoI fragmentof pJAM1006 This study blunt end ligated with a 9.94-kb NdeI-BlpI fragment (Fig. 4-4) of pJAM202; H. volcanii-E. coli shuttle expression plasmid with panB-his6 gene; PanB expressed with C-terminal His tag in H. Volcanii pJAM1022 Apr; pET22b-derived plasmid expressing smRSGFP- (Reuter and Maupin- SsrA in E. coli Furlow, 2004) pMCSG7 Apr; modified pET-30 Xa/LIC vector with inserted M. Donnelly tobacco etch virus (TEV) protease cleavage site (Stols et al., 2002) encoding region; Allows ligation-independent cloning (LIC) and expression of proteins from E. coli with cleavable N-terminal 6His-tags pJAM1060 Apr; 1.25-kb SspI-panB-BlpI-SspI PCR fragment This study generated (5’-CGGAATATT ATGTCACGCAGTC CATCTCTC-‘3 and 5’-GGAATATTGCTAAGCT CAGTACTGGTAGTCCGTG-‘3; SspI and BlpI sites in bold, BlpI italicized) from pJAM1006 blunt-end ligated into SspI digested pMCSG7; PanB with a cleavable N-terminal 6His-tag expressed in E. coli pJAM1059 Apr; Nvr; 1.32-kb NdeI-BlpI fragment of pJAM1060 This study ligated with a 9.94-kb NdeI-BlpI fragment of pJAM202; H. volcanii-E. coli shuttle expression plasmid with his6-tev-panB gene; PanB with a cleavable N-terminal 6His-tag expressed in H. volcanii pJAM1065 Apr; Nvr; 0.73-kb BamHI-BlpI fragment generated by This study PCR (5’-AAGGATCCATGAGTAAAGGAGAAG- ‘3 and 5’-GAGCTCAGCTTTTAGGCTTTG TATAG-‘3; BamHI and BlpI in bold) from pJAM1022 ligated into a 9.8-kb BamHI-BlpI fragment from pJAM202; smRS-GFP with a C- terminal Ala addition expressed in H. volcanii pJAM1083 Apr; Nvr; 0.11-kb BamHI-NcoI fragment generated This study by PCR (5’-GCTGACGAACTCTGAACCTATGA- ‘3 and 5’-CATAACCATGGATTGGAAGTA CAGG-‘3; NcoI site in bold) ligated into 10.45kb BamHI-NcoI fragment of pJAM1065; designed for simple cohesive end ligation (NcoI-BlpI) into the H. volcanii-E. coli shuttle expression plasmid with direct upstream 6His-TEV site coding region; Protein expression in H. volcanii with a cleavable N- terminal 6His-tag

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Table 4-1. Continued Strains and Phenotype or Genotype; oligonucleotides for PCR Source Plasmids amplification pJAM1084 Apr; Nvr; 1.23-kb NcoI-BlpI fragment generated by This study PCR (5’-CATACCATGGACATGATGACCGATA CTGTGG-‘3 and 5’-CAGCTCAGCTTACGCG AACGC-‘3; NcoI and BlpI sites in bold) ligated into a 10-kb fragment of pJAM1083; PanA expressed with a cleavable N-terminal 6His-tag in H. volcanii pJAM1085 Apr; Nvr; 1.24-kb NcoI-BlpI fragment generated by This study PCR (5’-GTAGCCATGGAGATGTCACGCAG TCC-‘3 and 5’-CTGCTCAGCTCAGTACTGG TAGTCCG-‘3; NcoI and BlpI sites in bold) ligated into a 10-kb fragment of pJAM1083; PanB expressed with a cleavable N-terminal 6His-tag in H. volcanii pJAM1086 Kmr; 1.3-kb NdeI-BlpI fragment of pJAM1084 This study ligated into a 5.1-kb NdeI-BlpI fragment of pET24b; PanA expressed with a cleavable N-terminal 6His-tag in H. volcanii

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Fig. 4-1. Amino acid sequence alignment of H. volcanii PanA and PanB. Identical residues are shaded in black. Functionally conserved and semi-conserved amino acid residues are shaded in grey. Dashes indicate gaps introduced in protein sequence alignment. Boxed residues are predicted with a >90% probability to form a coiled-coil conformation (see Materials and Methods). Consensus sequences of the Walker A and B boxes of the P-loop nucleotidase core are indicated below the alignment as G X2GXGKT and DEXD, respectively (where X is any amino acid residue). The AAA ATPase second region of homology or SHR motif [(T/S)-(N/S)-X5-DXA-X2-R-X2-RX-(D/E)] is also indicated. The N-terminal sequences of the PanA-His and PanA ∆1-40- His antigens were identical to residues 2 – 14 and 41 – 51 of the deduced primary sequence, respectively. MALDI-TOF Q-STAR detected the mass spectra of 11 tryptic fragments of the PanB-His antigen, which encompassed 36% of the primary amino acid sequence. The masses (Da) and corresponding residue numbers of PanB-His were as follows: 716.3148, 30- 34; 749.3711, 35-40; 1004.5522, 330-337; 1264.6408, 205-261; 1307.7720, 317-328; 1490.6807, 379-392; 1548.7975, 301-313; 1836.7691, 275-289; 2180.0845, 118-137; 2408.1564, 4-24; 2739.3637, 138-162.

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1 2 3 4 kDa 66

45 Anti-PanA Anti-PanB

Fig. 4-2. PanA and PanB are produced in H. volcanii. Cell lysate of stationary phase H. volcanii DS70 (10 µg) (lanes 2 and 4) as well as PanA-His (45 ng) (lane 1) and PanB-His (20 ng) (lane 3) purified from recombinant E. coli were separated by 7.5 % SDS-PAGE. Protein was analyzed by Western blot using anti-PanA (lanes 1 and 2) and anti-PanB (lanes 3 and 4) antibodies. Molecular mass standards are indicated to the right. Preimmune serum was included as a negative control (data not shown).

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Fig. 4-3. Plasmid map of pJAM1012. This demonstrates an example of an E. coli-H. volcanii shuttle vector utilizing a rRNA promoter from Halobacterium cutirubrum (HaP) and a T7 terminator (T7) to express genes recombinantly, in this case PanB-His, in H. volcanii. This plasmid also features an ampicillin resistance cassette for expression in E. coli (AmpR) and a novobiocin resistance cassette for expression in H. volcanii (NovR).

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Fig. 4-4. PanA and PanB associate in vivo in H. volcanii. IMAC purified PanA-His and PanB-His from E. coli and H. volcanii separated by 7.5% SDS-PAGE and visualized by Coomassie Blue staining (CB stain), or electro-blotted to PVDF membrane and analyzed by Western blot using either rabbit raised anti-PanA or anti-PanB primary antibody and goat raised, AP-conjugated anti-rabbit secondary antibody. Lane designations are as follows; m) marker a) PanA- His expressed from E.coli BL21 (DE3) pJAM642 b) PanB-His expressed from E.coli BL21(DE3) pJAM1006, a*) PanA-His expressed from H. volcanii DS70 pJAM650, b*) PanB-His expressed from H. volcanii DS70 pJAM1012. A) CB stained SDS-PAGE and Western blot analysis of initial IMAC purification of PanB-His from H. volcanii DS70 pJAM1012. Protein amounts loaded are as follows; b (70 ng), b* (0.4 µg) B) CB stained SDS-PAGE and Western blot analysis of PanA-His and PanB-His from E. coli and H. volcanii. Protein amounts loaded are as follows; a (1.1 µg), b (0.9 µg), a* (1.5 µg), b* (1.6 µg)

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Fig. 4-5. Gel filtration chromatographs of PanA-His and PanB-His proteins purified by IMAC and hydroxyapatite ionic exchange chromatography from recombinant H. volcanii. The combinations of strains and plasmids used are GG102 pJAM1012 (A), GG101 pJAM650 (B), DS70 pJAM1012 (C), and DS70 pJAM650 (D). 0.8 ml samples were applied to each run containing protein concentrations of 1.0, 0.8, 0.6, and 1.7 mg/ml, respectively. Fraction numbers are indicated on the x axes and the protein elution profile is indicated on the y axes by A280 nm. Peaks corresponding to proposed dodecamers (DD), hexamers (H), dimers (D), and monomers (M) are indicated and based upon molecular masses of 45.6 and 45.4 kDa for PanA and PanB respectively.

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Fig. 4-6. Purified Pan complexes from H. volcanii. Pan proteins were purified from H. volcanii, separated by SDS-PAGE gel (12%) and visualized by CB stain. Purification scheme included IMAC, hydroxyapatite, and gel filtration chromatography and were performed as described in methods. Lane designated M is a marker with indicated molecular weight standards. Lanes 1-4 are PanA-His or PanB-His purified from GG102 pJAM1012 (0.30 µg) (Lane 1), DS70 pJAM1012 (0.43 µg) (Lane 2), GG101 pJAM650 (0.4 µg) (Lane 3), and DS70 pJAM650 (0.4 µg) (Lane 4),

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Fig. 4-7. ATP-hydrolyzing activity of PanA-His purified from H. volcanii GG101 (pJAM650). Reaction conditions included 0.0052 mg·ml-1 PanA-His, 10 mM MgCl2, 1 mM ATP and unless otherwise indicated 100 mM Tris (open shapes) or MOPS (filled shapes) buffer pH 7.5, 2 M KCl (triangles) or NaCl (squares) incubated at 45°C. The y-axes are in specific ATP-hydrolyzing activity (µmol Pi·min-1·mg-1 protein). A) Reactions incubated at temperatures ranging from 20-75°C. B) Reactions performed using NaCl or KCl concentrations ranging from 0.5 to 3.5 M. C) Reactions buffered at a pH ranging from 6.3 to 10.3. Alternative buffers included 100 mM K-MES (x) or CAPSO (+).

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Fig. 4-8. SmRS-GFP-SsrA purification from recombinant E. coli. A. SDS-PAGE (12%) analysis of steps in the purification of smRS-GFP-SsrA including a marker (M), soluble lysate (L) (2.14 µg), IMAC fraction (IMAC) (3.65 µg), Hydroxyapatite fraction (H) (2.65 µg), and Superose 6 fraction (S6) (0.45 µg). B. Fraction S6 (smRS-GFP-SsrA in Buffer B) visualized using a Leica MZ75 dissecting microscope (excitation 480/40 nm and emission 510 nm).

CHAPTER 5 SUMMARY AND CONCLUSIONS

The purpose of this study was to gain a better understanding of energy-dependent proteolysis in archaea, especially in relation to the determinants of substrate recognition.

Among archaea, Haloferax volcanii was selected for use in this study due to its established genetic system, ease of culture, and synthesis of multiple 20S proteasome isoforms and Pan proteins. The results of this study offer a means by which to study energy-dependent proteolysis in haloarchaea using GFP variants and provide insight into the biochemical and physical properties of multiple Pan regulatory particles from H. volcanii.

This study demonstrates the production of an active GFP protein in an archaeon for its use as an in vivo reporter of proteolysis. Initially, E-GFP with a C-terminal His- tag was expressed and IMAC purified from recombinant H. volcanii. E-GFP containing

IMAC fractions displayed significant fluorescence in high salt buffer (2 M NaCl, 500 mM Imidazole, 20 mM Tris, pH 7.2). Despite its activity upon partial purification, fluorescence could not be detected in H. volcanii whole cells, presumably due to low cellular protein levels. Protein insolubility being suspect as the cause of its low abundance led to an attempt to express several soluble modified GFP proteins (smGFP, smBFP, and smRS-GFP) in H. volcanii. Subsequent Western blot analysis compared the soluble cell lysates of H. volcanii individually expressing E-GFP to those expressing each one of the smGFP proteins individually. Densitometry of the corresponding GFP bands

123 124 estimated the protein levels of the smGFP proteins to be about 10-fold greater than those of E-GFP.

H. volcanii expressing smRS-GFP displayed a uniform cell fluorescence (ex. 488 nm, em. 509 nm) in contrast to H. volcanii alone which displayed no visual fluorescence.

It has yet to be determined why H. volcanii expressing smGFP and smBFP does not display significant fluorescence compared to H. volcanii alone, but may be due to the increased innate cell fluorescence of H. volcanii at the shorter wavelengths of light used to stimulate fluorescence of these proteins (ex. 397 nm, em. 507 and ex. 385, em. 448, respectively). A growth curve of H. volcanii expressing smRS-GFP was compared to that of H. volcanii alone including time points that measured optical density as well as fluorescence. Both demonstrated a similar growth pattern, however, in contrast, H. volcanii expressing smRS-GFP displayed a concomitant rise in fluorescence during log phase and a plateau of fluorescence throughout stationary phase.

The apparent stability in the protein level of smRS-GFP allowed its utilization as a reporter protein. In this regard, a multitude of C-terminal additions, ranging from single amino acids to the eleven-residue SsrA motif, were added to smRS-GFP in an attempt to distinguish their effect on its cellular protein level. Protein levels of these smRS-GFP variants were measured by Western blot analysis or estimated by fluorescence readings throughout growth. The most severe effect resulted from the addition of the SsrA motif (AANDENYALAA) (smRS-GFP-SsrA), which altered smRS-

GFP intracellular protein levels below the point of detection by Western blot. This modification did not significantly alter its respective mRNA levels as they were comparable to those of unaltered smRS-GFP.

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It was determined by Western blot analysis that 20S proteasomes were at least

partially responsible for the degradation of smRS-GFP-SsrA by the addition of an

irreversible proteasome specific inhibitor, clasto-lactacystin β-lactone (clβl) to cell

cultures of H. volcanii expressing this protein. A similar study using the serine protease

inhibitor PMSF in place of clβl displayed no increase in smRS-GFP-SsrA levels,

indicating that cellular serine proteases do not degrade this substrate to a significant

extent. In vitro inhibition assays of 20S proteasomes purified from H. volcanii

demonstrated that clβl was a highly specific inhibitor having a Ki of about 40 nM.

Multiple mutations and truncations of the SsrA motif partially restored smRS-

GFP cellular levels from about 0.5% to about 30% of unaltered smRS-GFP levels,

presumably by suppressing their proteolytic recognition. Most notably, the restoration of

smRS-GFP proteins levels to about 30% of unaltered smRS-GFP resulted from the

addition of a sequence equal in length to that of SsrA although which included four

contiguous charged residues and replaced three of the five Ala residues

(AANDDKDLSNN). The addition of Ala residues to the C-terminus of smRS-GFP

appears partially responsible for fluctuations in protein levels. In fact, a single Ala

addition reduced smRS-GFP protein levels by about 90%. Surprisingly, the addition of

one of any of the 20 essential amino acids to the C-terminus of smRS-GFP negatively

affected the fluorescence of H. volcanii cultures expressing these reporter proteins,

although to varying degrees. Single residue additions to smRS-GFP that had the least impact on culture fluorescence compared to cultures expressing unaltered smRS-GFP were typically charged residues (Asp, Glu, and Lys) while additions having the most profound impact were usually hydrophobic amino acids (Ala, Ile, Val). It has yet to be

126 determined what factors are responsible for the variable fluorescence of H. volcanii cultures expressing each smRS-GFP reporter protein. It is possible that these proteins are differentially regulated via proteolysis by Pan/20S proteasomes and/or other cellular proteases of H. volcanii.

This study also represents the first attempt at Pan characterization from an organism which contains multiple Pan paralogs (PanA and PanB). PanA and PanB protein sequences were determined to be closely related with 60% identical and 78% similar amino acid residues. The N-terminal domains of PanA and PanB are predicted to adopt a coil-coiled conformation and in contrast to the central domain are not highly related (17% identity). Likewise, the C-terminal domains are also less similar than the central domains (49% identical). This divergence in the primary sequence of the N- and

C-termini, combined with the highly conserved central domain, suggests that PanA and

PanB mediate similar yet distinct functions in H. volcanii. It is possible that the PanA and

PanB proteins of H. volcanii recognize different substrate proteins and/or differ in their affinity for α1β and α1α2β 20S proteasome subtypes.

This study determined that H. volcanii forms at least three Pan complexes in vivo; two homo-oligomers consisting of entirely of PanA or PanB, and a hetero-oliogmer consisting of both proteins. Gel filtration elution profiles of these complexes suggest they form hexamers (about 270 kDa) and dodecamers (about 650 kDa), which may be in the form of a dimer of hexamers. The ATP-hydrolyzing activity of the 650 kDa homo- oligomeric PanA complex was analyzed in order to determine the optimal conditions for this enzyme. PanA-His hydrolyzed ATP to an optimal extent in 2.5 M KCl at 45 to 50°C in MOPS, buffered at a pH of 7.5 to 8.5. These data are very consistent with the growth

127

conditions of H. volcanii, which are neutrophilic halophiles whose optimal growth

temperature is around 42 to 45°C. These conditions also correlate well to the previously determined optimal peptide-hydrolyzing conditions of H. volcanii 20S proteasomes in terms of salt concentration and pH. The temperature preference of H. volcanii 20S proteasomes is slightly higher (60 to 65°C) than that of PanA. However, non-optimal proteasome activity at a lower temperature is not predicted to limit proteolysis, as several groups have suggested that the rate limiting step in protein degradation by ATPase- associated proteases, including Pan/20S proteasomes, is prior to peptide bond cleavage.

In addition H. volcanii grows in regions where temperatures rarely exceed 45°C, and therefore the optimal peptide-hydrolyzing activity is most likely an artifact of proteasomal evolution. Substrate degradation by H. volcanii Pan/20S proteasomes in vitro has yet to be demonstrated despite using several putative substrate proteins, including smRS-GFP-SsrA purified from recombinant E. coli. It is possible that the C- terminal His-tag of the Pan proteins is interfering with 20S proteasome association or α- ring gate-opening.

Future research will focus on elucidating the responsible factors of the variable protein levels of smRS-GFP reporter proteins in H. volcanii. Current efforts are underway to construct Pan proteins with removable N-terminal His-tags by way of a TEV protease cleavage site and α-subunits with gate deletions. These efforts may allow for Pan/20S proteasome degradation of substrates in vitro. The different Pan isoforms can be used in these assays in an attempt to distinguish possible differences in their ability to recognize substrates of 20S proteasomes from H. volcanii. These assays would also be useful in providing further evidence for putative substrate proteins of Pan/20S proteasomes, of

128 which a list is currently being compiled by proteome analysis of H. volcanii cultures deleted of Pan subunits or under various conditions such as in the presence of clβl.

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BIOGRAPHICAL SKETCH

Christopher John Reuter was born on April 16, 1978, in Belleville, Illinois, to parents Gregory and Debra Reuter. He has one younger sister, Sarah. In May 2001, he graduated with a Bachelor of Science degree in microbiology and cell science from The

University of Florida. During the summer of 2001, Christopher joined the lab of Dr. Julie

Maupin-Furlow and soon after, in August 2001, he was admitted to the graduate program of the Department of Microbiology and Cell Science at the University of Florida where he continues working toward completing his Ph.D. degree. In December 2002 he was married to his high school sweetheart, now Jennifer Reuter, who is currently carrying their first child.

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