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Contributions of the Individual b Subunits to the Function of the Peripheral Stalk of F1F0 ATP Synthase


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CONTRIBUTIONS OF THE INDIVIDUAL b SUBUNITS TO THE FUNCTION OF THE PERIPHERAL STALK OF F 1 F 0 ATP SYNTHASE By TAMMY WENG BOHANNON GRABAR A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2004

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Copyright 2004 by Tammy Weng Bohannon Grabar

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This document is dedicated to my husband, Chuck, and my daughter, Kaia.

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ACKNOWLEDGMENTS The work illustrated in this dissertation and my growth as a scientist could not have been accomplished without the guidance, encouragement and support of several people on both the professional and personal levels. The first person I would like to thank is my mentor, Dr. Brian Cain. He allowed me to join his laboratory when I was fresh out of college, even though I had no real experiences in a scientific lab. He exhibited extreme patience while teaching me everything from how to hold and operate a pipette to pouring agarose gels to cloning my own plasmids. His evident passion and excitement about science opened my eyes to a whole new world of opportunity. Prior to joining his laboratory, I had never even dreamed of joining graduate school and pursuing a PhD; therefore, I feel extreme gratitude and consider myself very fortunate to have joined his lab. Once I joined the lab, Dr. Cain allowed me the freedom to make my own initial scientific and experimental decisions, which was an excellent teaching method for me, but he was always there for guidance and support whenever it was needed. I would also like to thank him for being so involved in his lab. On any given day, I knew I could have his undivided attention if I needed to consult with him. Over the years Dr. Cain has spent a tremendous amount of time teaching me to think critically about scientific experiments, how to communicate my data and ideas to others, how to give a professional scientific presentation, and how to write scientific papers, and for those countless hours I thank him. iv

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I would also like to thank every person on my committee: Dr. Linda Bloom, Dr. Art Edison and Dr. Dan Purich from my department, and Dr. Julie Maupin-Furlow from the Microbiology and Cell Sciences Department. I have known Dr. Maupin-Furlow the longest. She taught one of the most challenging courses I took as an undergraduate. When an unexpected death occurred in my family, she was kind enough to allow me to postpone an exam without questioning my motive, which was very unusual for most of my professors while I was an undergraduate. Thanks to her, that was the only class in which my grade was not affected that semester. I would also like to thank her for her continual support in helping me get into graduate school and then subsequently taking the time to hike across campus to join my committee meetings. I would like to thank Dr. Purich for teaching me, in the middle of a physical biochemistry class, that sometimes we have to take some time off to go outside and get some fresh air. That was always an important lesson when endless hours in the lab led to careless mistakes. I also enjoyed his sometimes unusual stories and adventures that he had to share with me when I was spending entire days in the biochemistry library studying for exams. I would like to thank Dr. Edison for his constant support and encouragement. He has always been the first person who publicly and very kindly commended me after my journal club presentation. I believe his kind words of support through the years helped me to gain the courage I needed to believe in myself to really deliver a good presentation. I would also like to thank him for his eagerness to understand every aspect about my project. And last, but not least, I would like to thank Dr. Bloom. As a woman in my department and a new mother with a career in academia, she has been a wonderful role model. She has always had kind words and smiles to bestow on me. I would also like to thank her for v

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assisting me during my committee meetings when discussions of fluorescence started to go over my head. I would also like to take the time to thank everyone that I had the pleasure of working with in the lab. These are the people I spent countless hours with during the course of the day and held many scientific and personal conversations with, and I am happy to be able to call them my friends. I could not have spent the last five years with a better group of people. Drs. Tammy Otto and Debra Zies were members of the lab when I first joined and were the ones who taught me the ways of the lab. Dr. Michelle Gumz joined the graduate program and subsequently joined Dr. Cains lab the same time as I. I would like to thank Tammy, Debbie and Michelle for their scientific and personal support as well as sharing with me memories of pool barbeques, wedding showers and baby showers. Dr. Deepa Bhatt joined the lab as a postdoc during my graduate career. Her friendship and scientific guidance have been very valuable to me. I would like to thank her for giving me fantastic advice on all of my oral presentations and reviewing all of my papers. And finally, I would like to thank my family for all the encouragement, love and support they have unwaveringly offered over the years. I would like to thank my dad for always finding the positive in everything that was negative and always encouraging me to overcome the many obstacles that graduate school hurled towards me. He never lost faith in me, even when I was ready to give up. I would like to thank my mom for her tremendous support as well. She spent a lot of time and energy stocking my refrigerator and freezer full of meals when I found that I did not have the time to care for myself. She has spent many weeks and weekends at my home since my daughter was born so that I vi

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could have extra time to work on my dissertation. And last, but not least, I would like to thank my husband for his constant emotional support and belief in me. He has been with me through the thick and thin of graduate school and never once complained of my emotional torment when things were not going my way. I could not have accomplished this without him. vii

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TABLE OF CONTENTS Page ACKNOWLEDGMENTS.................................................................................................iv LIST OF TABLES............................................................................................................xii LIST OF FIGURES.........................................................................................................xiii ABBREVIATIONS.........................................................................................................xvi ABSTRACT.......................................................................................................................xx CHAPTER 1 BACKGROUND AND SIGNIFICANCE....................................................................1 Introduction...................................................................................................................1 Structure and Function of F 1 F 0 ATP Synthase.............................................................3 The Catalytic Core.................................................................................................6 The hexamer.............................................................................................8 The subunit................................................................................................12 The Rotor Stalk....................................................................................................12 The subunit................................................................................................14 The subunit................................................................................................16 The ring of c subunits...................................................................................21 The Stator Stalk...................................................................................................25 The a subunit................................................................................................27 The subunit................................................................................................33 The b subunit................................................................................................37 Subunit Equivalence...................................................................................................51 F 1 F 0 ATP Synthase Mechanism..................................................................................56 Proton Translocation: Driving Rotation..............................................................57 Coupling..............................................................................................................60 Catalysis: The Binding Change Mechanism.......................................................62 Genetic Expression and Assembly.............................................................................63 Summary.....................................................................................................................65 viii

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2 INTEGRATION OF UNEQUAL LENGTH b SUBUNITS INTO F 1 F 0 ATP SYNTHASE...............................................................................................................67 Introduction.................................................................................................................67 Materials and Methods...............................................................................................69 Materials..............................................................................................................69 Strains and Media................................................................................................70 Recombinant DNA Techniques...........................................................................70 Mutagenesis and Strain Construction..................................................................75 Crude Preparative Procedures.............................................................................78 Determination of Protein Concentration.............................................................80 Ni-Resin Purification...........................................................................................80 Assays of F 1 F 0 ATP Synthase Activity...............................................................82 Immunoblot Analysis..........................................................................................85 Results.........................................................................................................................88 HA-Epitope Tagged b Subunits...........................................................................88 Construction and growth characteristics of mutants....................................88 Effects of epitope tags..................................................................................90 Expression of different b subunits in the same cell......................................91 Ni-Resin Purification....................................................................................92 V5-Epitope Tagged b Subunits...........................................................................97 Construction and growth characteristics of mutants....................................97 Effects of epitope tags................................................................................100 Detections of heterodimers.........................................................................103 Formation of mixed length b subunits in F 1 F 0 ATP synthase...........................107 Discussion.................................................................................................................111 3 GENETIC COMPLEMENTATION BETWEEN MUTANT b SUBUNITS IN F 1 F 0 ATP SYNTHASE.....................................................................................................114 Introduction...............................................................................................................114 Materials and Methods.............................................................................................116 Materials............................................................................................................116 Strains and Media..............................................................................................117 Recombinant DNA Techniques.........................................................................117 Mutagenesis and Strain Construction................................................................118 Preparative Procedures......................................................................................120 Immunoblot Analysis........................................................................................120 Assays of F 1 F 0 ATP Synthase Activity.............................................................121 Results.......................................................................................................................121 Construction and Growth Characteristics of Mutants.......................................121 Heterodimer Formation of b arg36 Defective Subunits with b wt ..........................123 Heterodimer formation of b 153end-his Complemented with b wt-V5 ...................128 Heterodimer Formation of b +124-130-his Complemented with b wt-V5 ....................131 ix

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Mutual Complementation..................................................................................135 Discussion.................................................................................................................139 4 DEVELOPMENT OF CYSTEINE CHEMICAL MODIFICATIONS OF ALTERED b SUBUNITS............................................................................................................143 Introduction...............................................................................................................143 Materials and Methods.............................................................................................146 Materials............................................................................................................146 Strains and Media..............................................................................................146 Recombinant DNA Techniques.........................................................................147 Mutagenesis and Strain Construction................................................................148 Crude Preparative Procedures...........................................................................152 Assays of F 1 F 0 ATP Synthase Activity.............................................................152 Immunoblot Analysis........................................................................................153 Results.......................................................................................................................153 Construction and Growth Characteristics of Mutants.......................................153 Effects of Cysteine Mutations...........................................................................157 Discussion.................................................................................................................158 5 MUTAGENISIS OF THE AMINO AND CARBOXYL TERMINI OF THE b SUBUNIT IN F 1 F 0 ATP SYNTHASE.....................................................................161 Introduction...............................................................................................................161 Materials and Methods.............................................................................................165 Materials............................................................................................................165 Strains and Media..............................................................................................165 Recombinant DNA Techniques.........................................................................166 Mutagenesis and Strain Construction................................................................166 Crude Preparative Procedures...........................................................................168 Assays of F 1 F 0 ATP Synthase Activity.............................................................169 Results.......................................................................................................................169 Amino Terminal Mutations...............................................................................169 Construction and growth characteristics of mutants..................................169 Effects of amino terminal mutations..........................................................171 Carboxyl Terminal Mutations...........................................................................173 Construction and growth characteristics of mutants..................................173 Effects of carboxyl terminal mutation........................................................175 Discussion.................................................................................................................176 6 CONCLUSIONS AND FUTURE DIRECTIONS...................................................180 Conclusions...............................................................................................................180 Integration of Unequal Length b Subunits into F 1 F 0 ATP Synthase.................181 x

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Genetic Complementation between Mutant b Subunits in F 1 F 0 ATP synthase..........................................................................................................183 Development of Cysteine Chemical Modifications of Altered b Subunits.......185 Mutagenesis of the Amino and Carboxyl Termini of the b subunit in F 1 F 0 ATP Synthase.................................................................................................186 Future Directions......................................................................................................188 Complementing Mutant b Subunits...................................................................189 Function of F 1 F 0 ATP Synthase Incorporated with b Subunit Heterodimers....190 Positions of the Individual b Subunits in F 1 F 0 ATP Synthase...........................190 Length of the Peripheral Stalk in F 1 F 0 ATP Synthase Complexes Incorporated with Shortened and Lengthened b Subunits.............................191 Other Implications....................................................................................................195 APPENDIX A MUTAGENIC OLIGONULCEOTIDES.................................................................202 B DEVELOPING PROTOCOL FOR PURIFYING F 1 F 0 ATP SYNTHASE.............206 Purification of Enzyme Complexes Incorporated with b Subunit Heterodimers.....206 Culture...............................................................................................................206 Disruption of Bacteria.......................................................................................207 Ni-Resin Purification.........................................................................................209 V5-Epitope Iimmunoprecipitation.....................................................................210 Detection of Purified Enzyme Complexes........................................................210 Assays of F 1 F 0 ATP Synthase Activity.............................................................211 LIST OF REFERENCES.................................................................................................212 BIOGRAPHICAL SKETCH...........................................................................................237 xi

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LIST OF TABLES Table page 1-1. F 1 F 0 ATP synthase subunit equivalency....................................................................52 2-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing epitope tagged uncF(b) genes...................................................90 3-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing epitope tagged uncF(b) genes............................................122 4-1. Description of uncF(b) cysteine mutations.............................................................149 4-2. Description of the unc operon cysteine mutations 1 .................................................154 4-3. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing cysteine...............................................................................156 5-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing uncF(b) mutations at the amino terminus...........................170 5-2. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing uncF(b) insertions or deletions throughout the b subunit..175 6-1. Preliminary data of coexpressed mutant b subunits................................................190 A-1. Oligonucleotide sequences.....................................................................................203 A-2. Oligonucleotide description....................................................................................204 xii

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LIST OF FIGURES Figure page 1-1. Timeline of developing views of F 1 F 0 ATP synthase...................................................5 1-2. Space-filling structural model of Escherichia coli F 1 F 0 ATP synthase........................7 1-3. Structure of the subunit of E. coli F 1 F 0 ATP synthase.............................................19 1-4. Controversial models of the a subunit topology.........................................................29 1-5. Amino acid sequence of the E. coli F 1 F 0 ATP synthase b subunit.............................38 1-6. Gross structure of the E. coli F 1 F 0 ATP synthase and the domains of the b subunit......................................................................................................................40 1-7. Model for F 1 F 0 ATP synthase peripheral stalk orientation dependent upon the direction of rotation during ATP synthesis or hydrolysis........................................47 1-8. Speculative models for the b-like subunits.................................................................55 1-9. Model of proton translocation and torque generation in F 0 ........................................59 1-10. The binding change mechanism...............................................................................63 2-1. Oligonucleotides for epitope tags and mutagenesis of uncF(b).................................74 2-2. Construction of the single transcript expression system............................................77 2-3. Histidine and HA-epitope-tagged b subunit expression system.................................89 2-4. Western blot analysis of histidine and HA-epitope tagged b subunits.......................92 2-5. Investigation of detergent solubilization of F 1 F 0 ATP synthase complexes..............94 2-6. Ni-resin purification of F 1 F 0 expressing different length b subunits, treated with the cross-linker BS 3 ..................................................................................................95 2-7. Ni-resin purification of histidine and HA-epitope tagged F 1 F 0 treated with the cross-linker BS 3 ........................................................................................................96 2-8. Histidine and V5-epitope-tagged b subunit expression system..................................98 xiii

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2-9. ATP-driven energization of membrane vesicles prepared from uncF(b) gene mutants...................................................................................................................101 2-10. NADH-driven acidification of membrane vesicles prepared from uncF(b) mutants...................................................................................................................102 2-11. Ni-resin purification of F 1 F 0 ATP synthase treated with the cross-linker BS 3 .......106 2-12. Ni-resin purification of F 1 F 0 ATP synthase expressing unequal length b subunits...................................................................................................................108 2-13. Quantitation of b subunit heterodimeric F 1 F 0 .........................................................110 2-14. Interactions of b subunits of unequal lengths.........................................................112 3-1. Oligonucleotides for epitope tags and C-terminal truncation of uncF(b)................119 3-2. Ni-resin purification of F 1 F 0 ATP synthase incorporated with b arg36 subunit mutations................................................................................................................124 3-3. ATP-driven energization of membrane vesicles prepared from uncF(b) arg36 gene mutants...........................................................................................................127 3-4. Ni-resin purification of F 1 F 0 ATP synthase containing a b subunit carboxylterminal truncation.................................................................................................129 3-5. ATP-driven energization of membrane vesicles incorporated with F 1 F 0 ATP synthase containing a b subunit carboxyl-terminal truncation...............................130 3-6. Ni-resin purification of membranes incorporated with b +124-130-his subunit mutation..................................................................................................................132 3-7. ATP-driven energization of membrane vesicles incorporated with a defective b +124-130 subunit mutation........................................................................................133 3-8. Ni-resin purification of F 1 F 0 ATP synthase incorporated with complementing defective b subunits................................................................................................136 3-9. ATP-driven energization of membrane vesicles incorporated with F 1 F 0 ATP synthase containing complementing defective b subunits.....................................137 3-10. Interactions of defective b subunit with wild type b subunits found in intact F 1 F 0 ATP synthase complexes...............................................................................140 3-11. Model of F 1 F 0 ATP synthase incorporated with complementing defective b subunits ..................................................................................................................142 xiv

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4-1. Model of E. coli F 1 F 0 ATP synthase.........................................................................144 4-2. Oligonucleotides for cysteine mutagenesis of the unc operon.................................150 4-3. Expression plasmid of cysteine mutants...................................................................155 4-4. Western blot analysis of cysteine mutant b subunits of differing length.................158 4-5. Model of F 1 F 0 ATP synthase with cysteine substitutions in the b and subunits..159 5-1 Amino acid sequence and domains of the E. coli b subunit......................................162 5-2. Oligonucleotides for mutagenesis at the amino and carboxyl termini in the unc operon.....................................................................................................................167 5-3. ATP-driven energization of membrane vesicles prepared from b subunit membrane domain mutants....................................................................................172 5-4. Amino acid insertion and deletion analysis of the E. coli b subunit........................174 5-5. Mutations constructed throughout the b subunit......................................................177 6-1. Design of FRET experiments to measure the peripheral stalk.................................193 6-2. Model of rotation inhibition due to a fusion protein on the subunit.....................195 6-3. Sequence alignments of subunits b and b from various species with the b subunit of E. coli....................................................................................................200 B-1. Diagram of purification procedures for homogeneous heterodimeric b V5 /b his F 1 F 0 ATP synthase complexes...............................................................................208 xv

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ABBREVIATIONS ACMA, 9-amino-6-chloro-2-methoxyacridine ADP, adenosine-5-diphosphate ala, alanine AO, tegamineoxide WS-35 Ap, ampicillin Ap r ampicillin resistant asn, asparagine ADP, adenosine-5-diphosphate ATP, adenosine-5-triphosphate b +7-his seven amino acid insertion in the b subunit with a 6X histidine epitope tag at the amino terminus b 7-V5 seven amino acid deletion in the b subunit with a V5 epitope tag at the carboxyl terminus b ser84cys substitution of a cysteine for serine at amino acid position 84 in the b subunit -ME, -mercaptoethanol bp, base pair BS 3 bis(3-sulfo-N-hydroxysuccinimide ester) BCA, bicinchoninic acid BSA, bovine serum albumin Cm, chloramphenicol xvi

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Cm r chloramphenicol resistance cys, cysteine DACM, N-(7-dimethylamino-4-methyl-coumarinyl)-maleimide DCCD, dicyclohexylcarbodiimide 2 the second -helix in the epsilon () subunit ECD, 1-ethyl-3[3-dimethylamino]propyl carbodiimide ECL, enhanced electrochemiluminescence EDTA, ethylenediaminetetraacetic acid FPLC, fast polynucleotide liquid chromatography FRET, fluorescence resonance energy transfer g, gravitational force gln, glutamine glu, glutamate GFP, green fluorescent protein HA, peptide epitope of hemagglutinin protein of human influenza virus ICBR, Interdisciplinary Center for Biotechnology Research IPTG, isopropyl-1-thio--D-galactopyranoside kb, kilobase kD, kilodalton LB, Luria Bertani medium LBG, Luria Bertani media supplemented with 0.2% glucose LDAO, lauryldimethylamine oxide LSB, Laemmli sample buffer mG, milligram xvii

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mL, milliliter MOPS, 3-[N-morpholino]propanesulfonic acid NADH, -nicotineamide adenine dinucleotide, reduced form NFDM, nonfat dry milk Ni-CAM, high capacity nickel chelate affinity matrix NMR, nuclear magnetic resonance spectroscopy PAGE, polyacrylamide gel electrophoresis PBS, phosphate-buffered saline PBST, phosphate-buffered saline supplemented with 0.1% tween20 PCR, polymerase chain reaction P i inorganic phosphate P/O, number of ATPs made per 2 e transferred to oxygen PVDF, polyvinylidene fluoride rms, root mean square ser, serine SDS, sodium dodecyl sulfate (lauryl sulfate) TID, 3-(trifluoromethyl)-3-(m-[ 125 I]iodophenyl)diazirine TBS, tris-buffered saline thr, threonine TTBS, tris-buffered saline supplemented with 0.1% tween20 TD, taurodeoxycholate TE, tris[hydoxymethyl]aminomethane, ethylenediaminetetraacetic acid buffer, pH 8.0 T M tris[hydoxymethyl]aminomethane, magnesium sulfate buffer, pH 7.5 xviii

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Tm, melting temperature of double stranded DNA Tris, tris[hydoxymethyl]aminomethane g, microgram L, microliter V5, epitope found in the P and V proteins of the paramyxovirus, SV5 v/v, volume/volume wt, wild type w/v, weight/volume xix

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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 CONTRIBUTIONS OF THE INDIVIDUAL b SUBUNITS TO THE FUNCTION OF THE PERIPHERAL STALK OF F 1 F 0 ATP SYNTHASE By Tammy Weng Bohannon Grabar August 2004 Chair: Brian D. Cain Major Department: Biochemistry and Molecular Biology The universal molecule of biological energetics is adenosine triphosphate (ATP), and the enzyme involved in providing the majority of cellular ATP is F 1 F 0 ATP synthase. Enzymes in this family utilize the electrochemical gradient of protons across membranes to synthesize ATP from ADP and inorganic phosphate in a coupled reaction. The cytoplasmic F 1 and the membrane-bound F 0 sectors are linked by two stalk structures, the rotor stalk and the peripheral stalk. Proton conduction through the F 0 sector drives the rotation of the rotor stalk within the catalytic core, which is held steadfast by the peripheral stalk. In Escherichia coli, the subunit of F 1 and a parallel homodimer of identical b subunits constitute the peripheral stalk of F 1 F 0 ATP synthase. Work accomplished in this dissertation indicates that the bacterial enzyme does not require two identical b subunits to form the dimer. Two different length b subunits, with a size difference of at least 14 amino acids, were capable of forming the b dimer of an intact F 1 F 0 ATP synthase complex. Also, in work presented in this dissertation, a defective xx

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mutation in one region of the b subunit was overcome by dimer formation with a second b subunit that contained defective mutation in a different region but had a wild-type sequence in the region of the former defective b subunit. This mutual complementation between fully defective b subunits indicated that each of the two b subunits makes a unique contribution to the functions of the peripheral stalk, such that one mutant b subunit is making up for what the other is lacking. Interestingly, the equivalent of the bacterial b subunit in plants exists as two genetically different subunits, and the mammal counterpart exists as at least four subunits. This work suggests that the individual functions of the b subunits may be reflected in the fact that higher organisms evolved to encode multiple b-type subunits. xxi

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CHAPTER 1 BACKGROUND AND SIGNIFICANCE Introduction The premiere of Peter Mitchells chemiosmotic theory in 1961 eventually resulted in the major breakthrough of the characterization of F 1 F 0 ATP synthases. Basically, his theory stated that protons are pumped across energy transducing membranes, thereby creating an electrochemical gradient of protons (1). This proton gradient, also known as the proton-motive force, consists of two components: i) a chemical component due to the concentration gradient of protons and ii) an electrical component, or membrane potential, due to the positive charge of the protons (H + ). As a result, one side of the membrane is more positive than the other. The potential energy of this gradient can then be transduced to chemical energy or utilized to perform work when the protons diffuse back across the membrane from the higher to the lower potential (2). The protons can diffuse across the membrane through specific transmembrane proton conductors, which can synthesize adenosine 5-triphosphate (ATP) or co-transport solutes, and in the case of bacteria drive flagellar rotation. The ability to consume nutrients and convert them to energy is required of all living organisms, from microscopic bacteria to plants to humans. The universal molecule of biological energetics is ATP, and in almost all organisms, the central enzyme involved in providing the majority of cellular ATP is F 1 F 0 ATP synthase (3-6). F 1 F 0 ATP synthases are responsible for the production of ATP in the final step of processes called oxidative phosphorylation and photophosphorylation. They provide the bulk of cellular energy in 1

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2 the majority of eukaryotes and prokaryotes. The synthesis of ATP occurs at a rate of about 100 s -1 which maintains a concentration of about 3 mM ATP in Escherichia coli and greater concentrations in mitochondria and chloroplasts with no noticeable product inhibition (3). In eukaryotes, they are located in the inner mitochondrial membrane, or in the thylakoid membrane of chloroplasts. In most bacteria, F 1 F 0 ATP synthase is located in the cytoplasmic membrane. Enzymes in this family utilize the electrochemical gradient of protons across these membranes in order to synthesize ATP from ADP and inorganic phosphate (P i ) in a coupled reaction. In bacteria, the reaction of ATP synthases can be reversed if the situation of a dissipated electrochemical proton gradient arises. In this case, ATP derived from glycolysis can be hydrolyzed in order to pump protons across the membrane, creating a membrane potential. The membrane potential can then be utilized to drive other cellular processes such as the extrusion of sodium ions, nutrient uptake and flagellar rotation. An explosion of research concerning F 1 F 0 ATP synthase has occurred during the past few decades. In particular, a great deal of knowledge of the enzyme has been solved only in the past decade. A plethora of relatively recent reviews concerning every aspect of F 1 F 0 ATP synthase can be found in the special editions of Journal of Bioenergetics and Biomembranes (volume 32, 2000) and Biochimica et Biophysica Acta (volume 1458, 2000) as well as reviews authored by Noji and Yoshida (2001), Senior et al. (2002), Capaldi et al. (2002) and Weber and Senior (2004). This chapter will provide detailed explanations of what is currently known about F 1 F 0 ATP synthase including the mechanism of the enzyme as a whole as well as structure and functions of individual subunits, equivalence of the bacterial enzyme to its eukaryotic equivalents and genetic

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3 expression and assembly. The research presented in this dissertation primarily concerns the b subunit of the Escherichia coli (E. coli) F 1 F 0 ATP synthase. Hence the b subunit will be discussed extensively later in this chapter. Structure and Function of F 1 F 0 ATP Synthase The structure and function of F 1 F 0 ATP synthases are remarkably similar from bacteria to humans. In E. coli, the simplest form of the enzyme, F 1 F 0 ATP synthase is a complex enzyme composed of twenty-two polypeptides of eight different types with the stoichiometry of 3 3 ab 2 c 10 (Figure 1-2) (6, 7). The deduced molecular size is approximately 530 kDa. The structure of F 1 F 0 ATP synthase in chloroplasts is very similar with the exception that there are two isoforms of the b subunit. On the other hand, the mitochondrial enzyme is more complex, including an extra 7-9 small subunits which are thought to have roles in enzyme regulation (8-10). Discussion of F 1 F 0 ATP synthase is commonly divided into two portions, F 1 and F 0 The F 1 portion of the enzyme is composed of the cytoplasmic subunits, 3 3 and is responsible for the synthesis of ATP. The F 0 portion consists of the membrane-bound subunits, ab 2 c 10 and is responsible for the translocation of protons through the membrane. New insights concerning the functions and the intersubunit contacts have refined the way F 1 F 0 ATP synthase is perceived, dividing discussions of the enzyme into the catalytic core, the rotor (central) stalk and the stator (peripheral) stalk (3, 11-14). The catalytic core consists primarily of the 3 3 subunits, the rotor stalk consists of the c 10 subunits and the stator stalk consists of the b 2 subunits. Over the years, F 1 F 0 ATP synthase has received worldwide recognition as the tiniest rotary motor known to mankind (15, 16). Protons passing through the enzyme

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4 complex drive the rotation of the rotor c 10 subunits at about 100 Hz. This rotation, which is absolutely essential for the machinery of the enzyme, transmits energy over a distance greater than 100 by providing the means by which conformational changes in the F 1 catalytic core, 3 3 take place for the synthesis of ATP (3, 4). Structural studies of F 1 F 0 ATP synthase commenced in the early 1960s and persist to this day in pursuit of a complete high-resolution structure. Negative staining procedures in the early 1960s initially revealed the traditional tripartite features of the enzyme complex from sub-mitochondrial particles, consisting of what was referred to as the headpiece, stalk and basepiece (Figure 1-1) (17). Ten years later, the first electron micrograph of a detergent-solubilized F 1 F 0 ATP synthase was published, confirming the existing idea of a tripartite molecule (18). Appreciably, electron microscopy (EM) in combination with other biochemical data of isolated F 1 exposed a hexagonal arrangement of alternating subunits with a seventh mass found in the center of the array (Figure 1-1) (7, 19). Based on this premise it was first suggested that F 1 consisted of an alternating hexagonal array of three and three subunits with the and situated centrally (7). The idea did not gain favorable recognition for some twenty years until verified by x-ray crystallography (20, 21). Continuous improvements in EM technology led to numerous publications of various ATP synthases which defined the average overall dimensions of about 190 from top to bottom and about 37 assigned to the stalk structure (22-25). Using a combination of traditional biochemical, molecular biological and immunological techniques along with EM, many important discoveries were made that led to what we now understand of F 1 F 0 ATP synthase. The first direct evidence for rotation of the stalk appeared in 1990 (24), but was not followed by the visualization of the peripheral stalk,

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5 Figure 1-1. Timeline of developing views of F 1 F 0 ATP synthase. Electron microscopy and biochemical analysis from the early 1970s through the 1980s allowed visualization of the classical tripartite features of F 1 F 0 ATP synthase consisting of what was referred to as the headpiece, stalk and basepiece. Furthermore, the arrangement of the F 1 subunits were first proposed in 1974, though it did not gain favor until high resolution structure was obtained twenty years later. The first direct evidence for rotational catalysis appeared in 1990. Improved EM techniques showed the existence of a peripheral stalk, assigned the role of the stator to hold the F 1 sector in place against the proposed rotation, and a cap in the late 1990s. In 1994, the first high-resolution structure (2.8 ) of F 1 appeared, consisting of the 3 3 hexamer with partial structure of the subunit. Currently there is no high resolution structure for the entire F 1 F 0 ATP synthase complex though many of the subunits have been solved individually or in part by model polypeptides.

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6 assigned the task of the stator to hold F 1 in place against the rotation of the centrally located stalk, until many years later (26-29). Today there is still no high-resolution structure of the entire F 1 F 0 ATP synthase enzyme complex from any organism. X-ray crystallographic and NMR data of partial complex structures from rat, bovine, yeast and E. coli or model polypeptides deduced from nucleotide sequences have accumulated over the past twenty years to allow for a composite structural model with both highand low-resolution structures (Figure 1-2). Currently, complete high-resolution structures are available for the , and c subunits and partial structures for the and b subunit. There is currently no high-resolution data for the membrane-integral a subunit. The Catalytic Core The first high-resolution structure, resolved to 2.85 consisted of 3 3 of F 1 prepared from bovine heart under inhibited conditions and in the absence of P i and substoichiometric amounts of ADP (20). This major breakthrough was shortly followed by a 2.80 F 1 isolated from rat liver in the absence of the physiological cation, Mg 2+ (21). The arrangement of the subunits in the two structures obtained were exceptionally similar and confirmed Catteral and Pedersens proposal made two decades prior by showing the three and three subunits arranged alternatively with the amino and carboxyl-termini of the subunit, each forming an -helix, extending up through the center of the hexamer (Figure 1-2) The only difference in the structures, which was the occupancy of the three nucleotide binding sites located at the interfaces, was likely due to the difference in preparation conditions (7) or crystal quality. In the former,

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7 Figure 1-2. Space-filling structural model of Escherichia coli F 1 F 0 ATP synthase. The model is based on a composite of high and low resolution structures taken from E. coli, yeast and bovine F 1 F 0 ATP synthases. F 1 F 0 subunits included in the model were 3 3 ab 2 c 10 The subunits were color coded as follows: red; green; cyan; orange; yellow; b, blue; a, brighter yellow; c, darker blue. The direction of the arrow indicates the direction of rotation of c 10 during ATP synthesis. The yellow and blue cylinders represent the a subunit and portions of the b subunit that currently have no high-resolution structures from any species.

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8 one of the catalytic sites was empty and in the latter, all three active sites were occupied with nucleotides (20, 21). Since the occupancy state of the three catalytic sites has been of considerable debate, a more accurate depiction of the F 1 moiety, which would give some insight about the mechanism of ATP synthesis, must be attained from crystals obtained under physiological conditions. A more accurate depiction of the structures and roles of each individual subunits of the F 1 ATPase follows. The hexamer Homology. The subunit of the E. coli F 1 F 0 ATP synthase, product of the uncA gene, is the largest subunit consisting of 513 amino acids with a deduced molecular weight of 55,313 Da. The subunit, a product of the uncD gene, is a 459 amino acid subunit, with a molecular weight of 50,325 Da. Based on the primary sequences, the and subunits of E. coli F 1 have the most obvious homologies in the chlororplast and mitochondrial enzymes (30). The highest conserved subunit from the E. coli F 1 F 0 ATP synthase is the subunit with approximately 70% homology with the chloroplast and mitochondria equivalents (31). The subunits exhibit roughly 50% homology (31). A total of 6 nucleotide binding sites are housed at the interfaces, three catalytic contributed primarily by the subunit and three noncatalytic housed primarily by the subunit (32, 33). The nucleotide binding regions have sequence homologies with other proteins that bind nucleotide or phosphate, including secA protein, N-ethylmaleimide sensitive fusion protein, herpes simplex virus UL15, Ca 2+ -ATPase, H + /K + ATPase and Na + /K + ATPase (34-37). Furthermore, the nucleotide binding motif, GXXXXGKT/S, known as the Walker A motif, which was first identified in the and sequences of F 1 has been found to be conserved in the high-resolution structures of other proteins

PAGE 30

9 including p21 ras adenylate kinase, RecA, elongation factor Tu, and transducin(20, 38-42). Tertiary structure. The first high-resolution structure of bovine F 1 resolved at the atomic level (2.8 ) was solved by Walkers group a decade ago (20). It was found to be a flattened sphere approximately 80 high and 100 wide with the three and three subunits arranged as a hexamer of alternating subunits around a centrally located 90 long -helix formed by the subunit. A dimple 15 deep is located at the top of F 1 The amino-terminal regions of the and subunits were once thought to be in close proximity to the membrane due to labeling experiments (43). Contrary to this early data, the crystallographic data placed the amino-terminal regions on the top of the 3 3 hexamer over 100 away from the lipid bilayer. The folds of the and subunits were found to be nearly identical. They each consisted of a six-stranded -barrel at the amino terminus ( 19-95 9-82 ), a central domain containing the nucleotide-binding site ( 96-379 83-363 ) and a bundle of seven and six helices at the carboxyl termini of the and subunits, respectively ( 380-510 364-474 ) (20). The nucleotide binding domain consisted of a nine stranded -sheet with nine associated -helices, of which the -carbons of the seven -strands and the seven associated helices can be superimposed onto the RecA protein ATP binding site with an rms separation of 1.9 (20). The three catalytic sites were located at the interfaces of the 3 3 hexamer. In the original crystal structure, now commonly referred to as the reference structure, two of the three sites were occupied by nucleotide, containing MgADP ( DP site) and MgAMP-PNP ( TP site). The third site was empty and designated E . The DP and TP

PAGE 31

10 subunits were in similar, closed conformations whereas the E adopted an open conformation, differing from the other two by a large hinge motion of the carboxyl terminal domain of greater that 20 Subsequently, several high-resolution structures of crystals obtained under various nucleotide conditions gave the same overall structure of bovine F 1 with two nucleotides bound (two nucleotide structures) (44-49). The TP site was found to occasionally contain a diphosphate nucleotide, establishing that there is no requirement for TP to be occupied by a nucleotide triphoshate to produce a conformational change (48). A more recent structure of bovine F 1 solved by the Walker group at 2.0 showed all three catalytic sites bound by nucleotide (50). Both the TP and DP sites contained MgADP, adopting the closed conformation, whereas the site corresponding to the E site in previous structures contained MgADP+P i and adopted a half-closed conformation. It is thought that the DP site is actually the catalytic site. The structure of rat liver F 1 was solved (2.8 ) in the presence of physiological concentrations of nucleotides but in the absence of the physiological cation, Mg 2+ In this structure, all three nucleotide binding sites adopted strikingly similar conformations, analogous to the DP and TP of the previously reported structure of the bovine F 1 This structure had no indication of the open conformation and showed the presence of nucleotide in all three sites (21). The structure of an 3 3 complex from a thermophilic bacterium was solved in the absence of nucleotides and exhibited all three subunits in the open conformation, suggesting there is a correlation between the open conformation and the absence of nucleotide (51). A low resolution crystal structure (4.4 ) of the E. coli F 1 has been obtained by Capaldis group, in which the catalytic sites are thought to be very similar to that of the bovine structure; however, the occupancy state of the nucleotide binding sites

PAGE 32

11 was unclear (52). The frequent reports of two nucleotide structures have bewildered scientists due to the vast body of biochemical data from numerous laboratories, using a variety of techniques, which establish indisputably that all three catalytic sites are readily filled with nucleotide (32, 53). It is possible that the enzyme preferentially crystallizes in a ground state intermediate which may occur after the release of product, leaving one site empty and opened (54). Nevertheless, the accumulating structural data may be indicative of several intermediary steps that may form during the synthesis of ATP. A detailed account of the mechanism of ATP synthesis follows later in this chapter. The crystal structure does offer some insight of the chemical mechanism of ATP synthesis (20). In the subunit, 4.4 from the terminal phosphate of the bound nucleotide triphosphate, there is clearly a density for a water molecule hydrogen bonded to the carboxylate of glu188 This carboxylate is positioned to allow an inline nucleophilic attack of the water molecule on the terminal phosphate. The guanidinium of a neighboring residue, arg373 is thought to help stabilize the negative charge on the terminal phosphate during the transition state (20). This same arrangement can be found in the catalytic site of transducin(42). The crystal structure also provides some insight as to why the nucleotide binding sites in the subunit are noncatalytic. There is no spacial equivalent of the carboxylate of glu188 in the subunit. The spatial equivalence in the subunit is filled by a gln208 with the side chain pointed away from the terminal phosphate (20). The binding of the adenine to the noncatalytic site of the subunit is highly specific, unlike the subunit nucleotide binding site, which can accommodate GTP, ITP as well as ATP (55, 56). This specificity is due to several hydrogen bonds as well as the presence of the tyr368 close to the 2-position of the adenine ring in the

PAGE 33

12 subunit, while in the subunit the adenine is in contact with a hydrophobic interface (20). Though the binding sites in the subunits are highly specific, the roles remain obscure. The subunit The subunit plays an important role in the catalytic core. Interactions between the aminoand carboxyl-terminal -helices and the 3 3 subunits are responsible for the conformational changes that result in ATP catalysis. The subunit is a fundamental part of the rotor stalk. The Rotor Stalk Two narrow stalks, a centrally located stalk and a peripheral stalk, have been observed to link the catalytic core of F 1 and the membrane-bound proton translocating F 0 with about of 40-45 in between (27). The central stalk came into view three decades ago via EM and has since become widely referred to as the rotor stalk. The rotor stalk consists of the and subunits. The bottom of the rotor stalk is connected firmly to the F 0 ring of c subunits located in the membrane and the top extends 90 within the 3 3 hexamer of F 1 where it forms crucial interactions with both the and subunits (57-59). F 1 F 0 ATP synthase is an extraordinary enzyme due to its ability to couple potential energy, obtained from proton translocation through F 0 in the membrane, to the synthesis of chemical energy, over 100 away in F 1 by a rapid rotation of subunits. Although predicted by Boyer in the 1970s, evidence of rotation did not appear until the early 1990s. The X-ray crystal structure solved by Walkers group suggested that the subunit was the rotating subunit by suggesting it could distribute itself to all three subunits as opposed to just one (20). Consistent with this idea was inhibition of the F 1

PAGE 34

13 complex by crosslinking the subunit to one of the or subunits (60, 61) and recovery after photobleaching experiments (62, 63). More convincing evidence was provided when Duncan et al. crosslinked the subunit to an unlabeled subunit by disulfide bond and then mixed the complex with 35 S-labeled subunit (along with the and subunits). When the disulfide bond was broken and ATP was added, the subunit was observed to switch from labeled to unlabeled subunit (64). Finally, direct evidence was achieved in single molecule experiments by attachment of a fluorescent actin filament to the subunit and observance of unidirectional rotation of the actin filament upon addition of ATP (15). Direct observation of the rotating subunit was soon followed by observance of the rotation of the and c subunits at the same speed and direction, indicating that these three subunits rotate in synchrony, forming the central rotary machinery of the enzyme complex (65-67). Until very recently, rotation has only been observed in the direction of ATP hydrolysis. Direct evidence for the synthesis of ATP by F 1 has been shown by attaching a magnetic bead to the subunit of F 1 fixed to a glass surface and the rotating the bead, in the appropriate direction, using electrical magnets (68). The first structural information obtained for the subunit of E. coli was accomplished by nuclear magnetic resonance (NMR) studies (69) and is good agreement with the crystal structure solved at 2.3 (70). In all previous crystal structures of the rotor stalk, the portion of the rotor stalks subunit protruding from the F 1 hexamer and the subunit were disordered. Recently, the structure of the bovine homologs of the rotor stalk and subunits has been solved and refined to 2.4 (48). The structures of the E. coli and subunits are remarkably similar with that from bovine F 1 When

PAGE 35

14 comparing the structures of the and rotor stalk obtained under different conditions or from different sources, in combination with an overwhelming amount of biochemical and immunological evidence, it is clear that the domains of the two subunits undergo major shifts in position, which reflects its fundamental role in the synthesis of ATP (20, 44, 48, 71-80). The subunit In E. coli, the subunit is the third largest subunit of F 1 F 0 ATP synthase, encoded by the uncG gene as a 286 residue polypeptide with a deduced molecular weight of 31,563 Da. It plays an essential role in coupling proton transport to the synthesis of ATP. The first visualization of a portion of the subunit was a bovine F 1 partial structure solved in combination along with the 3 3 hexamer revealing three -helices (20). The 209-272 (residues 223-286 in the E. coli sequence) carboxyl terminus formed a long (90 ) -helix extending from the stalk structure seen by EM to about 15 from the top of the hexamer. The bottom half of this helix formed a left-handed anti-parallel coiled coil with a shorter -helix composed of the amino-terminal residues 1-45 (20). The two helices protruded about 30 from the bottom of F 1 An approximately 20 kink in the latter helix was produced by pro40 and a similar but less pronounced kink was induced by leu217 in the former (48). A third, much smaller -helix, composed of 73-90 (residues 83-99 in the E. coli sequence) was inclined at about 45 degrees from the larger helices and located directly under the F 1 hexamer. More recently, the complete structure of the bovine rotor stalk has been solved to 2.4 (48). The overall length of the stalk, from the carboxyl terminus of the subunit to the very bottom where it contacts the ring of c subunits, was 114 The portion that

PAGE 36

15 protrudes from the 3 3 hexamer, i.e. the part seen in electron micrographs, was 47 long and 54 wide at its largest cross-section. A completely new / domain, consisting of a five-strand -sheet (1-5) and six -helices (a-f), was identified in the complete structure of the subunit. Helices a and f extended into the 3 3 hexamer to form the antiparallel coiled coil discussed previously. Strands 1-3 along with helices b and c formed a Rossman fold that forms extensive interactions with the subunit as well as the ring of c subunits in the membrane (discussed below). This fold was linked to a -hairpin, formed by strands 4 and 5, by helix d. Overall, this / domain had a globular, oval shape with the dimensions 51 wide by 41 high. The positioning of the / domain at the base of the rotor stalk may provide stability to the structure of the rotor stalk during rotational catalysis (48). A low-resolution crystal structure of the E. coli F 1 F 0 ATP synthase subunit was solved to 4.4 (52). Upon comparison with the high-resolution bovine structure obtained one year later, a few differences were observed (48). Helices a and f (see above) were extended by and extra 12 and 20 residues, respectively. Four additional putative -helices, designated B and D-F, were found in the E. coli structure and had little agreement with the bovine subunit structure. E. coli helix B runs parallel with the bovine -strand 5 and may correspond to it. Helix D has no apparent equivalent in the bovine model and helices E and F appear to overlap with regions in the bovine (bacterial see Table 1-1) and (no equivalent in bacteria) subunits. The remainder of the E. coli structure appeared similar to the bovine structure. The crystal structure displays a strikingly asymmetrical F 1 due to differences in the domains of the and subunits and the interactions formed with the single subunit

PAGE 37

16 (20). The obvious asymmetric positioning of the coiled coil of the subunit is a key feature to the mechanics of the binding change mechanism of F 1 F 0 ATP synthase. Its large carboxyl terminus -helix passes through a hydrophobic sleeve formed by six proline-rich loops of the and subunits, undoubtingly resulting in the conformational changes occurring in the catalytic sites (20). In the E subunit (see above), several hydrogen bonds are formed with the subunit, which forms a catch, resulting in conformational changes. Specifically, residues arg254 and gln255 in the carboxyl terminal helix form hydrogen bongs with E-asp317 E-thr318 and E-asp319 Also, a second catch is formed between the carboxyl terminal domain of the T subunit and the short helix of the subunit. Hydrogen bonds formed between lys87 lys90 and ala80 with T-asp394 and T-glu398 This sequence of the subunit, DELSEED ( 394 400 ), is a portion of the binding site of amphipathic cationic inhibitors and putatively the ATPase inhibitor protein (81-83). Recently, mutations of residues involved in the catch loops were shown in inhibit ATP hydrolysis activity by the soluble F 1 -ATPase (84). Structural information suggests the two antiparallel coiled coil -helices of the subunit may unwind during rotational catalysis and the subunit rotates around the F 1 axis while undertaking a net translation of about 23 (85). It is likely that these gross changes observed in the structures revealed individual functional states of the enzyme complex during catalysis. The subunit The subunit of E. coli F 1 F 0 ATP synthase consists of 138 amino acids with a molecular weight of 15, 068 Da and is encoded by the uncC gene. The subunit has several putative functions in the F 1 F 0 ATP synthase complex including structural, inhibitory and coupling roles. Structurally, the binding of F 1 to F 0 has long been known

PAGE 38

17 to require the presence of the subunit, which had implicated it as part of stalk structure (86). In isolated F 1 and in isolated F 1 F 0 to a lesser extent, it has been shown to inhibit ATP hydrolysis activity (87-89). The removal of from isolated F 1 resulted in up to a 10-fold increase in ATP hydrolysis activity. Furthermore, a truncated version of the subunit lost all inhibitory functions but still promoted binding of F 1 to F 0 hence the inhibitory feature has been assigned to the extreme carboxyl terminus (90). It was speculated that it acts as an inhibitor by reducing the rate at which the product is released from the catalytic site (91). Also, the subunit, as part of the rotor stalk, plays a role in the coupling of proton translocation to the catalytic site. The subunits diversity of functions is supported by the findings that it produces several points of interactions with the (61, 74, 92), (61, 88, 89, 93, 94) and (74, 95-97) subunits of F 1 and the c (98, 99) subunits of F 0 An innovative set of experiments conducted by members of the Dunn laboratory made use of different sized fluorescent proteins, ranging from the 12 kDa cytochrome b 562 protein to the 30 kDa flavodoxin reductase protein with a 20 residue linker (100). The proteins were fused to the carboxyl-terminus of the subunit. Since the subunit is part of the rotor stalk, according to the concept of rotational catalysis, the fusion of a large protein at this site should sterically hinder rotation due to the presence of the peripheral stalk. Cells expressing the smaller cytochrome b 562 protein fused to the subunit grew on minimal media, indicating a functional F 1 F 0 ATP synthase complex. However, cells expressing the larger flavodoxin reductase protein fused to the subunit, though found in an intact enzyme, failed to grow. These results provided the first evidence, in vivo, supporting rotational catalysis.

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18 High-resolution structural data of the E. coli subunit was first solved by NMR followed by the X-ray crystal structures of the isolated and complexed subunits (69, 70, 101). The isolated subunit consisted of an 84 residue amino-terminal -sandwich domain and a 48 residue carboxyl-terminal helix-turn-helix domain in which the two-helices formed an antiparallel hairpin (70). The -sandwich consisted of two five-stranded -sheets folded as a rigid, flattened -barrel. The structure of the isolated subunit from E. coli was very similar to the F 1 complex isolated from bovine, which included the 3 3 subunits (48, 70). Superimposition of 127 of the amino acid C resulted in an rms deviation of 1.6 On the other hand, in the E. coli complex, resolved to 2.1 the subunit assumed a strikingly different conformation, in which the two -helices of the antiparallel hairpin at the carboxyl-terminus are wide apart and wrapped around the subunit (Figure 1-3A) (101). Subsequently, both conformations of the subunit have been trapped in E. coli F 1 F 0 ATP synthase by crosslinking experiments, confirming the existence of both in an intact enzyme complex (102). Furthermore, Capaldis group observed that when the carboxyl-terminal helices assume the hairpin conformation, bringing them closest to the F 0 sector, ATP hydrolysis was activated. Still, the enzyme was fully coupled in the direction of either hydrolysis or synthesis. In contrast, when the two helices were open, assuming a position closer to the F 1 sector, ATP hydrolysis was inhibited and the enzyme functioned only in the direction

PAGE 40

19 Figure 1-3. Structure of the subunit of E. coli F 1 F 0 ATP synthase. The residue numbers and subunit labels are color coded to match the subunits it represents. The subunit has been suggested to undergo large conformational changes during catalysis from an overwhelming amount of biochemical data. Two different structures have been obtained for the subunit, confirming the previous data. A) Superimposition of the C trace of the structure obtained from isolated subunit (-helices shown in red and the -sandwich shown in blue) with the structure obtained from the complex (yellow). B) Composite structural model. Rotor stalk is based on the crystal structure obtained from the complex. The DELSEED 2 and the -sandwich are indicated to show the close proximity of DELSEED and 2 as well as the relative distance between the 2 and the -sandwich.

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20 of ATP synthesis. This conformational switch of the subunit was therefore suggested to play a key role as a selective inhibitor of ATP hydrolysis and directional regulator of rotational catalysis by acting as a ratchet (102). Movement of the two -helices was consistent with other observations. Changes in the subunit conformation due to nucleotide occupancy in the catalytic sites has been observed in tryptic proteolysis experiments (89). Cysteine replacements in the carboxyl-terminal -helix ( 2 ) resulted in crosslinks with the and subunit (61, 103). More importantly, treatment with a zero-length crosslinker, 1-ethyl-3[3-dimethylamino]propyl carbodiimide (EDC), resulted in a high yield of crosslinks between the subunit and the DELSEED ( 380 386 ) region of the subunit ( DELSEED ) following ATP hydrolysis in the catalytic sites, but these interactions are disrupted upon the subsequent binding of ATP. Also, in a composite structure of F 1 F 0 ATP synthase incorporated with the E. coli complex as solved by Rodgers and Wilce, the sandwich was at least 10 away from the DELSEED region (Figure 1-3B) (101). The carboxyl-terminal 2 produces several points of interactions with the and as well as points of interactions with its own -sandwich domain (61, 74, 88, 89, 92-99). In order for the 2 to interact with the DELSEED 1 and the subunit -sandwich domain, it is clear from the structure that the subunit would be required to undergo large movements during the catalytic cycle. In E. coli, F 1 F 0 ATP synthase can act in two functional directions. In the case of a dissipated electrochemical gradient, the F 1 F 0 complex acts primarily as an ATPase in order to pump protons across the membrane to provide a gradient to drive various ion transport activities in the cell. Under severe conditions where cellular ATP levels are exceedingly low the enzyme acts predominantly in the direction of ATP synthesis.

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21 Therefore, one can imagine that the ability to selectively turn off ATP hydrolysis, while preserving ATP synthesis function, may be important for E. coli. In mitochondria the ability to control the F 1 F 0 ATP synthase complex is essential. It is believed to act exclusively in the direction of ATP synthesis and is strictly regulated (104). The ring of c subunits The c subunit is one of the three membrane-bound F 0 subunits of F 1 F 0 ATP synthase. Ten copies of the c subunit form a ring in the membrane that plays a crucial role in both proton translocation and rotation of the rotor stalk (105). It is the smallest subunit of the F 1 F 0 ATP synthase enzyme complex with 79 amino acids and a molecular weight of 8,256 Da and it is encoded by the uncE gene. Structure and topology. Early biochemical, genetic and immunological data had suggested the structure of the c subunit to be that of a helical hairpin with two lipophilic -helices (amino acids 1-41 and 50-79) separated by a hydrophilic loop (amino acids 42-49). Both of the putative transmembrane helices in the regions of c leu4-leu19 and c phe53-phe76 were vulnerable to chemical modification by the nonpolar photoreactive reagent 3-(trifluoromethyl)-3-(m-[ 125 I]iodophenyl)diazirine (TID), which is a hydrophobic carbene generator that is believed to react from the nonpolar region of the lipid bilayer, indicating that these regions were in fact in the hydrophobic phase of the bilayer (106). The loop region of the hairpin is substantially more polar and antibodies against it were shown to bind to F 1 -stripped inverted membrane vesicles suggesting that it resides in the cytoplasmic of the cell (107, 108). Both of the transmembrane helices are devoid of charged amino acids with the noteworthy exception of c asp61 in the center of the second helix, which undergoes a protonation and deprotonation cycle during proton translocation

PAGE 43

22 (discussed in F 1 F 0 ATP Synthase Mechanism below) Dicyclohexylcarbodiimide (DCCD) reacts specifically with c asp61 blocking proton translocation, and this reaction is blocked by the mutations c ala24ser or c ile28thr suggesting that the c subunit is folded in such a way so that the asp61 of the second helix is in close vicinity to residues 24 and 28 of the first helix (109, 110). This model was further supported by the ability to move the critical aspartate from residue 61 in the second helix to residue 24 in the first helix without disruption of enzyme function (111). Also, only one subunit in the ring of 10 c subunits need be modified by DCCD to inhibit activity, indicating that each one of the c subunits is consecutively involved in proton translocation (112, 113). Furthermore, modification of the c subunit by DCCD trapped the configuration of the subunit (discussed above), providing evidence for a connection between the c and subunits (89). Intersubunit contacts made by the c subunit are evident from mutational data and gives some insight to the topology. Mutations constructed in the polar loop region can disrupt the binding of F 1 to F 0 (114-117). Three conserved amino acids, c arg41 -c gln42 -c pro43 lie at the apex of the polar loop region and are predicted to interact with the F 1 subunit (98, 116). F 1 F 0 ATP synthase complexes with the uncoupling mutation, c gln42glu were found to be recoupled with a second site suppressor mutation in the subunit of F 1 glu31gly, val, or cys (98) and was shortly followed by the observance of disulfide bridge formation between the c subunit and the and subunits (99, 118). Also, switching the essential aspartate from residue 61 in the second helix of the c subunit to residue 24 of the first helix (discussed above) resulted in a functional F 1 F 0 ATP synthase complex though the cells were not as healthy compared to cells containing a wild-type enzyme complex (111). Eighteen third-site suppressor mutants were found that helped to

PAGE 44

23 optimize this c ala24asp,asp61gly defect, with only five laying on the c subunit and 13 in the a subunit, all near the a arg210 residue, which is required for proton translocation (further discussed below) (119, 120). Early models of the organization of F 0 suggested that the ring of c subunits were situated on the periphery, surrounding the centrally located a and b subunits, which rotated in the center of the ring (121). This model was proved wrong by high-resolution NMR data (122) and cross-linking experiments (123, 124) which indicated that the oligomer of c subunits are closely packed with a lipid filled core less than 25 wide. The individual c subunits are packed front-to-back such that the second helix of each is situated towards the exterior and the first helix is located on the interior, which renders the c asp61 exposed to the lipid environment. The uncommonly high pKa (7.1) of the c asp61 carboxyl side chain is likely due to this hydrophobic environment (125). Furthermore, scanning force and cryoelectron microscopy demonstrated that F 0 is asymmetrically arranged in the membrane (27, 126, 127). For these reasons, the a and b subunits are thought to be situated to the periphery of the ring of c subunits. High resolution structures of membrane-bound proteins were nonexistent for many decades past structural determination of soluble proteins and still prove difficult to this day due to their highly hydrophobic nature. The membrane intrinsic c subunit of E. coli, which was solved by NMR in an organic solvent (chloroform-methanol-water) in the 1990s, was one of the earliest high-resolution structures of a transmembrane helical protein (122, 128-130). Notable, the c subunit could be reconstituted from the organic solvent mixture with complete preservation of function; therefore, it was clearly not irreversibly denatured (113). As predicted two decades prior, the c subunit folds as a hairpin of two extended -helices with the c asp61 of the second helix packed less than 5

PAGE 45

24 from the c ala24 and c ile28 of the first helix (122). With the exception of c asp61 in the second helix, both helices consist entirely of nonpolar amino acids. The first helix is greatly enriched in glycines and alanines, which led to a smaller diameter. The -helical structure of the second helix is interrupted around c asp61 due to disrupted hydrogen bonds around c pro64 which cause the angle of the helical packing to change direction from there to the carboxyl terminus (122). A recent study, using parallax analysis of fluorescence quenching, the proton binding site c asp61 was found to be deeply embedded in the membrane at about 1.8 from the center of the bilayer (131). Stoichiometry. The stoichiometry of c subunits would be valuable in determining the number of protons transported per ATP synthesized and will directly relate to the P/O ratio of oxidative phosphorylation. However, the number of c subunits in F 0 had been a matter of controversy for many years. The number of c subunits in an F 1 F 0 ATP synthase complex was suggested to be between 9 and 14, but whether this number fluctuated based on the species or environmental conditions or whether it was a fixed number were the two prevailing arguments until just a few years prior. Based on a related family of vacuolar (V-type) ATPases, in which the proposed subunit c had evolved into a fused dimer of four transmembrane helices with a single proton-transporting glutamate in the center of the fourth helix, Fillingame et al. set out to genetically fuse the E. coli c subunit by introducing a flexible loop of similar length (123). The generated c-c dimers and c-c-c trimers resulted in functional enzyme complexes. In combination with crosslinking studies and normalization to the / content of the membranes, the favored stoichiometry was fixed to 12 c subunits per F 1 F 0 ATP synthase complex (132). More recently, the experiment was revised to include only trimers and tetramers of the c subunit (105).

PAGE 46

25 Partial activity was observed in complexes incorporated with eight (c 4 + c 4 ) or nine c (c 3 + c 3 ) subunits and crosslinked products of more than 10 c subunits were observed but did not purify in intact enzyme complexes. Crosslinking showed that the preferred stoichiometry of c subunits in intact E. coli F 1 F 0 ATP synthase was c 4 + c 3 + c 3 or 10 c subunits. This number is consistent with the c 10 oligomer found in the yeast crystal structure of a yeast F 1 F 0 ATP synthase consisting of 3 3 c 10 resolved to 3.9 (58). However, the preferred number may still vary in different species. The archaebacteria Methanococcus jannascjii ATP synthase has a natural c subunit trimer and therefore cannot incorporate the E. coli equivalent of c 10 in the membrane (133). With 10 c subunits present in the membrane 3.3 protons are required per ATP synthesized, which was compatible with the early experimentally determined ratio of 3 H + /ATP estimated from E. coli whole cells (134). This value also indicates a P/O value of 2.3 from NADH-linked substrates and 1.4 for succinate, also compatible with the predicted values of 3 and 2, respectively (135). The Stator Stalk The b and subunits were once believed to form the central, rotating stalk of F 1 F 0 ATP synthase. However, high resolution crystallographic data refuted this idea in the mid 1990s (20). The stator stalk did not come into view until improved EM technology observed a peripheral stalk in the late 1990s (26-29) and the visualization of the subunit, as a cap structure atop F 1 F 0 ATP synthase, soon followed (27-29). As its name implies, the role of the stator is to hold the 3 3 hexamer in place against the rotation of the subunit during rotational catalysis. Based on chemical crosslinking data, it is currently believed that the a subunit resides to the periphery of the ring of c subunits with

PAGE 47

26 the membrane-spanning domain of the b dimer situated to one side of the a subunit where it is in close proximity to both the a and c subunits. The b dimer extends out of the membrane and in a highly elongated conformation reaches to near the top of F 1 making contacts with theand subunits along the way and the subunit at its extreme carboxyl terminus. The stator stalk consists of the b subunit of F 0 and the subunit of F 1 Although the primary function of the a subunit of F 0 is considered to be a role in proton translocation along with the c 10 subunits, it nevertheless plays the part as a stator and will be discussed in this section. Structurally, the a subunit plays a role in both the formation of a dynamic interface with the ring of c subunits as well as the formation of a secure complex with the b dimer. Pursuit of a high-resolution structure for the a subunit remains a challenge to this day. The high-resolution structure of the region of the a subunit that forms the interface with the c 10 subunits is eagerly anticipated since it appears to be the crucial region for proton translocation. In regard to the stator stalk, partial structural information has been obtained for the E. coli b subunit membrane spanning domain and the subunit amino terminus by NMR studies (136, 137). X-ray crystallography has solved the structure of a model polypeptide based on the dimerization domain of the b subunit (138). The binding of F 1 to the membrane-bound F 0 requires both the and subunits, suggesting that each of the subunits are involved with the stalk structures of F 1 F 0 ATP synthase (87, 139, 140). In fact, the subunit forms an integral part of the peripheral stalk and the subunit functions as part of the central stalk (discussed below). The subunit of F 1 has been visualized seated at the very top of the F 1 3 3 hexamer by EM (11). However, recent

PAGE 48

27 evidence has suggested that the subunit may actually be positioned slightly to the side of F 1 in association with only a single subunit (Figure 1-2) (100, 141-143). The b subunit is the primary focus of this dissertation and will be discussed at length later in this chapter. The a subunit The E. coli a subunit is a large, extremely hydrophobic protein encoded by the uncB gene and consists of 271 amino acids with a molecular weight of 30,276 Da. All enzymes of the F 1 F 0 ATP synthase family contain an a subunit homolog with strong primary sequence homology even among evolutionarily diverse species (144). The most highly conserved region resides in the carboxyl-terminal one-third end, amino acid residues a 190-263 Notably, in the region that is involved in proton translocation, there is a remarkable conservation of the amino acid residues a leu207 a arg210 a leu211, a asn214 and a gln252 and an evident conservation of a glu219 and a his245 at the homologous positions in all a subunits from different species (144). The a arg210 is the most strictly conserved among all species and does not tolerate substitution with any other amino acid (further discussed below) (145-147). As mentioned above, there is no high-resolution structural data for the a subunit. Also, contradicting models exist concerning the number of transmembrane helices as well as the orientation in the membrane. Difficulty in studying the structure of the a subunit arises from its extreme hydrophobicity and the necessity to include the denaturant, trichloroacetate, in purification procedures. This is compounded by the fact that it cannot be expressed at high levels in E. coli and is not found in the membrane without the presence of both the b and c subunits (148-150). Furthermore, the a subunit is known to

PAGE 49

28 be a substrate of the protease FtsH, which will rapidly degrade the subunit if it is not in its native state (151). It was readily labeled with TID, which is a hydrophobic carbene generator that is believed to react from the nonpolar region of the lipid bilayer, but its solubility properties made it unsuitable for analysis as was done with the c and b subunit (106). Consequently, the amino acids in contact with the lipid phase of the bilayer were not identified. Due to difficulties in obtaining high-resolution structural data, much of what is known of the a subunit arises from mutational studies. Topology. Hydropathy analyses indicated five definite membrane-spanning regions and one putative membrane span (121, 140, 152, 153). Much of what is known of the a subunit structure and has come from the analysis of cysteine mutagenesis. Greater than 50 cysteine substitutions, which resulted in a functional F 1 F 0 ATP synthase, were used in two kinds of experiments (154-158). Various maleimide derivatives were used to search for the surface-accessible regions (154, 155). And double cysteine mutations were used to search for disulfide formation between a-a and a-c (153). The results supported the model in which the a subunit spans the membrane five times and the fourth span, which includes a arg210 is in contact with the second transmembrane -helix of the c subunit (Figure 1-4A). Additionally, residues that were originally thought to be located in the cytoplasm were not labeled, indicating that the six-membrane span model was incorrect (132, 159). The location of the amino-terminus of the a subunit has also been very controversial. A substantial amount of evidence indicates that the carboxyl terminus

PAGE 50

29 Figure 1-4. Controversial models of the a subunit topology. There is no high-resolution structural data for the a subunit. Mutagenesis, crosslinking and immunological experiments were used to study the topology. Roman numerals indicate the number of transmembrane helices. Small numbers indicate the relative position of the amino acid residue. Several crosslinking reactions were observed between the fourth helix of the a subunit (IV*) and the second helix of the c subunit in double cysteine mutants. Contradicting models exist for the topology of the a subunit. A) Model with five-transmembrane helices and the amino terminus residing in the periplasm. B) Model with six transmembrane helices and the amino terminus located in the cytoplasm.

PAGE 51

30 resides in the cytoplasm (154, 155, 160). This observation, in combination with the five-transmembrane helices, indicates that the amino-terminus should reside in the periplasmic space. Polyclonal antibodies against a peptide model of the extreme carboxyl-terminus as well as antibodies against epitope tags constructed at the carboxyl terminus of the a subunit revealed this region to be located in the cytoplasm (160). Moreover, cysteine substitutions at a 266 or a 277 were highly reactive on the cytoplasmic side of the membrane (154, 155). The orientation of the amino and carboxyl-termini was studied by gene fusion proteins and peptide-directed antibodies, revealing a cytoplasmic location of both termini (161, 162). Insertion of epitope tags at various positions also confirmed the cytoplasmic local of both termini, arguing in favor of the controversial six-transmembrane model of the a subunit (Figure 1-4B) (160). In the five-transmembrane model a stretch of about 37 amino acids at the amino-terminus resides in the periplasm with only one transmembrane helix, approximately up to residue a 66 present (Figure 1-4A) (154-156, 158). In the six-transmembrane model the amino-terminus resides in the cytoplasm with two transmembrane helices present before the first cytoplasmic loop, which range from approximately residues a 33-49 and a 54-70 (Figure 1-4B) (160). A series of a subunit amino-terminal truncations and internal deletions were constructed and the F 1 F 0 ATP synthase function was tested by growth on a succinate minimal media. Assembly of intact complexes was tested by membrane-associated ATPase activity and the presence of the a subunit was analyzed by immunoblot analysis (163). Four sections were found to be particularly interesting. The first 33 residues at the amino terminus were shown to be necessary for the insertion of the a subunit into the membrane. Two internal deletions, from residues a 91-99 and a 163-177 resulted in functional

PAGE 52

31 enzyme complexes, indicating that these regions were not important for function. A fourth deletion, from residues a 120-124 was concluded to be important for function, but not assemble because high levels of a subunit were found in the membrane, but the enzyme was not functional. The importance of the carboxyl-terminus was also analyzed by constructing a series of early termination codons (164). Sequence alignment of the a subunit demonstrates that many bacterial homologues contain glutamate and histidine residues at the extreme carboxyl-terminus (glu-glu-his in E. coli). However, truncation of the final four residues had no effect, and truncation of the final nine residues were tolerated at 25C, suggesting that the extreme carboxyl terminus of the a subunit did not significantly contribute to proton conduction or functional interactions with other subunits. Proton translocation. The first indication that the a subunit was directly involved in proton translocation appeared nearly two decades ago when mutations constructed in the a subunit (a ser206leu and a his245-tyr ) were found to affect F 0 -mediated proton pumping without influencing F 1 F 0 ATP synthase assembly (165). Since then, not including the cysteine mutations described above, more than 75 missense mutations have been constructed and analyzed in or near the conserved regions of the a subunit to chart the amino acids involved in proton translocation. In general, mutation of a conserved amino acid residue impaired F 0 -mediated proton translocation, but the severity of the defects varied (166). The only F 1 F 0 ATP synthase a subunit residue that is strictly conserved amongst all species, from bacteria to humans, and cannot endure any amino acid substitution, whether basic, acidic or nonpolar, was a arg210 (145-147). Mutations at this site abolished both

PAGE 53

32 ATP-driven proton pumping and passive F 0 -mediated proton translocation. Growth on succinate minimal media indicated no ATP synthesis by the mutants. The observed effects were shown not to be due to failure of F 1 F 0 ATP synthase to assemble because treatment with the detergent lauryldimethylamine oxide (LDAO) released F 1 from the prepared membranes and revealed abundant ATP hydrolysis activity. The presence of assembled F 1 F 0 ATP synthase complexes incorporated with an a arg210 mutant was later directly confirmed by Dr. James Gardner (167). Substitution with an alanine allowed passive F 0 -mediated proton translocation indicating that the proton channel was intact and suggested that the a arg210 is not obligatorily protonated or deprotonated during proton conduction (168). A second site suppressor mutation, a gln252arg which partially compensated for the a arg210gln mutation, was identified, and suggested to be in close proximity to each other with residence on the transmembrane helix 5 and 4, respectively, in the five-transmembrane model (121). The a 210 residue is thought to have a direct role in proton translocation. The orientation of the a subunits fourth transmembrane helix had been determined relative to the orientation of the c subunits second transmembrane -helix by crosslinking double cysteine mutants (157). Crosslinking data has positioned a 214 in close proximity to c 62 and c 65 and a 211 close to c 69 (157). This places the putative fourth helix of the a subunit in contact with the second helix of the c subunit. Models have the a 210 residue positioned near the center of the fourth helix at a level in the lipid bilayer very close to the essential c asp61 residue (14). Whether a 210 is directly protonated/deprotonated or controls protonation of the c asp61 residue remain unanswered (169). Insight from a high resolution structure of an intact F 0 is greatly desired and would provide extremely valuable answers to many of the unsolved questions.

PAGE 54

33 Single mutations at residues a 218 a 219 or a 245 were shown to have a considerable impact on F 0 -mediated proton conduction (144, 146, 170). When comparing amino acid sequences of various mitochondria, chloroplast and bacteria, there appears to be an instance of evolutionary covariation with these three amino acids (144). This suggests that when a mutation occurred in one of the three residues, it was accompanied by a second mutation to compensate for any loss in activity. This would cause the two residues to pass through evolution as a hereditary unit. Based on this observation, double mutants were constructed in the E. coli a subunit to imitate other lines of evolution (144, 171). Every double mutant studied resulted in functional F 1 F 0 ATP synthase complexes with considerably more activity than any of the single residue mutants. Due to the functional relationship, it is possible that these three amino acids are in close proximity to each other. A few other strongly conserved amino acid residues located on the fourth and fifth transmembrane helices are worth mentioning. Residues a asp214 and a gln252 were both strongly conserve but found nonessential, with the effects of mutations at these residues varied widely (146, 170, 172, 173). Models of the a subunit have these residues lining a water-filled proton channel. Recently, the aqueous accessibility of residues along transmembrane helices 2 and 5 has been shown to extend to both sides of the membrane (174). Also, a mutation at residue 217, a ala217arg blocked proton conduction and inhibited F 1 F 0 ATP synthase activity (167). The subunit The E. coli subunit is one of the F 1 subunits. It is discussed here because it is an essential part of the F 1 F 0 ATP synthase stator stalk. The simplest stator stalks occur in

PAGE 55

34 nonphotosynthetic prokaryotes and consist of a dimer of the F 0 b subunits (discussed in the following section) and a single F 1 subunit. The subunit displays a very low level of conservation across various species. It is a globular protein encoded by the uncH gene that consists of 177 residues with a molecular weight of 19,332 Da. It plays essential roles in both the binding of F 1 to F 0 as well as coupling of the catalytic activities of F 1 and F 0 (139, 175-180). Circular dichroism (CD) spectroscopy and sedimentation analysis studies performed on the subunit suggested a highly helical and elongated conformation (139). Structure of the subunit. A partial high-resolution structure of the subunit has been solved by NMR (137). During the purification procedure, a truncated form of the subunit was produced by a bacterial protease. It was revealed to be the amino-terminal 134 amino acids ( 1-134 ) by mass spectroscopy and N-terminal sequencing. The same sized subunit fragment was often seen in F 1 preparations and could be produced in isolated E. coli F 1 by treatment with trypsin without liberating the 1-134 from the F 1 complex (89). Furthermore, purified 1-134 stably binds to -free F 1 preparations. The high affinity of the 1-134 subunit for F 1 indicated that the conformation of the fragment was preserved during the purification procedure. NMR was performed on both the 1-134 fragment and the intact subunit; however, the quality of data for the intact subunit was not sufficient enough for structural analysis due to its propensity to aggregate at high concentrations. To date, the carboxyl-terminal 43 amino acid residues ( 135-177 ) of the subunit is the only portion of the F 1 sector not known at the atomic level. The amino-terminal 105 residues of the subunit formed a dense globular domain, while the region from residues 106-134 was mostly disordered with the exception of one

PAGE 56

35 -helix (137). The amino-terminal domain, 1-105 consisted of a six -helix bundle with the dimensions 45 x 20 x 30 Helices 1 ( 4-20 ) and 2 ( 24-38 ), and helices 5 ( 70-81 ) and 6 ( 88-104 ) organized into V-shapes that intercalated to form a core. Helices 3 ( 41-47 ) and 4 ( 53-64 ) were packed compactly against this four-helix core. Following this globularly packed domain there was a loop region followed by a seventh -helix ( 118-129 ). Comparison of the structural data for the intact subunit against that of the 1-134 fragment illustrated the same structure for residues 1-104, but the spectral shift of residues 105-134 was very different. It was possible that the carboxyl-terminal 42 residues missing from the 1-134 fragment affects this region of the subunit. subunit topology. Taken together with biochemical and immunological data, the structure revealed by NMR revealed that the subunit consists of two domains, an amino terminal domain, 1-104 and a carboxyl-terminal domain, 105-177 Under oxidizing conditions, two native cysteines present in the aminoand carboxyl-terminal domains of the intact subunit, cys64 and cys140 respectively, formed a disulfide bond. Furthermore, NMR data indicated some NOEs between the carboxyl-terminal -helix and the amino-terminal domain. The data indicated that there is probably a close interaction between the aminoand carboxyl-terminal domains of the intact subunit. Proteinase accessibility and immunological analysis were used to examine the topology of the subunit (89). The subunit was susceptible to trypsin digestion at the carboxyl-terminal 20 residues in isolated F 1 but not in intact F 1 F 0 ATP synthase, indicating a protection of the amino-terminal region by F 1 Deletion analysis of the carboxyl-terminal region also implied the importance of the subunit in binding F 1 to F 0

PAGE 57

36 Taken together, these observations suggested that the amino-terminal domain is predominantly involved in the binding of the subunit to F 1 and the carboxyl-terminal domain is involved in binding to F 0 The location of the subunit has had a history of being very controversial. Prior to the high resolution structure obtained by Abrahams et al. (1994), the b and subunits were expected to form part of the central stalk of the F 1 F 0 ATP synthase enzyme, which is now known to consist of the and subunits (20). Due to the dimensions, it seemed unlikely that the b and subunits could fit as part of the central stalk, which implied that they must form a separate connection between F 1 and F 0 Improving EM technology did not allow visualization of the second stalk structure at the periphery of the F 1 F 0 ATP synthase complex until many years later (27, 28). Prior to visualization by EM, several early crosslinking studies had been reported in the quest to find the location of the subunit binding on F 1 finding it to be on the subunit (89, 181-184). Notably, crosslinking the subunit to the subunit did not have a great impact on F 1 F 0 ATP synthase function, as would be expected if the subunit formed part of the stator stalk (185). High-resolution structure of the F 1 3 3 hexamer with a partial structure of the subunit had revealed a dimple in the top of the hexamer approximately 15 deep that was adjacent to the core space where the aminoand carboxyl-terminal -helices of the subunit resided (20). EM studies had revealed a cap structure at the very top of F 1 in both E. coli and mitochondrial complexes (27, 29, 186). It was thus believed that the subunit resided in the dimple of F 1 as the cap seen in the EM structures (187). This possibility was refuted when Prescott et al. demonstrated the ability to stably incorporate the green fluorescent protein (GFP), via varying length peptide linkers (0, 4 or 27 amino

PAGE 58

37 acids), to the carboxyl-terminus of the subunit without interrupting function of the enzyme complex. GFP forms a rigid, stable structure with the dimensions 24 wide and 48 high (188). This study indicated that the putative cap structure could not possibly occupy the entire dimple atop F 1 More recent evidence has suggested that the subunit may actually be positioned slightly to the side of F 1 in association with only a single subunit (Figure 1-2) (100, 141, 142, 189). The b subunit The b subunit is required for the normal assembly and function of F 1 F 0 ATP synthase (190). The E. coli F 1 F 0 ATP synthase has two identical b subunits, which form a homodimer, that are the product of a single gene (Figure 1-2). It is an elongated amphipathic polypeptide that crosses the membrane one time at its amino-terminus and has an extensive hydrophilic carboxyl-terminal domain. This pattern is characteristic of b-type subunits of ATP synthases, although the mitochondrial b has two consecutive membrane-spanning segments at the amino-terminus (191). Most ATP synthase b-type subunits consist of between 150 and 170 amino acid residues. The E. coli b subunit, encoded by the uncF gene, consists of 156 amino acid residues and has a deduced molecular weight of 17, 265 Da (Figure 1-5). Domains and Structure. Currently there is no high-resolution structure of the entire b subunit. Several factors probably contribute to the difficulty of structural analysis. The b dimer is a thin, highly extended, mostly -helical structure, its dimerization is comparatively weak and reversible (192), and it displays evidence of flexibility (193-195). This has led to alternative low-resolution approaches to study the structure of the b dimer such as circular dichroism (CD) spectroscopy, deletion analysis,

PAGE 59

38 Figure 1-5. Amino acid sequence of the E. coli F 1 F 0 ATP synthase b subunit. The E. coli b subunit is a 156 residue amphipathic polypeptide. The amino acid sequence and the four domains are shown. The transmembrane domain (b 1-22 ), tether domain (b 24-60 ), dimerization domain (b 63-122 ) and the -binding domain (b 123-156 ) are shown in blue, orange, green and red, respectively. The large purple stars indicate residues capable of forming high yields of b-b crosslinks upon cysteine substitution. The smaller purple stars indicate residues found to form low-yields of crosslinks. The arrows indicate positions crosslinked to other subunits of ATP synthase. High-resolution structures based on model polypeptides consisting of b 1-34 and b 62-122 (underlined residues) have been solved by NMR and crystallography, respectively. NMR analysis of residues 1-34 has revealed a -helical structure with a rigid 20 bend at positions 23-26. X-ray crystallography revealed a highly -helical structure with modeled into a right-handed coiled coil.

PAGE 60

39 analytical ultracentrifugation, chemical crosslinking, and the analysis of tendencies for disulfide bond formation. CD spectroscopy analysis has predicted the secondary structure of the b subunit to be approximately 80% -helical with about 14% -turn conformation (196). Although there is no high-resolution structure of the intact F 0 sector, an abundance of evidence suggests the necessity of the b subunit to exist in the dimeric state. The hydrophilic region of b, consisting of residues b 24-156 (also known as b sol ), has been expressed and shown to form highly extended dimers capable of binding to F 1 -ATPase in solution (197). Sedimentation equilibrium ultracentrifugation gives a molecular weight value of about 30,000 Da for b sol consistent with a dimer of two 15,000 Da b sol monomers (13). The existence of the dimeric state of the b subunits was confirmed by covalently cross-linking the two b subunits in the complex and verifying the activity of the enzyme (198). Furthermore, the ability of b to bind to F 1 was discovered to be directly proportional to the ability of b to form dimers, suggesting the necessity of the b dimer formation before the binding of F 1 to the complex (199). The dimerization of the b subunit has been shown to be relatively weak and reversible. The monomeric and dimeric forms of b sol were shown to exist in a dynamic equilibrium and the dimer was converted to the monomeric state at 40C (192). This same melting characteristic was observed with CD spectroscopy (200). Furthermore, the similar traits were observed in photosynthetic organisms, which encode two different b-type subunits, b and b (13). When the cytoplasmic regions of the b and b subunits from the cyanobacterium Synechocystis were expressed individually, the polypeptides were found to only exist in the monomeric state. However, when they were mixed together,

PAGE 61

40 the formation of the dimers was observed by chemical crosslinking and sedimentation equilibrium ultracentrifugation (13). Also, the dimers were observed to melt at 40C as was the case with the E. coli b subunit. The striking similarity between the E. coli b subunit and the photosynthetic organism b and b subunits indicate that the former is a good model in which to study the b subunit. Cross-linking and deletion analysis has led to the development of a four-domain model of the E. coli F 1 F 0 ATP synthase b subunit (Figures 1-5 and 1-6) (12). Amino acid Figure 1-6. Gross structure of the E. coli F 1 F 0 ATP synthase and the domains of the b subunit. The b subunit domains were described by Dunn et al. (12) The membrane-spanning domain roughly corresponds to amino acid residues 1-22, the tether domain is approximately residues 24-60, the dimerization domain is considered to be residues 60-122, and the F 1 -binding domain is roughly residues 123-156.

PAGE 62

41 residues b 1-22 corresponds to the hydrophilic membrane spanning domain. Residues b 24-60 roughly corresponds to the tether domain, which is the portion of the peripheral stalk often observed in electron micrographs. The dimerization domain, approximately b 60-122 is required for the dimerization of the two b subunits. And finally, the F 1 binding domain, roughly amino acid residues b 123-156 is required for the binding of F 1 to F 0 The amino-terminal membrane spanning domain, b 1-22 forms a single transmembrane span while the large remainder is a polar hydrophilic domain which extends above the cytoplasmic leaflet of the lipid bilayer and reaches towards the top of F 1 (Figure 1-6) (13, 191). A wealth of evidence has suggested this proposed topology for the b subunit. The amino terminal region was uniformly vulnerable to chemical modification by the a nonpolar photoreactive reagent, TID, which is a hydrophobic carbene generator that is believed to react from the nonpolar region of the lipid bilayer, indicating that this region was in fact in the hydrophobic phase of the bilayer (106). This observation was consistent with other labeling procedures including the labeling of b cys21 by hydrophobic nitrenes (201) or the hydrophobic maleimide N-(7-dimethylamino-4-methyl-coumarinyl)-maleimide (DACM) (148). Modification of b cys21 interfered with intersubunit interactions within F 0 Furthermore, reconstitution of the F 0 subunits upon labeling the b subunit with DACM resulted in reduced proton translocation as well as F 1 binding affinities (148). TID failed to label the region b asn2gln10 indicating that the first few residues at the amino-terminus protrude into the periplasm (106) Despite attempts by several laboratories, there is presently no high-resolution structure of the entire b subunit. Therefore, model polypeptides have been constructed in order to elucidate the structure of the b subunit by domain. A model polypeptide

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42 comparable to residues b 1-34 which contains the membrane-spanning domain, dissolved in a 4:4:1 v/v mixture of chloroform/methanol/water previously used for solving the structure of subunit c, has been solved by NMR (136). The data revealed an -helical monomeric structure with a 20 bend at residues 23-26 (KYVW) (Figure 1-5). The hydrophobic residues, b 4-22 formed an -helix, followed by the 20 bend, and then resumed with -helical structure from residues b 27-34 The bend was proposed to be positioned as the b subunit exits the membrane. A series of cysteine substitutions resulted in a high yield of crosslinks formed at residues b 2 b 6 and b 10 (Figure 1-5). A lower yield of crosslinks were observed to form at residues b 3 b 8 b 9 and b 11 (Figure 1-5). No crosslinks were observed when cysteines were substituted for residues 12-21. The observance of continuous crosslinks between residues b 6-11 were suggested to indicate a dynamic interaction between the contacting faces of the two b subunits in this region (132). These observations led to a dimeric model in which the extreme amino-termini of the b subunits crossed each other in close proximity at an angle of about 35 in the region of residues b 4-11 and then the two b subunits angled apart as they traverse the membrane towards the cytoplasmic side (Figure 1-5) (136). The region of the 20 bend, b 23-26 was suggested to change the direction of the second -helix, b 27-33 such that it would extend into the cytoplasm at an angle perpendicular to the plan of the membrane. This model was confirmed by a systematic mutational analysis of the membrane-spanning domain performed by Hardy et al. (202). The tether domain of the b subunit, roughly b 24-60 is the least defined part of the subunit domains from a structural point of view. It corresponds to the portion of the peripheral stalk often seen in electron micrographs and is called tether simply because

PAGE 64

43 it is the section of the b subunit that links the more defined membrane-spanning and dimerization domains Figure 1-6). There is no high resolution structure for this region of the b subunit. The NMR structure of b 1-34 described above extended slightly within this domain, revealing an -helix at least up through residue b 34 .(136) Also, a heptad repeat, extending from just outside of the membrane and continuing without interruption up to residue b ala79 suggests the structure to be a coiled coil (197, 203). Though crosslinking studies showed that the tether domains of the two b subunits are in parallel and in close proximity, this domain contributes little to the stability of the dimerization of the b subunits (192). Deletion constructs analyzed by sedimentation equilibrium experiments suggested that a form of the b subunit truncated in each end, b 53-122 was capable of forming dimers with an efficiency close to the complete cytoplasmic domain, b 24-156 indicating that the most pertinent intersubunit contacts of the b subunit was located within this central region, referred to as the dimerization domain (192). More recently, the dimerization domain has been refined to residues b 63-122 ; however, the amino-terminal boundary is likely to decrease even further due to the observation that deletion of residues b 54-64 or b 65-75 resulted in intact and functional F 1 F 0 ATP synthase complexes (Figure 1-6) (194). A crystal structure of a monomeric dimerization domain, based on a model polypeptide consisting of residues b 62-122 has recently been solved and refined to 1.55 (138). Based on an undecad repeat and crosslinking data, Dunn and coworkers have constructed a model in which the two -helices of the b 62-122 region formed a coiled-coil with a right-handed superhelical twist. A number of previous studies supported this coiled-coil arrangement. First, the shape of the b 53-122 polypeptide was consistent with a

PAGE 65

44 coiled-coil of similar length as determined from its frictional coefficient (1.60) in an ultracentrifuge and from NMR relaxation parameters (192). Secondly, small-angle X-ray scattering by b 52-122 in solution specified a maximum length to be about 95 consistent with the expected coiled-coil length (13). Thirdly, CD spectroscopy indicated that this polypeptide was 100% -helical and the similar intensities of the minima suggested the helices to be arranged in a coiled-coil (204). Fourthly, cysteine substitution and crosslinking studies suggested a periodicity consistent with a parallel coiled-coil (Figure 1-5) (204). Finally, b subunit sequence analysis of E. coli and other prokaryotes revealed a conservation of an undecad pattern, which is a distinctive characteristic of a right-handed coiled coil. Mutation of amino acid residue b arg-83 which interrupts the undecad repeat, markedly stabilized the dimer, as expected for the proposed two-stranded, right-handed coiled-coil structure. The carboxyl-terminal F 1 -binding domain, b 124-156 also referred to as the -binding domain, has a more globular conformation and is required for the binding of F 1 to F 0 (205, 206). Work accomplished by Futai and coworkers two decades ago revealed that truncation of the extreme carboxyl-terminus of the F 1 -binding domain by only a few amino acids resulted in assembly defects in F 1 F 0 ATP synthase (206). Subsequently, work performed by Dunn and coworkers demonstrated that the final two to four amino acids of the b subunit were necessary for binding the subunit of F 1 (205). An addition of a cysteine at the carboxyl-terminus was chemically crosslinked to a cysteine introduced at 158 A close association of the two b subunits in the F 1 -binding domain was indicated by crosslinks formed between cysteines individually substituted at positions b 124 b 125

PAGE 66

45 b 126 b 127 b 128 b 129 b 130 b 131 b 132 b 139 b 144 b 146 or b 156 (Figure 1-5) (198, 200, 207). Hydrodynamic evidence favors a folded structure for this domain of the b subunit as opposed to the highly elongated structure of the remainder of the b subunit. Also, several studies have shown that either a b ala128glu mutation, deletion of the last four residues, or cold temperature dramatically decreased the sedimentation coefficient, by 23%, suggesting that the F 1 -binding domain underwent a conformational change from a globular structure to a less folded more extended conformation (192, 205, 208). The mutation, b ala128glu may have caused an electrostatic repulsion that would cause the two b subunits to push apart. The carboxyl-terminal residues may form an amphipathic helix, so the deletion would have disrupted essential interactions. And cold temperatures have been shown to weaken hydrophobic interactions in proteins, suggesting the importance of the hydrophobic amino acids in the folding of this domain (13). Dunn et al. suggested that these observations implied that the carboxyl-terminus of the b subunit has a weakly folded structure in which the hydrophobic amino acids are arranged to impart structural stability and create hydrophobic patches on the surface (13). The folded conformation appears to be required for the exposed hydrophobic patches to interact with F 1 Mutagenesis. Several mutant searches and site-directed mutagenesis studies have been performed in the membrane-spanning domain of the b subunit. However, only a single mutation, b gly9asp located near the periplasmic side of the lipid bilayer, resulted in a defective proton pore in an intact F 1 F 0 ATP synthase complex (209). Second site suppressors of this mutation have been found in the a(a pro240ala or a pro240leu ) and c (c ala62ser ) subunits that partially repaired the defect, indicating an interaction between the b subunit and both the a and c subunits (210, 211). The membrane-spanning domain

PAGE 67

46 contains one charged residue, b lys23 but mutations generated at this site did not influence proton translocation, suggesting that this domain of the b subunit did not have a direct function in proton conduction, although it was required for the maintenance of a functional F 0 complex (212). A systematic mutational study of the membrane-spanning domain conducted by Hardy et al. was described above and a triple mutant, b N2A,T6A,Q10A is described in Chapter 5 of this dissertation (202). A couple notable characteristics can be attributed to the tether region of the b subunit. Relatively large deletions and insertions of up to 11 and 14 amino acids, respectively, were accommodated in this region and the altered b subunits still assembled into fully functional F 1 F 0 ATP synthase complexes (193, 194). In fact, decreases observed in enzyme activity paralleled the decrease of b subunit found in the membrane, suggesting that the alterations affected assembly of the enzyme, but not the function. Assuming -helical structure, an 11 amino acid deletion would shorten the b subunit tether region by approximately 16.5 and a duplication of 14 amino acids would increase the length by about 21 This implies that the b subunit is highly flexible and the altered b subunits may compensate for the lost or gained distance via that flexibility. The fact that the peripheral stalk must extend from within the membrane to up near the top of F 1 suggests that some part of the stalk must be flexible enough to stretch or straighten in the shortened b subunits, or in the case of the lengthened b subunits, bend to take up the slack. Prior to these observations, the b subunit was often viewed as a rigid, rod-like structural feature during rotational catalysis. Also, though the b subunit is the least conserved subunit of F 1 F 0 ATP synthase, gaps are rarely found in sequence alignments of numerous organisms (213, 214). Therefore, the ability to manipulate the

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47 Figure 1-7. Model for F 1 F 0 ATP synthase peripheral stalk orientation dependent upon the direction of rotation during ATP synthesis or hydrolysis. The subunits are color coded as follows: light blue; grey; dark blue; green; orange; a, yellow; b, red; c, cyan. The panel on the left indicates the orientation of the b 2 dimer if the enzyme is actively synthesizing ATP. The panel on the right represents the position of the b 2 dimer during ATP hydrolysis. The arrows indicate the direction of rotation of the rotor stalk subunits (c 10 ). The red cylinders indicate regions of the b subunits for which there is no high-resolution structure.

PAGE 69

48 length of the peripheral stalk was an unexpected surprise to the field. The length of the wild-type b subunit is probably the optimum length for assembly of F 1 F 0 ATP synthase. The apparent flexibility has been proposed to help alleviate torsional strain brought about by rotational catalysis (193). Another hypothesis concerning the flexibility of the tether domain is that this region of the b 2 dimer serves as a hinge, allowing reorientation of the stator depending on the direction of rotation as the enzyme carries out ATP synthesis or hydrolysis (Figure 1-7) (195, 202). Another important feature of the tether domain is the evolutionarily conserved b arg36 that has been implicated in a structural role influencing proton conduction through F 0 Mutational studies at this amino acid residue led to numerous defects from failure to assemble or function to uncoupling phenotypes (215). Amino acid substitutions, b arg26ile or b arg26glu resulted in assembled F 1 F 0 ATP synthase complexes that displayed defects in F 0 -mediated proton translocation or a disruption of coupling activity, respectively. Substitution with a cysteine at this residue led to a crosslink product with the a subunit of F 0 (216, 217). The close proximity of this conserved residue to the a subunit indicates that it may play a role in aligning the proton exit channel. Protein-protein interactions between the two b subunit dimerization domains have been shown to be essential for forming the peripheral stalk (13). Mutations at a conserved residue, b ala79 resulted in major F 1 F 0 ATP synthase assembly defects (203, 218). The b ala79 mutations were modeled in the b sol polypeptide to investigate the affects of the mutations. The model polypeptides were shown to retain -helical structure, but chemical crosslinking and sedimentation experiments suggested that the b ala79 mutants were incapable of forming dimers (199).

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49 In the F 1 -binding domain, a mutation was found, b ala128asp that had little effect on the dimerization of the b subunits but led to an assembly defect of F 1 F 0 ATP synthase (208). However, the mutant was found to have a reduced tendency to interact with F 1 and sedimentation equilibrium ultracentrifugation experiments revealed a 12% decrease in the sedimentation coefficient, indicating a structural perturbation (discussed above). The studies suggested that the b ala128 residue was not important in b subunit dimerization, but it had an important structural role in the F 1 -binding domain. Intersubunit interactions. The formation of disulfide bonds between cysteine residues introduced in the membrane-spanning domain of the b subunit and the a subunit as well as second site suppressors of the b gly9asp found in the a subunit (discussed above) strongly suggests an interaction between the two stator subunits (209-211). The b gly9asp mutation was also partially suppressed by a mutation in the c subunit (discussed above), but it is not known whether it is due to a direct interaction between the b and c subunits or if the suppression is mediated through the a subunit. The b subunit has also been shown to interact with the and subunits of F 1 (198, 216). The interaction of the F 0 b subunit with the F 1 subunit has been well documented (200, 205, 219, 220). The interaction is mediated by the carboxyl-termini of both subunits and is essential for the binding of F 1 and F 0 The critical role of the subunit mediating the interaction between the b subunit and the bulk of F 1 has been demonstrated by the inability of -depleted F 1 to bind to F 0 (219). Crosslinking the b and subunits via introduced cysteines did not affect F 1 F 0 ATP synthase activity, which was consistent with the proposed role of the b 2 peripheral stalk as a stator and demonstrated that the binteraction need not be dynamic (221). Though it is believed that the binding

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50 of F 1 to F 0 heavily relies on the binteraction, evidence of other subunit contacts may also influence binding. Furthermore, the binding of b to was shown to be relatively weak by analytical ultracentrifugation, indicating that other subunit contacts may contribute to binding (220). Many crosslink formations were found when cysteines were introduced into the b and or subunits (Figure 1-5). A cysteine introduced to the carboxyl-terminus of the b subunit has been shown to crosslink to cys90 (198). Also, cysteines positioned at b 92 or b 109 formed crosslinked products with the subunit or both the and subunits, respectively (216). These results confirm that the b subunit is proximal to the 3 3 hexamer, but a direct interaction has not been confirmed. Stator stalk function. The necessity of a stator in F 1 F 0 ATP synthase was recognized after the realization that rotation was a fundamental feature of ATP catalysis. The current view of the peripheral stalk is primarily that of the stator which forms a connection the a subunit and the 3 3 hexamer, holding these subunits in place against the rotation of the rotor subunits, c 10 The idea of a flexible stator stalk has led to other proposed features of the b subunit. The apparent flexibility of the b subunit has been suggested to transiently store energy during rotational catalysis which could be potentially be expended to force the conformational change that allows the release of ATP(222). Another model describes the flexibility of the tether region as a hinge that could reorient the b subunit when switching between ATP synthesis and hydrolysis (discussed above) (Figure 1-7) (202). However, there is no direct evidence of whether the b subunit is actually acting in a flexible manner. Finally, the b subunit has recently been suggested to influence the nucleotide binding sites in the subunits (223). In these experiments, a spin label was incorporated at residue 331. Upon addition of b sol the

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51 spectrum of this spin label was observed to change in a way that implied that the catalytic sites were in a more open conformation. These results indicate that the current view of the stator stalk as a structural feature may soon be revised such that the b subunit has a more direct role during rotational catalysis. Subunit Equivalence The E. coli ATP synthase complex has, by far, the simplest architecture of all the F 1 F 0 ATP synthase enzyme complexes. It is composed of twenty-two polypeptides of eight different types with the stoichiometry 3 3 ab 2 c 10 (Figure 1-2, Table 1-1) (112, 224). The 3 3 subunits comprise the F 1 sector and the ab 2 c 10 subunits comprise the F 0 sector. In the E. coli enzyme, all eight subunits are necessary for the function of F 1 F 0 ATP synthase (225, 226). Chloroplasts also have a relatively simple architecture with the exception that they have nine different subunits (227) due to the fact that the two b subunits are products from two different genes and are not identical (Figure 1-8). In contrast, the F 1 F 0 ATP synthase from mammalian mitochondria is composed of at least thirty-one polypeptides of sixteen different types with the F 1 stoichiometry of 3 3 and a much more complex F 0 consisting of a, b 2 c 10-14 d, e, f, g, (F 6 ) 2 A 6 L, OSCP, IF 1 (7, 228-230). Yeast mitochondria F 1 F 0 ATP synthase has an extra three subunits compared to the mammalian enzyme, stf1p, i, and k. The F 1 sector. In the F 1 sectors, homologues of the E. coli F 1 F 0 ATP synthase have been identified for the and subunits, based on amino acid sequence homology, in the chloroplast and mitochondrial enzymes (30). Based on the primary sequences, the highest conserved subunit from the E. coli F 1 F 0 ATP synthase is the subunit with approximately 70% homology with the chloroplast and mitochondria

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52 Table 1-1. F 1 F 0 ATP synthase subunit equivalency Mitochondria Bacteria Chloroplast Yeast Bovine Function catalytic site catalytic site rotor OSCP OSCP stator rotor stabilization? a a (or IV) a (or 6) a proton channel, stator b b (or I) b (or 4) b stator b (or II) 9 stator c c (or III) c (or 8) c proton channel, rotor d d stator? 8 A6L stator? e e ? f f ? g g ? h F6 stator inh1p IF1 inhibitor stf1p ? j ? k ? equivalents (31). The subunits exhibit roughly 50% homology (31). The nucleotide binding regions of these two subunits also have sequence homologies with other proteins that bind nucleotide or phosphate, including the E. coli secA protein, N-ethylmaleimide sensitive fusion protein, herpes simplex virus UL15, Ca 2+ -ATPase, H + /K + ATPase and Na + /K + ATPase (34-37). Furthermore, the nucleotide binding motif, GXXXXGKT/S, which was first identified in the and sequences of F 1 has been found to be conserved in the high-resolution structures of other proteins including p21 ras adenylate kinase, RecA, elongation factor Tu, and transducin(20, 38-42). Interestingly, the subunit in the chloroplast F 1 F 0 ATP synthase complex contains an insert of about 35 amino acids that is not present in the mitochondrial or

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53 nonphotosynthetic eubacteria (231). This loop contained two cysteine residues that were found to be reduced in the active enzyme complex during photosynthesis and oxidized to a disulfide bond in the inactive enzyme while in the dark (232). The E. coli subunit is unfortunately known as the subunit in the mitochondrial enzyme (233) and also shares primary sequence homology with the mitochondrial IF 1 inhibitor protein. The E. coli and bovine subunits share 60% sequence identity, which had suggested that they are functionally equivalent (191, 234). The availability of high resolution structures for these subunits has revealed that they are strikingly similar. Superimposition yields a 1.64 rms deviation (48). The bacterial subunit has been suggested to be an inhibitor of ATP hydrolysis, undergoing large, ratchet-like conformational changes to selectively switch off ATP hydrolysis (102). In mitochondria, this inhibitor action of the bacterial subunit is ascribed to the IF 1 protein of the F 0 sector. It was suggested that the bacterial subunit was separated into the two polypeptides in the mitochondrial enzyme complex, and IF 1 Finally, the bacterial and chloroplast subunits of F 1 share a significant sequence homology (234). The subunits equivalent in the mitochondrial F 1 F 0 ATP synthase is known as oligomycin-sensitivity-conferring protein (OSCP) (235-237). The carboxyl-terminal region of the mitochondrial b subunit was been demonstrated to bind to the OSCP subunit (E. coli subunit) through subunit interactions (Collinson et al., 1994) and chemical crosslinking analysis (Soubannier et al., 1999). The F 0 sector. The F 0 sectors of the F 1 F 0 ATP synthase family is by far more diverse than the F 1 sector with an additional eight different subunit types in mammalian and an extra ten different subunits in yeast mitochondria (Table 1-1). The E. coli a and c subunits are respectively equivalent to the chloroplast IV and III subunits and the

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54 mitochondrial ATPase-6 and ATPase-8 subunits. In every case, both subunits are hydrophobic proteins required for proton translocation (120, 140). Unlike the a and c subunits, the E. coli b subunit does not have any obvious homologues in chloroplasts or cyanobacteria F 1 F 0 ATP synthases. However, both of them have two distinct subunits with similar hydrophobic and hydrophilic residue distribution (238). These subunits are referred to as subunits b and b in cyanobacteria and subunits I and II in chloroplasts. It is believed that only one of each of these subunits is present in the F 1 F 0 ATP synthase complex, incorporating into the enzyme as a b-b heterodimer as opposed to the b subunit homodimer present in E. coli. No obvious homologue of the E. coli b subunit has been found in the mitochondrial F 1 F 0 ATP synthase, even upon analysis of sequence, function or three-dimensional structure (239). However, hydropathy plot analysis does indicate that the mammalian mitochondrial b subunit that may be analogous to the E. coli b subunit (240, 241). Proteolysis studies and crosslinking data supported the location of the mitochondria b subunit at a position analogous to the E. coli b subunit (242, 243). At least three other subunits may play the role of the E. coli b dimer including subunit 8 (or A6L), d and F6. The mitochondrial b subunit is believed to have two -helical transmembrane spans at its amino-terminus arranged in an antiparallel configuration. The extreme amino-terminus of the mitochondrial b subunit is thought to begin on the cytoplasmic side of the membrane, traverses the membrane to the periplasmic leaflet of the membrane as an -helix, then turn back and traverses the membrane again as it exits the membrane in the cytoplasm and reaches towards the top of F 1 (Figure 1-8). The remainder of what would be equivalent to the E. coli b dimer is highly speculative, though the overall shape and characteristics of the b8dF6 subunits favor this explanation

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55 (Figure 1-8). In this model, the mitochondrial subunit 8 contributes to a third membrane spanning region, and a combination of subunit d and subunit F6 forms the hydrophilic domain. The F 0 subunit known as the subunit in mitochondrial F 1 F 0 ATP synthase has no counterpart in the bacterial or chloroplast enzymes. It is a small polypeptide (50 amino acids) folded into a helix-loop-helix. It is believed to play a role in the stabilization of the central stalk and its absence in the bacterial and chloroplast enzymes may explain why Figure 1-8. Speculative models for the b-like subunits. Shown are models for the bacterial, chloroplast and mitochondrial b subunits. The membrane-spanning regions are indicated by the black lines. An abundance of evidence supports the parallel arrangement of the bacterial b 2 homodimer and the chloroplast bb heterodimer. The mitochondrial analogue of the b subunit is believed to consist of up to four polypeptides. The model shown for the mitochondrial b8dF 6 structure is highly speculative.

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56 the bacterial subunit of the F 1 sector (equivalent to the subunit in mitochondria) easily dissociates from the F 1 complex whereas this has not been observed for the mitochondrial enzyme. Mitochondrial F 0 consists of several additional subunits not found in the E. coli or chloroplast enzymes including the e, f and g subunits as well as an extra three in yeast, stf1p, j, and k subunits. E. coli F 1 F 0 ATP synthase as a model. Initial studies of the F 1 F 0 ATP synthase complex were achieved with enzymes isolated from mitochondria or chloroplast. Although the bacterial, chloroplast and mitochondrial enzymes differ in oligomeric complexity, the enzymes show acceptable overall structural resemblance and primary sequence homology that it is widely accepted that the mechanism of action is the same in all organisms (240, 244). Therefore, studies using the bacterial model became widely accepted since it was more versatile and offered a large range of research that could not be readily undertaken with the more complex organisms. Other advantages include ease of genetic techniques, the ability of bacteria to grow via glycolysis, which allowed characterization of defective F 1 F 0 ATP synthases, and ease of large-scale purification procedures due to a practically unlimited supply of bacteria. F 1 F 0 ATP Synthase Mechanism The overall function of F 1 F 0 ATP synthase can be divided into three distinct parts: proton conduction, coupling, and catalysis. The a and c subunits of F 0 are responsible for the translocation of protons through the membrane. The F 1 and subunits of the rotor stalk are responsible for coupling the energy acquired from the proton gradient to the F 1 catalytic sites. And the three catalytic sites located at the interfaces of the three and

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57 subunits are responsible for the synthesis of ATP or sometimes, as in the case of bacteria, ATP hydrolysis. All three functions must be tightly integrated for the production of ATP. Proton Translocation: Driving Rotation The demonstration that the electrochemical gradient of protons drives the rotation of bacterial flagella (245) in combination with Peter Mitchells chemiosmotic theory (2) began the search for evidence of rotation in F 1 F 0 ATP synthesis. At the same time, a model for proton transport was suggested by Cox et al. (212) and Boyer developed his ideas for the binding change mechanism (discussed below) (246). But an indication of rotational catalysis was not evident until the high-resolution crystal structure of F 1 became available (20). This was followed a few years later by the first direct observation of rotation when Noji et al. fixed the top of F 1 to a glass coverslip and attached a fluorescently labeled actin filament to the subunit (15). Upon addition of ATP, rotation of the actin filament was observed under an optical microscope at 0.2-10 revolutions per second. At low concentrations of ATP (<600 nM), the actin filaments were observed to rotate in a step-wise manner at 120 intervals, which reflects the three catalytic sites in the F 1 3 3 hexamer (247). Experiments, in which two phenylalanine residues in the nucleotide binding pocket were mutated to reduce the binding affinity of ATP, indicated that the binding and hydrolysis of ATP is initially accompanied by a 90 substep followed by a 30 substep attributed to product release (248, 249). These observations were consistent with the two proposed catches observed between the and subunits in the high resolution structure (see The subunit above) (20). The observance of the rotation of the and c subunits at the same speed and direction, indicating that these three subunits rotate in synchrony, forming the central rotary machinery of the enzyme

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58 complex (65-67). The concept of rotational catalysis with the rotation of the c 10 subunits relative to the 3 3 hexamer is now well accepted. Several models have been proposed for proton translocation. One of the earliest models suggested a series of side chains spanning the lipid bilayer formed a proton wire, involving amino acid residues c asp61 a arg210 a glu219 and a his245 in which the protons hop from one side chain to another until it passes through the membrane (212, 226, 250). Other models include a water-filled proton channel model, formed by the charged residues of the a and c subunits, in which hydronium ions (H 3 O + ) pass through the lipid bilayer (251) and a proton carrier model, in which a proton binds on the exterior of the membrane followed by a conformational change that brings the complex through the membrane and releases the proton on the outer surface (252, 253). The current prevailing model suggests that protons, in the form of H 3 O + enter a half channel created by the a subunit (Figure 1-9) (254, 255). The H 3 O + is then believed to protonate one of the c 10 subunits at residue c asp61 which is positioned near the center of the lipid bilayer (131). Besides forming the proposed half channel, the a subunit is thought to play another crucial role in the protonation of the c asp61 The pKa of the c asp61 carboxyl side chain is uncommonly high, which is likely due to its lipid environment (125). The essential a arg210 residue is thought to facilitate a pKa shift of the c asp61 carboxyl side chain to a lower pKa form during proton translocation (14, 132). Either the protonation of the carboxylate, or possibly the release of a proton from a previously protonated c asp61 into an exit channel housed by the a subunit, somehow promotes the generation of torque (Figure 1-9). The torque produced by proton translocation is believed to drive the rotation of the ring of c 10 subunits relative to the a and b subunits

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59 during rotational catalysis. Translocation of three to four protons generates enough torque energy to rotate the c 10 ring by 120 and results in the synthesis of a single molecule of ATP. Figure 1-9. Model of proton translocation and torque generation in F 0 Subunits included are (green), (orange), a (yellow), b 2 (purple), c 10 (blue). Protons (red) are traveling in the direction of ATP synthesis. Protons are believed to travel through a half channel housed in the a subunit as hydronium, H 3 O + An essential residue located near the center of the lipid bilayer in the c subunit, c asp61 is thought to be protonated as another proton exits through another exit half channel located on the cytoplasmic side of the membrane. The protonation/deprotonation drives the rotation of the c 10 ring. The model was drawn from schemes proposed by Junge (254) and Vik and Antonio (255).

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60 Coupling In F 1 F 0 ATP synthase, the mechanism of energy coupling requires both the rotor stalk and the peripheral stalk. Rotation of the ring of c 10 subunits consequently results in the rotation of the entire rotor stalk, which is essential in coupling the energy obtained from proton translocation to the synthesis of ATP in the catalytic sites of the 3 3 hexamer located over 100 away. The role of the peripheral stalk is to hold the hexamer in place while the aminoand carboxyl-terminal-helices of the subunit rotates within. In a fully functional and coupled enzyme, the kinetics of proton translocation and ATP synthesis are linked so that one cannot proceed without the other, and vise versa (256). Mutational studies suggested that the polar loop of the c 10 subunits was involved in the coupling function. In F 1 F 0 ATP synthases incorporated with the c gln42glu subunit mutant, F 1 was found to bind normally to the F 0 mutant, but the passive leakage of protons through this complex was not prevented as in must be in coupled enzyme complexes (257). Also, ATP was hydrolyzed normally by this mutant, but hydrolysis was not coupled to active proton translocation. A similar uncoupled phenotype was found in complexes incorporated with a c arg41lys mutant (258). Another mutation at the same site, c arg41his was found to prevent the binding of F 1 to F 0 Thus, this loop region in the c subunit appears to play essential roles in both the binding of F 1 and the coupling of proton translocation to ATP synthesis. Second site suppressor mutations to the c gln42glu mutation were found in the subunit, specifically, glu31gly,val,lys that recouped proton translocation and ATP synthesis (98). Crosslinking studies of cysteine double mutants found cross-linked products between cys31 and c cys40 c cys42 and c cys43 (99). Moreover, a functional contribution of the subunit in energy coupling was demonstrated

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61 by subunit mutants that uncoupled proton transport and ATP hydrolysis (259). Second site suppressor mutations were found in other regions of the subunit (260, 261). Cysteine substitutions formed crosslinks between the subunit, cys205 and c cys40 c cys42 and c cys43 (118). Crosslinking studies also provided evidence for an interaction between and (75, 77). In addition, the subunit has been cross-linked to both the F 1 and F 0 subunits, via introduced cysteines, indicating that it spans the entire length of the stalk with the subunit (60, 99). Combined, these observations had suggested that the coupling mechanism occurred by direct interactions of the c subunit loop regions and the and rotor stalk of F 1 which convey the proton gradient energy to the catalytic sites, probably by direct interactions. The crystal structure later confirmed these interactions (20). The crystal structure displayed a strikingly asymmetrical F 1 due to differences in the domains of the and subunits and the interactions formed with the single subunit (20). The obvious asymmetric positioning of the coiled coil of the subunit is a key feature to the mechanics of the binding change mechanism (discussed below) of F 1 F 0 ATP synthase. Its large carboxyl terminus -helix passes through a hydrophobic sleeve formed by six proline-rich loops of the and subunits, presumably resulting in the conformational changes occurring in the catalytic sites (20). In the E subunit (see above), several hydrogen bonds are formed with the subunit which forms a catch, resulting in conformational changes. Specifically, residues arg254 and gln255 in the carboxyl terminal helix form hydrogen bongs with E-asp317 E-thr318 and E-asp319 Also, a second catch is formed between the carboxyl terminal domain of the T subunit and the short helix of the

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62 subunit. Hydrogen bonds form between lys87 lys90 and ala80 within the DELSEED region, T-asp394 and T-glu398 Structural information suggests the two antiparallel coiled coil -helices of the subunit may unwind during rotational catalysis and the subunit rotates around the F 1 axis while undertaking a net translation of about 23 (85). It is likely that these gross changes observed in the structures revealed individual functional states of the enzyme complex during catalysis. Catalysis: The Binding Change Mechanism F 1 F 0 ATP synthases house three catalytic sites, located at the 3 3 interfaces, with the predominate sites positioned in the subunit and some contributions made by the subunits (20). The minimal complex capable of normal ATP hydrolysis activity is the 3 3 complex (262). Boyer predicted that ATP synthesis requires a chronological involvement of the three catalytic sites, each of which changes its binding affinity for the substrates and products as it continues through a cyclical mechanism, referred to as the binding change mechanism (246, 263). The mechanism of ATP synthesis, in terms of the 3 3 and subunits and the substrates, ADP and P i is described here (Figure 1-10). The principles of the binding change mechanism have become the most commonly used model for recounting the means of ATP synthesis by F 1 F 0 ATP synthase. The three distinct catalytic sites were described as the tight (T) site containing ATP, an empty open (O) site, and the loose (L) site illustrated with ADP and P i bound (Figure 1-10). According to Boyer, catalysis starts with the binding of ADP and P i at the open site. The energy input from proton translocation drives the rotation of the rotor stalk, which ultimately results in the conformational changes responsible for ATP synthesis such that the tight site is converted to an open site, the open site assumes a loose site conformation,

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63 and the loose sites becomes a tight sight. ATP is formed in the new tight site and the molecule of ATP that was found in the original tight site is released and the binding change mechanism starts fresh (Figure 1-10). Boyers mechanism included three proposals: i) only one site is actively synthesizing ATP at any given moment, ii) the reaction occurs reversibly at this site, and iii) energy input is required to bind the substrates, ADP and P i into the catalytic sites and to release the synthesized ATP, but not for the actual reaction to occur. Strong evidence for this model has come from the crystal structure of F 1 (20) and the observance of rotation via an attached fluorescent actin filament (discussed previously) (15, 65-67). Figure 1-10. The binding change mechanism. This is a simplified model of a more detailed enzymatic mechanism described by Weber and Senior (189) in which all three catalytic sites are transiently filled with nucleotide during ATP synthesis. Subunits included are (red), (green) and (pink). Genetic Expression and Assembly The E. coli F 1 F 0 ATP synthase is encoded in the 7 kb unc operon, which was cloned and sequenced in its entirety (224, 264). The genes encoded are called the uncB,

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64 uncE, uncF, uncH, uncA, uncG, uncD and uncC coding for the a, c, b, , and subunits, respectively. A single copy of each gene is transcribed into a single polycysternic mRNA transcript, from which multiple polypeptides can be translated (265-267). The synthesis of the correct number of subunits, resulting in the stoichiometry of 3 3 ab 2 c 10 is thought to occur by translational regulation (264, 268). The efficiency at which the individual subunits are synthesized were observed to be variable and roughly corresponded to the stoichiometry of the intact enzyme complex (269, 270). Far less is understood about the assembly of the complex F 1 F 0 ATP synthase enzyme compared to the structural and mechanistic studies. Some have proposed that no particular pathway is necessary based on the observation that the complex can be dissembled into individual subunits and then reconstituted in vitro (113, 148, 271). On the contrary, some believe that the assembly follows an integrated pathway in vivo (272). The idea of an assembly pathway appealed to many since it would prevent a newly assembled F 1 sectors from freely hydrolyzing cellular ATP in the cytoplasm, and newly assembled F 0 sectors from acting as open proton pores in the membrane. If assembly were a random event, the potential to create isolated F 1 and F 0 sectors would exist. Some evidence supporting the integrated pathway does exist. The subunit is thought to function as an inhibitor of ATP hydrolysis activity, undertaking large conformational changes to allow the enzyme to switch from ATP synthesis to ATP hydrolysis under conditions of low ATP or low proton gradient, respectively (102, 177, 273-276). Its inhibitory action may possibly act to inhibit free ATPase activity in the partially assembled state. Some have speculated that the binding of the F 1 subunits to F 0 may

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65 influence the opening of the F 0 proton channel in vivo (176, 277-279). The a subunit was observed to be absent from membranes of cells lacking the b or c subunits (150). Furthermore, work accomplished in our laboratory has showed that the b subunit monomer does not integrate into the membrane has no affinity for F 1 indicating that the formation of the b dimer in the membrane is an early event in F 1 F 0 ATP synthase assembly (199, 203, 218). Summary There is now ample evidence indicating that F 1 F 0 ATP synthases are composed of three functional parts, the catalytic core, the rotor stalk and the stator stalk. In the E. coli enzyme complex, the catalytic core consists of the 3 3 subunits and functions as an ATP synthase or an ATPase. The rotor stalk consists of the c 10 subunits and couples the energy of proton conduction to the synthesis of ATP by rotating within the 3 3 hexamer. Finally, the stator stalk consists of the ab 2 subunits and remains in a fixed position, anchored to the membrane by the a and b 2 subunits and to the 3 3 hexamer via interactions made by the subunit. Much of what was known of F 1 F 0 ATP synthase has been irrefutably confirmed by high resolution structures of partial complexes or model polypeptides. To date, a high resolution structure of the complete enzyme, or at least the complete F 0 is eagerly anticipated. Since the visualization of the peripheral stalk less than a decade ago a plethora of data characterizing the b 2 homodimer has emerged. The observation that the b 2 homodimer was likely not a rigid structure, and possibly more of an elastic structure, created the foundation for the work described in the following chapters. The work illustrated in Chapters 2, 3, 4, 5 and 6 of this dissertation will characterize the role of the

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66 peripheral stalks dimer of identical b subunits in the E. coli F 1 F 0 ATP synthase by using a combination of site-directed mutagenesis and biochemical methods. Chapter 2 demonstrates the ability of the E. coli b subunit to form heterodimers and the capability of F 1 F 0 ATP synthase complex to tolerate the incorporation of two different length b subunits with a size difference of at least 14 amino acids (195). Chapter 3 demonstrates the formation of b heterodimers including at least one and up to two defective b subunits and documents indisputable evidence that F 1 F 0 ATP synthases incorporated with b subunit heterodimers are functional (280). Furthermore, the work accomplished in the chapter indicates, for the first time, that each of the two b subunits makes a unique contribution to the functions of the peripheral stalk, such that one mutant b subunit is making up for what the other is lacking. Chapter 4 describes cysteine chemical modifications constructed in the subunit and shortened, lengthened and wild-type length b subunits. The unc operon expression plasmids generated in this study will be used in future fluorescent labeling experiments. Chapter 5 documents mutagenesis experiments conducted on the extreme aminoand carboxyl termini of the b subunit (202) (Bhatt et al., manuscript in preparation, 2004). Finally, Chapter 6 summarizes the conclusions of this study and suggests the future directions that the work described in this dissertation has offered.

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CHAPTER 2 INTEGRATION OF UNEQUAL LENGTH b SUBUNITS INTO F 1 F 0 ATP SYNTHASE Introduction F 1 F 0 ATP synthases provide the bulk of cellular energy production in both eukaryotes and prokaryotes (3, 5, 6). Enzymes in this family utilize the electrochemical gradient of protons across membranes in order to synthesize ATP from ADP and inorganic phosphate in a coupled reaction (16). In Escherichia coli, F 1 F 0 ATP synthase is a complex enzyme composed of approximately twenty-two polypeptides with the stoichiometry of 3 3 ab 2 c 10 (6, 7). The F 1 portion is composed of the subunits 3 3 and is responsible for enzymatic catalysis. The F 0 portion of the enzyme consists of the ab 2 c 10 subunits and is responsible for the translocation of protons through the membrane. Electron microscopy has shown that the F 1 and F 0 sectors are linked by two slender stalk structures (11). During ATP synthesis proton translocation drives the rotation of the central stalk, which consists of subunits within the 3 3 hexamer held stationary by the peripheral stalk. This rotation propagates the conformational changes in the active sites located at the interfaces driving catalytic activity (3, 15, 62, 65, 281, 282). The subunit of F 1 and a dimer of two identical b subunits from F 0 comprise the peripheral stalk acting as the stator. The subunit has been visualized seated at the top of the F 1 3 3 hexamer (187). However, recent evidence has suggested that the subunit may be positioned slightly to the side of F 1 in association with a single subunit (100, 141-143). 67

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68 The C-terminal region of the subunit is in direct contact with the extreme C-terminal end of the b dimer (3, 185, 200, 219, 220). The b subunit dimer constitutes the majority of the peripheral stalk stretching from within the membrane to near the top of F 1 (12). Dimerization of the b subunits is required for the normal assembly and function of F 1 F 0 ATP synthase (199). The two b subunits are believed to exist in parallel as an extended structure spanning from the periplasmic side of the membrane to near the top of F 1 Each has a N-terminal transmembrane domain, a tether domain extending from the surface of the membrane to the bottom of F 1 a dimerization domain and a -binding domain (13). The ability of b to bind to F 1 was proportional to the ability of b to form dimers, suggesting the necessity of the b dimer formation before the binding of F 1 to the complex (199). Presently, there is no high-resolution structure of the entire b subunit. A model polypeptide of the first 34 residues of the N-terminus has been solved by NMR, revealing a hydrophobic membrane-spanning -helix (136). A crystal structure of a monomeric dimerization domain, consisting of residues 62-122, has been solved and refined to 1.55 (138). Dunn and coworkers have constructed a model in which the two -helices of the b 62-122 region form a right-handed coiled coil. Much of the structural information on the b dimer has been gleaned from classical biochemical approaches such as CD-spectroscopy, crosslinking and sedimentation experiments (30, 166, 192, 196, 197, 203, 283). These studies revealed that the overall structure of the b subunit dimer is a highly extended conformation with approximately 80% -helix. Previous studies have shown that b subunits with deletions of up to eleven amino acids and insertions of up to fourteen amino acids, corresponding to approximately 16 and 21 respectively, formed functional F 1 F 0 complexes (193, 194). When b subunits

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69 with either a seven amino acid deletion or an insertion, b 7 or b +7 respectively, were incorporated into the F 1 F 0 ATP synthase complex, the properties of the enzymes were essentially wild type. These observations suggested that the role of the b dimer is more of a flexible structural feature. However, it was not known whether this flexibility extended to the dimerization of two b subunits of unequal lengths and their incorporation into an enzyme complex. In the present study an experimental system was developed to allow expression of two different b subunit genes and determine whether the differing b subunits were assembled into an F 1 F 0 ATP synthase complex. Here, we demonstrate that the F 1 F 0 ATP synthase complex can tolerate b subunits with a size difference of at least 14 amino acids. Materials and Methods Materials Molecular biology enzymes and mutagenic oligonucleotides were obtained from Invitrogen (Carlsbad, CA), Life Technologies, Inc. (Grand Island, NY), New England Biolabs (Beverly, MA) and Stratagene (La Jolla, CA). Reagents were obtained from Sigma (St. Louis, MO), BioRad Laboratories (Hercules, CA) and Fisher Scientific (Pittsburgh, PA). Plasmid purification kits were acquired from Qiagen Inc. (Valencia, CA). The anti-rabbit immunoglobulin horseradish peroxidase-linked whole antibody (from donkey), anti-mouse immunoglobulin horseradish peroxidase-linked whole antibody (from sheep), Hybond ECL Nitrocellulose membrane, electrochemiluminescence Western blotting reagents and high performance chemiluminescence film were purchased from Amersham Biosciences (Piscataway, NJ). Polyclonal antibodies against SDS-denatured b subunit (284, 285) were generously

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70 provided by Dr. Karlheinz Altendorf (Universitt Osnabrck, Osnabrck, Germany). Mouse monoclonal antibodies against the peptide epitope of hemagglutinin protein of human influenza virus (HA epitope tag) were purchased from Roche Molecular Biochemicals (Indianapolis, IN). Monoclonal antibodies against the epitope found in the P and V proteins of the paramyxovirus, SV5 (V5 epitope tag) were purchased from Invitrogen. Strains and Media The bacterial strains used to create the epitope tagged b subunits include the wild type b subunit expression plasmid, pKAM14, and plasmids used to express b subunits shortened or lengthened by 7 amino acids, pAUL3 and pAUL19, respectively, and have been described previously (193, 194, 203). The plasmids encoding the different uncF(b) genes were used to compliment E. coli strain KM2 (b) carrying a chromosomal deletion of the gene (218). All strains were streaked onto plates containing Minimal A media supplemented with succinate (0.2% w/v), to determine enzyme viability. Cells harvested for membrane preparation were grown in Luria Bertani media supplemented with glucose (0.2% w/v) (LBG). Isopropyl-1-thio--D-galactopyranoside (IPTG)(40 g/ml), ampicillin (Ap) (100g/ml), and chloramphenicol (Cm) (25 g/ml) were added to media as needed. All cultures were incubated at 37C for the appropriate duration. Recombinant DNA Techniques Plasmid purification. Plasmid DNA was purified with the Qiagen Mini-Prep and Maxi-Prep kits according to the protocols provided from the manufacturer. Mini-preps required 4 mL (high copy plasmid) or 6 mL (low copy plasmid) of an overnight bacterial culture carrying the desired plasmid. Maxi-preps required a 500 mL culture grown

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71 overnight. Final elution volumes for the mini-prep kit was 30 or 50 L, for low copy and high copy plasmids, respectively, and 200-250 L for the maxi-prep kit. Final concentrations of 0.5 and 1.0 g/l plasmid were routinely obtained with the Qiagen mini and maxi-prep kits. Digestions, ligations, and transformations. Restriction endonuclease digestions, ligations, and transformations were performed according to the recommendations of the manufacturers (New England Biolabs, Stratagene and Invitrogen). For analytical purposes, restriction endonuclease digestions were normally prepared in a total volume of 20 L, including plasmid DNA, enzyme, buffer, ddH 2 O and occasionally BSA, and then incubated for an hour at the temperature specified by the manufacturer. Ligations required two purified double-stranded DNA fragments, a vector and an insert, of known length and concentration (ng/L). DNA fragments were routinely separated in a 0.8 % agarose gel by electrophoresis and the appropriate sized fragment was excised and purified using a Qiagen, Inc. QIAquick Gel Extraction kit. The vector fragment contained the antibiotic resistance gene and the origin of replication. The insert typically contained the desired gene or a site specific mutation. Two control reactions and two ligation reactions were set up. The typical reaction was set up in 20-40 L and included vector, insert, ATP, T4 DNA ligase buffer, T4 DNA ligase and ddH 2 O. The first control reaction was a control for uncut plasmids, containing no insert and no ligase, and the second, containing no insert, was a control for the vectors ability to ligate with itself. The femptomolar concentration (fmol/L) was determined from the known size and measured concentration. Two ligation reactions were then set up, the first had 1 part vector to 3 parts insert and the second was 1:10. Typically 5-15 fmol of vector was used.

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72 The four reactions were incubated for 5 minutes at room temperature if T4 High Concentration DNA ligase was used or overnight at 16 C if T4 DNA ligase was used and then transformed into competent E. coli. Transformations were performed in one of three different E. coli strains. DH5 competent cells and XL10-Gold Ultracompetent cells were competent bacteria purchased from Invitrogen and Stratagene. These bacteria were used for purposes of plasmid preparations or to transform with a mutagenesis reaction such as Quikchange or ligations. The DH5 and XL10-Gold bacteria were stored at C. Basically, 25 L pre-aliquoted cells were thawed on ice and 1-5 L plasmid DNA was added and mixed by gentle stirring with the pipette tip. The bacteria were incubated on ice for 30 minutes, heat shocked at 42 C for 45 seconds, and then incubated on ice for 2 minutes. 1 mL LBG was added and then incubated at 37 C taped to a roller drum. The cells were harvested by centrifugation for 1 minute, the supernatant was discarded, the bacteria were resuspended in the remaining media, spread onto a LBG plate supplemented with the appropriate antibiotic and incubated at 37 C overnight. XL10-Gold cells required treatment with -mercaptoethanol (-ME) before transformation to increase the efficiency. 2 L of the provided -ME mix was added to 45 L pre-aliquoted cells and incubated on ice for 10 minutes prior to transformation with gentle swirling every 2 minutes. The transformation proceeded as described above. KM2 (b) was a strain of E. coli generated by a previous lab member (218) and maintained in our laboratory. This strain was used for purposes of plasmid expression and the study of the b subunit of F 1 F 0 ATP synthase. Before transformation, KM2 had to be made competent. Sterile technique was important since KM2 cells cannot be selected for by an antibiotic. All reagents and supplies were pre-chilled unless otherwise noted. A culture of KM2

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73 cells was grown overnight at 37 C in 5 mL LBG. The overnight culture (100 L) was inoculated into 10 mL pre-warmed (37 C) LBG and allowed to grow for 2-4 hours. The fresh culture was poured into sterile polypropylene tubes and incubated on ice for 10 minutes. The bacteria were then harvested by centrifugation (6,000 rpm in a ss-34 rotor) for 10 minutes. The supernatant was discarded and the tubes were inverted on a Kimwipe for 1 minute to allow excess LBG to drain. The bacteria pellet was resuspended in 10 mL cold, sterile 50 mM CaCl 2 and incubated on ice for 45-60 minutes. The bacteria were harvested as described above, resuspended in 1 mL of the 50 mM CaCl 2 and stored at 4 C until use. Generally, the competent KM2 cells could be used after two hours on ice but were most efficient for transformation at 24 hours and expired at 72 hours. For transformation, 100-200 L of cells were used as previously described. Site-directed mutagenesis. Site-directed mutagenesis was performed either by means of a Stratagene Quikchange XL kit or by ligation-mediated mutagenesis. Oligonucleotides containing the desired mutation(s) were designed to anneal to the same sequence on opposite strands of the plasmid (sense and antisense primers) (Appendix A) (Figure 2-1). When possible, a silent mutation was encoded to add or delete a restriction endonuclease recognition sequence to allow for easy screening of the mutation. Primers were optimally designed by ensuring the mutation was in the middle of the sequence, a cytosine (C) or guanine (G) flanked both ends of the sequence, and the melting temperature (T m ) was greater than or equal to 78 C. The T m was calculated as T m =81.5+0.41(%GC)-675/N-%mismatch, where N was the primer length (bases). When introducing insertions or deletions, "%mismatch" was dropped from the formula.

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74 Figure 2-1. Oligonucleotides for epitope tags and mutagenesis of uncF(b). Shown are the sense strands of the mutagenic primer pairs. Green and red codons specify translation start and stop sites, respectively. A) Mutagenic primers encoding the sequence of the desired epitope tag insertion along with a silent mutation that introduced a new endonuclease recognition sequence. Epitope tags were inserted as described in the Materials and Methods. The oligonucleotides sequences specifying the histidine, HA or V5 epitope (bold blue) tag are labeled with the corresponding amino acid. The SphI, NdeI and SacI restriction sites (underlined) were added along with the histidine, HA and V5 tags, respectively, to facilitate screening. B) Oligonucleotides designed to construct the one-plasmid expression system.

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75 Primers were not fast polynucleotide liquid chromatography (FPLC) or polyacrylamide gel electrophoresis (PAGE) purified as called for in the protocol. The reaction mixture consisted of 5 L Stratagenes 10X Pfu buffer, 3 L QuikSolution, 50 ng wild type plasmid, 125 ng each of the sense and antisense oligonucleotides, 1 L of 25 mM dNTPs, and 1 unit Pfu turbo polymerase. The total volume was brought up to 50 L with ddH 2 O and PCR was performed in a Perkin Elmer GeneAmp PCR System 2400 thermocycler according to the following cycling parameters: 95C 1 minute presoak; 18 cycles of 95C 50 seconds, 60C 50 seconds, 68C 1 minute per kb of plasmid length; 4C 7 min. Upon completion of the thermocycling, 1 L DpnI restriction endonuclease was added to the reactions in order to digest the methylated (nonmutated) plasmid DNA. The plasmids carrying the desired mutation were then transformed into competent DH5 cells, purchased from Life Technologies, and grown on LBG plates supplemented with the appropriate antibiotic. Plasmids carrying the desired mutation(s) were screened for by restriction endonuclease analysis and then the nucleotide sequences were directly determined by automated sequencing in the core facility of the University of Florida Interdisciplinary Center for Biotechnology Research (ICBR). Mutagenesis and Strain Construction Plasmids pKAM14 (b, Ap r ) (203), pAUL3 (b 7 Ap r ) (194) or pAUL19 (b +7 Ap r ) (193) were used to construct the epitope tagged b subunits. Epitope tags were inserted into each of the plasmids using the Stratagene Quikchange kit. A histidine epitope tag was inserted at the N-terminus by mutagenesis between the first and second codons of the uncF(b) gene to express b his or b +7-his (Appendix A) (Figure 2-1). Plasmids pTAM37 (b his ) and pTAM35 (b +7-his ) were created by digesting both of the recombinant histidine

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76 tagged b subunit plasmids with PstI and NdeI and subsequently ligating the genes into a plasmid conferring the chloramphenicol resistance gene and the pACYC184 origin of replication (Table 2-1). Likewise, an HA epitope tag was inserted at the N-terminus by mutagenesis between the first and second codons of the uncF(b) gene to generate plasmids pTAM36 (b HA ) or pTAM34 (b 7-HA ). A V5 epitope tag was added to the C-terminus by site-directed mutagenesis before the termination codon of the uncF(b) gene to express b V5 or b 7-V5 from plasmids pTAM46 and pTAM47 (Figure 2-1). The recombinant HA-tagged and V5-tagged b subunit plasmids included the ampicillin resistance gene and the pUC18 origin of replication. Unique restriction enzyme sites SphI, NdeI and SacI were constructed near the histidine, HA and V5 epitope tag sequence, respectively, for an initial detection of the insertions, and then the nucleotide sequence was subsequently confirmed by automated sequencing in the ICBR core facility. Additionally, a set of plasmids was designed in order to express two different-tagged b subunits from a single transcript (Figure 2-2). As an example, a unique restriction enzyme site, SphI, was created in conjunction with the histidine tag between the Shine Dalgarno sequence and the first codon of uncF(b) to create pTAM35 (Figure 2-2A). In a separate site-directed mutagenesis reaction, an SphI site was added to the pTAM34 plasmid downstream of the HA-tagged b subunit and behind another favorable Shine Dalgarno sequence (Figure 2-2B). The two plasmids were digested with SphI and BstEII restriction endonucleases. The 3.2 kb vector fragment from pTAM34 and the 623 bp insert fragment from pTAM35 were isolated and then excised from a 0.8 % (w/v) agarose gel and purified with a QIAquick gel extraction kit. The vector and insert fragments were then allowed to ligate overnight at 16 C. It was then crucial to mutate an

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77 Figure 2-2. Construction of the single transcript expression system. A) A unique restriction enzyme site, SphI, was created in conjunction with the histidine tag between the Shine Dalgarno sequence and the first codon of uncF(b) to create pTAM3. B) An SphI site was added to pTAM34 downstream of the HA-tagged b subunit and behind another favorable Shine Dalgarno sequence. The two plasmids were digested with SphI and BstEII. The 3.2 kb vector fragment from pTAM34 and the 623 bp insert fragment from pTAM35 were ligated. Additional mutagenesis (see Materials and Methods) resulted in C) a 3.7 kb plasmid, pTAM40, which expressed b 7-HA and b +7-his from the same promoter and included the ampicillin resistance gene and the pUC18 origin of replication. In similar constructions, plasmids pTAM41 (b wt-HA and b +7-his ) and pTAM42 (b wt-HA and b wt-his ) were created.

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78 intrinsic added start codon, which is in the SphI recognition sequence, to prevent a missense mutation. The mutagenic oligonucleotide was designed to accomplish three tasks in one reaction: 1) mutate the ATG found in the SphI recognition sequence, 2) add a new Shine Dalgarno sequence in a favorable position from the true start codon, and 3) mutate the GTG start codon to a more favorable ATG start site (Figure 2-1). The resulting 3.7 kb plasmid, pTAM40, expressed b 7-HA and b +7-his from the same promoter and included the ampicillin resistance gene and the pUC18 origin of replication (Figure 2-2C). In similar constructions, plasmids pTAM41 (b wt-HA and b +7-his ) and pTAM42 (b wt-HA and b wt-his) were created (Table 2-1). Throughout this dissertation, the insertion or deletion and the epitope tag are indicated after the plasmid name for clarity, for example plasmid pTAM35 (b +7-his ). Each plasmid and the control plasmids pKAM14 (b) and pBR322 were expressed in the E. coli cell line KM2 (b) for study, so that the only b subunits in the cells were the product of the plasmid genes. The two plasmid expression system successfully allowed expression of various combinations of histidine tagged and V5-tagged b subunits in the same cell (Figure 2-7). Appropriate antibiotics were added to the growth medium, and in the case of the coexpressed plasmids, both ampicillin and chloramphenicol were added to select for cells expressing both plasmids. Crude Preparative Procedures Inverted membrane vesicles from KM2(b) strains expressing the desired b subunit epitope tagged F 1 F 0 ATP synthase complex were prepared for activity assays, Ni-resin purification and Western Blot analysis. Unless otherwise noted, all reagents, rotors and materials were kept at 4 C. The membrane preparations were prepared by inoculating a

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79 10 mL starter culture, grown overnight, into a 2 L Erlenmeyer flask containing 500 mL LBG, supplemented with the appropriate antibiotic, ampicillin (Ap) and/or chloramphenicol (Cm). Similarly, 1 mL of a starter culture was inoculated into a nephalo flask containing 50 mL LBG (Ap and/or Cm). The bacteria were grown at 37 C in a New Brunswick Scientific incubator shaker (220 rpm) and the turbidity was monitored using a Klett-Summerson photoelectric colorimeter. IPTG (40 M) was added when the turbidity reached 75 Klett units and the cells were collected when the turbidity reached 150 Klett units. The bacteria were harvested by centrifugation for 10 minutes at 8,000Xg in a Sorvall GSA rotor. The pellets were rinsed once with TM buffer (50 mM tris-HCl, 10 mM MgSO 4 pH 7.5) and then resuspended in a final volume of 10 mL TM buffer. DNaseI (10 mg/mL) was added to a final concentration of 10 g/mL and the bacteria were broken by passing through a French Pressure Cell one time at 14,000 psi. Cellular debris and unbroken cells were removed by centrifuging twice at 10,000Xg for 10 minutes. Membranes were then collected by ultracentrifugation at 150,000Xg in a Beckman 70.1 Ti rotor for 1.5 hours. The membrane pellets were rinsed once with TM buffer and then resuspended in TM buffer to a final volume of 2 mL using a 2 mL Wheaton tissue grinder. For the purposes of Western blot analysis or Ni-resin purification, one ultracentrifugation step sufficed. However, activity analysis required an additional ultracentrifugation step in order to remove nonspecifically bound ATPases. In this case, the membrane pellets were resuspended in a final volume of 10 mL using a 10 mL Wheaton tissue grinder and the ultracentrifugation step was duplicated.

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80 Determination of Protein Concentration Total membrane protein concentrations were determined by the bicinchoninic acid (BCA) assay (286). The membrane preparations were diluted 1:10 for this assay. For each sample, triplicates of 10 L, 20 L and sometimes 30 L of the diluted samples were aliquoted for the assay. Bovine serum albumin (BSA) was used to generate a standard curve. A stock of 1 mg/mL BSA was prepared and stored in microcentrifuge tubes at C. The exact BSA concentration was determined from the OD 280 (1.0 mg/mL BSA OD 280 =0.667). The BSA was aliquoted in triplicates ranging from 0 to 100 g (0, 5, 10, 20, 40, 60, 80, and 100 L) of protein. All of the samples were incubated in 3 mL of standard working reagent (SWR), consisting of 50 parts solution A (1% BCA-Na 2 2% Na 2 CO 3 H 2 O, 0.16% sodium potassium tartrate, 0.95% NaHCO 3 pH 11.25 and filtered) to one part solution B (4 % CuSO4H 2 O) plus 1% sodium dodecyl sulfate (SDS) mixed fresh as needed, for 30 minutes at 37 C and then for 10 minutes at room temperature. The absorbance at 562 nm was recorded for each sample with an LKB Biochrom Ultrospec II spectrophotometer. The standard curve was generated from the BSA OD readings, with a typical correlation coefficient of at least 0.997, and then protein concentrations were determined using linear regression. A typical concentration range of the total membrane protein prepared from 500 mL of media was 5-20 g/L. Ni-Resin Purification Ni-resin purification was achieved using the High Capacity Nickel Chelate Affinity Matrix (Ni-CAM) purchased from Sigma. It became necessary to optimize conditions and the ratio of protein to packed Ni-resin volume for each type of experiment. Determining the optimal ratio of protein to Ni-resin was critical because F 1 F 0 ATP

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81 synthase complexes with two histidine-tagged b subunits bind with a higher affinity than the complexes containing a b subunit heterodimer, which has only one histidine epitope tag. For the purposes of Western blot analysis, a total of 5 mg of membrane protein was brought up to 1 mL with final concentrations of 0.2% tegamineoxide WS-35, 0.15 M NaCl, and 1 mM imidazole. The purification procedure was accomplished using the trial scale miniprep method essentially described by the manufacturer. The 1 mL of clarified membrane protein was divided up equally into 5 microcentrifuge tubes, containing 100 L of packed Ni-resin that had been washed one time with 500 L equilibration/wash buffer (50 mM NaH 2 PO 4 H 2 0, 0.3 M NaCl, 10 mM imidazole, 0.2% tegamineoxide WS-35, pH8.0), and allowed to mix on a nutator for 5 minutes. The samples were centrifuged for 30 seconds at 5,000Xg and the supernatant was discarded. The resin was washed 8 times with 1 mL wash buffer by gently mixing for 1 minute, centrifuging for 30 seconds at 5,000Xg and discarding the supernatant. The histidine-tagged proteins were eluded by mixing 50 L elution buffer (50 mM NaH 2 PO 4 H 2 0, 0.3 M NaCl, 250 mM imidazole, 0.2% tegamineoxide WS-35, pH8.0) in each microcentrifuge tube for 1 minute, centrifugation, and then the supernatant was extracted. The elution step was repeated one more time to recover more of the target protein. Finally, the target protein was pooled together and then concentrated from 500 L to about 50 L by centrifugation with a Millipore Amicon Bioseparations Microcon YM-10 (14,000Xg for 30 minutes). To control for possible enzyme disruption during the solubilization and purification procedures, the tagged wild-type length b subunits were either mock treated or treated with the homobifunctional crosslinker, bis(3-sulfo-N-hydroxysuccinimide ester) (BS 3 ), after the addition of detergent. Membrane preparations were chemically crosslinked by

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82 treating with 1mM BS 3 for 30 minutes at room temperature. The cross-linking reaction was stopped by addition of 100 mM ethanolamine HCl, pH 7.5, for 10 minutes. For the purposes of densitometric analysis, a total of 15 mg membrane protein was solubilized and clarified as described above. The clarified protein was divided into 10 samples and then Ni-resin purified (1 ml packed resin volume divided into 10 microcentrifuge tubes) as described previously, and the final eluate was concentrated with a Microcon YM-10. Attempts to purify the heterodimers with the batch method (15 mg membrane protein to 1 mL packed resin volume in a 15 mL corning conical tube, undivided) proved unsuccessful; therefore division into several microcentrifuge tubes was necessary. Assays of F 1 F 0 ATP Synthase Activity Growth on a minimal succinate medium was used as an initial, in vivo, assay for enzyme viability. ATP hydrolysis activity was assayed by the acid molybdate method (146), which measures the release of P i from ATP, in order to determine the specific activity of the epitope tagged F 1 F 0 ATP synthase enzyme complexes. Membranes were assayed to determine the linearity with respect to time and enzyme concentration. Prior to the assay, all supplies and reagents were distributed, labeled and placed on ice or 37 C as specified below. For each membrane sample, duplicates of 60 g membrane protein were incubated in 4.0 mL of the reaction buffer (50 mM Tris-HCl, 1 mM MgCl 2 pH9.1) at 37 C. Stop buffer, which acts to prevent further hydrolysis of ATP, was prepared as needed and consisted of 1.3 part ddH 2 O to 0.6 part HCl/molybdate solution (2.5% NH 4 Mo 4 O 24 H 2 O, 4.0 N HCl) to 0.4 part 10% SDS. The stop buffer was divided into 2 mL aliquots in 13x100 mm disposable borosilicate glass tubes and kept on ice throughout

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83 the assay. Tubes of stop buffer were required for each time point of each sample, phosphate standard curve, and ATP only control. Prior to the beginning of the time course, two measures are taken to subtract background phosphate. First, a zero time point was taken by removing 435 L of the reaction buffer, containing the membranes before the addition of ATP, and adding it to 2 mL of stop buffer. Also, to account for spontaneous hydrolysis, ATP was incubated in 4 mL of reaction buffer with no membrane protein and then 435 L was removed and added to stop buffer. The time course was started by the addition of 80 L ATP (0.15 M in 25 mM Tris-HCl, pH 7.5) to the reaction buffer and additional time points were taken at 2, 5, 7, 10 and 12 minutes. A phosphate standard curve was generated by preparing duplicates of 1 mL reaction buffer containing 0, 0.02, 0.1, 0.2, 0.4 and 0.6 mol phosphate (KH 2 PO 4 ), and then adding 435 L of each to 2 mL stop buffer. Once the time course and the standard curve were completed, all of the samples in stop buffer were removed from the ice and allowed to come to room temperature. The inorganic phosphate concentration was determined by adding 100 L of Eikonogen solution (1 M NaHSO 3 0.1 M Na 2 SO 3 0.01 M 4-amino-3-hydroxy-1-naphthalenesulfonic acid), diluted 1:10, and incubating at room temperature for 30 minutes to allow color development. The absorbance of each time point at 700 nm was recorded with an LKB Biochrom Ultrospec II spectrophotometer and the amount of free phosphate in each sample was determined using linear regression. The rate of ATP hydrolysis was determined and the apparent specific activity was expressed as mol P i /min/mg membrane protein. Membrane energization was detected by the fluorescence quenching of 9-amino-6-chloro-2-methoxyacridine (ACMA) (271). First, F 1 F 0 ATP synthase-mediated ATP

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84 driven proton pumping activity in inverted membrane vesicles prepared from the epitope-tagged mutants was used as an indication of coupled activity. Acidification of the inverted membrane vesicles upon addition of ATP was directly examined by fluorescence quenching. Membrane protein (250 g) was suspended in 3 ml of assay buffer (50 mM MOPS, 10 mM MgCl 2 pH 7.3) in a quartz cuvette. The fluorescent dye ACMA was added to a final concentration of 1 M, and fluorescence was monitored with a Perkin-Elmer LS-3B Fluorescence Spectrometer with excitation at 410 nm and emission at 490 nm and directly recorded with a Perkin-Elmer GP-100 Graphics Printer. After several seconds, the fluorescent emission stabilized and was manually set to 85% for the purpose of the scale. The emission was graphically recorded for 1.5 minutes, ATP was added to a final concentration of 0.4mM and the emission was recorded for and additional 10-12 minutes. The level of -nicotinamide adenine dinucleotide, reduced form, (NADH)-driven fluorescence quenching was monitored for all membrane preparations to demonstrate that the vesicles were intact and closed. Membrane protein (250 g) was suspended in 3 ml of assay buffer (50 mM MOPS, 10 mM MgCl 2 pH 7.3). ACMA was added to a final concentration of 1 M, and fluorescence was recorded excitation set at 410 nm and emission set at 490 nm. After several seconds, the fluorescent emission stabilized and was manually set to 95%. The emission was recorded for 1 minute, 5 L NADH (0.1 mM) was added and the emission was continually recorded. Over time, membrane vesicle acidification peaked and the fluorescence quenching reached a maximum. As the fluorescence began to rise, 5 L of 0.3 mM KCN was added and the emission was recorded for a total time of about 10-15 minutes.

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85 Immunoblot Analysis Electrophoresis and transfer. Proteins were mixed with 2X Laemmli sample buffer (LSB) (62.5 mM tris-HCl, pH 6.8, 2% w/v SDS, 720 mM -mercaptoethanol, 20% glycerol, and 0.1% Bromophenol blue) and incubated for 5 minutes at 95 C. The proteins were then loaded on either a 15 cm 15% tris-glycine SDS gel or a purchased SDS-PAGE 15% Tris-HCl Ready Gel purchased from BioRad Laboratories and then transferred onto a nitrocellulose (anti-b or anti-V5 antibody incubation) or polyvinylidene fluoride (PVDF) (anti-HA antibody incubation) microporous membrane by electroblot. The proteins were separated in electrode buffer (25 mM tris, 192 mM glycine, and 0.1% SDS) by electrophoresis in a Mini-PROTEAN II cell (small gels) or a PROTEAN II (large gels). When it was necessary to distinguish two different tagged b subunits from the same immunoblot, a maximal separation was required. The proteins were run on a 15 cm 15% polyacrylamide tris-glycine gel at 100 mamp, with current held constant, until the dye reached the stacker and then at 24 mamp for 12 hours. It was necessary to double the amperage when two gels were running at the same time. The protein transfer was accomplished in transfer buffer (25 mM tris, 192 mM glycine, 20% methanol) at 4 C at either 100 V, 0.25 amp limit for 1 hour (small gels) or 100 V, 0.36 amp limit for 4 hours (large gels). Anti-b subunit antibody. The b subunit antibody incubation was performed essentially as described previously by Tamarappoo et al. (287). The current lab stock of anti-b subunit antibodies were diluted 1:40 (long-term stock) and 1:40,000 (short-term stock) for storage on a non-frost-free shelf at C and required and additional 1:40 dilution of the short-term stock for the working concentration. Upon completion of

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86 electrophoresis to a nitrocellulose membrane, the protein was visualized, for loading comparison, with fast green stain (50% methanol, 10% glacial acetic acid, 0.01% fast green) by incubation at room temperature with gentle shaking on a Bellco Biotechnology Orbital Shaker for 15 minutes followed by three 2-5 minute (depending on strength of stain) destain washes (50% methanol, 10% glacial acetic acid). The fast green stain was recycled for repetitive use and the destain was discarded. The membrane was washed three times for 5 minutes with tris-buffered saline (TBS) (10 mM tris-HCl, 150 mM NaCl, pH7.2) supplemented with 0.1% polyoxyethylenesorbitan monolaurate (tween 20) (TTBS) and then blocked for 1 hour at room temperature or overnight at 4 C in TBS supplemented with 5% nonfat dry milk (NFDM). The primary antibody incubation was performed with the anti-b subunit antibody (1:40) in TTBS supplemented with 2% BSA for 1 hour followed by three 5 minute washes with TTBS. The anti-b antibody was restored at C and thawed for reuse up to 3-5 times. Finally, secondary antibody incubation was performed with horseradish peroxidase-linked donkey anti-rabbit antibody (1:50,000 in TTBS-BSA), washed 4 times in TTBS (two 5 minute washes, two 10 minute washes) and then the antibody was detected by enhanced chemiluminescence (ECL). Signals were visualized on high performance chemiluminescence film using a Kodak X-Omat. Typical exposure times for the anti-b antibody ranged from 1-5 minutes for strong signals and 30-60 minutes for weaker signals. Anti-HA antibody. The HA-epitope tag antibody was performed essentially as described by the manufacturer followed by a secondary antibody incubation with horseradish peroxidase-linked anti-mouse (from sheep) antibody (1:10,000). Several initial attempts to visualize the HA-epitope tagged b subunit on a nitrocellulose

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87 membrane failed to give a clean signal. It was essential to enhance the sensitivity of the antibody by transfer onto a PVDF membrane for a higher signal to noise ratio. Before electrotransfer, it was crucial to prepare the membrane by immersion in 100% methanol for 15 seconds ddH 2 O for 2 minutes and then transfer buffer for 5 minutes. The wash buffer consisted of a phosphate buffered saline (PBS) (1% NaCl, 0.025% KCl, 0.18% Na 2 HPO 4 0.03% KH 2 PO 4 pH 7.4) supplemented with 0.1% tween 20 (PBST), membranes were blocked in PBS with 5% NFDM, and the primary and secondary antibody incubation was performed in PBST supplemented with 2.5% NFDM. Typical exposure times for the anti-HA antibody were about 1 hour. Occasionally, a stronger detection reagent, ECL Plus (Amersham Biosciences), was required for detection. ECL Plus is an extremely sensitive detection system; therefore it was necessary to follow protocol carefully, using optimal primary and secondary antibody conditions. The detection reagents were brought to room temperature before opening and then solutions A and B were mixed in a 40:1 ration. The final volume required was 0.1 mL per cm 2 of membrane. Excess wash buffer was allowed to drip off the membrane and then the detection solution was pipetted onto the membrane and incubated at room temperature for 5 minutes. Finally, it was critical to drain off excess detection reagent by touching the corner of the membrane onto a kimwipe before exposure to film. Anti-V5 antibody. The V5-epitope tag antibody incubation was performed as described by the manufacturer followed by secondary antibody incubation with horseradish peroxidase-linked sheep anti-mouse antibody (1:10,000). The wash buffer consisted of TTBS, membranes were blocked in TBS with 5% NFDM, and the primary and secondary antibody incubation was performed in TBS supplemented with 2.5%

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88 NFDM. Typical exposure times for the anti-V5 antibody were about 30 seconds to 1 minute for membrane proteins and about 1 hour to overnight to visualize Ni-resin purified proteins. Results HA-Epitope Tagged b Subunits To investigate whether it is possible for two b subunits of unequal length to dimerize to form the peripheral stalk in a functional enzyme complex, a collection of plasmids expressing two different epitope tagged b subunits were generated by site-directed mutagenesis. Initially a histidine-epitope tag and a HA-epitope tag were employed. Construction and growth characteristics of mutants The first approach developed to express two different b subunits in the same cell was a two plasmid expression system (Figure 2-2). A total of four plasmids were constructed expressing b wt-his (Cm r ), b +7-his (Cm r ), b wt-HA (Ap r ) or b 7-HA (Ap r ) (Table 2-1). The histidine and HA epitope tags were needed to facilitate enzyme purification and subunit detection on Western blot, respectively. Coexpression of both b subunits was selected for by media that contained both ampicillin and chloramphenicol. A previous analysis of F 1 F 0 ATP synthase complexes with b subunits shortened and lengthened by 7 amino acids found the mutants to be essentially wild-type (193, 194). Functional F 1 F 0 ATP synthase complexes have been studied with epitope tags positioned on many of the subunits, however, an epitope tag on the b subunit had not been attempted. It was possible that the epitope tags would affect enzyme assembly or function. The effects of the added epitope tags were studied by the ability of the plasmids to complement the E. coli strain KM2 (b) (218). Growth on succinate minimal

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89 medium was used as an initial qualitative gauge of enzyme activity in vivo since E. coli strains lacking F 1 F 0 ATP synthase cannot derive energy from nonfermentable carbon sources. In each case, the strains expressing the histidine and HA-epitope tagged b subunits grew comparably to the wild type strain (Table 2-1). Figure 2-3. Histidine and HA-epitope-tagged b subunit expression system. We employed an epitope-tag system in order to determine whether b subunits of unequal length can interact to form the dimer in an intact and functional enzyme complex. B) Both b 7-HA and b +7-his were expressed together in KM2 (b) cells using a two-plasmid expression system. Plasmid pTAM34 was designed to express high levels of HA-tagged b 7 This plasmid contains the genes conferring ampicillin resistance and the pUC18 origin of replication. Plasmid pTAM35 includes the chloramphenicol resistance gene, the pACYC184 origin of replication and expresses histidine-tagged b +7 In similar constructions, plasmids pTAM37 (b wt-his Cm r ) and pTAM36 (b wt-HA Ap r ), or pTAM35 (b +7-his Cmr) and pTAM46 (b wt-HA Ap r ) were developed for coexpression experiments.

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90 Table 2-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing epitope tagged uncF(b) genes Strains Description Growth 1 Specific activity 2 KM2/pKAM14 (+) b wt Ap r +++ 1.72 0.04 KM2/pBR322 (-) b, Ap r 0.50 0.01 KM2/pTAM37 b wt-his Cm r +++ 1.53 0.08 KM2/pTAM35 b +7-his Cm r +++ 1.35 0.03 KM2/pTAM36 b wt-HA Ap r +++ 1.48 0.03 KM2/pTAM34 b 7-HA Ap r +++ 1.39 0.04 KM2/pTAM37/pTAM36 b wt-his + b wt-HA Cm r & Ap r +++ nd KM2/pTAM35/pTAM36 b +7-his + b wt-HA Cm r & Ap r +++ nd KM2/pTAM35/pTAM34 b +7-his + b 7-HA Cm r & Ap r +++ 1.41 0.04 KM2/pTAM42 b wt-his + b wt-HA Ap r +++ nd KM2/pTAM41 b +7-his + b wt-HA Ap r +++ nd KM2/pTAM40 b +7-his + b 7-HA Ap r +++ nd KM2/pTAM46 b wt-V5 Ap r +++ 1.63 0.02 KM2/pTAM47 b 7-V5 Ap r +++ 1.40 0.05 KM2/pTAM37/pTAM46 b wt-his + b wt-V5 Cm r & Ap r +++ 1.70 0.03 KM2/pTAM35/pTAM46 b +7-his + b wt-V5 Cm r & Ap r +++ 1.58 0.02 KM2/pTAM35/pTAM47 b +7-his + b 7-V5 Cm r & Ap r +++ 1.53 0.03 1 E. coli strains were grown aerobically on succinate minimal medium. Colony size was scored after 72-hr incubation at 37C as: +++, 1.0 mm; ++, 0.3-0.5 mm; +, ~0.1 mm; -, no growth. 2 ATPase activities were measured as described under Materials and Methods. Units of specific activity = mol of PO 4 released per mg of protein/min S.D. Units were calculated from the slope of the line based on three measurements with incubations for 12 minutes. Effects of epitope tags Since F 1 has little affinity for the membrane in the absence of intact F 0 total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly. Under conditions of high pH, F 1 can be released from the influence of F 0 (146), so the amount of ATPase activity in the solution was used as a measure of the amount of intact enzyme complex located in the membrane vesicles. The histidine and HA-epitope tags had a slight, but not vitally significant affect on enzyme assembly. Membrane preparations with a b wt-HA or b 7-HA incorporated into the F 1 F 0 ATP synthase complex had specific activities of about 86% and 81% of the wild type strain, respectively (Table 2-1). The latter value was comparable to the effect of the seven

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91 amino acid deletion in the absence of the epitope tag. Similar activities were observed in membrane vesicles isolated from cells with a histidine epitope tag was incorporated onto the enzyme. F 1 F 0 ATP synthase complexes with a b wt-his or b 7-his dimer had specific activities of about 90% and 79% of the wild type strain, respectively (Table 2-1). Expression of different b subunits in the same cell The ability to express two different b subunits in the same cell in roughly equal quantities was crucial to the success of the experiment. Therefore it was necessary to establish whether two separate plasmids could be used to direct production of two different b subunits. Two b subunits, expressed from pTAM35 (b 7-his ) and pTAM34 (b 7-HA ), were expressed in the same KM2 (b) cell and their ability to exist in the same cell was determined by Western blot analysis (Figure 2-4A). As expected, immunoblot analysis of crude membrane preparations using anti-b antibodies showed the presence of the b subunit in all strains complemented with an epitope-tagged b subunit (Figure 2-4A, Lanes 3-6). As shown in the first lane, KM2 cells did not express b subunit. After transforming with plasmid pKAM14 (b wt ), the KM2 cells expressed the wild type b subunit (Lane 2). The third and forth lanes represented KM2 cells expressing the b 7-HA and b +7-his subunits, respectively. In lane 5, individual membrane preparations representing lanes 3 and 4 were mixed together. Finally, lane 6 represented the KM2 expressing both the short and long b subunits in the same cell. This figure demonstrated our ability to separate the short and long subunits enough to be sufficiently distinguishable. Notably, the KM2 cells successfully expressed the two different b subunits in roughly equal quantities from the same cell.

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92 Figure 2-4. Western blot analysis of histidine and HA-epitope tagged b subunits. A) KM2 (b) cells were transformed with pKAM14 (b wt ), pTAM34 (b 7-HA), pTAM35 (b +7-his ), or both pTAM34 and pTAM35. Crude membrane preparations were made and proteins separated on a 15% polyacrylamide Tris-HCl-SDS BioRad Ready gel. The proteins were transferred to a nitrocellulose membrane and probed with anti-b anitbodies. Lane 5 represents a mixture of membranes in lanes 3 and 4. B) The crude membrane preparations were solubilized with 0.2% LDAO and then subjected to Ni-resin purification under native conditions (see Materials and Methods). The purified proteins were then separated with SDS-PAGE and Western blotted with anti-b antibodies. Ni-Resin Purification Initial experiments utilizing the two plasmid expression system approach indicated that the two different b subunits were incorporated into F 1 F 0 ATP synthase in a segregated manner, suggesting that the b subunit dimer must form in parallel during F 1 F 0 ATP synthase assembly. The two b subunits, expressed from pTAM35 (b 7-his ) and pTAM34 (b 7-HA ), were expressed in the same KM2 (b) cell and their ability to exist in the same F 1 F 0 ATP synthase enzyme complex was determined by Ni-resin purification followed by Western blot analysis (Figure 2-4B). Upon Ni-resin purification, only enzyme containing the histidine epitope tagged b subunit should have been present. As

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93 anticipated, KM2 cells did not express b subunit. Western blot analysis confirmed that b subunits without a histidine epitope tag were washed off the resin during the wash steps (Figure 2-4B, Lanes 2 and 3) whereas b +7-his was retained by the Ni-resin (Figure 2-4B, Lane 4). In lane 5, when individual membrane preparations of KM2 individually expressing either the short or long subunit were mixed together, as expected, only the histidine epitope tagged b subunit was purified from the Ni-resin. In the KM2 cells expressing both b +7-his and b 7-HA only the histidine-tagged long b subunit appeared, suggesting the different b subunits were segregated into separate enzyme complexes. However, there were two important caveats: the enzyme complex may have dissociated before coming off the Ni-resin, and a difference of fourteen amino acids may have simply been too much for the stability of the enzyme. Both issues were studied. First, two additional detergents to solublize the membrane were investigated. The capability of the detergents to allow the enzyme to come off the resin intact was investigated by cross-linking the b subunits after solubilization and Ni-resin purification and then visualized by Western blots. Secondly, the size difference of the two b subunits was decreased by testing for the ability of the b +7-his subunit to dimerize with a b wt-HA subunit. Membrane preparations of KM2 (b) expressing b +7-his were solubilized with 0.2% solutions of tegamineoxide WS-35 (AO), taurodeoxycholate (TD), lauryldimethylamine oxide (LDAO) or SDS and allowed to incubate at room temperature for 30 minutes. AO, TD and LDAO were detergents commonly used to solubilize membrane proteins. LDAO was likely to dissociate F 1 from F 0 In theory, however, F 0 should have remained intact through the purification. The solubilized proteins were then mock treated (-) or treated (+) with the homobifuctional crosslinker, BS 3 The cross-linking reaction was stopped by

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94 Figure 2-5. Investigation of detergent solubilization of F 1 F 0 ATP synthase complexes. Membrane preparations of KM2 (b) expressing b +7-his were solubilized with 0.2% solutions of detergents: tegamineoxide WS-35 (AO), taurodeoxycholate (TD), lauryldimethylamine oxide (LDAO) or SDS. The solubilized proteins were then mock treated (-) or treated (+) with 1mM bis(3-sulfo-N-hydroxysuccinimide ester) (BS 3 ) for 20 minutes at room temperature. The cross-linking reaction was stopped by addition of 100 mM ethanolamine HCl, pH 7.5 for 10 minutes. The products were separated by SDS-PAGE, transferred to nitrocellulose, and probed with anti-b antibodies. addition of ethanolamine HCl, pH 7.5 and then the products were separated by SDS-PAGE, transferred to nitrocellulose, and probed with anti-b antibodies for Western blot analysis (Figure 2-5). The b +7-his dimers were detected in membrane preparations treated with BS 3 after solubilization, therefore, the crosslinked product showed that the b +7-his subunit dimer was stable during solubilization with AO, TD and LDAO (Figure 2-5, Lanes 4,6, and 8), but not with SDS (Figure 2-5, Lane 10), which was expected to denature the proteins. Once the detergent conditions were determined to be safe for the enzyme complex, the ability of the different b subunits, b +7-his and b 7-HA to heterodimerize was reevaluated by a slightly different approach. This time, membrane preparations were treated with the crosslinker, BS 3 prior to Ni-resin purification to ensure no disruption of the dimers (Figure 2-6). Immunoblot analysis of crude membrane

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95 preparations using anti-b antibodies showed the presence of the b subunit in all strains complimented with a b or an epitope-tagged b subunit (Figure 2-6A, Lanes 2-5). As expected, only b subunits with a histidine tag were retained by Ni-resin purification (Figure 2-6A, Lanes 6-11). F 1 F 0 complexes with only the HA-tagged b subunits were completely washed off the resin (Figure 2-6A, Lanes 6-7). To detect a heterodimer formation, immunoblot analysis using the anti-HA antibody was performed (Figure 2-6B). When the two plasmids were coexpressed in the same cell the anti-HA antibody did not detect the presence of b 7-HA after Ni-resin purification, indicating the two different Figure 2-6. Ni-resin purification of F 1 F 0 expressing different length b subunits, treated with the cross-linker BS 3 Crude membrane preparations of KM2 expressing the different b subunits were cross-linked (Materials and Methods). The products were then solubilized and subjected to Ni-resin purification before separation by SDS-PAGE and subsequent Western blot analysis. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody. B) Proteins were transferred to a PVDF membrane and probed using an anti-HA mouse monoclonal antibody.

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96 b subunits, b +7-his and b 7-HA did not dimerize. Similar experiments were performed with a seven amino acid size difference as opposed to the fourteen amino acid difference just attempted (data not shown). The immunoblot analysis gave identical results to the previous data, b +7-his and b wt-HA dimerization was not detected, suggesting the two different b subunits had incorporated into F 1 F 0 ATP synthase in a segregated manner. However, further experimentation utilizing this experimental approach indicated that dimerization was not detected between two wild type length b subunits, b wt-his and b wt-HA Figure 2-7. Ni-resin purification of histidine and HA-epitope tagged F 1 F 0 treated with the cross-linker BS 3 Crude membrane preparations of KM2 expressing the different b subunits were cross-linked (Materials and Methods). The products were then solubilized and subjected to Ni-resin purification before separation by SDS-PAGE and subsequent Western blot analysis. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody. B) Proteins were transferred to a PVDF membrane and probed using an anti-HA mouse monoclonal antibody.

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97 rendering the previous data worthless (Figure 2-7). In order to conclude that the different b subunits formed F 1 F 0 ATP synthase complexes in a segregated manner, it was necessary to detect an interaction between two wild type length b subunits. Several factors may have contributed to the failure of the first experimental system: unknown interactions between the HAand histidine-epitope tag may have hindered an interaction between the two b subunits, the anti-HA antibody was not sensitive enough or the signal-to-noise ratio was too small for detection, or expression from two different plasmids may have placed an early, detrimental affect on the experimental system. The latter was considered first. Dimerization of the b subunits is thought to be an early event in enzyme assembly, making the requirement for immediate dimerization of b via co-translation of a single b subunit transcript feasible. Therefore, three plasmids were designed to express two different tagged b subunits from a single transcript (Figure 2-3C, and Table 2-1). All three plasmids, pTAM40, pTAM41 and pTAM42, successfully expressed two different b subunits, nevertheless, an interaction between b wt-his and b wt-HA was not found. V5-Epitope Tagged b Subunits Several attempts to visualize an interaction between two different wild type length b subunits with a histidine and a HA-epitope tag failed. Attention was then focused on the types of epitope tags used and we decided to replace the HA tag with a V5-epitope tag. Construction and growth characteristics of mutants In a third attempt to investigate whether it is possible for two b subunits of unequal length to dimerize to form the peripheral stalk in a functional enzyme complex, a collection of plasmids expressing histidine or V5 epitope-tagged b subunits were

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98 Figure 2-8. Histidine and V5-epitope-tagged b subunit expression system. A) Both b 7-V5 and b +7-his were expressed together in KM2 (b) cells using a two-plasmid expression system. Plasmid pTAM47 was designed to express high levels of V5-tagged b 7 This plasmid contains the genes conferring ampicillin resistance and the pUC18 origin of replication. Plasmid pTAM35 includes the chloramphenicol resistance gene, the pACYC184 origin of replication and expresses histidine-tagged b +7 In similar constructions, plasmids pTAM37 (b wt-his Cm r ) and pTAM46 (b wt-V5 Apr), or pTAM35 (b +7-his Cm r ) and pTAM46 (b wt-V5 Apr) were developed for coexpression experiments. B) Line diagram of the different b subunits that were coexpressed in the same cell. Interactions between b wt-his / b wt-V5 b +7-his / b wt-V5 and b +7-his /b 7-V5 were investigated.

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99 generated by site-directed mutagenesis. Again, the epitope tags were needed to facilitate enzyme purification and subunit detection on a Western blot, respectively. In order to express two different b subunits in the same cell, we first used the two-plasmid expression system (Figure 2-8A). A total of four plasmids were constructed expressing b wt-his (Cm r ), b +7-his (Cm r ), b wt-V5 (Ap r ) or b 7-V5 (Ap r ) (Table I). The deletion removed the segment from Leu54-Ser60, and the insertion resulted in duplication of the same series of amino acids. Previous work had shown that F 1 F 0 ATP synthase complexes with b subunits shortened and lengthened by 7 amino acids were essentially wild-type (193, 194). However, it was possible that the epitope tags would affect enzyme assembly or function. This was particularly a concern at the C-terminus where small deletions at the extreme C-terminal end of the subunit had been shown to inhibit F 1 F 0 ATP synthase function (206, 288). Addition of the fourteen amino acid V5-epitope tag to the C-terminus of b might have impinged on enzyme assembly. The effects of the added epitope tags were studied by the ability of the plasmids to complement the E. coli strain KM2 (b) (218). Growth on succinate minimal medium was used as an initial qualitative gauge of enzyme activity in vivo since E. coli strains lacking F 1 F 0 ATP synthase cannot derive energy from nonfermentable carbon sources. In each case, the strains expressing the epitope-tagged b subunits grew comparably to the wild type strain (Table I). Hence, even though deletion of as few as two amino acids affected the ability of the b dimer to interact with the F 1 subunit (41,42), addition of fourteen amino acids to the C-terminus did not interfere with the interaction of the b and subunits.

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100 Effects of epitope tags Since F 1 has little affinity for the membrane in the absence of intact F 0 total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly. Under conditions of high pH, F 1 can be released from the influence of F 0 (146), so the amount of ATPase activity in the solution was used as a measure of the amount of intact enzyme complex located in the membrane vesicles. The V5-epitope tag did not have a significant affect on enzyme assembly. Membrane preparations with a b wt-V5 or b 7-V5 incorporated into the F 1 F 0 ATP synthase complex had specific activities of about 95% and 82% of the wild type strain, respectively (Table I). The latter value was comparable to the effect of the seven amino acid deletion in the absence of the epitope (194). A slightly greater decrease in specific activity was reproducibly observed in membrane vesicles isolated from cells when a histidine epitope tag was incorporated onto the b wt or b +7 subunit, about 90% and 79% of wild type, respectively. Nevertheless, abundant activity was retained, suggesting very limited effects resulting from addition of the epitope tags. Furthermore, comparable activities were observed in samples when the histidine-tagged and V5-tagged b subunits were coexpressed. F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the epitope-tagged mutants was used as an indication of coupled activity. Acidification of inverted membrane vesicles was examined by fluorescence of ACMA (Figure 2-9). The level of NADH-driven fluorescence quenching was monitored for all membrane preparations to demonstrate that the vesicles were intact and closed. The levels of NADH-driven fluorescence quenching were strong and directly comparable in every case (Figure 2-10). Membranes isolated from cells with a V5 epitope tag

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101 Figure 2-9. ATP-driven energization of membrane vesicles prepared from uncF(b) gene mutants. Cell membrane vesicles were prepared by differential centrifugation (see Materials and Methods). Membrane protein (200 g) was suspended in 3 ml of assay buffer (50 mM MOPS, 10 mM MgCl 2 pH 7.3). The fluorescent dye ACMA was added to a final concentration of 1 M, and fluorescence was recorded with excitation at 410 nm and emission at 490 nm. ATP was added as indicated to a final concentration of 1mM. The samples for each trace have been labeled according to the amino acid insertion or deletion and the epitope tag, so the strains used as the sources of the samples were as follows: b wt KM2/pKAM14; b wt-V5 KM2/pTAM46; b 7-V5 KM2/pTAM47; b wt-his KM2/pTAM37; b +7-his KM2/pTAM35; b wt-his / b wt-V5 KM2/pTAM37/pTAM46; b +7-his /b wt-V5 KM2/pTAM35/pTAM46; b +7his/b 7-V5 KM2/ pTAM35/pTAM47.

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102 Figure 2-10. NADH-driven acidification of membrane vesicles prepared from uncF(b) mutants. Membrane protein (250 g) was suspended in 3 ml of assay buffer (50 mM MOPS, 10 mM MgCl 2 pH 7.3). The fluorescent dye ACMA was added to a final concentration of 1 M, and fluorescence was recorded with excitation at 410 nm and emission at 490 nm. 5 mL NADH (0.1 mM) was added and the emission was continually recorded. Over time, membrane vesicle acidification peaked and the fluorescence quenching reached a maximum. As the fluorescence began to rise, 5 mL of 0.3 mM KCN was added and the emission was recorded for a total time of about 10-15 minutes. The samples for each trace have been labeled according to the amino acid insertion or deletion and the epitope tag, so the strains used as the sources of the samples were as follows: b wt KM2/pKAM14; b wt-V5 KM2/pTAM46; b 7-V5 KM2/pTAM47; b wt-his KM2/pTAM37; b +7-his KM2/pTAM35; b wt-his / b wt-V5 KM2/pTAM37/pTAM46; b +7-his /b wt-V5 KM2/pTAM35/pTAM46; b +7his/b 7-V5 KM2/ pTAM35/pTAM47.

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103 incorporated onto the b wt or b 7 subunit, KM2/pTAM46 or KM2/pTAM47, respectively, displayed a very slight reduction in coupled activity (Figure 2-9). The reduction in coupled activity correlated very well with the minor reduction in F 1 -ATP hydrolysis activity. A larger reduction in coupled activity, of about 20-25%, was observed in membrane vesicles isolated from cells when a histidine epitope tag was incorporated onto the b wt or b +7 subunit, KM2/pTAM37 or KM2/pTAM35, respectively. The decrease in coupled activity also paralleled with the reduction seen in F 1 -ATP hydrolysis activity. Moreover, the coupled activities observed in membranes isolated from cells coexpressing histidine-tagged and V5-tagged b subunits were, as expected, intermediary between the V5-tagged species and the histidine-tagged species. The larger reduction seen in the cells expressing a histidine epitope tag arose in part from the plasmid vector. The moderate-copy plasmid vector pACYC184 was used to express the histidine-tagged b subunits. When histidine-tagged b subunits were expressed from the pUC18 plasmid vector, both coupled activity and ATP hydrolysis activity were similar to untagged b subunits (data not shown). Hence the reductions in activity observed in membranes from the histidine-tagged strains reflected reduced amounts of intact F 1 F 0 ATP synthase incorporated into the membranes. For the purposes of this study, the important parameters were the presence of an intact and functional F 1 F 0 ATP synthase enzyme complex and the ability to distinguish the two different tagged b subunits via Western blot. Detections of heterodimers Dimerization of the b subunits is thought to occur early during enzyme assembly, making the requirement for immediate dimerization of b via co-translation of a single b subunit transcript, feasible. Therefore, it was necessary to establish whether a two

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104 plasmid expression system could be utilized in order to direct production of two different b subunits that will dimerize to form an intact F 1 F 0 enzyme complex. Two wild-type length b subunits, expressed from plasmids pTAM37 (b wt-his Cm r ) and pTAM46 (b wt-V5 Ap r ), were expressed in the same cell and studied for their ability to dimerize. Under the appropriate conditions, the histidine-tagged b subunits allowed for the easy purification of enzyme from a crude membrane preparation using a nickel affinity resin. Via immunoblot analysis, the presence or absence of the V5 epitope tag provided a means to determine whether the two different tagged b subunits are interacting to form a heterodimer. Immunoblot analysis of crude membrane preparations using anti-b antibodies showed the presence of the b subunit in all strains complemented with an epitope-tagged b subunit (Figure 2-11A, Lanes 2-4). As expected, only b subunits with a histidine tag were retained by Ni-resin purification (Figure 2-11A, Lanes 5-12). Importantly, F 1 F 0 complexes with only the V5-tagged b subunits were completely removed from the resin during the wash steps (Figure 2-11A, Lanes 7-8). To detect a heterodimer formation consisting of wild-type length b subunits, b wt-his and b wt-V5 expressed from two plasmids, immunoblot analysis using the anti-V5 antibody was performed (Figure 2-11B). When the two plasmids were coexpressed in the same cell the anti-V5 antibody detected the presence of b wt-V5 after Ni-purification, indicating that the expression system was successful and the tags do not hinder hetero-dimerization (Figure 2-11B, lanes 11-12). To confirm that the F 1 F 0 ATP synthase complex did not rupture during the solubilization and purification procedures, the tagged b subunits were either mock treated or treated with the homobifunctional crosslinker, BS 3 after the addition of detergent. As

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105 expected, crosslinked b subunits were readily observed only in samples that had undergone BS 3 treatment and survived Ni-resin purification (Figure 2-11A, Lanes 6, 10, 12). BS 3 chemical crosslinking had also demonstrated that the b +7-his subunit dimer was stable during solubilization with tegamineoxide WS-35, taurodeoxycholate and lauryldimethylamine oxide, but not with sodium dodecyl sulfate (SDS) (Figure 2-5). As an added precaution to ensure there was no nonspecific aggregation of b subunits after membrane solubilization or during Ni-resin purification, two independent membrane preparations were mixed together (Figure 2-11, Lanes 9-10). Membrane vesicles derived from strains KM2/pTAM37 (b wt-his ) and KM2/pTAM46 (b wt-V5 ) were mixed, the membranes were solubilized with 0.2% tegamineoxide and allowed to incubate at room temperature for 30 minutes, and the treated with BS 3 before performing Ni-resin purification. Immunoblot analysis with anti-b antibodies showed normal levels of histidine-tagged b subunit upon Ni-resin purification (Figure 2-11A, Lanes 9-10). No V5-epitope was present after Ni-resin purification indicating nonspecific aggregation was not a factor in the observed results (Figure 2-11B, Lanes 9-10). Therefore, any observed interactions of b subunits must have been due to heterodimers integrating within an intact F 1 F 0 ATP synthase complex in a cell expressing both b subunits. Importantly, immunoblotting using the anti-V5 antibody clearly detected V5 epitope tagged b subunits in Ni-resin purified samples when the two were coexpressed (Figure 2-11B, Lanes 11-12).

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106 Figure 2-11. Ni-resin purification of F 1 F 0 ATP synthase treated with the cross-linker BS 3 Crude membrane preparations of KM2 expressing the normal length b subunits with epitope tags were mock treated (-) or treated (+) with 1mM bis(3-sulfo-N-hydroxysuccinimide ester) (BS 3 ) for 20 minutes at room temperature. The cross-linking reaction was stopped by addition of 100 mM ethanolamine HCl, pH 7.5 for 10 minutes. The products were then solubilized with 0.2% tegamineoxide WS-35 and subjected to Ni-resin purification before separation on a 15% polyacrylamide Tris-glycine SDS gel and subsequent Western blot analysis. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody. B) Proteins were transferred to a nitrocellulose membrane and probed using an anti-V5 mouse monoclonal antibody.

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107 Formation of mixed length b subunits in F 1 F 0 ATP synthase The two plasmid expression system was used to study whether unequal length b subunits could form a dimer or, alternatively, whether the b subunits dimerize and incorporate into enzyme complexes in a segregated manner. In order to determine if there is a direct protein-protein interaction between the different length b subunits, we investigated the ability of b +7-his to retain an interaction with a b wt-V5 or a b 7-V5 subunit following Ni-purification (Figure 2-12). All membranes prepared from strains expressing a b subunit were readily detectable and distinguishable by size on an immunoblot (Figure 2-12, Lanes 1-7). Only b subunits with a histidine tag were retained by Ni-resin purification (Figure 2-12A, Lanes 8-12). Immunoblot analysis using an anti-V5 antibody was performed on the membrane preparations and Ni-purified products (Figure 2-12B). The V5-epitope tag was detected only in membrane vesicles derived from KM2 strains expressing either the V5-tagged b subunit or the coexpressed V5-tagged and his-tagged b subunits (Figure 2-12B, Lanes 1-7). As expected, upon Ni-resin purification, the V5 epitope was not detected in samples containing only histidine-tagged b subunit (Figure 2-12B, Lane 8). Likewise, samples containing only the V5-tagged b subunits were not detected upon Ni-resin purification, signifying that they were efficiently removed from the resin during wash steps (Figure 2-12B, Lanes 9-10). Finally, we investigated the ability of b +7-his to dimerize to form intact F 1 F 0 ATP synthase complexes with b wt-V5 or b 7-V5 The b wt-V5 was detected with anti-V5 antibodies and was readily observed to dimerize with the b +7-his indicating a b wt-his /b wt-V5 interaction (Figure 2-12B, Lane 11). To a much lesser extent, the b 7-V5 subunit was also distinguishable with anti-V5 antibodies (Figure 2-12B, Lane 12). The data indicated that the F 1 F 0 ATP synthase enzyme

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108 complex could tolerate b subunit heterodimers with a size difference of at least 14 amino acids. Detection of b subunit heterodimers led directly to two additional questions. How many b subunit heterodimer F 1 F 0 ATP synthase complexes were formed? Were they active? We had not yet purified a heterodimeric complex to homogeneity so the direct assay has not been performed. However, an indication of activity could be assessed based on the percentage of homodimeric and heterodimeric F 1 F 0 complexes. Therefore, Figure 2-12. Ni-resin purification of F 1 F 0 ATP synthase expressing unequal length b subunits. IPTG-induced cells were lysed in a French pressure cell, and membrane vesicles were isolated by differential ultra-centrifugation. The crude membrane preparation was solubilized with 0.2% tegamineoxide WS-35 and subjected to Ni-resin purification before separation on a 15% polyacrylamide Tris-glycine SDS gel and subsequent Western blot analysis. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody. B) Proteins were transferred to a nitrocellulose membrane and probed using an anti-V5 mouse monoclonal antibody.

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109 membranes were prepared from strains KM2/pTAM37/pTAM46 (b wt-his /b wt-V5 ), KM2/pTAM35/pTAM46 (b +7-his /b wt-V5 ) and KM2/pTAM35/pTAM47 (b +7-his /b 7-V5 ). All membranes prepared from strains expressing an epitope-tagged b subunit were readily detectable and distinguishable by size on an immunoblot of a 15 cm 15% SDS-PAGE gel (Figure 2-13A). Densitometry was used to determine the relative amounts of histidine-tagged and V5-tagged b subunits in samples coexpressing the different subunits (Figure 2-13B). Relative amounts of the histidineand V5-tagged b subunits were determined from crude membrane preparations that contained the different coexpressed b subunits. In each case, expression of the various length b his and b V5 subunits led to nearly equal incorporation into F 1 F 0 ATP synthase complexes (Figure 2-13B). Relative amounts of the b his and b V5 subunits were also determined from Ni-resin purified samples coexpressing both b subunits. Upon Ni-resin purification, all of the b V5 subunit seen on a Western blot must necessarily be incorporated into the enzyme complex as a heterodimer along with a b his subunit. Hence, when the two different wild-type length b subunits were coexpressed, there was at least 15% heterodimer formation (Figure 2-13B). It is possible that further manipulation of subunit expression and optimization of the purification procedure might result in a higher percentage of heterodimers found in the membranes. Somewhat lower percentages of heterodimer formation were found when b +7-his was coexpressed with b wt-V5 or b 7-V5 (Figure 5B). Based on the percentage of heterodimer formation and the activities observed when the different b subunits were coexpressed in the same cell (Figure 2-13, Table 2-1), it is likely that the heterodimeric F 1 F 0 ATP synthase species are functionally active.

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110 Figure 2-13. Quantitation of b subunit heterodimeric F 1 F 0 Membranes were prepared from IPTG-induced cells by differential centrifugation. A total of 15 mg membrane protein was solubilized, Ni-resin purified (1 ml packed resin volume), concentrated with a Microcon YM-10 and separated on a 15 cm 15% polyacrylamide Tris SDS gel to allow maximal separation. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody. This Western blot was deliberately overexposed to allow easy visualization of the V5-tagged b subunits in the Ni-resin purification lanes. B) Densitometric analysis of lower exposures was used to determine the relative amounts of histidine-tagged and V5-tagged b subunits in samples coexpressing the different subunits. Reletive amounts of b subunits were determined for crude membrane preparations and Ni-resin purified samples coexpressing both b subunits. The percent of heterodimer formation was calculated based on three independent membrane preparations as well as three separate exposure times.

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111 Discussion In the present chapter, we have developed an expression system that facilitates production, purification and detection of b subunits of unequal lengths within E. coli. The experiments involved an epitope tag system that allowed us to determine if the different b subunits segregated into homodimers, or alternatively, if a heterodimer of long and short b subunits can be incorporated into an F 1 F 0 ATP synthase complex. The histidine and V5 epitope tags introduced into the b subunits did not appreciably affect enzyme assembly or function. Expression of two different wild-type length b subunits led to three distinct F 1 F 0 ATP synthase complexes in the same cell; 1) a homodimer of histidine-tagged b subunits, 2) a homodimer of V5-tagged b subunits and 3) a heterodimer consisting of a histidine-tagged b and a V5-tagged b subunit. More importantly, three different F 1 F 0 ATP synthase complexes were present even when the b subunits were not of identical length. We observed dimerization of b subunits between b +7-his and both b wt-V5 and b 7-V5 This demonstrates that b subunits that differ in length by at least 14 amino acids can be incorporated into an enzyme complex. Given that the tether domain is likely an helix, the difference in length between the b subunits would be approximately 21 Dimerization could occur in two ways. First, assuming that the transmembrane domains were in parallel, the hydrophilic domains could be out of register. Alternatively, we favor a conformation in which both the transmembrane domains and the dimerization domains as defined by Dunn and coworkers (13) exist in parallel. This would require that a section of the tether domain in the longer b subunit be out of contact with the shorter b subunit (Figure 2-14). It is likely that a parallel alignment of the dimerization domain is required for enzyme assembly.

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112 Figure 2-14. Interactions of b subunits of unequal lengths. An epitope-tag system novel to F 1 F 0 ATP synthase studies was established in order to determine whether b subunits of unequal length could interact to form the dimer in an intact and functional enzyme complex. The b 7 subunit was expressed with a V5-epitope tag at its C-terminus (represented by the s shaped red line) and b +7 was expressed with a histidine tag at its N-terminus (represented by the curved oragnge line). The lightning bolt indicates a deletion and the orangte patch represents an insertion of amino acids. The histidine tag allowed for purification on a Nickel affinity column and the V5-tag allowed us to detect the presence of the b 7 subunit after Ni-purification using Western blot analysis. Expression of the two different epitope-tagged b subunits in the same cell leads to three distinct interactions within an F 1 F 0 ATP synthase complex (1) a homodimer of b 7-V5 (2) a homodimer of b +7-his and (3) a heterodimer consisting of both the shortened and lengthened b subunit.

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113 Recent electron microscopy and NMR studies have revealed a distinctive 20 bend in the b dimer within the tether domain (27, 136, 196). The research presented here suggests the possibility of straightening or further bending of the two b subunits within the peripheral stalk and lends support to the concept of a flexible peripheral stalk. This raises the question, why should the tether domain be so flexible as to allow insertions, deletions and dimerization of b subunits of unequal lengths? If one views the peripheral stalk to be a rope-like structure linking F 1 to F 0 then its position holding F 1 against the rotation of the central stalk would not be expected to be the same for counterclockwise and clockwise rotation. Flexibility of the tether domain might facilitate reorienting the peripheral stalk to act as a stator for rotation in either direction during ATP synthesis and ATP hydrolysis. The ability to generate and purify F 1 F 0 complexes with two genetically different b subunits provides a potentially useful experimental tool. It is now feasible to specifically label a single b subunit within a purified complex. This will facilitate biochemical modification experiments and the use of physical methods.

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CHAPTER 3 GENETIC COMPLEMENTATION BETWEEN MUTANT b SUBUNITS IN F 1 F 0 ATP SYNTHASE Introduction In the F 1 F 0 ATP synthase enzyme complex, the peripheral stalk consists of a parallel dimer of identical b subunits. Dimerization of the b subunits is thought to be an early event necessary for enzyme assembly and function (199). The two b subunits exist in an extended -helical conformation, spanning from the periplasmic side of the membrane to near the top of F 1 However, due to the asymmetric nature of the enzyme complex the two b subunits cannot participate in identical protein-protein interactions with the other subunits. In the amino-terminal membrane spanning region, the peripheral stalk contacts a single a subunit. Similarly, at the carboxyl end of the peripheral stalk, the two b subunits interact with a single subunit. The b dimer has been studied by a variety of traditional biochemical approaches such as CD spectroscopy, cross-linking, and sedimentation experiments (30, 166, 192, 196, 197, 203, 283). Limited structural information is available from studies of polypeptides modeling functionally defined domains of the b subunit. Dmitriev et al. studied the amino-terminal membrane spanning domain by NMR using a 34 amino acid polypeptide, revealing an -helix with a 20 bend at pro-27 and pro-28 (136). A systematic mutagenesis approach supported the model described in the NMR paper suggesting that the extreme amino-termini of the two b subunits were in close contact and then the subunits flare apart as they traverse the membrane (136, 202). X-ray crystallography of a model polypeptide reflecting residues 114

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115 62-122, corresponding to the dimerization domain, showed an extended, highly -helical structure (138). The structures of the tether domain contributing to the segment between the membrane surface and the bottom of F 1 and the F 1 binding domain in the carboxyl terminal region have yet to be determined. Previously we showed that b subunits with deletions and insertions in the tether region of up to eleven and fourteen amino acids, respectively, formed functional F 1 F 0 complexes (193, 194). These observations suggested that flexibility is an inherent characteristic of the peripheral stalk. This apparent flexibility also extended to the dimerization of the b subunits. When two b subunits with unequal length tether domains were expressed together, F 1 F 0 ATP synthase complexes containing heterodimeric peripheral stalks were assembled (195). F 1 F 0 complexes were able to tolerate the incorporation of two different b subunits with a size difference of at least fourteen amino acids. Although activity evidence suggested the heterodimeric F 1 F 0 complexes were likely functional, the earlier study was not design to rigorously demonstrate that these were active. A major problem that plagued all previous mutagenesis studies of the b subunit was that mutations constructed in the uncF(b) gene affected both subunits of the b homodimer. One missense mutation led to two amino acid replacements. Our ability to express two different b subunits in the same cell and detect b subunit heterodimer formation provided an approach to study the individual functional roles of the b subunits. In the present study, three different b subunit mutations were studied. Mutation of an evolutionarily conserved arginine, b arg36 to an isoleucine or glutamate has previously been shown to result in an intact, completely defective F 1 F 0 ATP synthase complex

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116 (215). Second, deletion of the last four amino acids at the C-terminus has been shown to affect the ability of the b dimer to form a stable interaction with the subunit F 1 resulting in a major defect in F 1 F 0 ATP synthase assembly (205, 206). Thirdly, insertions or deletions constructed in a cytoplasmic region of the b dimer that contains a stretch of hydrophobic amino acids, b !24-130 (VAILAVA) has been shown to completely affect the ability of the b subunit to form a stable dimer and insert into the membrane. Here, we demonstrate functional activity for F 1 F 0 ATP synthase complexes containing a heterodimeric peripheral stalk, with major defects occurring in the different domains of each b subunit. Materials and Methods A thorough account of many of the procedures used in Chapter 4 can be found in previous chapters. Many of the techniques, including recombinant DNA techniques, site directed mutagenesis, western blotting, as well as assays of protein concentration and F 1 F 0 ATP synthase activity have been described in detail in Chapter 2. A detailed description of purification of F 1 F 0 ATP synthases containing b subunit heterodimers to homogeneity can be found in Chapter 3. Materials Molecular biology enzymes and mutagenic oligonucleotides were obtained from Invitrogen (Carlsbad, CA), Life Technologies, Inc. (Grand Island, NY), New England Biolabs (Beverly, MA) and Stratagene (La Jolla, CA). Reagents were obtained from Sigma (St. Louis, MO), BioRad Laboratories (Hercules, CA) and Fisher Scientific (Pittsburgh, PA). Plasmid purification kits were acquired from Qiagen Inc. (Valencia, CA). The anti-rabbit immunoglobulin horseradish peroxidase-linked whole antibody (from donkey), anti-mouse immunoglobulin horseradish peroxidase-linked whole

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117 antibody (from sheep), Hybond ECL Nitrocellulose membrane, electrochemiluminescence Western blotting reagents and high performance chemiluminescence film were purchased from Amersham Biosciences (Piscataway, NJ). Polyclonal antibodies against SDS-denatured b subunit (284, 285) were generously provided by Dr. Karlheinz Altendorf (Universitt Osnabrck, Osnabrck, Germany. Monoclonal antibodies against the epitope found in the P and V proteins of the paramyxovirus, SV5 (V5 epitope tag) were purchased from Invitrogen. The Seize Primary Immunoprecipitation Kit was purchased from Pierce Biotechnology (Rockford, IL). Strains and Media The wild type b subunit expression plasmid, pKAM14, and plasmids used to express arg36 mutatations b subunits have been described previously (203, 215). The plasmids encoding the uncF(b) gene were used to complement E. coli strain KM2 (b) carrying a chromosomal deletion of the gene (218). All strains were streaked onto plates containing minimal A media supplemented with succinate (0.2% w/v), to determine enzyme viability. Cells harvested for membrane preparation were grown in Luria Broth supplemented with glucose (0.2% w/v) (LBG). Isopropyl-1-thio--D-galactopyranoside (IPTG)(40 g/ml), ampicillin (Ap) (100g/ml), and chloramphenicol (Cm) (25 g/ml) were added to media as needed. All cultures were incubated at 37C for the appropriate duration. Recombinant DNA Techniques Plasmid DNA was purified with the Qiagen Mini-Prep and Maxi-Prep kits. Restriction endonuclease digestions, ligations, and transformations were performed

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118 according to the recommendations of the manufacturers (New England Biolabs, Stratagene and Life Technologies, Inc.). Site-directed mutagenesis was performed either by means of a Stratagene Quikchange kit or by ligation-mediated mutagenesis. DNA fragments were separated in 0.8 % agarose gel by electrophoresis and purified using a Qiagen, Inc. QIAquick Gel Extraction kit. Plasmid sequences were determined by automated sequencing in the core facility of the University of Florida ICBR. Mutagenesis and Strain Construction Plasmid pKAM14 (b, Ap r ) (203) was used to construct b subunits with a deletion of the last four amino acids. Plasmids pKAM14 (b, Ap r ) (203), pTLC11 (b arg36ile Ap r ), or pTLC15 (b arg36glu Ap r ) (215) were used to construct the epitope tagged b subunits. Epitope tags were inserted into each of the plasmids using the Stratagene Quikchange kit. A four amino acid carboxyl-terminal truncation was accomplished by deletion of the final four codons of the uncF(b) gene to express b 153end (Appendix A) (Figure 3-1A). The restriction endonuclease recognition sequence for HindIII was constructed near the deleted sequence for an initial detection of the truncation. A histidine epitope tag was inserted at the N-terminus by mutagenesis between the first and second codons of the uncF(b) gene to express b his b 153end-his or b +124-130-his (Figure 3-1B). All of the recombinant histidine-tagged b subunit plasmids were then digested with PstI and NdeI and subsequently ligated into a plasmid conferring the chloramphenicol resistance gene and the pACYC184 origin of replication (Table 3-1). A V5 epitope tag was added to the C-terminus by site-directed mutagenesis before the termination codon of the uncF(b) gene to express b V5 b arg36ile-V5 or b arg36glu-V5 (Figure 3-1B). The recombinant V5-tagged b subunit plasmids included the ampicillin resistance gene and the pUC18 origin

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119 of replication. Unique restriction enzyme sites SphI and NdeI were constructed near the histidine and V5 epitope tag sequence, respectively, for an initial detection of the insertions, and then the nucleotide sequence was subsequently confirmed by automated sequencing in the ICBR core facility. Throughout the paper, the mutation and the epitope Figure 3-1. Oligonucleotides for epitope tags and C-terminal truncation of uncF(b). Shown are the sense strands of the mutagenic primer pairs. Mutagenesis was accomplished as described in the Materials and Methods. Green and red codons specify translation start and stop sites, respectively. The amino acids encoded in the sequences are labeled above the codons. A) Mutagenic primer encoding a four amino acid deletion (codons shown in purple) along with a silent mutations that introduced a new endonuclease recognition sequence, HindIII (underlined). B) Oligonucleotides designed to insert a histidine or V5-epitope tag (bold blue). The SphI and SacI restriction sites (underlined) were added along with the histidine and V5 epitope tag to facilitate screening.

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120 tag are indicated along with the plasmid name for clarity, for example plasmid pTAM51 (b 153end-his ). Each plasmid and the control plasmids pKAM14 (b) and pBR322 were expressed in the E. coli cell line KM2 (b) for study, so that the only b subunits in the cells were the product of the plasmid genes. The two plasmid expression system allowed expression of various combinations of histidine tagged and V5-tagged b subunits in the same cell (Figures 2C, 4C, 6C, 8C). Appropriate antibiotics were added to the growth medium, and in the case of the coexpressed plasmids, both ampicillin and chloramphenicol were added to select for cells expressing both plasmids. Preparative Procedures Inverted membrane vesicles from KM2 (b) strains expressing the desired b subunits were prepared essentially as described previously (194). Protein concentrations were determined by the bicinchoninic acid (BCA) assay (289). Ni-resin purification was achieved using the High Capacity Nickel Chelate Affinity Matrix (Ni-CAM) purchased from Sigma. A total of 5 mg of membrane protein was brought up to 1 mL with final concentrations of 0.2% tegamineoxide WS-35, 0.15 M NaCl, and 1 mM imidazole. The purification procedure was accomplished using the batch method as described by the manufacturer. Immunoblot Analysis Proteins were loaded on a 15% tris-glycine SDS gel and transferred onto nitrocellulose by electroblot. The b subunit antibody incubation was performed essentially as described previously (195), using a 1:25,000 dilution of the anti-b subunit antibodies. Secondary antibody incubation was performed with horseradish peroxidase-linked donkey anti-rabbit antibody (1:50,000), and the antibody was detected by

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121 enhanced chemiluminescence. The V5-subunit antibody incubation was performed as described by the manufacturer followed by a secondary antibody incubation with horseradish peroxidase-linked sheep anti-mouse antibody (1:10,000). Signals were visualized on high performance chemiluminescence film using a Kodak X-Omat. Assays of F 1 F 0 ATP Synthase Activity Growth on a minimal succinate medium was used as an initial, in vivo, assay for enzyme viability. ATP hydrolysis activity was assayed by the acid molybdate method (146). Membranes were assayed in buffer (50 mM Tris-HCl, 1 mM MgCl2, pH 9.1) to determine the linearity with respect to time and enzyme concentration. Membrane energization was detected by the fluorescence quenching of 9-amino-6-chloro-2-methoxyacridine (ACMA) (271). Results Construction and Growth Characteristics of Mutants To investigate the function of individual b subunits in a F 1 F 0 ATP synthase enzyme complex, plasmids expressing defective b subunits with either a histidine or a V5 epitope-tag were generated by site-directed mutagenesis. A total of five plasmids were constructed expressing the b wt-his b 153end-his b wt-V5 b arg36ile-V5 or b arg36glu-V5 subunits (Table 3-1). The epitope tags were needed to facilitate enzyme purification and subunit detection on a Western blot, respectively. In order to express two different b subunits in the same cell, we used the two-plasmid expression system described previously in Chapter 2(195). Enzyme complexes incorporated with a histidine or V5 epitope-tagged b subunit homodimer have been studied previously and were shown to result in a functional enzyme complex (195). E. coli strain KM2 (b) was used as the host because the uncF(b) gene has been deleted, eliminating any background b subunit.

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122 Table 3-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing epitope tagged uncF(b) genes Strains Description Growth 1 Specific activity 2,3 b, Ap r KM2/pKAM14 (+) +++ 1.18 0.05 wt b, Ap r KM2/pBR322 (-) 0.24 0.03 b, Cm r KM2/pTAM37 +++ 1.08 0.08 wt-his KM2/pTAM51 b, Cm 153end-his 0.22 0.05 r b, Ap r KM2/pTAM46 +++ 1.22 0.04 wt-V5 KM2/pTAM53 b, Ap arg36ile-V5 r 0.80 0.02 b arg36glu-V5 Ap r KM2/pTAM54 0.75 0.08 b, Cm KM2/pTAM38 +124-130-his 0.21 0.04 r b + b KM2/pTAM37/pTAM46 wt-V5 +++ 1.16 0.03 wt-his KM2/pTAM51/pTAM46 b + b 153end-his wt-V5 +++ 1.00 0.05 KM2/pTAM37/pTAM53 +++ 1.12 0.04 b wt-his + b arg36ile-V5 KM2/pTAM37/pTAM54 b + b wt-his +++ 1.10 0.05 arg36glu-V5 KM2/pTAM38/pTAM54 b +124-130-his + b wt-V5 +++ 1.20 0.03 KM2/pTAM51/pTAM53 b 153end-his + b arg36ile-V5 + 0.51 0.02 KM2/pTAM51/pTAM54 b 153end-his + b arg36glu-V5 0.37 0.07 1 E. coli strains were grown aerobically on succinate minimal medium. Colony size was scored after 72-hr incubation at 37C as: +++, 1.0 mm; ++, 0.3-0.5 mm; +, ~0.1 mm; -, no growth. 2 ATPase activities were measured as described under Materials and Methods. Units of specific activity = mol of PO 4 released per mg of protein/min S.D. Units were calculated from the slope of the line based on three measurements with incubations for 12 minutes. 3 Previously reported ATPase activities resulted from an abbreviated membrane preparation protocol. Activities reported here resulted from an additional wash step to remove nonspecifically bound ATPases. Previous analyses of FF ATP synthase complexes incorporated with the mutant b, b, b or b subunits found the mutants to be completely defective (205, 206, 215). Therefore, our ability to form FF complexes containing b heterodimers offered an approach to study activity in a complex with only a single functional b subunit protein. First, the effects of the b, b, b and b mutations were studied for dimerization with a wild type epitope tagged b subunit in order to complement the E. coli strain KM2 (b) (218). Growth on succinate minimal medium was used as an initial qualitative gauge of enzyme activity in vivo since E. coli strains lacking FF ATP synthase cannot derive energy from nonfermentable carbon sources (Table 3-1). As expected, when any one of the mutants was expressed 1 0 arg36ile arg36glu 153end +124-130-his 1 0 arg36ile-V5 arg36glu-V5 153end-his +124-130-his 1 0

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123 alone in the cells no growth was detected. In each case, the strains coexpressing the mutant epitope-tagged b subunits with a wild-type epitope-tagged b subunit grew comparably to the wild type strain. Heterodimer Formation of b arg36 Defective Subunits with b wt Although two unequal length b subunits formed a heterodimer in an intact F 1 F 0 ATP synthase complex (195), it was possible that a single amino acid substitution could affect dimerization, preventing assembly of a complex. To consider whether a V5 epitope-tagged b arg36 subunit, b arg36ile-V5 or b arg36glu-V5 could form a heterodimer with a wild type histidine epitope-tagged b subunit, b wt-his each pair of subunits were expressed in strains KM2/pTAM37/pTAM53 and KM2/pTAM37/pTAM54 (b) (Figure 3-2A and B). The b subunits were detected in membranes prepared from strains expressing a b subunit by immunoblot analysis (Figure 3-2A, Lanes 1-9). However, only b subunits in complexes with at least one histidine tag were retained by Ni-resin purification (Figure 3-2A, lanes 10-13). Immunoblot analysis using an anti-V5 antibody was performed on the membrane preparations and Ni-purified products (Figure 3-2B). The V5-epitope tag was detected only in membrane vesicles derived from KM2 strains expressing either the V5-tagged b subunit or the coexpressed V5-tagged and his-tagged b subunits (Figure 3-2B, Lanes 1-9). When only V5-tagged b subunits were expressed alone in a cell, no F 1 F 0 complexes were recovered from the Ni-resin purification (Figure 3-2B, Lane 10). Finally, we investigated the ability of b wt-his to dimerize to form intact F 1 F 0 ATP synthase complexes with b arg36ile-V5 or b arg36glu-V5 Both b arg36ile-V5 and b arg36glu-V5 were detected with anti-V5 antibodies after Ni-resin purification (Figure 3-2B, Lanes 12-13). The only mechanism for recovery of defective V5-tagged b subunits was through

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124 Figure 3-2. Ni-resin purification of F 1 F 0 ATP synthase incorporated with b arg36 subunit mutations. Cells expressing recombinant b subunits were lysed in a French pressure cell, and membrane vesicles were isolated by differential ultracentrifugation. The crude membrane preparation was solubilized with 0.2% tegamineoxide WS-35 and subjected to Ni-resin purification before separation on a 15% polyacrylamide Tris-glycine SDS gel for subsequent anti-b, or on a mini BioRad 15% Ready Gel for anti-V5 Western blot analysis (see Materials and Methods). A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody. B) Proteins were transferred to a nitrocellulose membrane and probed using an anti-V5 mouse monoclonal antibody. C) Line diagram of the different b dimers found in the cell. Green and red lines represent the histidine and V5 epitope tags, respectively. Orange represents the b subunit and the blue star represents the b arg36 mutation.

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125 dimerization with b wt-his indicating formation of heterodimeric F 1 F 0 ATP synthase complexes. Therefore, coexpression of the two different b subunits in the same cell led to three distinct interactions within an intact F 1 F 0 ATP synthase complex (Figure 3-2C): 1) a homodimer of b wt-his 2) a homodimer of b arg36ile-V5 or b arg36glu-V5 and 3) a heterodimer consisting of both the defective and wild type b subunit. Since F 1 displays reduced affinity for the membrane in the absence of intact F 0 total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly. Under conditions of high pH, F 1 can be released from the influence of F 0 (146), so the amount of ATPase activity in the solution was used as a measure of the amount of intact enzyme complex located in the membrane vesicles. Previous data indicated minimal affects on specific activity due to the epitope tags (195). Confirming these observations, the V5-epitope tag did not have an apparent affect on enzyme assembly. Membrane preparations with a b wt-V5 incorporated into the F 1 F 0 ATP synthase complex had virtually the same specific activity of the wild type strain (Table 3-1). A small decrease in specific activity was reproducibly observed in membrane vesicles isolated from cells when a histidine epitope tag was incorporated onto the b wt about 89% of the wild type strain. Furthermore, comparable activities were observed in samples when the histidine-tagged and V5-tagged b wt subunits were coexpressed. Also verifying previous data (215), when b arg36ile-V5 or b arg36glu-V5 was expressed alone in strains KM2/pTAM53 and KM2/pTAM54, the membranes retained abundant activity, about 60% and 54%, of the wild type strain, respectively (Table 3-1), indicating considerable amounts of intact F 1 F 0 ATP synthase complexes found in the membranes. When b arg36ile-V5 or b arg36glu-V5 were coexpressed with b wt-his by construction of strains

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126 KM2/pTAM37/pTAM53 or KM2/pTAM37/pTAM54 specific activities of about 94% and 91%, respectively, were observed indicating a quantity of intact F 1 F 0 ATP synthase complexes approaching the wild type strain levels. F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the epitope-tagged mutants was used as an indication of coupled activity. Acidification of inverted membrane vesicles was examined by fluorescence of ACMA (Figure 3-3). Membranes isolated from cells with a V5 epitope tag incorporated onto the b wt subunit, KM2/pTAM46 (b wt-V5 ), reproducibly displayed a very small reduction in coupled activity, correlating very well with the F 1 -ATP hydrolysis activity. Consistent with previous observations, a larger reduction in coupled activity of about 20% was observed in membrane vesicles isolated from strain KM2/pTAM37 (b wt-his ). The coupled activities observed in membranes isolated from cells coexpressing histidine-tagged and V5-tagged b subunits were intermediary between the V5-tagged species and the histidine-tagged species. Strains expressing either b arg36ile-V5 or b arg36glu-V5 displayed no ATP-dependent proton pumping activity as expected. Significantly, when either one of the b arg-36 mutants was coexpressed with b wt-his KM2/pTAM37/pTAM53 (b wt-his /b arg36ile-V5 ) or KM2/pTAM37/pTAM54 (b wt-his /b arg36glu-V5 ), the coupled activity observed was essentially the same as strain KM2/pTAM37/pTAM46 (b wt-his /b wt-V5 ). When expressed alone, the strains with b arg-36-V5 mutants had no activity and the strains expressing b wt-his displayed less activity than the b wt-his /b arg-36-V5 mutant enzyme. As a consequence, it was most likely that the F 1 F 0 ATP synthase complexes with either the b wt-his /b arg36ile-V5 or b wt-his /b arg36glu-V5 peripheral stalks were active. To demonstrate that the

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127 Figure 3-3. ATP-driven energization of membrane vesicles prepared from uncF(b) arg36 gene mutants. Membrane vesicles were prepared by differential centrifugation. Membrane protein (200 g) was suspended in 3 ml of assay buffer (50 mM MOPS, 10 mM MgCl 2 pH 7.3). The fluorescent dye ACMA was added to a final concentration of 1 M, and fluorescence was recorded with excitation at 410 nm and emission at 490 nm. ATP was added as indicated to a final concentration of 1mM. The samples for each trace have been labeled according to the b subunit mutation and the epitope tag, so the strains used as the sources of the samples were as follows: b, KM2/pBR322; b wt KM2/pKAM14; b wt-V5 KM2/pTAM46; b wt-his KM2/pTAM37; b arg36ile-V5 KM2/pTAM53; b arg36glu-V5 KM2/pTAM54; b wt-his /b wt-V5 KM2/pTAM37/pTAM46; b wt-his /b arg36ile-V5 KM2/pTAM37/pTAM53; b wt-his /b arg36glu-V5 KM2/ pTAM37/pTAM54.

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128 membrane vesicles were intact and closed, the level of NADH-driven fluorescence quenching was monitored for all membrane preparations. The levels of NADH-driven fluorescence quenching were strong and directly comparable in every case (data not shown, for a representative figure, see Figure 2-10). Heterodimer formation of b 153end-his Complemented with b wt-V5 Deletion of the last four amino acids at the C-terminus has been shown to dramatically affect the ability of the b dimer to form a stable interaction with the subunit F 1 resulting in a major F 1 F 0 ATP synthase assembly defect (205, 206). Immunoblot analysis was performed in order to detect a heterodimer interaction between b 153end and b wt (Figure 3-4A and B). Coexpression of pTAM51 (b 153end-his ) and pTAM46 (b wt-V5 ) in the same cell resulted in the appearance of a weak signal representing b wt-V5 in Ni-resin purified material (Figure 3-4B, Lane 8). Although there was no attempt to be quantitative in this assay, the appearance of the band suggested relatively inefficient assembly of these heterodimers. Nevertheless, when the two different b subunits are expressed in the same cell, three distinct F 1 F 0 complexes were assembled (Figure 3-4C). These included a homodimer of b wt-V5 in an intact F 1 F 0 ATP synthase complex, a homodimer of b 153end-his in a partially assembled defective enzyme, and a heterodimeric F 1 F 0 ATP synthase consisting of both subunits. Apparently, the b wt-V5 subunit stabilized the b 153end-his subunit within an intact F 1 F 0 ATP synthase complex. Total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly (Table 3-1). As expected, membranes with enzyme complexes incorporated with b 153end-his displayed a specific activity similar to the negative control, indicating virtually no interaction with F 1 Coexpression of pTAM51

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129 Figure 3-4. Ni-resin purification of F 1 F 0 ATP synthase containing a b subunit carboxyl-terminal truncation. Membrane preparation and Ni-resin purification and Western blot analysis were accomplished as described in Materials and Methods. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody or B) an anti-V5 mouse monoclonal antibody. C) line diagram of the different b dimers found in the cell.

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130 Figure 3-5. ATP-driven energization of membrane vesicles incorporated with F 1 F 0 ATP synthase containing a b subunit carboxyl-terminal truncation. Membrane preparation and ATP-driven proton pumping was accomplished as described in Materials and Methods. The samples for each trace have been labeled according to the b subunit mutation and the epitope tag, so the strains used as the sources of the samples were as follows: b, KM2/pBR322; b wt KM2/pKAM14; b wt-V5 KM2/pTAM46; b wt-his KM2/pTAM37; b 153end-his KM2/pTAM51; b wt-his /b wt-V5 KM2/pTAM37/pTAM46; b wt-V5 /b 153end-his KM2/pTAM46/pTAM51.

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131 (b 153end-his ) and pTAM46 (b wt-V5 ) resulted in a specific activity of about 81% of the wild type strain (Table 3-1). An indication of coupled activity was shown by F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the epitope-tagged mutants (Figure 3-5). Membranes containing only the b 153end-his subunit exhibited no coupled activity as expected. Membranes isolated from cells coexpressing pTAM51 (b 153end-his ) and pTAM46 (b wt-V5 ) displayed a slight reduction in coupled activity, of about 20%, compared to membranes from cells coexpressing b wt-V5 and b wt-his It is feasible that the observed reductions in enzymatic activities were entirely due to formation of the mutant homodimeric species; this could not be directly demonstrated by these methods due to background activity from b wt-V5 homodimeric F 1 F 0 ATP synthases present in the membranes. Heterodimer Formation of b +124-130-his Complemented with b wt-V5 It has been shown by Dr. Deepa Bhatt that insertions or deletions constructed in a hydrophobic stretch corresponding to amino acids 124-130 (VAILAVA) of the b subunit results in loss of enzyme function (Bhatt et al., manuscript in preparation). Immunoblot analysis was performed in order to detect a heterodimer interaction between b+124-130-his and bwt-V5 (Figure 3-6A and B). The b subunit was detected only in membranes prepared from strains expressing a functional b subunit (Figure 3-6A, Lanes 1-6). No b subunit was present in membranes when KM2 (b) was complemented with plasmid pTAM38 (b+124-130-his). Immunoblot analysis using an anti-V5 antibody was performed on the membrane preparations and Ni-purified products (Figure 3-6B). Interestingly, coexpression of pTAM38 (b+124-130-his) and pTAM46 (bwt-V5) in the same cell resulted in the appearance of a signal representing bwt-V5 in a Ni-resin purified sample (Figure 3-6B,

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132 Figure 3-6. Ni-resin purification of membranes incorporated with b +124-130-his subunit mutation. Membrane preparation and Ni-resin purification and Western blot analysis were accomplished as described in Materials and Methods. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody or B) an anti-V5 mouse monoclonal antibody. C) line diagram of the different b dimers found in the cell.

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133 Figure 3-7. ATP-driven energization of membrane vesicles incorporated with a defective b +124-130 subunit mutation. Membrane preparation and ATP-driven proton pumping was accomplished as described in Materials and Methods. The samples for each trace have been labeled according to the b subunit mutation and the epitope tag, so the strains used as the sources of the samples were as follows: b, KM2/pBR322; b wt KM2/pKAM14; b wt-V5 KM2/pTAM46; b wt-his KM2/pTAM37; b +124-130-his KM2/pTAM38; b wt-his /b wt-V5 KM2/pTAM37/pTAM46; b wt-V5 /b +124-130-his KM2/pTAM46/pTAM38.

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134 Lane 10). The only mechanism for recovery of b wt-V5 was through dimerization with b +124-130-his indicating that the b wt-his subunit rescued some of the defective b subunit and formed heterodimeric F 1 F 0 ATP synthase complexes. Therefore, coexpression of the two different b subunits in the same cell led to two distinct interactions within an intact F 1 F 0 ATP synthase complex (Figure 3-6C): 1) a homodimer of b wt-V5 and 2) a heterodimer consisting of both the defective and wild type b subunit. F 1 -ATP hydrolysis activity of total membrane protein was used as a test of F 1 F 0 ATP synthase assembly (Table 3-1). As expected, membranes with enzyme complexes incorporated with b +124-130-his displayed a specific activity similar to the negative control, indicating virtually no interaction with F 1 Coexpression of pTAM38 (b 124-130-his ) and pTAM46 (b wt-V5 ) resulted in a specific activity similar to the wild type strain (Table 3-1). It was likely that the F 1 F 0 ATP synthase complexes incorporated with the b 124-130-his /b wt-V5 were intact. An indication of coupled activity was shown by F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the epitope-tagged mutants (Figure 3-7). Membranes containing only the b 124-130-his subunit exhibited no coupled activity as expected. Membranes isolated from cells coexpressing pTAM38 (b 124-130-his ) and pTAM46 (b wt-V5 ) displayed virtually the same amount of coupled activity as the membranes from cells coexpressing b wt-V5 and b wt-his Since coexpression of the two different b subunits in the same cell led to only two distinct interactions, it was likely that the F 1 F 0 ATP synthase complexes incorporated with the b 124-130-his /b wt-V5 were functional.

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135 Mutual Complementation In order to look for mutual complementation of b subunits with defects in different functional domains, b arg36ile-V5 and b arg36glu-V5 were coexpressed with b 153end-his in the absence of a functional wild type b subunit. Significantly, when b 153end-his and b arg36ile-V5 were coexpressed in the same cell, KM2/pTAM51/pTAM53, small colonies were present after 5 days of growth. Expression of either b arg36ile-V5 or b 153end-his alone in KM2 (b) results in a completely defective enzyme complex, any F 1 F 0 ATP synthase activity must necessarily have come from a heterodimer formed from two defective b subunits. Therefore, the b arg36ile and b 153end mutations qualified as intergenic second site suppressor mutations. To determine whether there was a direct protein-protein interaction between two mutant subunits, we investigated the ability of b 153end-his to form an interaction with b arg36ile-V5 or b arg36glu-V5 by the standard immunoblot (Figure 3-8A and B). All of the controls provided on the first immunoblot were included and displayed similar results. An additional control was included to ensure complete solubilization of membrane vesicles was achieved and nonspecific aggregation of the b subunits was not a concern (Figure 3-8A and B, Lanes 10-11). Two independent membrane preparations were mixed together. Membrane vesicles derived from strains KM2/pTAM51 (b 153end-his ) and KM2/pTAM53 (b arg36ile-V5 ) or KM2/pTAM54 (b arg36glu-V5 ) were mixed and the membranes were solubilized with 0.2% tegamineoxide WS-35 prior to Ni-resin purification. Immunoblot analysis with anti-b antibodies showed normal levels of histidine-tagged b subunit upon Ni-resin purification (Figure 3-8A, Lanes 10-11). No V5-epitope was present after Ni-resin purification, indicating that neither non-specific

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136 Figure 3-8. Ni-resin purification of F 1 F 0 ATP synthase incorporated with complementing defective b subunits. Membranes preparation, Ni-resin purification and Western blot analysis was accomplished as described in Materials and Methods. A) Proteins were transferred to a nitrocellulose membrane and probed using a polyclonal anti-b antibody or B) an anti-V5 mouse monoclonal antibody. C) line diagram of the different b dimers found in the cell. Three interactions, b arg36mut-V5 /b arg36mut-V5 and b 153end-his /b 153end-his homodimers as well as b arg36mut-V5 /b 153end-his heterodimers, existed in the cell.

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137 Figure 3-9. ATP-driven energization of membrane vesicles incorporated with F 1 F 0 ATP synthase containing complementing defective b subunits. Membrane preparation and ATP-driven proton pumping was accomplished as described in Materials and Methods. The samples for each trace have been labeled according to the b subunit mutation and the epitope tag, so the strains used as the sources of the samples were as follows: b, KM2/pBR322; b wt KM2/pKAM14; b 153end-his KM2/pTAM51; b arg36ile-V5 KM2/pTAM53; b arg36glu-V5 KM2/pTAM54; b wt-his /b wt-V5 KM2/pTAM37/pTAM46; b 153end-his /b arg36ile-V5 KM2/pTAM51/pTAM53; b 153end-his /b arg36glu-V5 KM2/pTAM51/pTAM54.

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138 aggregation was nor incomplete solubilization was a factor in the observed results (Figure 3-8B, lanes 10-11). Therefore, in cells expressing two different b subunits, any observed interactions of b subunits must have been due to heterodimers integrating into an intact F 1 F 0 ATP synthase complex. Importantly, immunobotting using the anti-V5 antibody clearly detected V5 epitope-tagged b subunits in Ni-resin purified samples when two different b subunits, b 153end-his and b arg36ile-V5 or b arg36glu-V5 were coexpressed (Figure 3-8B, Lanes 12-13). The data indicated that dimerization between the two defective b subunits could occur and the F 1 F 0 ATP synthase complex accepted incorporation of a b heterodimer with mutations affecting two different domains. Expression of two different defective b subunits yielded three distinct F 1 F 0 ATP synthase complexes including the b arg36ile-V5 or b arg36glu-V5 homodimeric F 1 F 0 a partially assembled b 153end-his homodimer complex, and the heterodimeric F 1 F 0 ATP synthase complexes. Total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly (Table 3-1). Membrane vesicles isolated from strains KM2/pTAM51/pTAM53 (b 153end-his /b arg36ile-V5 ) and KM2/pTAM51/pTAM54 (b 153end-his /b arg36glu-V5 ) yielded membrane associated specific activities of about 29% and 14%, respectively, which was intermediate between the specific activities if either of the mutants were expressed alone in the cell. An indication of coupled activity was obtained by F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the epitope-tagged mutants (Figure 3-9). As expected, membranes from all three strains expressing defective b subunits by themselves showed no proton pumping activity. Membranes isolated from cells expressing b 153end-his /b arg36ile-V5 or b 153end-his /b arg36glu-V5 KM2/pTAM51/pTAM53

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139 or KM2/pTAM51/pTAM54, respectively, displayed coupled ATP-driven proton pumping of about 20% and 16%. Since either homodimer leads to a completely defective enzyme, any coupled activity observed necessarily came from an intact heterodimeric F 1 F 0 ATP synthase. The data indicated that F 1 F 0 ATP synthase complexes incorporated with heterodimers of two mutant b subunits were indeed functional. Furthermore, the results demonstrated that the roles of the two b subunits in the peripheral stalk were not equivalent. Discussion Historically the b subunit dimer has been viewed as a single functional unit. However, the asymmetric nature of the F 1 F 0 ATP synthase enzyme complex suggested that the functional role of each b subunit should not necessarily be considered equivalent. Protein-protein contacts made by one b subunit cannot be made by the other. In the present work, a unique expression system that facilitates the production, purification and detection of F 1 F 0 ATP synthase complexes incorporated with a b subunit heterodimer was utilize in order to study the functional roles of the two b subunits. The experiments involved a two-plasmid expression system that directed production of two different b subunits in the same cell and an epitope tag system that allowed detection of a b subunit heterodimeric F 1 F 0 ATP synthase species. Three regions of the b subunit were considered. An evolutionarily conserved arginine, b arg36 located near the interface between the membrane and tether domains had been found to be crucial for F 1 F 0 ATP synthase function (215). Enzyme complexes incorporated with a b arg36ile-V5 or b arg36glu-V5 were found to be intact yet functionally defective. Second, the C-terminal last four amino acids had been shown to be essential for the F 1 binding domain (206, 288).

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140 Enzyme complexes with a b 153end-his were found to be only partially assembled. Thirdly, insertions and deletions in a hydrophobic stretch of amino acids in the b subunit corresponding to amino acids 124-130 (VAILAVA) resulted in a complete loss of enzyme function. The b dimer was not found in membranes when cells expressed only the b +124-130 subunit. Heterodimerization was detected in cells expressing either arg36 mutation, b arg36ile-V5 or b arg36glu-V5 with b wt-his b 153end-his /b wt-V5 and b +124-130-his /b wt-V5 (Figure 3-10). Figure 3-10. Interactions of defective b subunit with wild type b subunits found in intact F 1 F 0 ATP synthase complexes. An epitope-tag system was used in order to determine if a defective b subunit could form a dimer with a wild type b subunit. A V5 epitope tag was constructed at the C-terminus of the b subunit (shown in red) or a histidine epitope tag was placed at the N-terminus of the b subunit. Four mutant b subunits were studied and all were found to form a dimer with a wild type b subunit (1) b arg36ile-V5 (2) b arg36glu-V5 (3) b 153end-his and (4) b +124-130-his

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141 Dimerization was observed in membrane preparations from cells expressing both b 153end-his /b arg36ile-V5 and b 153end-his /b arg36glu-V5 More importantly, enzyme complexes incorporated with the mutant heterodimers were functionally active, suggesting that each of the b subunits were complementing the other to form an intact and functional enzyme complex. This observation demonstrated unambiguously that F 1 F 0 ATP synthase complexes containing b heterodimers were active and provided evidence that each of the individual b subunits provide specialized functions within the peripheral stalk. Clearly, each of the mutant b subunits compensates for what the other is lacking. This raises a question concerning the relative positions of the individual b subunits of the peripheral stalk. In order for F 1 F 0 ATP synthase containing the two different mutant b subunits to be intact and functional, it is likely that the b arg36ile-V5 (or b arg36glu-V5 ) subunit must be positioned such that its extreme C-terminus forms the appropriate contacts with the subunit of F 1 Similarly, the b 153end-his subunit must be positioned so that its b arg36 makes the appropriate contacts with the F 0 subunits (Figure 3-11). Incorrect positioning of the mutant b subunits during assembly might be expected to lead to an inactive or partially assembled enzyme complex.

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142 Figure 3-11. Model of F 1 F 0 ATP synthase incorporated with complementing defective b subunits. The b arg36ile subunit was expressed with a V5 epitope tag at its carboxyl terminus (represented by the S-shaped red line) and the b 153end subunit was expressed with a histidine tag at its amino terminus (represented by the orange curved line). Complexes incorporated with a b subunit heterodimer resulted in a functional enzyme complex. The b arg36ile-V5 subunit provides an intact F 1 binding domain for interaction with the subunit, and the b 153end-his subunit carries out the interaction with the a subunit.

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CHAPTER 4 DEVELOPMENT OF CYSTEINE CHEMICAL MODIFICATIONS OF ALTERED b SUBUNITS Introduction F 1 F 0 ATP synthases utilize the electrochemical gradient of protons across membranes to synthesize ATP from ADP and P i (3, 290, 291). In E. coli, F 1 F 0 ATP synthase is a multimeric enzyme composed of twenty-two polypeptides of eight different types (Figure 4-1). The F 1 portion is composed of the subunits 3 3 and is responsible for catalysis. The F 0 portion of the enzyme consists of the ab 2 c 10 subunits and conducts proton translocation through the membrane. Two stalk structures link the F 1 and F 0 sectors. Proton translocation drives the rotation of the central stalk, known as the rotor, within the stationary 3 3 hexamer, resulting in conformational changes within the catalytic sites (3, 15, 16, 63, 64). A second stalk structure, known as the peripheral stalk, consists of the and b subunits (Figure 4-1). The role of the peripheral stalk is thought to be that of a stator, holding the hexamer in place against the rotation of the central stalk. The subunit can be chemically fixed to a single subunit without loss of enzyme activity, which would be expected if was part of the peripheral stalk linking F 1 and F 0 supporting the stator concept (185). The carboxyl termini of the and b subunits have been shown to be in direct contact (3, 142, 200, 219, 220). 143

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144 Figure 4-1. Model of E. coli F 1 F 0 ATP synthase. The peripheral stalk of F 1 F 0 ATP synthase is composed of the subunit of F 1 and the homodimer of b subunits of F 0 Shown in orange and red are the and b subunits, respectively. It has been shown previously that insertions and deletions of amino acids in the tether region of the b subunit do not affect enzyme activity. However, it was not known how the insertions or deletions were accommodated in the context of an intact enzyme. An understanding of the structure of the peripheral stalk can be obtained by investigating the apparent length of the peripheral stalk in shortened, lengthened and wild type b subunits.

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145 A dimer of two b subunits is the major component of the peripheral stalk of F 1 F 0 ATP synthase and is necessary for normal assembly and function. The b dimer spans the cytoplasmic membrane and reaches towards the topside of F 1 where it meets the subunit (Figure 4-1) (100, 141-143). A substantial amount of evidence suggests the b dimer to be at least 80% -helical, with 14% -turn, and exist in a parallel, elongated conformation, spanning about 45 from the top of the membrane to the bottom of the 3 3 hexamer or about 100 to the top of the hexamer (12, 13, 166, 196, 292). Four domains comprise the b subunits: the amino-terminal membrane spanning, tether, dimerization and the carboxyl terminal -binding domains (Figure 4-5) (13). Despite the previous model of the b subunit dimer, describing it as a rigid structural feature of F 1 F 0 ATP synthase, accumulating evidence suggests a more flexible stalk model (193-195, 202). It has been shown by Dr. Paul Sorgen that F 1 F 0 ATP synthase retains sufficient levels of activity upon relatively large deletions or insertions in the b subunit. An eleven amino acid deletion and a fourteen amino acid insertion in the region of the b subunit spanning the tether domain and the beginning of the dimerization domain was accommodated by the enzyme (Figure 4-5). Assuming -helical structure, this 21 insertion and 16 deletion corresponds to well over a third of the length spanning from the top of the membrane to F 1 or right under a quarter of the length spanning towards the top of F 1 Furthermore, four amino acid insertions and deletions were successfully accommodated throughout most of the dimerization and -binding domains of the b subunit (manuscript in preparation). In contrast to the previously accepted role of the b subunit as a rigid stator, these observations suggest that the role of the b dimer is more of a flexible or elastic structural feature during rotational catalysis. In the present

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146 chapter, cysteine chemical modifications were created in the and b subunits to provide reactive thiols groups for future labeling studies. Materials and Methods A more thorough account of many of the procedures used in Chapter 6 can be found in previous chapters. Many of the techniques, including recombinant DNA techniques, site directed mutagenesis, crude membrane preparation procedures, western blotting, as well as assays of protein concentration and F 1 F 0 ATP synthase activity have been described in detail in Chapter 2. Materials Molecular biology enzymes and mutagenic oligonucleotides were obtained from Invitrogen (Carlsbad, CA), Life Technologies, Inc. (Grand Island, NY), New England Biolabs (Beverly, MA) and Stratagene (La Jolla, CA). Reagents were obtained from Sigma (St. Louis, MO), BioRad Laboratories (Hercules, CA) and Fisher Scientific (Pittsburgh, PA). Plasmid purification kits were acquired from Qiagen Inc. (Valencia, CA). The anti-rabbit immunoglobulin horseradish peroxidase-linked whole antibody (from donkey), Hybond ECL Nitrocellulose membrane, electrochemiluminescence Western blotting reagents and high performance chemiluminescence film were purchased from Amersham Biosciences (Piscataway, NJ). Polyclonal antibodies against SDS-denatured b subunit (284, 285) were generously provided by Dr. Karlheinz Altendorf (Universitt Osnabrck, Osnabrck, Germany. Strains and Media The bacterial strains used to create the cysteine chemical modifications in the b subunit include the wild type b subunit expression plasmid, pKAM14 (b, Ap r ), and plasmids used to express b subunits shortened or lengthened by 11 amino acids, pAUL5

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147 (b 11 Ap r ) and pAUL47 (b +11 Ap r ), respectively, and have been described previously (193, 194, 203). The plasmids encoding the uncF(b) gene were used to compliment Escherichia coli (E. coli) strain KM2 (b) carrying a chromosomal deletion of the gene. Plasmid pJLG1 (a, c, b, Cm r ), generated by Dr. James Gardner in our lab, was used as a temporary vector in order to facilitate cloning due to its extra unique restriction enzyme sites. The wild type unc operon expression plasmid, pAES9 (abc, Cm r ), was used to create a cysteine-less F 1 F 0 ATP synthase complex and has been described previously (178). The plasmids encoding the unc operon (abc) were used to compliment E. coli strain 1100BC (abc) carrying a chromosomal deletion for the entire operon. All strains were streaked onto plates containing Minimal A media supplemented with succinate (0.2% w/v), to determine enzyme viability. Cells harvested for membrane preparation were grown in Luria Broth supplemented with glucose (0.2% w/v) (LBG). Isopropyl-1-thio--D-galactopyranoside (IPTG)(40 g/ml), ampicillin (Ap) (100g/ml), and chloramphenicol (Cm) (25 g/ml) were added to media as needed. All cultures were incubated at 37C for the appropriate duration. Recombinant DNA Techniques Plasmid DNA was purified with the Qiagen Mini-Prep and Maxi-Prep kits. Restriction endonuclease digestions, ligations, and transformations were performed according to the recommendations of the manufacturers (New England Biolabs, Stratagene and Life Technologies, Inc.). Site-directed mutagenesis was performed either by means of a Stratagene Quikchange kit or by ligation-mediated mutagenesis. DNA fragments were separated in 0.8 % agarose gel by electrophoresis and purified using a

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148 Qiagen, Inc. QIAquick Gel Extraction kit. Plasmid sequences were determined by automated sequencing in the core facility of the University of Florida ICBR. Mutagenesis and Strain Construction Plasmids pKAM14 (b, Ap r ) (203), pAUL5 (b 11 Ap r ) (194) or pAUL47 (b +11 Ap r ) (193) and pJLG1 (a, c, b, Cm r ) (167) were used to construct cysteine mutant b subunits. Mutagenesis was performed in three different subunits; therefore, the ultimate goal was to express the entire unc operon containing the desired cysteine mutations from one plasmid. Plasmid pJLG1 (a, c, b, Cm r ) was used as a temporary vector in order to facilitate cloning due to its extra unique restriction enzyme sites. b subunit mutations. In order to construct the b subunit mutations, plasmids pAUL5 (b 11 Ap r ), pAUL47 (b +11 Ap r ) and pJLG1 (a, c, b, Cm r ) were first digested with restriction enzymes PpuMI and NarI in order to move the b subunit deletion and insertion into the pJLG1 plasmid. The resulting 204 and 270 bp cassettes isolated from pAUL5 (b 11 Ap r ) and pAUL47 (b +11 Ap r ), respectively, were subsequently ligated into the pJLG1 vector, resulting in pTAM1 (b 11 Cm r ) and pTAM3 (b +11 Cm r ) (Table 4-1). In order to site specifically label the b subunit in future experiments, a native amino acid was mutated to a cysteine. Two sites were chosen for replacement, b ser84 and b gly43 located respectively above and below the region of length alteration. Furthermore, the b subunit had a native cysteine, b cys20 located in the membrane-spanning portion of the subunit, which was mutated to a serine. The mutations were created in each of the plasmids using the Stratagene Quikchange kit. Sense and antisense mutagenic oligonucleotides were created for each of the desired b subunit mutations (Appendix A) (Figure 4-2A). First, the native cysteine was abolished by mutagenesis of codon 20

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149 Table 4-1. Description of uncF(b) cysteine mutations Plasmid Description 1, 2 b Length b cys20ser b gly43cys b ser84cys Growth 3 pKAM14 b wt Ap r wt +++ pBR322 b, Ap r NA NA NA pTAM1 b 11 Cm r 11 nd pJLG1 4 b wt Cm r wt nd pTAM3 b +11 Cm r +11 nd pTAM8 b 11-cys20ser, gly43cys 11 +++ pTAM9 b cys20ser, gly43cys wt +++ pTAM10 b +11-cys20ser, gly43cys +11 +++ pTAM11 b 11-cys20ser, ser84cys wt +++ pTAM12 b cys20ser, ser84cys 11 +++ pTAM13 b +11-cys20ser, ser84cys +11 +++ 1 Description as found in text. 2 Plasmids pTAM1-pTAM13 all encode the uncB(a) and c genes, but for the sake of brevity, only the b subunit mutations are described. 3 Plasmids were transformed into E. coli strain KM2 (b) and grown aerobically on succinate minimal medium. Colony size was scored after 72-hr incubation at 37C as: +++, 1.0 mm; ++, 0.3-0.5 mm; +, ~0.1 mm; -, no growth. 4 Plasmid courtesy of Dr. James Gardner. of the uncF(b) gene to express b cys20ser b 11-cys20ser and b +11-cys20ser Then, a cysteine was introduced by mutagenesis of codon 43 (b gly43cys ) or codon 84 (b ser84cys ) of the uncF(b) gene to generate a set of six plasmids: pTAM9 (b cys20ser, gly43cys ), pTAM8 (b 11-cys20ser, gly43cys ), pTAM10 (b +11-cys20ser, gly43cys ), pTAM12 (b cys20ser, ser84cys ), pTAM11 (b 11-cys20ser, ser84cys ) and pTAM13 (b +11-cys20ser, ser84cys ) (Table 4-1). The existing unique restriction enzyme sites recognized by SnaBI, Bsp1285I and XbaI were abolished by silent mutation near the encoded b cys20ser b gly43cys and b ser84cys mutations, respectively, for an initial detection of mutations. Subsequently, the nucleotide sequence was subsequently confirmed by automated sequencing in the ICBR core facility (Figure 4-2A). The recombinant altered b subunits included the chloramphenicol resistance gene and the pACYA184 origin of replication.

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150 Figure 4-2. Oligonucleotides for cysteine mutagenesis of the unc operon. Shown are the sense strands of the mutagenic primer pairs. The desired mutations are shown in color. Bold script indicates a change in nucleotide. Restriction enzyme recognition sequences that were silently added or abolished are underlined. Mutations were introduced as described in the Materials and Methods. A) Blue indicates the cysteine mutations constructed in uncF(b). The SnaBI, Bsp1285I and XbaI restriction sequences were knocked out along with the b cys20ser b gly43cys and b ser84cys respectively, to facilitate screening. B) Red indicates the cysteine mutations generated in uncH(). The ClaI sequence was knocked out and the EcoRI site was generated along with cys140ser for screening purposes. C) Green indicates the cysteine mutation made in unc(). The BamHI sequence was added to screen for the cys90ser mutation.

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151 and subunit mutations. The subunit has two indigenous cysteines, cys64 and cys140 which have been shown to be highly reactive with maleimide reagents (293). We planned to chemically label both sites specifically, so plasmids were constructed which expressed only one of the two cysteines by mutating one, either cys64ser or cys140ser to a serine. Also, another cysteine located within the subunit of F 1 has been shown to be highly reactive to maleimide reagents. This cysteine was mutated to a serine without loss of enzyme function (198). Plasmid pAES9 (abc, Cm r ) was used to construct the cysteine mutant and subunits. Due to the difficulty of performing Quikchange on a 10.9 kb plasmid, it was necessary temporarily move the genes encoding the and subunits into plasmid pBluescript (pBS) by digesting pAES9 (abc, Cm r ) with the restriction enzymes BamHI and EcoRI, flanking the genes of interest, and then ligating the 3.8 kb cassette into pBS, pTAM2. Sense and antisense mutagenic oligonucleotides were created for each of the desired and subunit mutations (Figure 4-2B and C). First, each of the native cysteines found in the subunit were substituted with serines in two different plasmids by site-directed mutagenesis of codon 64 or 140 of the unc () gene to express cys64ser or cys140ser Then the native cysteine found in the subunit was changed to a serine by mutagenesis of codon 90 of the unc () gene to express cys90ser The ClaI sequence was silently knocked out and the EcoRI site was generated along with cys140ser (Figure 4-2B) and the BamHI sequence was added along with the cys90ser mutation (Figure 4-2C) for initial screening purposes and then the nucleotide sequence was subsequently confirmed by automated sequencing in the ICBR core facility. The genes encoding the and subunits with the generated cysteine

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152 mutations were then shuttled back into pAES9 (abc, Cm r ) to create plasmids pTAM20 ( cys90ser cys64ser abc, Cm r ) and pTAM21 ( cys90ser cys140ser abc, Cm r ). The recombinant plasmids included the chloramphenicol resistance gene and the pACYA184 origin of replication. Construction of intact mutant unc operons. The final step of plasmid construction was to move each of the previously generated b subunit cysteine mutants (Table 4-1) into both pTAM20 ( cys90ser cys64ser abc, Cm r ) and pTAM21 ( cys90ser cys140ser abc, Cm r ), generating plasmids pTAM22-pTAM33 (Table 4-2). Each of the plasmids encoded the entire unc operon with the cys90ser and b cys20ser mutations as well as every possible combination of shortened, lengthened or wild type length b subunits, b gly43cys or b ser84cys and cys64ser or cys64ser The final products included 12 plasmids that encoded 12 genetically different F 1 F 0 ATP synthase complexes as well as the chloramphenicol resistance gene and the pACYA184 origin of replication (Table 4-2). Crude Preparative Procedures Inverted membrane vesicles from KM2 (b) or 1100BC (abc) strains expressing the desired cysteine mutations were prepared essentially as described in Chapter 2 (194). Bacteria were grown in 500 mL LBG supplemented with the appropriate antibiotic, harvested, and passed through a French Pressure Cell one time at 14,000 psi. Membranes were then collected by differential centrifugation. Protein concentrations were determined by the bicinchoninic acid (BCA) assay (286). Assays of F 1 F 0 ATP Synthase Activity Growth on a minimal succinate medium was used as an initial, in vivo, assay for enzyme viability. ATP hydrolysis activity was assayed by the acid molybdate method

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153 (146). Membranes were assayed in buffer (50 mM Tris-HCl, 1 mM MgCl2, pH9.1) to determine the linearity with respect to time and enzyme concentration. Immunoblot Analysis Proteins were loaded on a 15% tris-glycine SDS gel and transferred onto nitrocellulose by electroblot. The b subunit antibody incubation was performed essentially as described previously, using a 1:25,000 dilution of the anti-b subunit antibodies. Secondary antibody incubation was performed with horseradish peroxidase-linked donkey anti-rabbit antibody (1:50,000), and the antibody was detected by enhanced chemiluminescence. Signals were visualized on high performance chemiluminescence film using a Kodak X-Omat. Results Construction and Growth Characteristics of Mutants Single cysteines, which will provide reactive thiols for site-specific chemical modification, were strategically placed within the F 1 F 0 ATP synthase enzyme complex. Several cysteine mutations were generated in the unc operon in a multi-step construction scheme to ultimately result in 12 plasmids encoding different combinations of six different amino acid substitutions as well as b subunits of altered length (Table 4-2) (Figure 4-3). A previous analysis of F 1 F 0 ATP synthase complexes with b subunits shortened and lengthened by 11 amino acids found the mutants to be essentially wild-type when expressed in the presence of 40 M IPTG (193, 194). Functional F 1 F 0 ATP synthase complexes have been studied with many different cysteine mutations;

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154 Table 4-2. Description of the unc operon cysteine mutations 1 Plasmid 2 Length 3 cys90ser b cys20ser b gly43cys b ser84cys cys140ser cys64ser pAES9 wt pBR322 NA NA NA NA NA NA pTAM22 11 pTAM23 wt pTAM24 +11 pTAM25 11 pTAM26 wt pTAM27 +11 pTAM28 11 pTAM29 wt pTAM30 +11 pTAM31 11 pTAM32 wt pTAM33 +11 1 Plasmids were designed to contain various combinations of the cysteine mutations in the b and subunits. Symbols: -, plasmid does not contain the listed mutation; plasmid encodes the mutation listed above. 2 All plasmids, with the exception of pBR322, encoded the entire unc operon and conferred chloramphenicol resistance. 3 Length of the tether domain of the b subunit. Symbols: empty plasmid vector; wt, normal length;11, eleven amino acid deletion; +11, eleven amino acid insertion.

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155 Figure 4-3. Expression plasmid of cysteine mutants. F 1 F 0 ATP synthase complexes with site-specific cysteine mutations were designed to be expressed from a single plasmid encoding the entire unc operon. Plasmid pTAM24 is one of twelve of the constructed plasmids. The name of the unc gene is listed followed by the subunit it expresses in parenthesis. The black stars represents codons that expressed a native cysteine that had been substituted with a serine. The red star represents a glycine to cysteine substitution at codon 43 of the uncF(b) gene. The green dot represents an 11 amino acid insertion. Plasmid pTAM24 expresses the entire unc operon with the following mutations: b +11, cys20ser, gly43cys cys140ser and cys90ser The plasmids confer chloramphenicol resistance and the pACYA184 origin of replication. Similar constructions include those listed in Table 4-2. however, it was necessary to determine whether the mutations generated in the present study would have affect activity. The effects of the cysteine mutations were studied by the ability of the plasmids to complement either the E. coli strain KM2 (b) (Table 4-1) or 1100BC (abc) (Table 4-3). Growth on succinate minimal medium was used

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156 as an initial qualitative gauge of enzyme activity in vivo since E. coli strains lacking F 1 F 0 ATP synthase cannot derive energy from nonfermentable sources. In each case, the strains expressing the different cysteine mutations grew comparable to the wild type strain (Table 4-3). Table 4-3. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing cysteine Strains 1 Description 2 Growth 3 Activity 4 pAES9 a, b, c, Cm r +++ 100% pBR322 empty vector, Ap r 12% pAUL5 5 b 11 Ap r +++ 90% pAUL47 5 b +11 Ap r +++ 87% pTAM22 cys90ser cys140ser b 11, cys20ser gly43cys +++ 89% pTAM23 cys90ser cys140ser b cys20ser gly43cys +++ 95% pTAM24 cys90ser cys140ser b +11, cys20ser gly43cys +++ 87% pTAM25 cys90ser cys140ser b 11, cys20ser ser84cys +++ 88% pTAM26 cys90ser cys140ser b cys20ser ser84cys +++ 101% pTAM27 cys90ser cys140ser b +11, cys20ser ser84cys +++ 89% pTAM28 cys90ser cys64ser b 11, cys20ser gly43cys +++ 92% pTAM29 cys90ser cys64ser b cys20ser gly43cys +++ 96% pTAM30 cys90ser cys64ser b +11, cys20ser gly43cys +++ 90% pTAM31 cys90ser cys64ser b 11, cys20ser ser84cys +++ 93% pTAM32 cys90ser cys64ser b cys20ser ser84cys +++ 99% pTAM33 cys90ser cys64ser b +11, cys20ser ser84cys +++ 91 % 1 With the exception of pAUL5 and pAUL47, all plasmids were expressed in the 1100BC cell line. Plasmids pAUL5 and pAUL47 encode only the b subunit and were expressed in the KM2 cell line. 2 The description for plasmids pTAM22-pTAM33 only list the subunits with cysteine mutations for brevity; however, these plasmids also encode the wild type genes for the , a and c subunits. 3 E. coli strains were grown aerobically on succinate minimal medium supplemented with 40 M IPTG. Colony size was scored after 72-hr incubation at 37C as: +++, 1.0 mm; ++, 0.3-0.5 mm; +, ~0.1 mm; -, no growth. 4 E. coli strains were grown in LBG supplemented with the appropriate antibiotics and 40 M IPTG. ATPase activities were measured as described under Materials and Methods. Activity is listed as a percentage of the wild type strain (pAES9), which was set to 100%. Units were calculated from the slope of the line based on three measurements with incubations for 12 minutes. 5 Plasmids courtesy of Dr. Paul Sorgen.

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157 Effects of Cysteine Mutations Membrane associated ATP hydrolysis activity. F 1 has very little affinity for the membrane in the absence of intact F 0 therefore, total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly. The cysteine mutations had very little, if any at all, affect on enzyme assembly. In general, membranes with the cysteine mutations incorporated into the F 1 F 0 ATP synthase complex had specific activities ranging from about 88 to 95% of the wild types strain (Table 4-3). These values were comparable to the effect of the eleven amino acid deletion and insertions in the absence of the cysteine mutations. In fact, membranes with F 1 F 0 ATP synthase complexes incorporated with b subunits that were wild type in length, yet carried all the cysteine mutations, ranged from 95 to 100% of the wild type strain. It was likely that the slight decrease in ATP hydrolysis activity was due to the change in length in the b subunits and not due to the cysteine mutations. Western blot analysis. If the b subunit of F 1 F 0 ATP synthase does not dimerize and become incorporated into an intact F 1 F 0 ATP synthase complex, it is generally turned over and therefore absent from membrane preparations. Therefore the presence of the b subunit in the membrane preparations was detected by Western blot analysis. Proteins from membrane preparations generated from E. coli strain 1100BD (abc) complemented with pAES9 (abc), pTAM31 ( cys90ser cys64ser ,, a, b 11, cys20ser ser84cys c), pTAM32 ( cys90ser cys64ser ,, a, b cys20ser ser84cys c) and pTAM33 ( cys90ser cys64ser ,, a, b +11, cys20ser ser84cys c) were separated by SDS-PAGE and detected with anti-b antibodes (Figure 4-4). The b subunit expressed from pTAM32 and pAES9 migrated to the same molecular weight as expected. The size

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158 difference in the b subunit is clear between the 11 and +11, confirming the expression of unequal length b subunits. Figure 4-4. Western blot analysis of cysteine mutant b subunits of differing length. 1100DBC cells were transformed with plasmids pAES9 (abc), pTAM31 ( cys90ser cys64ser ,, a, b 11, cys20ser ser84cys c), pTAM32 ( cys90ser cys64ser ,, a, b cys20ser ser84cys c) and pTAM33 ( cys90ser cys64ser ,, a, b+11, cys20ser ser84cys c). Proteins from a crude membrane preparation were separated on a 15% polyacrylamide Tris-HCl-SDS BioRad Ready gel and then transferred to a nitrocellulose membrane in order to probe with anti-b antibodies. Discussion In the current chapter, we have developed a set of twelve unc operon expression plasmids that encode amino acid substitutions to void the F 1 F 0 ATP synthase complex of all known reactive thiols as well as generate strategically placed cysteines. Cysteines were chosen because the thiol side chain is highly reactive and can be modified by maleimide reagents. The cysteine mutations did not affect enzyme assembly or function. Plasmids were designed to express a single cysteine at one of two locations in the subunit as well as one position, either above or below the site of insertion or deletion, in the b subunit (Figure 4-5). The idea for establishing a single reactive cysteine in both F 1 and F 0 is to use them as targets for labeling with fluorescence compounds. F 1 can readily be stripped from F 0

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159 Figure 4-5. Model of F 1 F 0 ATP synthase with cysteine substitutions in the b and subunits. The red stars indicate the approximate location of generated cysteines. The region colored green is the approximate location of the eleven amino acid insertion (b +11 ) or deletion (b 11 ). A set of twelve plasmids was designed to express one cysteine in the and unequally length b subunits. each cysteine labeled separately with a "donor" and "acceptor" fluorescent compound, and then stoichiometrically reconstituted under conditions of high ionic strength (203). With a single fluorescent compound located above and below the region of insertion or deletion in the tether domain of the b subunit, physical measurements can be calculated and compared. Comparing the length of the wild type, shortened and lengthened peripheral stalks will give some insight on the overall possible conformation of the altered F 1 F 0 ATP synthase complex. The F 1 F 0 ATP synthase enzyme complex must, in some way, adapt to the shortened or lengthened b subunits. This can occur in at least one of two ways: 1) distortion of F 1 to accommodate the change in length of the peripheral stalk or 2) distortion of the peripheral stalk itself. If the former was true, F 1 would become distorted by bending of the central stalk or compression of the 3 3 hexamer and the rigid stalk hypothesis would gain favor. Since F 1 F 0 ATP synthase function requires the ability of the central stalk to rotate freely while making specific interactions within the catalytic

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160 hexamer, this mechanism is highly unlikely. If the latter situation were true, distortion would be limited to the b dimer and the flexible stalk hypothesis would hold true. This could occur by straightening or increasing the naturally occurring 20 bend in the shortened or lengthened peripheral stalk, respectively, or by stretching or compressing the secondary structure in the b dimer. The use of FRET will provide evidence as to which of the hypotheses is true. One major limitation plagued this line of research. F 1 F 0 ATP synthase is incorporated with a homodimer of b subunits, hence two b subunits, each with a reactive cysteine. Since FRET requires only one b subunit be labeled, a system necessarily had to be developed to allow purification of F 1 F 0 ATP synthase complexes with two genetically different b subunits: one cysteine-less and one with a single cysteine. The system utilizes epitope tags placed on the b subunits and is discussed in detail in Chapter 2. Future FRET studies will use this epitope-tagged scheme in order to express F 1 F 0 ATP synthase complexes with a b dimer containing only one cysteine.

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CHAPTER 5 MUTAGENISIS OF THE AMINO AND CARBOXYL TERMINI OF THE b SUBUNIT IN F 1 F 0 ATP SYNTHASE Introduction Two stalk structures have been shown to link the F 1 and F 0 sectors (11, 187). The central, or "rotary" stalk, extends within the 3 3 hexamer. Protons diffuse through the membrane, via the a and c subunits of F 0 resulting in the rotation of the central stalk within the hexamer, ultimately generating ATP (4, 16, 189). The peripheral stalk, or "stator", is positioned to the side of the F 1 F 0 ATP synthase complex and reaches from the periplasmic leaflet of the membrane up to near the top of F 1 in an -helical highly extended conformation (12, 13, 166, 204, 294), where it makes contact with a single subunit (100, 141-143). The primary function of the peripheral stalk is to prevent the rotation of the 3 3 hexamer against the rotation of the central stalk. The peripheral stalk consists of the subunit of F 1 and the b subunit dimer of F 0 (198, 288), which have been shown to be in direct contact with each other at their carboxyl termini (3, 4, 200, 219, 220). The b subunit dimer is the major component of the peripheral stalk, anchoring F 1 to the membrane. Despite attempts by several other laboratories, there is presently no high-resolution structure of the entire b subunit. Therefore, model polypeptides have been constructed in order to elucidate the structure of the b subunit by domain. Four domains comprise the b subunits: the amino terminal membrane-spanning, tether, dimerization and the carboxyl terminal -binding domains (Figure 5-1) (12, 13). A model polypeptide comparable to residues 1-34, which contains 161

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162 the membrane-spanning domain, has been solved by nuclear magnetic resonance (NMR). The data revealed an -helical monomeric structure with a 20 bend at residues 23-26 (136). A series of crosslinking studies led to a dimeric model in which the extreme Figure 5-1 Amino acid sequence and domains of the E. coli b subunit. The b subunit is a 156 residue, 17,264 Dalton amphipathic polypeptide. Dunn and coworkers have defined four domains in the b subunit: the membrane spanning (blue), tether (orange), dimerization (green) and -binding domains (red). However, the boundaries of the dimerization domain are continually being refined. Dr. Paul Sorgen demonstrated that F 1 F 0 ATP synthase could retain sufficient levels of activity upon relatively large deletions or insertions of up to eleven and fourteen amino acids, respectively, in the tether and beginning of the dimerization regions. The black lines indicate the amino acids deleted and the cyan indicates the amino acids duplicated that resulted in a functional F 1 F 0 ATP synthase. Red lines indicate amino acids deleted (below sequence) or inserted (above sequence) that resulted in a loss of enzyme function.

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163 amino-termini of the b subunits crossed each other in close proximity, the two b subunits then angled apart as they traverse the membrane towards the cytoplasmic side (136). The tether domain, contained within residues 23 and approximately 60, does not add significant stability to the dimerization of the b subunits and its role is the least distinct portion of the b subunit. This is the region of the b subunit that can be seen in electron micrographs (EM) of intact F 1 F 0 ATP synthase complexes (11, 187). There is currently no high-resolution structure for this domain. The structural conformation of the tether domain is suggested to be that of an -helical coiled-coil based on the heptad repeat which extends from the membrane surface to residue b Ala79 (138, 203). It has been shown by Dr. Paul Sorgen that F 1 F 0 ATP synthase retains sufficient levels of activity upon relatively large deletions or insertions of up to eleven and fourteen amino acids, respectively, within the b subunit tether domain, suggesting that this domain contributes a significant degree of flexibility to the peripheral stalk (Figure 5-1) (193, 194). The dimerization domain is currently defined as residues 63-122 and is required for the b subunit to form a dimer. A crystal structure of a monomeric polypeptide, modeled from residues 62-122, has recently been solved and refined to 1.55 (138). Dunn and coworkers have constructed a model in which the two -helices of the b 62-122 region form a right handed coiled coil. Finally, the carboxyl-terminal region, residues 123-156, forms the -binding domain. The extreme two to four amino acids of the carboxyl terminus are significantly important for the ability of the b subunit to form an interaction with the subunit. Deletion of these amino acids has been known to inhibit F 1 F 0 ATP synthase function (205, 206).

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164 This chapter focuses on the extreme amino and carboxyl termini of the b subunit. At the amino end, several mutational studies have been conducted in the membrane-spanning region; however, very few were known to produce a deficient F 1 F 0 ATP synthase complex. A single amino acid mutation, b gly9asp has been suggested to have a strong negative influence proton conduction by the a and c subunits (209). A systematic mutational analysis of the membrane domain, performed by Andrew Hardy in our laboratory, supports the model predicted by Dmitriev et al. It appears the extreme amino termini of the b subunit dimer are in close contact with each other, accounting for most of the important b-b interactions in the membrane domain, and as the dimer traverses the membrane the two b subunits flares apart (202). Three amino acids that have been shown to exhibit the strongest crosslinking efficiency in the membrane-spanning domain were replaced with alanines (b asn2ala, thr6ala, gln10ala ), yielding a defecting F 1 F 0 ATP synthase complex. Here, we demonstrate that mutation of only a single amino acid, at positions 2, 6 or 10, does not significantly affect the function of the enzyme. Several insertions and deletions can be tolerated in the tether region of the b subunit dimer (Figure 5-1) (193, 194). Evidence that the b subunit may form specific interactions with an subunit of F 1 suggested that length alterations may not be tolerated in regions of b running parallel to the F 1 hexamer (198). Other members in our laboratory are currently studying a series of insertions and deletions throughout the dimerization and -binding domains. At the carboxyl terminal end of the b subunit, mutation of the last amino acid to a cysteine, b leu156cys and chemical crosslinking forms an uncoupled F 1 F 0 ATP synthase complex (198). Furthermore, deletion of as few as two to four amino acids yields a completely defective enzyme (205, 206). However, an

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165 insertion of amino acids had not been attempted. Here, we demonstrate that a four amino acid insertion at the b subunit carboxyl terminus does not affect the activity of the enzyme. Materials and Methods Many of the procedures utilized in Chapter 7 can be found in detail in the previous chapters. Many of the techniques, including recombinant DNA techniques, site directed mutagenesis, western blotting, as well as assays of protein concentration and F 1 F 0 ATP synthase activity have been described in detail in Chapter 2. Materials Molecular biology enzymes and mutagenic oligonucleotides were obtained from Invitrogen (Carlsbad, CA), Life Technologies, Inc. (Grand Island, NY), New England Biolabs (Beverly, MA) and Stratagene (La Jolla, CA). Reagents were obtained from Sigma (St. Louis, MO), BioRad Laboratories (Hercules, CA) and Fisher Scientific (Pittsburgh, PA). Plasmid purification kits were acquired from Qiagen Inc. (Valencia, CA). Strains and Media The wild type b subunit expression plasmid, pKAM14, has been described previously (193, 194, 203). The plasmids encoding the uncF(b) gene were used to compliment E. coli strain KM2 (b) carrying a chromosomal deletion of the gene (218). All strains were streaked onto plates containing Minimal A media supplemented with succinate (0.2% w/v), to determine enzyme viability. Cells harvested for membrane preparation were grown in Luria Broth supplemented with glucose (0.2% w/v) (LBG). Isopropyl-1-thio--D-galactopyranoside (IPTG)(40 g/ml) and ampicillin (Ap)

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166 (100g/ml) were added to the media as needed. All cultures were incubated at 37C for the appropriate duration. Recombinant DNA Techniques Plasmid DNA was purified with the Qiagen Mini-Prep and Maxi-Prep kits. Restriction endonuclease digestions, ligations, and transformations were performed according to the recommendations of the manufacturers (New England Biolabs, Stratagene and Life Technologies, Inc.). Site-directed mutagenesis was performed either by means of a Stratagene Quikchange kit or by ligation-mediated mutagenesis. DNA fragments were separated in 0.8 % agarose gel by electrophoresis and purified using a Qiagen, Inc. QIAquick Gel Extraction kit. Plasmid sequences were determined by automated sequencing in the core facility of the University of Florida Interdisciplinary Center for Biotechnology Research (ICBR). Mutagenesis and Strain Construction Plasmid pKAM14 (b, Ap r ) (203) was used to construct all the b subunit mutants. The mutations were created in each of the plasmids using the Stratagene Quikchange kit. Amino terminal mutations. An alanine scan at the amino terminus of the b subunit, producing b asn2ala, thr6ala, gln10ala was performed by Andrew Hardy, resulting in the expression of a defective F 1 F 0 ATP synthase complex. Here, the individual sites were mutated, one at a time, to determine if the defect was the result of any particular amino acid. Sense and antisense mutagenic oligonucleotides were designed for each of the desired b subunit mutations (Appendix A) (Figure 5-2A). Three separate site-directed mutagenesis reactions were performed at codons 2, 6 and 10 of the uncF(b) gene in order to express mutant b subunits from plasmids pTAM43 (b asn2ala ), pTAM44 (b thr6ala ) and

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167 Figure 5-2. Oligonucleotides for mutagenesis at the amino and carboxyl termini in the unc operon. Shown are the sense strands of the mutagenic primer pairs. The amino terminus mutations are depicted in orange and the carboxyl terminus insertion is shown in blue. Bold script indicates a change in nucleotide. Green and red indicate start and stop codons, respectively. Restriction enzyme sequences that were silently added are underlined. Mutations were introduced as described in the Materials and Methods. A) The DraI, StuI and MfeI restriction sequences were added along with the b asn2ala b thr6ala and b asn10ala respectively, to facilitate screening. B) The codons expressing the last four amino acids of the b subunit (VAEL) were duplicated or deleted in order to insert or remove four amino acids at the extreme carboxyl terminus. The SacI or HindIII restriction sequences were added, respectively, along with the mutations for screening purposes.

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168 pTAM45 (b gln10ala ) (Table 5-1). The restriction enzyme sites recognized by DraI, StuI and MfeI were silently added near the encoded b asn2ala b thr6ala and b gln10ala mutations (Figure 5-2A), respectively, for an initial detection of mutations, and then the nucleotide sequence was subsequently confirmed by automated sequencing in the ICBR core facility. The recombinant mutant b subunits included the ampicillin resistance gene and the pUC18 origin of replication. Carboxyl terminal mutation. The b subunit dimer can tolerate relatively large insertions and deletions in the tether region (193, 194). Insertions have been generated throughout the dimerization and -binding domains, and the corresponding deletions are currently under construction by other members of the laboratory. Here, a four amino acid carboxyl-terminal truncation was accomplished by deletion of the final four codons of the uncF(b) gene to express b 153end from plasmid pTAM51 (Figure 5-2B). Likewise, a four amino acid insertion was constructed by duplication of the final four codons of the uncF(b) gene to express b +153-156 (Figure 5-2B). The restriction endonuclease recognition sequence for HindIII or SacI, respectively, was constructed near the deleted sequence for an initial detection of the truncation and deletion, and then the nucleotide sequence was later established by automated sequencing in the ICBR core facility (Figure 5-2B). The recombinant lengthened b subunit included the pUC18 origin of replication and conferred ampicillin resistance. Crude Preparative Procedures Inverted membrane vesicles from KM2 (b) or 1100BC (abc) strains expressing the desired cysteine mutations were prepared essentially as described in Chapter 2 (194). Bacteria were grown in 500 mL LBG supplemented with the

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169 appropriate antibiotic, harvested, and passed through a French Pressure Cell one time at 14,000 psi. Membranes were then collected by differential centrifugation. Protein concentrations were determined by the bicinchoninic acid (BCA) assay (286). Assays of F 1 F 0 ATP Synthase Activity Growth on a minimal succinate medium was used as an initial, in vivo, assay for enzyme viability. ATP hydrolysis activity was assayed by the acid molybdate method (146). Membranes were assayed in buffer (50 mM Tris-HCl, 1 mM MgCl2, pH9.1) to determine the linearity with respect to time and enzyme concentration. Membrane energization was detected by the fluorescence quenching of 9-amino-6-chloro-2-methoxyacridine (ACMA) (271). Results This chapter focuses on the extreme amino and carboxyl termini of the b subunit. Work accomplished at the amino terminal was an extension of studies accomplished by Andrew Hardy. Work accomplished at the carboxyl terminal end was a contribution to an insertion and deletion analysis, currently in progress, of the b subunit dimerization and -binding domains. Amino Terminal Mutations Mutations simultaneously effecting amino acids b asn2 b thr6 and b gln10 produces a completely deficient F 1 F 0 ATP synthase complex (202). To investigate the mutations individually, the sites were each mutated one by one and then the effects were studied. Construction and growth characteristics of mutants A previous analysis of F 1 F 0 ATP synthase complexes incorporated with b subunits including three mutations at codons 2, 6 and 10 (b asn2ala, thr6ala, gln10ala ) found the

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170 Table 5-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing uncF(b) mutations at the amino terminus Strains Description Growth 1 Activity 2 KM2/pKAM14 (+) b wt Ap r +++ 1.46 0.03 KM2/pBR322 (-) b, Ap r 0.17 0.02 KM2/pAWH24 3 b asn2ala, thr6ala, gln10ala Ap r 0.83 0.09 KM2/pTAM43 b asn2ala Ap r +++ 1.49 0.09 KM2/pTAM44 b thr6ala Ap r +++ 1.13 0.07 KM2/pTAM45 b gln10ala Ap r +++ 1.15 0.04 1 E. coli strains were grown aerobically on succinate minimal medium supplemented with 40 M IPTG. Colony size was scored after 72-hr incubation at 37C as: +++, 1.0 mm; ++, 0.3-0.5 mm; +, ~0.1 mm; -, no growth. 2 E. coli strains were grown in LBG supplemented with the appropriate antibiotics and 40 M IPTG. ATPase activities were measured as described under Materials and Methods. Units were calculated from the slope of the line based on three measurements with incubations for 12 minutes. 3 Work accomplished by Andrew W. Hardy. mutant to be completely defective; however, membrane-associated ATP hydrolysis activity and immunoblot analysis revealed that the triple mutant was forming intact F 1 F 0 ATP synthase complexes, suggesting that the mutant was incapable of performing coupled proton translocation (202). As a result of these observations, the alanine substitutions were studied individually. The amino acids expressed by codons 2, 6 and 10 were individually mutated to express an alanine, resulting in plasmids pTAM43 (b asn2ala ), pTAM44 (b thr6ala ), and pTAM45 (b gln10ala ) (Table 5-1). The effects of the mutations were studied by the ability of the plasmids to compliment E. coli strain KM2 (b) (218). Growth on succinate minimal media was used as an initial qualitative gauge of enzyme activity in vivo since E. coli strains lacking F 1 F 0 ATP synthase cannot derive energy from nonfermentable sources. In each case, the strains expressing the different alanine substitutions grew comparably to the wild type strain (Table 5-1).

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171 Effects of amino terminal mutations Since F 1 has little affinity for the membrane in the absence of intact F 0 total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly. Under conditions of high pH, F 1 can be released from the influence of F 0 (146), so the amount of ATPase activity in the solution was used as a measure of the amount of intact enzyme complex located in the membrane vesicles. The b asn2ala substitution, expressed from KM2/pTAM43, had no effect on enzyme assembly (Table 5-1). The b thr6ala and b gln10ala expressed from KM2/pTAM44 and KM2/pTAM45, had a slight, but not vitally significant effect on enzyme assembly. Membranes with a b thr6ala or b gln10ala incorporated into the F 1 F 0 ATP synthase complex had specific activities of about 77% of the wild type strain. F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the mutants was used as an indication of coupled activity. Acidification of inverted membrane vesicles was examined by fluorescence of ACMA (Figure 5-3). The level of NADH-driven fluorescence quenching was monitored for all membrane preparations to demonstrate that the vesicles were intact and closed. The levels of NADH-driven fluorescence quenching were strong and directly comparable in every case (data not shown, for a representative figure, see Figure 2-10). Membranes isolated from cells expressing b asn2ala KM2/pTAM43, displayed an insignificant reduction, approximately 1.6 %, in coupled activity, correlating with the wild-type-like F 1 -ATP hydrolysis activity (Figure 5-2 and Table 5-1). A larger reduction in coupled activity, of about 11%, was observed in membrane vesicles isolated from cells in which a b the6ala or b gln10ala expressed from KM2/pTAM44 or KM2/pTAM45, respectively, was

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172 Figure 5-3. ATP-driven energization of membrane vesicles prepared from b subunit membrane domain mutants. Cell membrane vesicles were prepared by differential centrifugation (see Materials and Methods). Membrane protein (200 g) was suspended in 3 mL of assay buffer (50 mM MOPS, 10 mM MgCl 2 pH 7.3). The fluorescent dye ACMA was added to a final concentration of 1 mM and fluorescence was recorded with excitation at 410 nm and emission at 490 nm. ATP was added as indicated to a final concentration of 1 mM. The samples for each trace have been labeled according to the mutation, so the strains used are as follows: b, membranes from strain KM2/pBR322; b wt KM2/pKAM14; b asn2ala, thr6ala, gln10ala KM2/pAWH24; b asn2ala KM2/pTAM43; b thr6ala KM2/pTAM44; b gln10ala KM2/pTAM45.

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173 incorporated into F 1 F 0 ATP synthase (Figure 5-3). The decrease in coupled activity paralleled the reduction seen in F 1 -ATP hydrolysis activity (Table 5-1). Carboxyl Terminal Mutations Since the original insertions and deletions constructed by Dr. Paul Sorgen in the in the tether region, a collaborative effort has been made by other members of the laboratory to study the remainder of the b subunit (Figure 5-4). Insertions and deletions constructed in a hydrophobic region of the b subunit in the F 1 binding domain b 124-130 was accomplished by Dr. Deepa Bhatt. Four amino acid duplications, scattered throughout the dimerization and F 1 binding domains, were constructed by an undergraduate in the laboratory Stephanie Cole and the corresponding deletions are currently underway by another undergraduate, Megan Greenlee. Here, a four amino acid insertion and deletion was constructed at the extreme carboxyl terminus of the b subunit. Construction and growth characteristics of mutants The C-terminal region of the b dimer is in direct contact with the extreme C-terminal end of the subunit (3, 185, 200, 219, 220). Small deletions at the extreme C-terminal end of the b subunit had been shown to inhibit F 1 F 0 ATP synthase function (206, 288). Two plasmids were generated to study the affects of deleting or duplicating the final four amino acids of the b subunit. Plasmids pTAM51 and pTAM52 were expressed in the E. coli KM2 (b) cell line to express b 153end or b +153-156 respectively (Table 5-2). Since E. coli strains lacking functional F 1 F 0 ATP synthase cannot derive energy from nonfermentable sources, growth on succinate minimal medium was used as an initial determination of enzyme activity in vivo. As expected, KM2 cells complemented with

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174 Figure 5-4. Amino acid insertion and deletion analysis of the E. coli b subunit. Dunn and coworkers have defined four domains in the b subunit: the membrane spanning (blue), tether (orange), dimerization (green) and -binding domains (pink). Dr. Paul Sorgen demonstrated that F 1 F 0 ATP synthase could retain sufficient levels of activity upon relatively large deletions or insertions in the tether and beginning of the dimerization regions. An analysis of the remainder of the b subunit is currently underway. Lines drawn below the sequence correspond to deletions, lines drawn above the sequence correspond to a duplication (insertion). The purple lines indicate the amino acids deleted and the cyan indicates the amino acids duplicated that resulted in a functional F 1 F 0 ATP synthase. Red lines indicate amino acids deleted (below sequence) or inserted (above sequence) that resulted in a loss of enzyme function. Grey lines indicate deletions that are currently under construction. plasmid pTAM51 (b 153end ) failed to grow on a succinate medium (Table 5-2). In contrast, the strain expressing b +153-156 KM2/pTAM52, grew comparably to the wild type strain. Hence, even though deletion of only four amino acids affected the ability of the b

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175 dimer to interact with the F 1 subunit, addition of four amino acids to the carboxyl terminus did not interfere with the interaction of the b and subunits. Effects of carboxyl terminal mutation Total membrane associated F 1 -ATP hydrolysis activity was used as a test of F 1 F 0 ATP synthase complex assembly (Table 5-2). As expected, membranes with enzyme complexes incorporated with b 153end displayed a specific activity similar to the negative control, indicating virtually no interaction with F 1 On the other hand, membranes with Table 5-2. Aerobic growth properties and membrane-associated ATP hydrolysis activity of mutants expressing uncF(b) insertions or deletions throughout the b subunit Strains Description Growth 1 % Activity 2 KM2/pKAM14 (+) b wt Ap r +++ 100 KM2/pBR322 (-) b, Ap r 13 KM2/pSC1 3 b +59-62 Ap r +++ 82 KM2/pSC2 3 b +73-76 Ap r +++ 89 KM2/pSC3 3 b +90-93 Ap r +++ 103 KM2/pSC4 3 b +102-105 Ap r +++ 90 KM2/pMMG2 4 b 102-105 Ap r +++ nd KM2/pDB20 5 b 124-125 Ap r 12 KM2/pDB21 5 b +124-125 Ap r 19 KM2/pDB22 5 b +124-127 Ap r 19 KM2/pDB23 5 b +124-130 Ap r (+) 21 KM2/pDB24 5 b 2x(+124-125) Ap r (+) 16 KM2/ pDB25 5 b 3x(+124-125) Ap r (+) 11 KM2/pSC5 3 b +143-146 Ap r +++ 87 KM2/pMMG1 4 b 143-146 Ap r +++ nd KM2/pTAM52 b +153-156 Ap r +++ 108 KM2/pTAM51 b 153-156-his Ap r 12 1 E. coli strains were grown aerobically on succinate minimal medium supplemented with 40 M IPTG. Colony size was scored after 72-hr incubation at 37C as: +++, 1.0 mm; ++, 0.3-0.5 mm; +, ~0.1 mm; -, no growth; (+), sporadic colonies from reversions. 2 E. coli strains were grown in LBG supplemented with the appropriate antibiotics and 40 M IPTG. ATPase activities were measured as described under Materials and Methods. Units were calculated from the slope of the line based on three measurements with incubations for 12 minutes. Value presented as percentage (%) of the wild type control (KM2/pKAM14). 3 Work accomplished by Stephanie Cole. 4 Work accomplished by Megan Greenlee. 5 Work accomplished by Dr. Deepa Bhatt.

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176 enzyme complexes incorporated with b +153-156 displayed a specific activity similar to the wild type strain. An indication of coupled activity was shown by F 1 F 0 ATP synthase-mediated ATP-driven proton pumping activity in membrane vesicles prepared from the mutants (data not shown, work accomplished by Stephanie Cole). Membranes containing the b 153end subunit exhibited no coupled activity as expected. Membranes isolated from cells expressing pTAM52 (b +153-156 ) exhibited coupled activity comparable to the wild type strain. The levels of NADH-driven fluorescence quenching were strong and comparable in every case (data not shown). The coupled activities observed reflected the observations from the F 1 -ATP hydrolysis activities (Table 5-2). Discussion In a collaborative effort, members of the laboratory have conducted an extensive mutational study of the entire length of the b subunit (Figure 5-5). At the amino terminal end of the b subunit, Andrew Hardy conducted a systematic mutational analysis of the membrane-spanning domain. His work established that there were specific sequence requirements in the b subunit membrane spanning domain and supported a model in which the extreme amino-termini of the two b subunits are in close contact with one another, accounting for most of the important b-b interactions in the membrane domain, and then flares apart as they cross the membrane (202). Here, an alanine scan was performed on three amino acids that had been shown to exhibit the strongest crosslinking efficiency in the membrane-spanning domain (b asn2ala, thr6ala, gln10ala ), yielding a defecting yet intact F 1 F 0 ATP synthase complex (202). It appears that there were adequate b-b interactions to support assembly of the enzyme, but the proton channel of F 0

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177 Figure 5-5. Mutations constructed throughout the b subunit. Shown are the approximate locations of the expressed b subunit mutants constructed and studied in our laboratory. Red indicates mutants constructed and studied in the present chapter. Shown in blue is the mutant created and studied in Chapter 3. The purple stars indicate mutants constructed and studied by Stephanie Cole. The blue stars indicate mutants constructed and studied by Megan Greenlee. The orange stars indicate a select few of the mutants constructed and studied by Dr. Deepa Bhatt. The green star designates the region of insertions and deletions studied by Dr. Paul Sorgen. The cyan star shows a mutant constructed and studied by Andrew Hardy. Growth indicates the ability of the strain to compliment the E. coli KM2 (b) cell line and grow on minimal succinate media. Symbols are as follows: +++, colonies 1.5 mm; -, no growth; (+), sporatic colonies from reversions.

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178 was not functional, suggesting that the b subunit membrane spanning domain may have a role in allowing the proton channel of the a and c subunits to align correctly (202). Here, we demonstrated that mutation of only a single amino acid, at positions 2, 6 or 10 (b asn2ala, b thr6ala or b gln10ala ), did not significantly affect the function of the enzyme. In fact, the minimal loss of enzyme activity associated from any of the single mutations did not nearly add up to the total defect of all three mutations combined. The results support the idea that although no single amino acid is necessary for the arrangement of a functional proton channel, several sites contribute synergistically to the formation of a functional F 1 F 0 ATP synthase enzyme complex. In the tether region of the b subunit, Dr. Paul Sorgen had constructed a series of insertions and deletions and observed that the altered b subunits were capable of forming the peripheral stalk of a functional F 1 F 0 ATP synthase complex (193, 194). It was not known whether similar insertions or deletions could be accommodated in other regions of the b subunit where crucial interactions with F 1 subunits are believed to occur. Dr. Deepa Bhatt observed that neither insertions nor deletions were tolerated in a small stretch of hydrophobic amino acids found in the F 1 binding domain, b 124130 (Figure 5-5). Four amino acid insertions, as well as the corresponding deletions, scattered throughout the remainder of the soluble portion of the b dimer were constructed by Stephanie Cole and Megan Greenlee and were found to result in a functional F 1 F 0 ATP synthase complex (Figure 5-5). In the present chapter, we confirmed that a four amino acid truncation at the extreme carboxyl terminus of the b subunit (b 153end ) results in a partially assembled defective enzyme complex. However, duplication of the last four amino acids (b +153-156 ) had no appreciable affect on enzyme assembly or function.

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179 The research presented here suggests the importance of the extreme amino and caborxyl termini. Although it was believed that the dimerization domain provided the majority of the protein-protein interactions by allowing the formation of the b subunit dimer (12, 13), our studies indicated that other regions of the b subunit are required for normal assembly of a functional F 1 F 0 ATP synthase complex due to contacts made with other subunits in the enzyme. Though the three amino acid substitutions at the amino terminus allowed dimerization and assembly of an intact F 1 F 0 ATP synthase, it appears that its sequence is crucial for the correct alignment of the proton channel formed by the a and c subunits. Likewise, the final four amino acids at the carboxyl terminus is not critical for the dimerization and partial assembly of the enzyme, however it is essential for the binding of F 1 due to its contacts with the subunit.

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CHAPTER 6 CONCLUSIONS AND FUTURE DIRECTIONS Conclusions The work presented in the preceding chapters had a considerable impact on the way the b 2 homodimer of the E. coli F 1 F 0 ATP synthase is viewed and may provide some implications concerning the genetically dissimilar b-type subunit heterodimers found in higher organisms. A previous analysis of the b subunit performed in Dr. Cains laboratory had suggested that the b 2 dimer has a flexible characteristic in its tether region (193, 194). A two plasmid expression system novel to the studies of the E. coli b subunit was developed in Chapter 2 in order to determine if the apparent flexibility extended to the dimerization of the b subunit. This work provided interesting information concerning the tether region of the peripheral stalk. The two plasmid expression system was also utilized in Chapter 3 to study the function of individual b subunits in the peripheral stalk by exploiting b subunits with known defective mutations. From this work, a model has been developed concerning the positioning of the two b subunits relative to the F 1 F 0 ATP synthase complex and the intersubunit contacts made by the individual b subunits. On another note, Chapter 4 describes single cysteine substitutions that were generated in different length b subunits with the ultimate goal of studying the apparent flexibility of the b 2 dimer. And finally, Chapter 5 describes mutagenesis work performed on the extreme aminoand carboxyl-termini of the b 2 dimer in contribution to ongoing experiments performed by others in the laboratory. The following sections will summarize and discuss the implications of the results from Chapters 2, 3, 4 and 5 and 180

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181 then discuss the future directions that will be taken as a direct result of the work accomplished in this dissertation. Chapter 2: Integration of Unequal Length b Subunits into F 1 F 0 ATP Synthase The work presented in Chapter 2 stemmed from observations made by Dr. Paul Sorgen in our laboratory that b subunits with deletions of up to eleven amino acids and insertions of up to fourteen amino acids, corresponding to approximately 16 and 21 respectively, formed intact and functional F 1 F 0 ATP synthase complexes (193, 194). This work had suggested that the tether region of the b subunit possesses a certain degree of flexibility. However, it was not known whether this flexibility extended to the dimerization of two b subunits of unequal lengths and their incorporation into an enzyme complex. An experimental system was developed to allow expression of two different b subunit genes and determine whether the differing b subunits were assembled into an F 1 F 0 ATP synthase complex. The experiments involved an epitope tag system that allowed us to determine if the different b subunits segregated into homodimers, or alternatively, if a heterodimer of long and short b subunits can be incorporated into an F 1 F 0 ATP synthase complex. The histidine and V5 epitope tags introduced into the b subunits did not appreciably affect enzyme assembly or function. Expression of two different wild-type length b subunits led to three distinct F 1 F 0 ATP synthase complexes in the same cell; 1) a homodimer of histidine-tagged b subunits, 2) a homodimer of V5-tagged b subunits and 3) a heterodimer consisting of a histidine-tagged b and a V5-tagged b subunit. More importantly, three different F 1 F 0 ATP synthase complexes were present even when the b subunits were not of identical length. We observed dimerization of b

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182 subunits between b +7-his and both b wt-V5 and b 7-V5 This demonstrates that b subunits that differ in length by at least 14 amino acids can be incorporated into an enzyme complex. Given that the tether domain is likely an helix, the difference in length between the b subunits would be approximately 21 Dimerization could occur in two ways. First, assuming that the transmembrane domains were in parallel, the hydrophilic domains could be out of register. Alternatively, we favor a conformation in which both the transmembrane domains and the dimerization domains as defined by Dunn and coworkers (13) exist in parallel. This would require that a section of the tether domain in the longer b subunit be out of contact with the shorter b subunit (Figure 2-14). It is likely that a parallel alignment of the dimerization domain is required for enzyme assembly. Recent electron microscopy and NMR studies have revealed a distinctive 20 bend in the b dimer within the tether domain (27, 136, 196). The research presented here suggests the possibility of straightening or further bending of the two b subunits within the peripheral stalk and lends support to the concept of a flexible peripheral stalk. This raises the question, why should the tether domain be so flexible as to allow insertions, deletions and dimerization of b subunits of unequal lengths? If one views the peripheral stalk to be a rope-like structure linking F 1 to F 0 then its position holding F 1 against the rotation of the central stalk would not be expected to be the same for counterclockwise and clockwise rotation. Flexibility of the tether domain might facilitate reorienting the peripheral stalk to act as a stator for rotation in either direction during ATP synthesis and ATP hydrolysis (Figure 1-7). The ability to generate and purify F 1 F 0 complexes with two genetically different b subunits provides a potentially useful experimental tool. It is now feasible to specifically

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183 label a single b subunit within a purified complex. This will facilitate biochemical modification experiments and the use of physical methods in future experiments. Chapter 3: Genetic Complementation between Mutant b Subunits in F 1 F 0 ATP synthase The work documented in Chapter 3 directly stemmed from the experimental system developed in Chapter 2. Our ability to genetically express two different b subunits in the same cell and detect heterodimer formation provided us with the unique opportunity to study each of the b subunits in the peripheral stalk as an individual b subunit rather than a dimer forming a single entity. A major problem that had plagued all previous mutagenesis studies of the b subunit was that mutations constructed in the uncF(b) gene affected both subunits of the b homodimer. One missense mutation led to two amino acid replacements. However, the asymmetric nature of the F 1 F 0 ATP synthase enzyme complex suggested that the functional role of each b subunit should not necessarily be considered equivalent. Protein-protein contacts made by one b subunit cannot be made by the other. The work presented Chapter 3 exploited three b subunit mutations that had already been characterized and shown to result in completely defective F 1 F 0 ATP synthase complexes when expressed in both b subunits. An evolutionarily conserved arginine, b arg36 located near the interface between the membrane and tether domains had been found to be crucial for F 1 F 0 ATP synthase function (215). Enzyme complexes incorporated with a b arg36ile-V5 or b arg36glu-V5 were found to be intact yet functionally defective. Second, the C-terminal last four amino acids had been shown to be essential for the F 1 binding domain (206, 288). Enzyme complexes with a b 153end-his were found to be only partially assembled. Thirdly, insertions and deletions in a hydrophobic stretch of amino acids in the b subunit corresponding to residues b 124-130 (VAILAVA) resulted in

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184 a complete loss of enzyme function. The b subunit dimer was not found in membranes when cells expressed only the b +124-130 subunit. Heterodimerization was detected in cells expressing either arg36 mutation, b arg36ile-V5 or b arg36glu-V5 with b wt-his b 153end-his /b wt-V5 and b +124-130-his /b wt-V5 (Figure 3-10). Dimerization was also observed in membrane preparations from cells expressing both b 153end-his /b arg36ile-V5 and b 153end-his /b arg36glu-V5 More importantly, enzyme complexes incorporated with the mutant heterodimers were functionally active, suggesting that each of the b subunits were complementing the other to form an intact and functional enzyme complex. This observation demonstrated unambiguously that F 1 F 0 ATP synthase complexes containing b heterodimers were active and provided evidence that each of the individual b subunits provide specialized functions within the peripheral stalk. Clearly, each of the mutant b subunits compensates for what the other is lacking. The work accomplished in this chapter raises a question concerning the relative positions of the individual b subunits of the peripheral stalk. In order for F 1 F 0 ATP synthase containing the two different mutant b subunits to be intact and functional, it is likely that the b arg36ile-V5 (or b arg36glu-V5 ) subunit must be positioned such that its extreme C-terminus forms the appropriate contacts with the subunit of F 1 Similarly, the b 153end-his subunit must be positioned so that its b arg36 makes the appropriate contacts with the F 0 subunits (Figure 3-11). Incorrect positioning of the mutant b subunits during assembly might be expected to lead to an inactive or partially assembled enzyme complex. An answer to this question will be a goal discussed in Future Directions.

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185 Chapter 4: Development of Cysteine Chemical Modifications of Altered b Subunits The work described in Chapter 4 was performed in order to examine the nature of the apparent flexibility of the tether region. It has been shown by Dr. Paul Sorgen that an eleven amino acid deletion and a fourteen amino acid insertion in the the b subunit spanning the tether domain and the beginning of the dimerization domain was accommodated by the enzyme (Figure 4-5). Assuming -helical structure, this 21 insertion and 16 deletion corresponds to well over a third of the length spanning from the top of the membrane to F 1 or right under a quarter of the length spanning towards the top of F 1 In contrast to the previously accepted role of the b subunit as a rigid stator, these observations suggested that the role of the b dimer is more of a flexible or elastic structural feature during rotational catalysis. In Chapter 4, cysteine chemical modifications were created in the and b subunits to provide reactive thiols groups for future labeling and measurement studies. A set of twelve unc operon expression plasmids that encode amino acid substitutions were developed to void the F 1 F 0 ATP synthase complex of all known reactive thiols as well as generate strategically placed cysteines. Cysteines were chosen because the thiol side chain is highly reactive and can be modified by maleimide reagents. The cysteine mutations did not affect enzyme assembly or function. Plasmids were designed to express a single cysteine at one of two locations in the subunit, cys64 or cys140 as well as one position, either above or below the site of insertion or deletion in the b subunit, b cys84 or b cys43 (Figure 4-5). The idea for establishing a single reactive cysteine in both F 1 and F 0 is to use them as targets for labeling with fluorescence compounds (see Future Directions). One major limitation plagued this line of research. F 1 F 0 ATP synthase is incorporated with a homodimer of b

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186 subunits, hence two b subunits, each with a reactive cysteine. Since only one b subunit was desired to be labeled, a system necessarily had to be developed to allow purification of F 1 F 0 ATP synthase complexes with two genetically different b subunits: one cysteine-less and one with a single cysteine. With the epitope-tag system developed in Chapter 2, this should not be a problem for future studies. Chapter 5: Mutagenesis of the Amino and Carboxyl Termini of the b subunit in F 1 F 0 ATP Synthase In a collaborative effort, members of the laboratory have conducted an extensive mutational study of the entire length of the b subunit (Figure 5-5). The work illustrated in Chapter 5 was achieved in order to contribute to two other projects in the lab. The chapter focuses on the extreme aminoand carboxyl-termini of the b subunit. At the amino end, a systematic mutational analysis of the membrane domain was performed by Andrew Hardy in our laboratory. His work established that there were specific sequence requirements in the b subunit membrane spanning domain and supported a model in which the extreme amino-termini of the two b subunits are in close contact with one another, accounting for most of the important b-b interactions in the membrane domain, and then flares apart as they cross the membrane (202). Three amino acids that have been shown to exhibit the strongest crosslinking efficiency in the membrane-spanning domain were replaced with alanines (b asn2ala, thr6ala, gln10ala ), yielding a defecting F 1 F 0 ATP synthase complex. It appeared that there were adequate b-b interactions to support assembly of the enzyme, but the proton channel of F 0 was not functional, suggesting that the b subunit membrane spanning domain may have a role in allowing the proton channel of the a and c subunits to align correctly (202). Work performed in contribution to this dissertation demonstrated that mutation of only a single amino acid, at positions 2, 6 or

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187 10 (b asn2ala, b thr6ala or b gln10ala ), did not significantly affect the function of the enzyme. In fact, the minimal loss of enzyme activity associated from any of the single mutations did not nearly add up to the total defect of all three mutations combined. The results support the idea that although no single amino acid is necessary for the arrangement of a functional proton channel, several sites contribute synergistically to the formation of a functional F 1 F 0 ATP synthase enzyme complex. In the other line of research described in Chapter 5, mutations were constructed at the extreme carboxyl-terminus of the b subunit in contribution to an extensive insertion and deletion analysis. In the tether region of the b subunit, Dr. Paul Sorgen had constructed a series of insertions and deletions and observed that the altered b subunits were capable of forming the peripheral stalk of a functional F 1 F 0 ATP synthase complex (193, 194). It was not known whether similar insertions or deletions could be accommodated in other regions of the b subunit where crucial interactions with F 1 subunits are believed to occur. Dr. Deepa Bhatt observed that neither insertions nor deletions were tolerated in a small stretch of hydrophobic amino acids found in the F 1 binding domain, b 124130 (Figure 5-5). Four amino acid insertions, as well as the corresponding deletions, scattered throughout the remainder of the soluble portion of the b dimer were constructed by Stephanie Cole and Megan Greenlee and were found to result in a functional F 1 F 0 ATP synthase complex (Figure 5-5). Work described in this dissertation confirmed that a four amino acid truncation at the extreme carboxyl terminus of the b subunit (b 153end ) results in a partially assembled defective enzyme complex. However, duplication of the last four amino acids (b +153-156 ) had no appreciable affect on enzyme assembly or function.

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188 The research presented here suggests the importance of the extreme amino and carboxyl termini. Although it was believed that the dimerization domain provided the majority of the protein-protein interactions by allowing the formation of the b subunit dimer (12, 13), our studies indicated that other regions of the b subunit are required for normal assembly of a functional F 1 F 0 ATP synthase complex due to contacts made with other subunits in the enzyme. Though the three amino acid substitutions at the amino terminus allowed dimerization and assembly of an intact F 1 F 0 ATP synthase, it appears that its sequence is crucial for the correct alignment of the proton channel formed by the a and c subunits. Likewise, the final four amino acids at the carboxyl terminus is not critical for the dimerization and partial assembly of the enzyme, however it is essential for the binding of F 1 due to its contacts with the subunit. Future Directions One of the major objectives for the field as a whole is to obtain a high-resolution structure of an intact F 1 F 0 ATP synthase complex, or at least a complete F 0 The high resolution structure of F 1 confirmed many existing views and it was also a valuable tool used to assist a plethora of experimental planning that concerned F 1 We believe the same will hold true for F 0 A high-resolution structure will help to unveil many of the mysteries of F 0 such as proton translocation, and the mechanism in which the c 10 ring rotates while the adjacent a and b subunits remain in place. Although the work illustrated in this dissertation has contributed a great deal of novel information concerning the b subunit of the E. coli F 1 F 0 ATP synthase, it also left many open questions for future investigators. The following section discusses some of the future directions that directly resulted from work accomplished in this dissertation.

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189 Complementing Mutant b Subunits The data presented in Chapter 3 leads to the immediate question, do additional pairs of mutant b subunits exist that can mutually complement each other? The asymmetric nature of the F 1 F 0 ATP synthase complex suggests that only one b subunit must make the appropriate contact within the enzyme. However, this trait may sometimes be restricted by the quaternary structure of the b subunit dimer. In other words, due to the coiled coil arrangement of the two b subunits, one region of a particular b subunit may make a contact near the amino terminal end and then that same b subunit may also align to make an additional important contact up towards the carboxyl terminal end. Pairs of plasmids encoding a mutant uncF(b) gene, which expresses a defective b subunit, are currently being cotransformed into KM2 (b) cells and studied for their ability to grow on succinate. Since the expression system requires one plasmid to carry the ampicillin resistance gene and the second to carry the chloramphenicol resistance gene, an undergraduate in the laboratory, Stacia Howard, is currently cloning some of the other uncF(b) gene mutants in the laboratorys possession into chloramphenicol resistant plasmids. Table 6-1 describes preliminary evidence of ATP-driven proton pumping of various combinations of mutant b subunits. Indeed it appears that at least one pair of b mutants, b N2A, T6A, Q10A /b arg83pro do not mutually suppress each other (Table 6-1). Furthermore, it appears that some of the complementing pairs have more activity than the other. Whether the apparent reduction in activity is due to a smaller percentage of heterodimeric F 1 F 0 or whether the assembled heterodimeric species are still impaired will require purification of the heterodimeric complexes to homogeneity (described next).

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190 Table 6-1. Preliminary data of coexpressed mutant b subunits. b +124-130 b 153end b arg83pro b arg36ile *** *** b arg36glu *** *** *** b N2A, T6A, Q10A * b +124-125 nd nd *** b +124-127 * b 124-127 nd b ala79lys *** *** *** Defective b subunits were coexpressed in the KM2 (b) cell line. Membrane preparation and preliminary ATP-driven proton pumping were accomplished as described in Chapter 2 Materials and Methods. Symbols: ***, proton-pumping activity at least 20% of samples prepared from wild type strains; *, <10% activity; -, no activity; nd, no data. Function of F 1 F 0 ATP Synthase Incorporated with b Subunit Heterodimers Work discussed in Chapters 2 and 3 have provided evidence that F 1 F 0 ATP synthase complexes can be incorporated with b subunit heterodimers. Although it was unambiguously demonstrated that the heterodimeric F 1 F 0 ATP synthase complexes were functional, a direct confirmation of activity will be pursued by performing activity assays on homogeneous enzyme preparations. These assays will also allow us to directly determine the activity of the complementing, double b subunit mutants (Chapter 3) relative to wild type F 1 F 0 complexes. A detailed account of a developing protocol for purifying the F 1 F 0 ATP synthase complexes with b subunit heterodimers can be found in Appendix B. Positions of the Individual b Subunits in F 1 F 0 ATP Synthase The two b subunits of F 1 F 0 ATP synthase have traditionally been viewed as a single functional entity. However, work described in Chapter 3 indicated that this may not be the case. F 1 F 0 ATP synthase complexes incorporated with a heterodimeric b subunit in which each b subunit contains a defective mutation were functional. This observation suggested that the individual b subunits have specialized roles within the enzyme

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191 complex. Furthermore, the asymmetric nature of the enzyme suggests that the two b subunits cannot participate in the exact same interactions. For example, the b subunit cannot have the same contacts with the single a in the membrane-spanning region and likewise with the subunit at the carboxyl-terminus. The unique experimental system described in Chapters 2 and 3 will be used to investigate the individual roles and positions of each b subunit. The experiments will heavily rely on the use of defective b subunits along with very efficient and well characterized chemical crosslinking techniques. Length of the Peripheral Stalk in F 1 F 0 ATP Synthase Complexes Incorporated with Shortened and Lengthened b Subunits FRET. Relatively large insertions and deletions in the tether region of the b subunit were accommodated by the F 1 F 0 ATP synthase complex and allowed retention of function (193, 194). Comparing the length of the wild type, shortened and lengthened peripheral stalks will give some insight on the overall possible conformation of the altered F 1 F 0 ATP synthase complex. The F 1 F 0 ATP synthase enzyme complex must, in some way, adapt to the shortened or lengthened b subunits. This can occur in at least one of two ways: i) distortion of F 1 to accommodate the change in length of the peripheral stalk or ii) distortion of the peripheral stalk itself. If the former was true, F 1 could become distorted by bending of the central stalk or compression of the 3 3 hexamer and the rigid stalk hypothesis would gain favor. Since F 1 F 0 ATP synthase function requires the ability of the central stalk to rotate freely while making specific interactions within the catalytic hexamer, this mechanism is highly unlikely. If the latter situation were true, distortion would be limited to the b dimer and the flexible stalk hypothesis would hold true. This could occur by straightening or increasing the naturally occurring

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192 20 bend in the shortened or lengthened peripheral stalk, respectively, or by stretching or compressing the secondary structure in the b dimer. The use of fluorescence resonance energy transfer (FRET) will provide evidence as to which of the hypotheses is true. The flexible stalk hypothesis suggests that the overall conformation of the F 1 F 0 ATP synthase complex is maintained regardless of the alterations in the length of the b subunit and the changes in length are accommodated by changes in the tether region. If the flexible stalk hypothesis holds true, two predictions can be made based on the relative measured length of the shortened, lengthened and wild type length b subunits: i) the altered length of the tether region will not effect on the measured distance between a fixed site in F 1 and a site in the dimerization domain of the b subunit and ii) the distance between a fixed site in F 1 and a site below the tether region alteration will vary by only a few due to the bending or straightening of the tether region. On the other hand, if the rigid stalk hypothesis holds true a change in distance from a fixed site in F 1 and a site below the alteration would result in an approximately 16 difference between the wild type length b and b 11 The length of the peripheral stalks with altered length b subunits will be measured by FRET. FRET requires the introduction of single donor and acceptor fluorophores into a purified F 1 F 0 ATP synthase complex. A common approach to performing FRET is to advantageously place a single cysteine at each end of the distance to be measured. Fluorescent maleimide derivatives are highly and specifically reactive with cysteine side chains. In order to perform FRET, single cysteines were strategically placed within the subunit of F 1 and b subunit of F 0 This work has been accomplished and is discussed in Chapter 4. These cysteines will be used to donorand acceptor-label F 1 and F 0 with

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193 Figure 6-1. Design of FRET experiments to measure the peripheral stalk. The fluorophores will be added to the thiol side chains of the cysteines generated in Chapter 4. The stars indicate the approximate positions of the d cys140 b cys84 and b cys43 The green shaded area of the b subunit represents the location of the insertions or deletions. maleimide reagents for the FRET studies (Figure 6-1). F 1 can readily be stripped from F 0 each cysteine labeled separately with a "donor" and "acceptor" fluorescent compound, and then stoichiometrically reconstituted under conditions of high ionic strength (203). With a single donor fluorophore positioned in the subunit and a single acceptor fluorescent compound located above or below the region of insertion or deletion in the tether domain of the b subunit, physical measurements can be calculated and compared (Figure 6-1).

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194 One major limitation plagued this line of research. F 1 F 0 ATP synthase is incorporated with a homodimer of b subunits, hence two b subunits, each with a reactive cysteine. Since FRET requires only one b subunit be labeled, a system necessarily had to be developed to allow purification of F 1 F 0 ATP synthase complexes with two genetically different b subunits: one cysteine-less and one with a single cysteine. The system utilizes epitope tags placed on the b subunits and was discussed in detail in Chapter 2. Future FRET studies will use this epitope-tagged scheme in order to express F 1 F 0 ATP synthase complexes with a b dimer containing only one cysteine. Alternative approach. Another approach for the investigation of a potentially flexible peripheral stalk may also be attempted in future experiments. An innovative set of experiments conducted by members of the Dunn laboratory made use of different sized fluorescent proteins, ranging from the 12 kDa cytochrome b 562 protein to the 30 kDa flavodoxin reductase protein with a 20 residue linker (100). The proteins were fused to the subunit of the central rotor stalk. If the fusion protein were small enough, one would expect the central stalk to rotate freely. If the fusion protein were too bulky, one could imagine that the protein would run into the peripheral stalk during rotational catalysis. As the fusion proteins became larger, an inhibition of activity resulted due to the bulky fluorescent protein running into the peripheral stalk during rotation. The growth curve of the bacteria was also monitored and revealed that the larger fusion proteins inhibited growth more than the smaller proteins. If this same technique were to be applied to F 1 F 0 ATP synthases incorporated with b 11 and compared to the activity of complexes incorporated with b +14 conclusions could be made concerning the flexibility of the peripheral stalk. For example, if the 20 bend in the b subunit were straightened in

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195 Figure 6-2. Model of rotation inhibition due to a fusion protein on the subunit. A natural 20 bend in the b subunit has been observed near the surface of the membrane (136). Shortening the b subunit by 11 amino acids (left panel) or lengthing by 14 amino acids(right panel) may be accompanied by the straightening or further bending, respectively, of this 20 bend. Genetic fusion proteins on the subunit have been described by Cipriano et al. (represented by the yellow oval) (100). The larger the fusion protein, the greater the inhibition of activity due to the inability of the central stalk to rotate during catalysis. We propose a model in which F 1 F 0 ATP synthase complexes incorporated with shorten or lengthened b subunits may accommodate smaller or larger fusion proteins, respectively. the shortened b subunit, inhibition of growth might become apparent with the smaller fusion proteins (Figure 6-2). Conversely, in complexes incorporated with the lengthened b subunit, the tether region may bend further, allowing the larger fusion proteins to pass between the central and peripheral stalks. Comparison of the growth properties of the two extreme tolerable lengths of the b subunit could lend support to the flexible stalk model. Other Implications How does the work presented in this dissertation affect the way in which the b subunit homologues of F 1 F 0 ATP synthase are currently viewed? This work suggests that the importance of the b subunit dimer may be reflected in the fact that higher organisms

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196 evolved to encode multiple b subunit equivalents. The b subunit is the least conserved subunit of F 1 F 0 ATP synthase. In E. coli, the peripheral stalk of the F 1 F 0 ATP synthase complex exists as a dimer of identical b subunits expressed from a single gene. However, the equivalent of the bacterial b subunit in photosynthetic bacteria and plants exists as two different subunits, referred to as b and b, and the mammal counterpart exists as at least four subunits, expressed from two and four separate genes, respectively (Figure 1-8). The second non-identical b-type subunit found in photosynthetic organisms has been assumed to appear in order to gain additional functions in connection with photophosphorylation, though there is no direct evidence of this proposal (295, 296). Here, I would like to propose a different view concerning the multiple b-type subunit genes found in higher organisms. Work described in this dissertation suggested that the bacterial enzyme does not require two identical b subunits to form the dimer. Two different length b subunits, with a size difference of at least14 amino acids, were capable of forming the b dimer of an intact F 1 F 0 ATP synthase complex. Interestingly, the b and b subunits of photosynthetic organisms are rarely of equal length, with size differences ranging from about 4 to 40 amino acids due to extensions at either the aminoor carboxyl-termini or sometimes due to gaps found within the center of a b subunit. Also, in work presented in this dissertation, a defective mutation in one region of the b subunit has been shown to be overcome by dimer formation with a second b subunit that contained defective mutation in a different region but had a wild-type sequence in the region of the former defective b subunit. This observation has implications concerning the two individual b subunits in which each b subunit makes individual contacts within the enzyme complex. I believe the presence of the two b subunits are required for the

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197 structural stability of the peripheral stalk. However, I would like to take this observation a step further and ask the following question: Why do bacteria encode only one b subunit gene, whereas higher organisms encode multiple b-type subunit genes, if single amino acid mutations are capable of destroying the enzyme function? One could speculate that bacteria do not require a functional F 1 F 0 ATP synthase in order to survive. Though growth would be slow, bacteria can persist via glycolysis until environmental pressures either cause death, a reversion of the uncF(b) back to a wild-type gene, or a second site suppressor mutation. Higher organisms, on the other hand, do not have the luxury of solely surviving from glycolysis due to their higher energy needs. Selective pressures may have caused higher organisms to evolve to encode two different b subunit genes, where important amino acids may occur in only the b subunit or the b subunit. This is reflected in my finding that the b or the b subunits in plants have only about 15-20% identity with the E. coli b subunit, but the amino acids conserved in the chloroplast b and b subunits generally occur at different residues, suggesting that the b dimer, as a whole, actually contains a larger number of conserved residues that are involved in making the functional contacts (Figure 6-3A, C). In fact, when the sequences of the b and b sequences were manually merged together, such that identical or similar amino acids replaced the nonconserved residues, the percent identity and similarity of the fictional merged b subunit nearly doubled (Figure 6-3B, C). It appears the actual number of conserved residues is actually higher than believed to be, the difference being, the conserved residue need only to be in one or the other b subunit. This proposal may account for why the b-type subunits are the least conserved of all the F 1 F 0 ATP synthase. If only one of the b-type subunits is required to make the appropriate contacts, mutations

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198 occurring in the other b-type subunits of the organism would not have a dramatic affect on the enzyme function.

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199

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200 Figure 6-3. Sequence alignments of subunits b and b from various species with the b subunit of E. coli. The E. coli sequence is entirely in bold script. The blue star indicates residues that are similar across all of the species. The red arrow indicates residues that are identical across all of the species. The hyphen indicates gaps. Sequences were randomly chosen from the National Center for Biotechnology Information (NCBI) database. Residues indicated in red bold script in the b and b subunits of the various species are identical to the corresponding residue in the E. coli b subunit. Residues indicated in blue are similar to the E. coli b subunit. In parenthesis are the percent identity followed by the percent similarity. A) Sequence alignment of subunits b and b of other species with the b subunit of E. coli. B) The b and b subunits of each species were merged together so that the identical or most similar amino acid (relative to E. coli) replaced nonconserved amino acids. C) Graphical representation of sequence alignments.

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201

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APPENDIX A MUTAGENIC OLIGONULCEOTIDES Many of the mutations created during the course of this dissertation were via site-directed mutagenesis, performed either by means of a Stratagene Quikchange XL kit or by ligation-mediated mutagenesis. For the Quikchange XL procedure, oligonucleotides containing the desired mutation(s) were designed to anneal to the same sequence on opposite strands of the plasmid (sense and antisense primers). When possible, a silent mutation was encoded to add or delete a restriction endonuclease recognition sequence to allow for easy screening of the mutation. Primers were optimally designed by ensuring the mutation was in the middle of the sequence, a cytosine (C) or guanine (G) flanked both ends of the sequence, and the melting temperature (T m ) was greater than or equal to 78 C. The T m was calculated as T m =81.5+0.41(%GC)-675/N-%mismatch, where N was the primer length (bases). When introducing insertions or deletions, "%mismatch" was dropped from the formula. For a detailed description of the reaction, see Chatper 2 Recombinant DNA Techniques under Materials and Methods. The resulting plasmids that carried the desired mutation were then transformed into competent DH5 cells, purchased from Life Technologies, and grown on LBG plates supplemented with the appropriate antibiotic. Plasmids carrying the desired mutation(s) were screened for by restriction endonuclease analysis and then the nucleotide sequences were directly determined by automated sequencing in the core facility of the University of Florida Interdisciplinary Center for Biotechnology Research (ICBR). A description of the oligonucleotides follows. All oligonucleotides are listed in the 5 to 3 direction. 202

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203 Table A-1. Oligonucleotide sequences. Primer Sequence TB1 5 ctaaatagaagcatgctgctgtgcaccaccaccaccaccacaatcttaacgcaacaatcctcggc 3 TB2 5 gccgaggattgttgcgttaagattgtggtggtggtggtggtgcacagcagcatgcttctatttag 3 TB3 5 ctaaatagaggcattgtgctatgtacccatatgacgtgccggactacgcgaatcttaacgcaacaatcctcggc TB4 5 gccgaggattgttgcgttaagattcgcgtagtccggcacgtcatatgggtacatagcacaatgcctctatttag TB5 5 ctgtaaggagggaggggcatgcgtctgaattttattacg 3 TB6 5 cgtaataaaattcagacgcatgcccctccctccttacag 3 TB7 5 ggagggagggggaggctgctgtgc 3 TB8 5 gcacagcagcctccccctccctcc 3 TB10 5 cgtctgtcacgcaaagttaagctgaattcgaaaattgataagtctgtaatggcagg 3 TB11 5 cctgccattacggacttatcaattttcgaattcagcttaactttgcgtgacagacg 3 TB12 5 ctgttcgttctgttctccatgaagtacgtttggccgcc 3 TB13 5 ggcggccaaacgtacttcatggagaacagaacgaacag 3 TB14 5 gaaattgctgactgccttgcttccgcagaacgagcacataag 3 TB15 5 ccttatgggctcgttctgcggaagcaaggcagtcagcaatttc 3 TB16 5 gcgaacaaacgccgctgccagattctcgacgaagc 3 TB17 5 gcttcgtcgagaatctggcagcggcgtttgttcg 3 TB18 5 gccgagtcgtttatcgcagtttctggtgagcaactggac 3 TB19 5 ccagttgctcaccagaaaccgcgataaacgactc 3 TB20 5 ctgtacgtgatgttcgctgtcgc 3 TB21 5 gctcgcccctacgccaaagcagc 3 TB23 5 ggcatgaaagttaagtctactggccggatcctggaag 3 TB24 5 cttccaggatccggccagtagacttaactttcatgcc 3 TB25 5 ctgctgcgatggaaaaacgtctgtcac 3 TB27 5 gggagggggaggcagatatgcaccaccacc 3 TB28 5 ggtggtggtgcatatctgcctccccctccc 3 TB29 5 gtcctgccagcgttctacactttggtg 3 TB30 5 gaggcattgtgctgtgtggccgccattaatggc 3 TB31 5 gccattaatggcggccacacagcacaatgcctc 3 TB32 5 gaggcattgtgctgtgcaccaccaccaccaccaccaccaccaccactggccgccattaatggc 3 TB33 5 gccattaatggcggccagtggtggtggtggtggtggtggtggtggtgcacagcacaatgcctc 3 TB36 5 ggagggaggggctgatgcaccaccac 3 TB37 5 gtggtgcatcagcccctccctcc 3 TB38 5 cgtggataaacttgtcgctgagctcggtaaaccgatcccgaacccg ctgctgggtctggactctacctaaggagggaggggctgatgtct 3 TB39 5catcagcccctccctccttaggtagagtccagacccagcagcgg gttcgggatcggtttaccgagctcagcgacaagtttatccacg 3 TB405 catcgtggataagctttaaggagggaggggctg 3 TB415 cagcccctccctccttaaagcttatccacgatg 3 TB40+ 5 gataaacttgtcgctgaactggtcgctgagctctaaggagggaggg 3 TB41+ 5 ccctccctccttagagctcagcgaccagttcagcgacaagtttatc 3

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204 Table A-2. Oligonucleotide description. Primer Conc. 1 Sense 2 Description T m 3 TB1 0.892 S TB2 0.606 A addition of histidine-epitope tag (HHHHHH) to the N-terminus of the b subunit; +SphI 85 TB3 0.329 S TB4 0.414 A addition of HA-epitope tag (YPYDVPDYA) to the N-terminus of the b subunit; +NdeI 85 TB5 2.846 S TB6 2.236 A addition of SphI site 3 of the b subunit for cloning purposes; + SphI 78 TB7 1.951 S TB8 1.593 A mutation of SphI site 3 of the b subunit and addition of a favorable ATG prior to next gene; -SphI 78 TB10 0.308 S TB11 0.650 A cys140ser mutation; +EcoRI; -ClaI 80 TB12 0.977 S TB13 1.538 A b cys20ser mutation; -SnaBI 79 TB14 1.151 S TB15 1.446 A b gly43cys mutation; -Bsp1285I 77 TB16 1.100 S TB17 1.290 A b ser84cys mutation; -XbaI 75 TB18 1.366 S TB19 1.242 A cys64ser mutation 82 TB20 2.684 S sequences b in the sense (not a good one) 75 TB21 2.422 S sequences in the sense direction 81 TB23 1.380 S TB24 1.640 A cys90ser mutation; +BamHI 77 TB25 5.016 S sequences in the sense direction 78 TB27 0.886 S TB28 1.235 A mutations to create Shine Delgarno sequence upstream of the 2 nd b gene in mix plasmids 76 TB29 5.279 A sequences b in the inverse complement direction 79 TB30 4.334 S TB31 4.608 A deletion of residues 1-24 of b subunit to create b sol 85 TB32 8.301 S TB33 7.618 A addition of histidine-epitope tag (HHHHHHHHHH) to the N-terminus of the b sol subunit 78 TB36 0.439 S TB37 0.477 A mutations to create a better Shine Delgarno sequence upstream of the 2 nd b gene in mix plasmids 78 TB38 12.33 S TB39 9.310 A addition of V5-epitope tag (GKPIPNPLLGLDST) to the C-terminus of the b subunit; +SacI 79 TB404.522 S TB414.891 A deletion of last 4 amino acids at the C-terminus of the b subunit; +HindIII 76 TB40+ 1.109 S TB41+ 0.983 A duplication of last 4 amino acids at the C-terminus of the b subunit; +SacI 78 1 Concentration of stock oligonucleotide in g/l, stored in the -20C. 2 Direction of the primer: S, sense; A, antisense. 3 Melting temperature in C.

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APPENDIX B DEVELOPING PROTOCOL FOR PURIFYING F 1 F 0 ATP SYNTHASE Purification of Enzyme Complexes Incorporated with b Subunit Heterodimers The purification procedure described below has been attempted one time; therefore a detailed account of the current protocol is given here. Once the particulars of the procedure are optimized, the same protocol will be applied to all the F 1 F 0 ATP synthases incorporated with different combinations of heterodimeric mutant b subunits. For the sake of simplicity, the different b subunits are simply referred to as b V5 and b his Culture Inverted membrane vesicles from KM2(b) strains expressing the desired b subunits will be prepared essentially as described previously and optimized for large-scale membrane preparations (199, 297). One of the advantages of studying a bacterial enzyme is the practically unlimited amount of bacteria that may be grown and harvested. The following can be scaled up or down to meet the experimental needs. The bacteria will grown by inoculating 20 mL starter culture, grown overnight, into a 2 L Erlenmeyer flask holding 1 L LBG supplemented with both ampicillin (Ap) (100 g/mL) and chloramphenicol (Cm) (25 g/mL). Similarly, 1 mL of the starter culture will be inoculated into a nephalo flask containing 50 mL LBG (Ap and/or Cm) to monitor growth. For the purposes of the large-scale purification, a total of 10 L LBG-Ap-Cm, contained in 10 flasks, will be grown. The bacteria will be grown at 37 C in a New Brunswick Scientific incubator shaker (220 rpm) and the turbidity monitered using a Klett-Summerson photoelectric colorimeter. IPTG (40 M) will be added when the 206

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207 turbidity reaches 75 Klett units and the cells will subsequently be collected when the turbidity reaches 150 Klett units. The bacteria will be harvested by centrifugation for 10 minutes at 8,000Xg in a Sorvall GSA rotor. The pellets will be rinsed once with TM buffer (50 mM tris-HCl, 10 mM MgSO 4 pH 7.5), weighed, and then stored at C until the following day. Typically, a 10 L culture will yielded approximately 25 g bacteria. Disruption of Bacteria The 25 g frozen bacteria will be allowed to thaw at room temperature and resuspended in a final volume of 200 mL TM buffer. Resuspension requires rigorous vortexing, until the bacterial clumps dislodge from the walls of the bottles, and then continuously pipetted with a 25 mL pipette until no cell clumps are apparent. It is important to sufficiently break all clumps of bacteria in order to efficiently break open the cells. The bacteria will be treated with lysozyme in order to disrupt the bacterial cell wall. Prior to lysozyme treatment, the cells will be prepared by dissolving 0.91 g ethylenediamine tetra acetic acid (EDTA) in the suspension. The 200 mL bacterial suspension should be stirred with a magnetic stir bar at room temperature throughout the lysozyme treatment. 200 mL of 200 mM Tris-HCl, pH 7.5 and 1 M sucrose will be added and stirred for 3 minutes. Lysozyme (0.6 mg per g cells) will be added and stirred for 5 minutes. The bacterial cell walls may then be broken by osmotically shocking with 400 mL ddH 2 O. The suspension will be stirred for an additional 5-10 minutes. Hereafter, all samples, reagents and equipment should be kept at 4 C unless otherwise specified. The bacteria will be harvested by centrifugation and the viscous supernatant may be discarded. The pellet will be resuspended in a final volume of 125 mL TM buffer by pipetting. Notably, in a trial attempt at purification, the bacterial pellet was extremely

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208 Figure B-1. Diagram of purification procedures for homogeneous heterodimeric b V5 /b his F 1 F 0 ATP synthase complexes. The b subunit dimers represents intact F 1 F 0 ATP synthase. Basically, membrane preparations will be solubilized in 0.2% tegamineoxide WS-35. The solubilized proteins will then be Ni-resin purified in order to remove the homodimeric b V5 /b V5 F 1 F 0 complexes. This will be followed by a V5 immunoprecipitation step to remove the homodimeric b his /b his F 1 F 0 complexes, leaving only the heterodimeric b V5 /b his complexes in solution. A battery of tests will then be run in order to determine the success of the purification procedures and then ultimately activity assays will be performed.

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209 viscous and resuspension required about 30 minutes. DnaseI (10 mg/mL) will be added to help decrease the viscosity during resuspension. The bacteria will be broken by a single pass through the French Pressure Cell at 20,000 psi. Cellular debris and unbroken cells will be removed by centrifuging twice at 10,000Xg for 10 minutes each. Membranes will then be collected by ultracentrifugation at 150,000Xg in a Beckman 70.1 Ti rotor for 1.5 hours. The membranes pellets will be rinsed once with Tm buffer and then resuspended in a final volume of 30 mL Tm buffer using a 30 mL Wheaton tissue grinder. Total membrane protein concentration will be determined by the bicinchoninic acid (BCA) assay (Markwell et al., 1978). Ni-Resin Purification The crude membrane preparation described above will contain all membrane proteins, including homodimeric b V5 /b V5 F 1 F 0 complexes, homodimeric b his /b his F 1 F 0 complexes and heterodimeric b V5 /b his F 1 F 0 complexes (Figure B-1). The membrane preparation will be solubilized in 0.2% tegamineoxide WS-35 and then the homodimeric b his /b his F 1 F 0 complexes and heterodimeric b V5 /b his F 1 F 0 complexes will be isolated using the Ni-CAM resin purification procedure (see Chapter 2 Materials and Methods). The homodimeric b V5 /b V5 F 1 F 0 complexes will be eliminated at this step due to the lack of a histidine epitope tag. Preliminary experiments showed that Ni-CAM resin overloaded with membrane protein only produced the homodimeric b his /b his F 1 F 0 complexes and the heterodimeric complexes were washed away. This can be explained by the fact that the heterodimeric b V5 /b his complexes only contain one histidine epitope tag where the homodimeric b his /b his enzyme complexes are incorporated with two histidine epitope tags. Therefore, an optimization of the ratio of membrane protein to Ni-CAM resin will be

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210 necessary. The optimized ratio will lie somewhere around 1.0 mg membrane protein to 0.2 mL packed Ni-CAM resin. V5-Epitope Iimmunoprecipitation The mixture of the homodimeric b his /b his F 1 F 0 ATP synthase complexes and the heterodimeric b V5 /b his F 1 F 0 complexes will then be separated by a second epitope-mediated purification (Figure B-1). Immunoprecipitation of the heterodimeric b V5 /b his F 1 F 0 will be accomplished by immobilizing mouse monoclonal anti-V5 antibody (purchased from Invitrogen) on a resin using the Seize Primary Immunoprecipitation Kit (Purchased from Pierce Biotechnology). This kit has been successfully used several times by Dr. Michelle Gumz in our laboratory and consistently gave high yields of purified proteins. Detection of Purified Enzyme Complexes Once the purified heterodimeric bV5/bhis F1F0 ATP synthase complexes are eluted from the V5-affinity resin, the eluant will be tested for the presence of intact and pure enzyme complexes (Figure B-1). The product will be ran on a 10-18% gradient SDS PAGE gel and then silver stained in order to examine the subunit content of the purified product. All eight subunits should be distinguishable in the stained gel, indicating the presence of intact F1F0 ATP synthase complexes. The homogeneity of the purified product will then be examined by immunoblot analysis. The bV5 and bhis were separable on a 15 cm 15% SDS PAGE gel when ran at 100 mamp, with current held constant, until the dye reached the stacker and then at 24 mamp for 12 hours (see Figure 2-13 for a representative figure). It was necessary to double the amperage when two gels were running at the same time. Upon transfer to a nitrocellulose membrane, immunoblot analysis using anti-b antibodies will be performed in order to determine if the

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211 heterodimeric bV5/bhis F1F0 has been purified to homogeneity. Densitometry will be used to determine the relative amounts of the bV5 and bhis subunits. If the product is indeed pure, the bV5 and bhis subunits should be present in a 1:1 ratio. Assays of F1F0 ATP Synthase Activity Once the purified heterodimeric bV5/bhis F1F0 ATP synthase complexes are obtained, the battery of traditional activity assays will be performed. ATP hydrolysis assays will be performed by the standard procedure described in Chapter 2 (Materials and Methods) in both the presence and absence of DCCD. ATP hydrolysis activity will be used both as an assay of yield of intact complexes and of functional coupling. DCCD is a c subunit specific inhibitor of F1F0 ATP hydrolysis; therefore, DCCD sensitivity can be used as an indication of functional coupling. ATP-driven proton pumping assays will require reconstitution of the solubilized F1F0 ATP synthase into liposomes. This will be accomplished by a protocol described by Aggeler et al. (60) and modified by Dr. James Garder for use in our laboratory (167, 297). ATP-driven proton pumping will then be assayed by the standard method routinely used in the laboratory (Chapter 2, Materials and Methods).

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LIST OF REFERENCES 1. Mitchell P, Keilin's respiratory chain concept and its chemiosmotic consequences. Science 1979, 206(4423):1148-1159. 2. Mitchell P, Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biol Rev Camb Philos Soc 1966, 41(3):445-502. 3. Senior AE, Nadanaciva S, Weber J, The molecular mechanism of ATP synthesis by F1F0-ATP synthase. Biochim Biophys Acta 2002, 1553(3):188-211. 4. Senior AE, Weber J, Happy motoring with ATP synthase. Nat Struct Mol Biol 2004, 11(2):110-112. 5. Nath S, The molecular mechanism of ATP synthesis by F1F0-ATP synthase: a scrutiny of the major possibilities. Adv Biochem Eng Biotechnol 2002, 74:65-98. 6. Fillingame RH, Divall S, Proton ATPases in bacteria: comparison to Escherichia coli F1F0 as the prototype. Novartis Found Symp 1999, 221:218-229; discussion 229-234. 7. Pedersen PL, Ko YH, Hong S, ATP synthases in the year 2000: evolving views about the structures of these remarkable enzyme complexes. J Bioenerg Biomembr 2000, 32(4):325-332. 8. Belogrudov GI, Tomich JM, Hatefi Y, Membrane topography and near-neighbor relationships of the mitochondrial ATP synthase subunits e, f, and g. J Biol Chem 1996, 271(34):20340-20345. 9. Devenish RJ, Prescott M, Roucou X, Nagley P, Insights into ATP synthase assembly and function through the molecular genetic manipulation of subunits of the yeast mitochondrial enzyme complex. Biochim Biophys Acta 2000, 1458(2-3):428-442. 10. Velours J, Paumard P, Soubannier V, Spannagel C, Vaillier J, Arselin G, Graves PV, Organisation of the yeast ATP synthase F(0):a study based on cysteine mutants, thiol modification and cross-linking reagents. Biochim Biophys Acta 2000, 1458(2-3):443-456. 11. Wilkens S, F1F0-ATP synthase-stalking mind and imagination. J Bioenerg Biomembr 2000, 32(4):333-339. 212

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213 244(2):279-282. 12. Dunn SD, McLachlin DT, Revington M, The second stalk of Escherichia coli ATP synthase. Biochim Biophys Acta 2000, 1458(2-3):356-363. 13. Dunn SD, Revington M, Cipriano DJ, Shilton BH, The b subunit of Escherichia coli ATP synthase. J Bioenerg Biomembr 2000, 32(4):347-355. 14. Fillingame RH, Jiang W, Dmitriev OY, Jones PC, Structural interpretations of F(0) rotary function in the Escherichia coli F(1)F(0) ATP synthase. Biochim Biophys Acta 2000, 1458(2-3):387-403. 15. Noji H, Yasuda R, Yoshida M, Kinosita K, Jr., Direct observation of the rotation of F1-ATPase [see comments]. Nature 1997, 386(6622):299-302. 16. Noji H, Yoshida M, The rotary machine in the cell, ATP synthase. J Biol Chem 2001, 276(3):1665-1668. 17. Fernandez-Moran H, Cell-membrane ultrastructure. Low-temperature electron microsopy and x-ray diffraction studies of lipoprotein components in lamellar systems. Circulation 1962, 26:1039-1065. 18. Kagawa Y, Reconstitution of oxidative phosphorylation. Biochim Biophys Acta 1972, 265(3):297-338. 19. Catterall WA, Coty WA, Pedersen PL, Adenosine triphosphatase from rat liver mitochondria. 3. Subunit composition. J Biol Chem 1973, 248(21):7427-7431. 20. Abrahams JP, Leslie AG, Lutter R, Walker JE, Structure at 2.8 A resolution of F1-ATPase from bovine heart mitochondria [see comments]. Nature 1994, 370(6491):621-628. 21. Bianchet MA, Hullihen J, Pedersen PL, Amzel LM, The 2.8-A structure of rat liver F1-ATPase: configuration of a critical intermediate in ATP synthesis/hydrolysis. Proc Natl Acad Sci U S A 1998, 95(19):11065-11070. 22. Soper JW, Pedersen PL, Isolation of an oligomycin-sensitive ATPase complex from rat liver mitochondria. Methods Enzymol 1979, 55:328-333. 23. Boekema EJ, Berden JA, van Heel MG, Structure of mitochondrial F1-ATPase studied by electron microscopy and image processing. Biochim Biophys Acta 1986, 851(3):353-360. 24. Gogol EP, Lucken U, Capaldi RA, The stalk connecting the F1 and F0 domains of ATP synthase visualized by electron microscopy of unstained specimens. FEBS Lett 1987, 219(2):274-278. 25. Tsuprun VL, Orlova EV, Mesyanzhinova IV, Structure of the ATP-synthase studied by electron microscopy and image processing. FEBS Lett 1989,

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214 active and inactive forms of protooncogenic ras proteins. 1990, 247(4945):939-945. 26. Bottcher B, Schwarz L, Graber P, Direct indication for the existence of a double stalk in CF0F1. J Mol Biol 1998, 281(5):757-762. 27. Wilkens S, Capaldi RA, Electron microscopic evidence of two stalks linking the F1 and F0 parts of the Escherichia coli ATP synthase. Biochim Biophys Acta 1998, 1365(1-2):93-97. 28. Wilkens S, Capaldi RA, Solution structure of the epsilon subunit of the F1-ATPase from Escherichia coli and interactions of this subunit with beta subunits in the complex. J Biol Chem 1998, 273(41):26645-26651. 29. Karrasch S, Walker JE, Novel features in the structure of bovine ATP synthase. J Mol Biol 1999, 290(2):379-384. 30. Futai M, Noumi T, Maeda M, ATP synthase (H+-ATPase): results by combined biochemical and molecular biological approaches. Annu Rev Biochem 1989, 58:111-136. 31. Futai M, Noumi T, Maeda M, Molecular genetics of F1-ATPase from Escherichia coli. J Bioenerg Biomembr 1988, 20(1):41-58. 32. Weber J, Senior AE, Catalytic mechanism of F1-ATPase. Biochim Biophys Acta 1997, 1319(1):19-58. 33. Senior AE, Catalytic sites of Escherichia coli F1-ATPase. J Bioenerg Biomembr 1992, 24(5):479-484. 34. Sato K, Mori H, Yoshida M, Mizushima S, Characterization of a potential catalytic residue, Asp-133, in the high affinity ATP-binding site of Escherichia coli SecA, translocation ATPase. J Biol Chem 1996, 271(29):17439-17444. 35. Weidenhaupt M, Bruckert F, Satre M, Identification of the Dictyostelium discoideum homolog of the N-ethylmaleimide-sensitive fusion protein. Gene 1998, 207(1):53-60. 36. Yu D, Weller SK, Herpes simplex virus type 1 cleavage and packaging proteins UL15 and UL28 are associated with B but not C capsids during packaging. J Virol 1998, 72(9):7428-7439. 37. Ovchinnikov Yu A, Luneva NM, Arystarkhova EA, Gevondyan NM, Arzamazova NM, Kozhich AT, Nesmeyanov VA, Modyanov NN, Topology of Na+,K+-ATPase. Identification of the extraand intracellular hydrophilic loops of the catalytic subunit by specific antibodies. FEBS Lett 1988, 227(2):230-234. 38. Milburn MV, Tong L, deVos AM, Brunger A, Yamaizumi Z, Nishimura S, Kim SH, Molecular switch for signal transduction: structural differences between Science

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215 nucleotide-free alpha 3 beta 3 subcomplex of F1-ATPase from the thermophilic Bacillus PS3 is a symmetric trimer. Structure 1997, 5(6):825-836. 39. Shen H, Yao BY, Mueller DM, Primary structural constraints of P-loop of mitochondrial F1-ATPase from yeast. J Biol Chem 1994, 269(13):9424-9428. 40. Story RM, Weber IT, Steitz TA, The structure of the E. coli recA protein monomer and polymer. Nature 1992, 355(6358):318-325. 41. Kjeldgaard M, Nissen P, Thirup S, Nyborg J, The crystal structure of elongation factor EF-Tu from Thermus aquaticus in the GTP conformation. Structure 1993, 1(1):35-50. 42. Noel JP, Hamm HE, Sigler PB, The 2.2 A crystal structure of transducin-alpha complexed with GTP gamma S. Nature 1993, 366(6456):654-663. 43. Aggeler R, Zhang YZ, Capaldi RA, Labeling of the ATP synthase of Escherichia coli from the head-group region of the lipid bilayer. Biochemistry 1987, 26(22):7107-7113. 44. Abrahams JP, Buchanan SK, Van Raaij MJ, Fearnley IM, Leslie AG, Walker JE, The structure of bovine F1-ATPase complexed with the peptide antibiotic efrapeptin. Proc Natl Acad Sci U S A 1996, 93(18):9420-9424. 45. van Raaij MJ, Abrahams JP, Leslie AG, Walker JE, The structure of bovine F1-ATPase complexed with the antibiotic inhibitor aurovertin B. Proc Natl Acad Sci U S A 1996, 93(14):6913-6917. 46. Orriss GL, Runswick MJ, Collinson IR, Miroux B, Fearnley IM, Skehel JM, Walker JE, The deltaand epsilon-subunits of bovine F1-ATPase interact to form a heterodimeric subcomplex. Biochem J 1996, 314 ( Pt 2):695-700. 47. Braig K, Menz RI, Montgomery MG, Leslie AG, Walker JE, Structure of bovine mitochondrial F(1)-ATPase inhibited by Mg(2+) ADP and aluminium fluoride. Structure Fold Des 2000, 8(6):567-573. 48. Gibbons C, Montgomery MG, Leslie AG, Walker JE, The structure of the central stalk in bovine F(1)-ATPase at 2.4 A resolution. Nat Struct Biol 2000, 7(11):1055-1061. 49. Lobau S, Weber J, Senior AE, Nucleotide occupancy of F1-ATPase catalytic sites under crystallization conditions. FEBS Lett 1997, 404(1):15-18. 50. Menz RI, Walker JE, Leslie AG, Structure of bovine mitochondrial F(1)-ATPase with nucleotide bound to all three catalytic sites: implications for the mechanism of rotary catalysis. Cell 2001, 106(3):331-341. 51. Shirakihara Y, Leslie AG, Abrahams JP, Walker JE, Ueda T, Sekimoto Y, Kambara M, Saika K, Kagawa Y, Yoshida M, The crystal structure of the

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216 during catalysis by F1-ATPase. 1995, 92(24):10964-10968. 52. Hausrath AC, Gruber G, Matthews BW, Capaldi RA, Structural features of the gamma subunit of the Escherichia coli F(1) ATPase revealed by a 4.4-A resolution map obtained by x-ray crystallography. Proc Natl Acad Sci U S A 1999, 96(24):13697-13702. 53. Senior AE, ATP synthesis by oxidative phosphorylation. Physiol Rev 1988, 68(1):177-231. 54. Weber J, Senior AE, ATP synthase: what we know about ATP hydrolysis and what we do not know about ATP synthesis. Biochim Biophys Acta 2000, 1458(2-3):300-309. 55. Issartel JP, Dupuis A, Garin J, Lunardi J, Michel L, Vignais PV, The ATP synthase (F0-F1) complex in oxidative phosphorylation. Experientia 1992, 48(4):351-362. 56. Perlin DS, Latchney LR, Wise JG, Senior AE, Specificity of the proton adenosinetriphosphatase of Escherichia coli for adenine, guanine, and inosine nucleotides in catalysis and binding. Biochemistry 1984, 23(21):4998-5003. 57. Nakamoto RK, Ketchum CJ, al-Shawi MK, Rotational coupling in the F0F1 ATP synthase. Annu Rev Biophys Biomol Struct 1999, 28:205-234. 58. Stock D, Leslie AG, Walker JE, Molecular architecture of the rotary motor in ATP synthase [see comments]. Science 1999, 286(5445):1700-1705. 59. Capaldi RA, Schulenberg B, Murray J, Aggeler R, Cross-linking and electron microscopy studies of the structure and functioning of the Escherichia coli ATP synthase. J Exp Biol 2000, 203 Pt 1:29-33. 60. Aggeler R, Haughton MA, Capaldi RA, Disulfide bond formation between the COOH-terminal domain of the beta subunits and the gamma and epsilon subunits of the Escherichia coli F1ATPase. Structural implications and functional consequences. J Biol Chem 1995, 270(16):9185-9191. 61. Aggeler R, Capaldi RA, Nucleotide-dependent movement of the epsilon subunit between alpha and beta subunits in the Escherichia coli F1F0-type ATPase. J Biol Chem 1996, 271(23):13888-13891. 62. Sabbert D, Engelbrecht S, Junge W, Intersubunit rotation in active F-ATPase. Nature 1996, 381(6583):623-625. 63. Sabbert D, Engelbrecht S, Junge W, Functional and idling rotatory motion within F1-ATPase. Proc Natl Acad Sci U S A 1997, 94(9):4401-4405. 64. Duncan TM, Bulygin VV, Zhou Y, Hutcheon ML, Cross RL, Rotation of subunits Escherichia coliProc Natl Acad Sci U S A

PAGE 238

217 65. Kato-Yamada Y, Noji H, Yasuda R, Kinosita K, Jr., Yoshida M, Direct observation of the rotation of epsilon subunit in F1-ATPase. J Biol Chem 1998, 273(31):19375-19377. 66. Sambongi Y, Iko Y, Tanabe M, Omote H, Iwamoto-Kihara A, Ueda I, Yanagida T, Wada Y, Futai M, Mechanical rotation of the c subunit oligomer in ATP synthase (F0F1): direct observation [see comments]. Science 1999, 286(5445):1722-1724. 67. Panke O, Gumbiowski K, Junge W, Engelbrecht S, F-ATPase: specific observation of the rotating c subunit oligomer of EF(o)EF(1). FEBS Lett 2000, 472(1):34-38. 68. Itoh H, Takahashi A, Adachi K, Noji H, Yasuda R, Yoshida M, Kinosita K, Mechanically driven ATP synthesis by F1-ATPase. Nature 2004, 427(6973):465-468. 69. Wilkens S, Dahlquist FW, McIntosh LP, Donaldson LW, Capaldi RA, Structural features of the epsilon subunit of the Escherichia coli ATP synthase determined by NMR spectroscopy [see comments]. Nat Struct Biol 1995, 2(11):961-967. 70. Uhlin U, Cox GB, Guss JM, Crystal structure of the epsilon subunit of the proton-translocating ATP synthase from Escherichia coli. Structure 1997, 5(9):1219-1230. 71. Weiss MA, McCarty RE, Cross-linking within a subunit of coupling factor 1 increases the proton permeability of spinach chloroplast thylakoids. J Biol Chem 1977, 252(22):8007-8012. 72. Moroney JV, McCarty RE, Reversible uncoupling of photophosphorylation by a new bifunctional maleimide. J Biol Chem 1979, 254(18):8951-8955. 73. Moroney JV, Andreo CS, Vallejos RH, McCarty RE, Uncoupling and energy transfer inhibition of photophosphorylation by sulfhydryl reagents. J Biol Chem 1980, 255(14):6670-6674. 74. Aggeler R, Chicas-Cruz K, Cai SX, Keana JF, Capaldi RA, Introduction of reactive cysteine residues in the epsilon subunit of Escherichia coli F1 ATPase, modification of these sites with tetrafluorophenyl azide-maleimides, and examination of changes in the binding of the epsilon subunit when different nucleotides are in catalytic sites. Biochemistry 1992, 31(11):2956-2961. 75. Aggeler R, Capaldi RA, Cross-linking of the gamma subunit of the Escherichia coli ATPase (ECF1) via cysteines introduced by site-directed mutagenesis. J Biol Chem 1992, 267(30):21355-21359. 76. Aggeler R, Cai SX, Keana JF, Koike T, Capaldi RA, The gamma subunit of the Escherichia coli F1-ATPase can be cross-linked near the glycine-rich loop region

PAGE 239

218 Biochemistry 1980, 19(3):526-531. of a beta subunit when ADP + Mg2+ occupies catalytic sites but not when ATP + Mg2+ is bound. J Biol Chem 1993, 268(28):20831-20837. 77. Aggeler R, Capaldi RA, ATP hydrolysis-linked structural changes in the N-terminal part of the gamma subunit of Escherichia coli F1-ATPase examined by cross-linking studies. J Biol Chem 1993, 268(20):14576-14578. 78. Moroney JV, McCarty RE, Light-dependent cleavage of the gamma subunit of coupling factor 1 by trypsin causes activation of Mg2+-ATPase activity and uncoupling of photophosphorylation in spinach chloroplasts. J Biol Chem 1982, 257(10):5915-5920. 79. Turina P, Capaldi RA, ATP hydrolysis-driven structural changes in the gamma-subunit of Escherichia coli ATPase monitored by fluorescence from probes bound at introduced cysteine residues. J Biol Chem 1994, 269(18):13465-13471. 80. Gogol EP, Johnston E, Aggeler R, Capaldi RA, Ligand-dependent structural variations in Escherichia coli F1 ATPase revealed by cryoelectron microscopy. Proc Natl Acad Sci U S A 1990, 87(24):9585-9589. 81. Bullough DA, Ceccarelli EA, Roise D, Allison WS, Inhibition of the bovine-heart mitochondrial F1-ATPase by cationic dyes and amphipathic peptides. Biochim Biophys Acta 1989, 975(3):377-383. 82. Zhuo S, Paik SR, Register JA, Allison WS, Photoinactivation of the bovine heart mitochondrial F1-ATPase by [14C]dequalinium cross-links phenylalanine-403 or phenylalanine-406 of an alpha subunit to a site or sites contained within residues 440-459 of a beta subunit. Biochemistry 1993, 32(9):2219-2227. 83. Jackson PJ, Harris DA, Sites of protein-protein interaction on the mitochondrial F1-ATPase inhibitor protein. Biochem J 1986, 235(2):577-583. 84. Greene MD, Frasch WD, Interactions among gamma R268, gamma Q269, and the beta subunit catch loop of Escherichia coli F1-ATPase are important for catalytic activity. J Biol Chem 2003, 278(51):51594-51598. 85. Hausrath AC, Capaldi RA, Matthews BW, The conformation of the epsilonand gamma-subunits within the Escherichia coli F(1) ATPase. J Biol Chem 2001, 276(50):47227-47232. 86. Sternweis PC, The epsilon subunit of Escherichia coli coupling factor 1 is required for its binding to the cytoplasmic membrane. J Biol Chem 1978, 253(9):3123-3128. 87. Sternweis PC, Smith JB, Characterization of the inhibitory (epsilon) subunit of the proton-translocating adenosine triphosphatase from Escherichia coli.

PAGE 240

219 269(14):10221-10224. 88. Mendel-Hartvig J, Capaldi RA, Catalytic site nucleotide and inorganic phosphate dependence of the conformation of the epsilon subunit in Escherichia coli adenosinetriphosphatase. Biochemistry 1991, 30(5):1278-1284. 89. Mendel-Hartvig J, Capaldi RA, Nucleotide-dependent and dicyclohexylcarbodiimide-sensitive conformational changes in the epsilon subunit of Escherichia coli ATP synthase. Biochemistry 1991, 30(45):10987-10991. 90. Kuki M, Noumi T, Maeda M, Amemura A, Futai M, Functional domains of epsilon subunit of Escherichia coli H+-ATPase (F0F1). J Biol Chem 1988, 263(33):17437-17442. 91. Dunn SD, Zadorozny VD, Tozer RG, Orr LE, Epsilon subunit of Escherichia coli F1-ATPase: effects on affinity for aurovertin and inhibition of product release in unisite ATP hydrolysis. Biochemistry 1987, 26(14):4488-4493. 92. Aggeler R, Ogilvie I, Capaldi RA, Rotation of a gamma-epsilon subunit domain in the Escherichia coli F1F0-ATP synthase complex. The gamma-epsilon subunits are essentially randomly distributed relative to the alpha3beta3delta domain in the intact complex. J Biol Chem 1997, 272(31):19621-19624. 93. Tozer RG, Dunn SD, The epsilon subunit and inhibitory monoclonal antibodies interact with the carboxyl-terminal region of the beta subunit of Escherichia coli F1-ATPase. J Biol Chem 1987, 262(22):10706-10711. 94. Bragg PD, Hou C, Role of minor subunits in the structural asymmetry of the Escherichia coli F1-ATPase. Biochem Biophys Res Commun 1990, 166(1):431-435. 95. Cox GB, Cromer BA, Guss JM, Harvey I, Jeffrey PD, Solomon RG, Webb DC, Formation in vivo, purification and crystallization of a complex of the gamma and epsilon subunits of the F0F1-ATPase of Escherichia coli. J Mol Biol 1993, 229(4):1159-1162. 96. Tang C, Capaldi RA, Characterization of the interface between gamma and epsilon subunits of Escherichia coli F1-ATPase. J Biol Chem 1996, 271(6):3018-3024. 97. Sawada K, Watanabe H, Moritani-Otsuka C, Kanazawa H, Subunit interactions of Escherichia coli F1-ATPase: mutants of the gamma subunits defective in interaction with the epsilon subunit isolated by the yeast two-hybrid system. Arch Biochem Biophys 1997, 348(1):183-189. 98. Zhang Y, Oldenburg M, Fillingame RH, Suppressor mutations in F1 subunit epsilon recouple ATP-driven H+ translocation in uncoupled Q42E subunit c mutant of Escherichia coli F1F0 ATP synthase. J Biol Chem 1994,

PAGE 241

220 61 with dicyclohexylcarbodiimide. J Biol Chem 1991, 266(31):20934-20939. 99. Zhang Y, Fillingame RH, Subunits coupling H+ transport and ATP synthesis in the Escherichia coli ATP synthase. Cys-Cys cross-linking of F1 subunit epsilon to the polar loop of F0 subunit c. J Biol Chem 1995, 270(41):24609-24614. 100. Cipriano DJ, Bi Y, Dunn SD, Genetic fusions of globular proteins to the epsilon subunit of the Escherichia coli ATP synthase: Implications for in vivo rotational catalysis and epsilon subunit function. J Biol Chem 2002, 277(19):16782-16790. 101. Rodgers AJ, Wilce MC, Structure of the gamma-epsilon complex of ATP synthase. Nat Struct Biol 2000, 7(11):1051-1054. 102. Tsunoda SP, Rodgers AJ, Aggeler R, Wilce MC, Yoshida M, Capaldi RA, Large conformational changes of the epsilon subunit in the bacterial F1F0 ATP synthase provide a ratchet action to regulate this rotary motor enzyme. Proc Natl Acad Sci U S A 2001, 98(12):6560-6564. 103. Capaldi RA, Aggeler R, Wilkens S, Gruber G, Structural changes in the gamma and epsilon subunits of the Escherichia coli F1F0-type ATPase during energy coupling. J Bioenerg Biomembr 1996, 28(5):397-401. 104. Harris DA, Das AM, Control of mitochondrial ATP synthesis in the heart. Biochem J 1991, 280 ( Pt 3):561-573. 105. Jiang W, Hermolin J, Fillingame RH, The preferred stoichiometry of c subunits in the rotary motor sector of Escherichia coli ATP synthase is 10. Proc Natl Acad Sci U S A 2001, 98(9):4966-4971. 106. Hoppe J, Brunner J, Jorgensen BB, Structure of the membrane-embedded F0 part of F1F0 ATP synthase from Escherichia coli as inferred from labeling with 3-(Trifluoromethyl)-3-(m-[125I]iodophenyl)diazirine. Biochemistry 1984, 23(23):5610-5616. 107. Girvin ME, Hermolin J, Pottorf R, Fillingame RH, Organization of the F0 sector of Escherichia coli H+-ATPase: the polar loop region of subunit c extends from the cytoplasmic face of the membrane. Biochemistry 1989, 28(10):4340-4343. 108. Hensel M, Deckers-Hebestreit G, Schmid R, Altendorf K, Orientation of subunit c of the ATP synthase of Escherichia coli--a study with peptide-specific antibodies. Biochim Biophys Acta 1990, 1016(1):63-70. 109. Hoppe J, Schairer HU, Sebald W, Identification of amino-acid substitutions in the proteolipid subunit of the ATP synthase from dicyclohexylcarbodiimide-resistant mutants of Escherichia coli. Eur J Biochem 1980, 112(1):17-24. 110. Fillingame RH, Oldenburg M, Fraga D, Mutation of alanine 24 to serine in subunit c of the Escherichia coli F1F0-ATP synthase reduces reactivity of aspartyl

PAGE 242

221 111. Miller MJ, Oldenburg M, Fillingame RH, The essential carboxyl group in subunit c of the F1F0 ATP synthase can be moved and H(+)-translocating function retained. Proc Natl Acad Sci U S A 1990, 87(13):4900-4904. 112. Hermolin J, Fillingame RH, H+-ATPase activity of Escherichia coli F1F0 is blocked after reaction of dicyclohexylcarbodiimide with a single proteolipid (subunit c) of the F0 complex. J Biol Chem 1989, 264(7):3896-3903. 113. Dmitriev OY, Altendorf K, Fillingame RH, Reconstitution of the Fo complex of Escherichia coli ATP synthase from isolated subunits. Varying the number of essential carboxylates by co-incorporation of wild-type and mutant subunit c after purification in organic solvent. Eur J Biochem 1995, 233(2):478-483. 114. Fraga D, Fillingame RH, Conserved polar loop region of Escherichia coli subunit c of the F1F0 H+-ATPase. Glutamine 42 is not absolutely essential, but substitutions alter binding and coupling of F1 to F0. J Biol Chem 1989, 264(12):6797-6803. 115. Fraga D, Fillingame RH, Essential residues in the polar loop region of subunit c of Escherichia coli F1F0 ATP synthase defined by random oligonucleotide-primed mutagenesis. J Bacteriol 1991, 173(8):2639-2643. 116. Fraga D, Hermolin J, Fillingame RH, Transmembrane helix-helix interactions in F0 suggested by suppressor mutations to Ala24-->Asp/Asp61-->Gly mutant of ATP synthase subunit. J Biol Chem 1994, 269(4):2562-2567. 117. Miller MJ, Fraga D, Paule CR, Fillingame RH, Mutations in the conserved proline 43 residue of the uncE protein (subunit c) of Escherichia coli F1F0-ATPase alter the coupling of F1 to F0. J Biol Chem 1989, 264(1):305-311. 118. Watts SD, Zhang Y, Fillingame RH, Capaldi RA, The gamma subunit in the Escherichia coli ATP synthase complex (ECF1F0) extends through the stalk and contacts the c subunits of the F0 part. FEBS Lett 1995, 368(2):235-238. 119. Fillingame RH, Subunit c of F1F0 ATP synthase: structure and role in transmembrane energy transduction. Biochim Biophys Acta 1992, 1101(2):240-243. 120. Fillingame RH, H+ transport and coupling by the F0 sector of the ATP synthase: insights into the molecular mechanism of function. J Bioenerg Biomembr 1992, 24(5):485-491. 121. Hatch LP, Cox GB, Howitt SM, The essential arginine residue at position 210 in the alpha subunit of the Escherichia coli ATP synthase can be transferred to position 252 with partial retention of activity. J Biol Chem 1995, 270(49):29407-29412.

PAGE 243

222 from Methanococcus jannaschii has six predicted transmembrane helices but only 122. Girvin ME, Rastogi VK, Abildgaard F, Markley JL, Fillingame RH, Solution structure of the transmembrane H+-transporting subunit c of the F1F0 ATP synthase. Biochemistry 1998, 37(25):8817-8824. 123. Jones PC, Fillingame RH, Genetic fusions of subunit c in the F0 sector of H+-transporting ATP synthase. Functional dimers and trimers and determination of stoichiometry by cross-linking analysis. J Biol Chem 1998, 273(45):29701-29705. 124. Jones PC, Jiang W, Fillingame RH, Arrangement of the multicopy H+-translocating subunit c in the membrane sector of the Escherichia coli F1F0 ATP synthase. J Biol Chem 1998, 273(27):17178-17185. 125. Assadi-Porter FM, Fillingame RH, Proton-translocating carboxyl of subunit c of F1Fo H(+)-ATP synthase: the unique environment suggested by the pKa determined by 1H NMR. Biochemistry 1995, 34(49):16186-16193. 126. Singh S, Turina P, Bustamante CJ, Keller DJ, Capaldi R, Topographical structure of membrane-bound Escherichia coli F1F0 ATP synthase in aqueous buffer. FEBS Lett 1996, 397(1):30-34. 127. Wilkens S, Dunn SD, Capaldi RA, A cryoelectron microscopy study of the interaction of the Escherichia coli F1-ATPase with subunit b dimer. FEBS Lett 1994, 354(1):37-40. 128. Girvin ME, Fillingame RH, Helical structure and folding of subunit c of F1F0 ATP synthase: 1H NMR resonance assignments and NOE analysis. Biochemistry 1993, 32(45):12167-12177. 129. Girvin ME, Fillingame RH, Hairpin folding of subunit c of F1Fo ATP synthase: 1H distance measurements to nitroxide-derivatized aspartyl-61. Biochemistry 1994, 33(3):665-674. 130. Girvin ME, Fillingame RH, Determination of local protein structure by spin label difference 2D NMR: the region neighboring Asp61 of subunit c of the F1F0 ATP synthase. Biochemistry 1995, 34(5):1635-1645. 131. von Ballmoos C, Meier T, Dimroth P, Membrane embedded location of Na+ or H+ binding sites on the rotor ring of F1F0 ATP synthases. Eur J Biochem 2002, 269(22):5581-5589. 132. Fillingame RH, Jones PC, Jiang W, Valiyaveetil FI, Dmitriev OY, Subunit organization and structure in the F0 sector of Escherichia coli F1F0 ATP synthase. Biochim Biophys Acta 1998, 1365(1-2):135-142. 133. Ruppert C, Kavermann H, Wimmers S, Schmid R, Kellermann J, Lottspeich F, Huber H, Stetter KO, Muller V, The proteolipid of the A(1)A(0) ATP synthase

PAGE 244

223 F0F1-ATPase: a requirement for arginine at position 210 of the subunit. Biochim Biophys Acta 1987, 894(3):399-406. two proton-translocating carboxyl groups. J Biol Chem 1999, 274(36):25281-25284. 134. Kashket ER, Stoichiometry of the H+-ATPase of growing and resting, aerobic Escherichia coli. Biochemistry 1982, 21(22):5534-5538. 135. Lehninger AL, Nelson DL, Cox MM, Lehninger principles of biochemistry. In., 3rd edn. New York: Worth Publishers; 2000. 136. Dmitriev O, Jones PC, Jiang W, Fillingame RH, Structure of the membrane domain of subunit b of the Escherichia coli F0F1 ATP synthase. J Biol Chem 1999, 274(22):15598-15604. 137. Wilkens S, Dunn SD, Chandler J, Dahlquist FW, Capaldi RA, Solution structure of the N-terminal domain of the delta subunit of the E. coli ATPsynthase [letter]. Nat Struct Biol 1997, 4(3):198-201. 138. Del Rizzo PA, Bi Y, Dunn SD, Shilton BH, The "second stalk" of Escherichia coli ATP synthase: structure of the isolated dimerization domain. Biochemistry 2002, 41(21):6875-6884. 139. Sternweis PC, Smith JB, Characterization of the purified membrane attachment (beta) subunit of the proton translocating adenosine triphosphatase from Escherichia coli. Biochemistry 1977, 16(18):4020-4025. 140. Fillingame RH, Molecular Mechanics of ATP Synthesis by F1F0-Type H+-Transporting ATP Synthases. In: The Bacteria. Edited by Krulwich TA, vol. XII. New York: Academic Press; 1990: 345-391. 141. Prescott M, Nowakowski S, Gavin P, Nagley P, Whisstock JC, Devenish RJ, Subunit gamma-green fluorescent protein fusions are functionally incorporated into mitochondrial F1F0-ATP synthase, arguing against a rigid cap structure at the top of F1. J Biol Chem 2003, 278(1):251-256. 142. Weber J, Wilke-Mounts S, Senior AE, Identification of the F1-binding surface on the delta-subunit of ATP synthase. J Biol Chem 2003. 143. Weber J, Muharemagic A, Wilke-Mounts S, Senior AE, F1Fo-ATP synthase: Binding of delta subunit to a 22-residue peptide mimicking the N-terminal region of alpha subunit. J Biol Chem 2003. 144. Hartzog PE, Cain BD, Second-site suppressor mutations at glycine 218 and histidine 245 in the alpha subunit of F1F0 ATP synthase in Escherichia coli. J Biol Chem 1994, 269(51):32313-32317. 145. Lightowlers RN, Howitt SM, Hatch L, Gibson F, Cox GB, The proton pore in the Escherichia colia

PAGE 245

224 synthase is important for the structure or assembly of the membrane sector F(o). Arch Biochem Biophys 1999, 368(1):193-197. 146. Cain BD, Simoni RD, Proton translocation by the F1F0 ATPase of Escherichia coli. Mutagenic analysis of the a subunit. J Biol Chem 1989, 264(6):3292-3300. 147. Eya S, Maeda M, Futai M, Role of the carboxyl terminal region of H(+)-ATPase (F0F1) a subunit from Escherichia coli. Arch Biochem Biophys 1991, 284(1):71-77. 148. Schneider E, Altendorf K, All three subunits are required for the reconstitution of an active proton channel (F0) of Escherichia coli ATP synthase (F1F0). Embo J 1985, 4(2):515-518. 149. von Meyenburg K, Jorgensen BB, Michelsen O, Sorensen L, McCarthy JE, Proton conduction by subunit a of the membrane-bound ATP synthase of Escherichia coli revealed after induced overproduction. Embo J 1985, 4(9):2357-2363. 150. Hermolin J, Fillingame RH, Assembly of F0 sector of Escherichia coli H+ ATP synthase. Interdependence of subunit insertion into the membrane. J Biol Chem 1995, 270(6):2815-2817. 151. Akiyama Y, Kihara A, Ito K, Subunit a of proton ATPase F0 sector is a substrate of the FtsH protease in Escherichia coli. FEBS Lett 1996, 399(1-2):26-28. 152. Vik SB, Dao NN, Prediction of transmembrane topology of F0 proteins from Escherichia coli F1F0 ATP synthase using variational and hydrophobic moment analyses. Biochim Biophys Acta 1992, 1140(2):199-207. 153. Wang S, Vik SB, Single amino acid insertions probe the alpha subunit of the Escherichia coli F1F0-ATP synthase. J Biol Chem 1994, 269(4):3095-3099. 154. Valiyaveetil FI, Fillingame RH, Transmembrane topography of subunit a in the Escherichia coli F1F0 ATP synthase. J Biol Chem 1998, 273(26):16241-16247. 155. Long JC, Wang S, Vik SB, Membrane topology of subunit a of the F1F0 ATP synthase as determined by labeling of unique cysteine residues. J Biol Chem 1998, 273(26):16235-16240. 156. Wada T, Long JC, Zhang D, Vik SB, A novel labeling approach supports the five-transmembrane model of subunit a of the Escherichia coli ATP synthase. J Biol Chem 1999, 274(24):17353-17357. 157. Jiang W, Fillingame RH, Interacting helical faces of subunits a and c in the F1F0 ATP synthase of Escherichia coli defined by disulfide cross-linking. Proc Natl Acad Sci U S A 1998, 95(12):6607-6612. 158. Patterson AR, Wada T, Vik SB, His(15) of subunit a of the Escherichia coli ATP

PAGE 246

225 159. Vik SB, Long JC, Wada T, Zhang D, A model for the structure of subunit a of the Escherichia coli ATP synthase and its role in proton translocation. Biochim Biophys Acta 2000, 1458(2-3):457-466. 160. Jager H, Birkenhager R, Stalz WD, Altendorf K, Deckers-Hebestreit G, Topology of subunit a of the Escherichia coli ATP synthase. Eur J Biochem 1998, 251(1-2):122-132. 161. Lewis MJ, Chang JA, Simoni RD, A topological analysis of subunit alpha from Escherichia coli F1F0-ATP synthase predicts eight transmembrane segments. J Biol Chem 1990, 265(18):10541-10550. 162. Yamada H, Moriyama Y, Maeda M, Futai M, Transmembrane topology of Escherichia coli H(+)-ATPase (ATP synthase) subunit a. FEBS Lett 1996, 390(1):34-38. 163. Lewis MJ, Simoni RD, Deletions in hydrophilic domains of subunit a from the Escherichia coli F1F0-ATP synthase interfere with membrane insertion or F0 assembly. J Biol Chem 1992, 267(5):3482-3489. 164. Vik SB, Lee D, Marshall PA, Temperature-sensitive mutations at the carboxy terminus of the alpha subunit of the Escherichia coli F1F0 ATP synthase. J Bacteriol 1991, 173(14):4544-4548. 165. Cain BD, Simoni RD, Impaired proton conductivity resulting from mutations in the a subunit of F1F0 ATPase in Escherichia coli. J Biol Chem 1986, 261(22):10043-10050. 166. Cain BD, Mutagenic analysis of the F0 stator subunits. J Bioenerg Biomembr 2000, 32(4):365-371. 167. Gardner JL, Cain BD, Amino acid substitutions in the a subunit affect the epsilon subunit of F1F0 ATP synthase from Escherichia coli. Arch Biochem Biophys 1999, 361(2):302-308. 168. Valiyaveetil FI, Fillingame RH, On the role of Arg-210 and Glu-219 of subunit a in proton translocation by the Escherichia coli F0F1-ATP synthase. J Biol Chem 1997, 272(51):32635-32641. 169. Rastogi VK, Girvin ME, Structural changes linked to proton translocation by subunit c of the ATP synthase [see comments]. Nature 1999, 402(6759):263-268. 170. Hatch LP, Cox GB, Howitt SM, Glutamate residues at positions 219 and 252 in the a-subunit of the Escherichia coli ATP synthase are not functionally equivalent. Biochim Biophys Acta 1998, 1363(3):217-223.

PAGE 247

226 171. Cain BD, Simoni RD, Interaction between Glu-219 and His-245 within the a subunit of F1F0-ATPase in Escherichia coli. J Biol Chem 1988, 263(14):6606-6612. 172. Hartzog PE, Cain BD, Mutagenic analysis of the a subunit of the F1F0 ATP synthase in Escherichia coli: Gln-252 through Tyr-263. J Bacteriol 1993, 175(5):1337-1343. 173. Kuo PH, Nakamoto RK, Intragenic and intergenic suppression of the Escherichia coli ATP synthase subunit a mutation of Gly-213 to Asn: functional interactions between residues in the proton transport site. Biochem J 2000, 347 Pt 3:797-805. 174. Angevine CM, Herold KA, Fillingame RH, Aqueous access pathways in subunit a of rotary ATP synthase extend to both sides of the membrane. Proc Natl Acad Sci U S A 2003, 100(23):13179-13183. 175. Smith JB, Sternweis PC, Heppel LA, Partial purification of active delta and epsilon subunits of the membrane ATPase from escherichia coli. J Supramol Struct 1975, 3(3):248-255. 176. Angov E, Ng TC, Brusilow WS, Effect of the delta subunit on assembly and proton permeability of the F0 proton channel of Escherichia coli F1F0 ATPase. J Bacteriol 1991, 173(1):407-411. 177. Jounouchi M, Takeyama M, Chaiprasert P, Noumi T, Moriyama Y, Maeda M, Futai M, Escherichia coli H(+)-ATPase: role of the delta subunit in binding Fl to the Fo sector. Arch Biochem Biophys 1992, 292(2):376-381. 178. Stack AE, Cain BD, Mutations in the delta subunit influence the assembly of F1F0 ATP synthase in Escherichia coli. J Bacteriol 1994, 176(2):540-542. 179. Hazard AL, Senior AE, Mutagenesis of subunit delta from Escherichia coli F1F0-ATP synthase. J Biol Chem 1994, 269(1):418-426. 180. Hazard AL, Senior AE, Defective energy coupling in delta-subunit mutants of Escherichia coli F1F0-ATP synthase. J Biol Chem 1994, 269(1):427-432. 181. Tozer RG, Dunn SD, Column centrifugation generates an intersubunit disulfide bridge in Escherichia coli F1-ATPase. Eur J Biochem 1986, 161(2):513-518. 182. Bragg PD, Hou C, Effect of disulfide cross-linking between alpha and delta subunits on the properties of the F1 adenosine triphosphatase of Escherichia coli. Biochim Biophys Acta 1986, 851(3):385-394. 183. Bragg PD, Hou C, Chemical crosslinking of alpha subunits in the F1 adenosine triphosphatase of Escherichia coli. Arch Biochem Biophys 1986, 244(1):361-372.

PAGE 248

227 2000, 267(10):3040-3048. 184. Lill H, Hensel F, Junge W, Engelbrecht S, Cross-linking of engineered subunit to ()3 in chloroplast F-ATPase. J Biol Chem 1996, 271(51):32737-32742. 185. Ogilvie I, Aggeler R, Capaldi RA, Cross-linking of the delta subunit to one of the three alpha subunits has no effect on functioning, as expected if delta is a part of the stator that links the F1 and F0 parts of the Escherichia coli ATP synthase. J Biol Chem 1997, 272(26):16652-16656. 186. Bottcher B, Bertsche I, Reuter R, Graber P, Direct visualisation of conformational changes in EF(0)F(1) by electron microscopy. J Mol Biol 2000, 296(2):449-457. 187. Wilkens S, Zhou J, Nakayama R, Dunn SD, Capaldi RA, Localization of the delta subunit in the Escherichia coli F1F0 ATPsynthase by immuno electron microscopy: the delta subunit binds on top of the F1. J Mol Biol 2000, 295(3):387-391. 188. Ormo M, Cubitt AB, Kallio K, Gross LA, Tsien RY, Remington SJ, Crystal structure of the Aequorea victoria green fluorescent protein. Science 1996, 273(5280):1392-1395. 189. Weber J, Senior AE, ATP synthesis driven by proton transport in F1F0-ATP synthase. FEBS Lett 2003, 545(1):61-70. 190. Schneider E, Altendorf K, Subunit b of the membrane moiety (F0) of ATP synthase (F1F0) from Escherichia coli is indispensable for H+ translocation and binding of the water-soluble F1 moiety. Proc Natl Acad Sci U S A 1984, 81(23):7279-7283. 191. Walker JE, Saraste M, Gay NJ, E. coli F1-ATPase interacts with a membrane protein component of a proton channel. Nature 1982, 298(5877):867-869. 192. Revington M, McLachlin DT, Shaw GS, Dunn SD, The dimerization domain of the b subunit of the Escherichia coli F1F0-ATPase. J Biol Chem 1999, 274(43):31094-31101. 193. Sorgen PL, Bubb MR, Cain BD, Lengthening the second stalk of F1F0 ATP synthase in Escherichia coli. J Biol Chem 1999, 274(51):36261-36266. 194. Sorgen PL, Caviston TL, Perry RC, Cain BD, Deletions in the second stalk of F1F0-ATP synthase in Escherichia coli. J Biol Chem 1998, 273(43):27873-27878. 195. Grabar TB, Cain BD, Integration of b subunits of unequal lengths into F1F0-ATP synthase. J Biol Chem 2003, 278(37):34751-34756. 196. Greie JC, Deckers-Hebestreit G, Altendorf K, Secondary structure composition of reconstituted subunit b of the Escherichia coli ATP synthase. Eur J Biochem

PAGE 249

228 197. Dunn SD, The polar domain of the b subunit of Escherichia coli F1F0-ATPase forms an elongated dimer that interacts with the F1 sector. J Biol Chem 1992, 267(11):7630-7636. 198. Rodgers AJ, Capaldi RA, The second stalk composed of the band delta-subunits connects F0 to F1 via an alpha-subunit in the Escherichia coli ATP synthase. J Biol Chem 1998, 273(45):29406-29410. 199. Sorgen PL, Bubb MR, McCormick KA, Edison AS, Cain BD, Formation of the b subunit dimer is necessary for interaction with F1ATPase. Biochemistry 1998, 37(3):923-932. 200. Rodgers AJ, Wilkens S, Aggeler R, Morris MB, Howitt SM, Capaldi RA, The subunit delta-subunit b domain of the Escherichia coli F1F0 ATPase. The b subunits interact with F1 as a dimer and through the delta subunit. J Biol Chem 1997, 272(49):31058-31064. 201. Hoppe J, Montecucco C, Friedl P, Labeling of subunit b of the ATP synthase from Escherichia coli with a photoreactive phospholipid analogue. J Biol Chem 1983, 258(5):2882-2885. 202. Hardy AW, Grabar TB, Bhatt D, Cain BD, Mutagenesis studies of the F1F0 ATP synthase b subunit membrane domain. J Bioenerg Biomembr 2003, 35(5):389-397. 203. McCormick KA, Deckers-Hebestreit G, Altendorf K, Cain BD, Characterization of mutations in the b subunit of F1F0 ATP synthase in Escherichia coli. J Biol Chem 1993, 268(33):24683-24691. 204. Revington M, Dunn SD, Shaw GS, Folding and stability of the b subunit of the F(1)F(0) ATP synthase. Protein Sci 2002, 11(5):1227-1238. 205. McLachlin DT, Bestard JA, Dunn SD, The b and subunits of the Escherichia coli ATP synthase interact via residues in their C-terminal regions. J Biol Chem 1998, 273(24):15162-15168. 206. Takeyama M, Noumi T, Maeda M, Futai M, F0 portion of Escherichia coli H+-ATPase. Carboxyl-terminal region of the b subunit is essential for assembly of functional F0. J Biol Chem 1988, 263(31):16106-16112. 207. McLachlin DT, Dunn SD, Dimerization interactions of the b subunit of the Escherichia coli F1F0-ATPase. J Biol Chem 1997, 272(34):21233-21239. 208. Dunn SD, Bi Y, Revington M, A re-examination of the structural and functional consequences of mutation of alanine-128 of the b subunit of Escherichia coli ATP synthase to aspartic acid. Biochim Biophys Acta 2000, 1459(2-3):521-527.

PAGE 250

229 coli ATP synthase. J Biol Chem 1998, 273(15):8646-8651. 209. Porter AC, Kumamoto C, Aldape K, Simoni RD, Role of the b subunit of the Escherichia coli proton-translocating ATPase. A mutagenic analysis. J Biol Chem 1985, 260(13):8182-8187. 210. Kumamoto CA, Simoni RD, Genetic evidence for interaction between the a and b subunits of the F0 portion of the Escherichia coli proton translocating ATPase. J Biol Chem 1986, 261(22):10037-10042. 211. Kumamoto CA, Simoni RD, A mutation of the c subunit of the Escherichia coli proton-translocating ATPase that suppresses the effects of a mutant b subunit. J Biol Chem 1987, 262(7):3060-3064. 212. Cox GB, Fimmel AL, Gibson F, Hatch L, The mechanism of ATP synthase: a reassessment of the functions of the b and a subunits. Biochim Biophys Acta 1986, 849(1):62-69. 213. Tiburzy HJ, Berzborn RJ, Subunit II (b') and not subunit I (b) of photosynthetic ATP synthases is equivalent to subunit b of the ATP synthases from nonphotosynthetic eubacteria. Evidence for a new assignment of b-type F0 subunits. Z Naturforsch [C] 1997, 52(11-12):789-798. 214. Blair A, Ngo L, Park J, Paulsen IT, Saier MH, Jr., Phylogenetic analyses of the homologous transmembrane channel-forming proteins of the F0F1-ATPases of bacteria, chloroplasts and mitochondria. Microbiology 1996, 142 ( Pt 1):17-32. 215. Caviston TL, Ketchum CJ, Sorgen PL, Nakamoto RK, Cain BD, Identification of an uncoupling mutation affecting the b subunit of F1F0 ATP synthase in Escherichia coli. FEBS Lett 1998, 429(2):201-206. 216. McLachlin DT, Coveny AM, Clark SM, Dunn SD, Site-directed cross-linking of b to the alpha, beta, and a subunits of the Escherichia coli ATP synthase. J Biol Chem 2000, 275(23):17571-17577. 217. Long JC, DeLeon-Rangel J, Vik SB, Characterization of the first cytoplasmic loop of subunit a of the Escherichia coli ATP synthase by surface labeling, cross-linking, and mutagenesis. J Biol Chem 2002, 277(30):27288-27293. 218. McCormick KA, Cain BD, Targeted mutagenesis of the b subunit of F1F0 ATP synthase in Escherichia coli: Glu-77 through Gln-85. J Bacteriol 1991, 173(22):7240-7248. 219. Sawada K, Kuroda N, Watanabe H, Moritani-Otsuka C, Kanazawa H, Interaction of the delta and b subunits contributes to F1 and F0 interaction in the Escherichia coli F1F0-ATPase. J Biol Chem 1997, 272(48):30047-30053. 220. Dunn SD, Chandler J, Characterization of a b2delta complex from Escherichia

PAGE 251

230 association with the F1-moiety. 1998, 37(19):6911-6923. 221. McLachlin DT, Dunn SD, Disulfide linkage of the b and delta subunits does not affect the function of the Escherichia coli ATP synthase. Biochemistry 2000, 39(12):3486-3490. 222. Cherepanov DA, Mulkidjanian AY, Junge W, Transient accumulation of elastic energy in proton translocating ATP synthase. FEBS Lett 1999, 449(1):1-6. 223. Kersten MV, Dunn SD, Wise JG, Vogel PD, Site-directed spin-labeling of the catalytic sites yields insight into structural changes within the F0F1-ATP synthase of Escherichia coli. Biochemistry 2000, 39(13):3856-3860. 224. Walker JE, Saraste M, Gay NJ, The unc operon. Nucleotide sequence, regulation and structure of ATPsynthase. Biochim Biophys Acta 1984, 768(2):164-200. 225. Dunn SD, Heppel LA, Properties and functions of the subunits of the Escherichia coli coupling factor ATPase. Arch Biochem Biophys 1981, 210(2):421-436. 226. Schneider E, Altendorf K, Bacterial adenosine 5'-triphosphate synthase (F1F0): purification and reconstitution of F0 complexes and biochemical and functional characterization of their subunits. Microbiol Rev 1987, 51(4):477-497. 227. Fromme P, Graber, P., Salnikow, J., FEBS Lett 1987, 218:27-30. 228. Collinson IR, Fearnley IM, Skehel JM, Runswick MJ, Walker JE, ATP synthase from bovine heart mitochondria: identification by proteolysis of sites in F0 exposed by removal of F1 and the oligomycin-sensitivity conferral protein. Biochem J 1994, 303 ( Pt 2):639-645. 229. Walker JE, Collinson IR, The role of the stalk in the coupling mechanism of F1F0-ATPases. FEBS Lett 1994, 346(1):39-43. 230. Lutter R, Saraste M, van Walraven HS, Runswick MJ, Finel M, Deatherage JF, Walker JE, F1F0-ATP synthase from bovine heart mitochondria: development of the purification of a monodisperse oligomycin-sensitive ATPase. Biochem J 1993, 295 ( Pt 3):799-806. 231. Miki J, Maeda M, Mukohata Y, Futai M, The gamma-subunit of ATP synthase from spinach chloroplasts. Primary structure deduced from the cloned cDNA sequence. FEBS Lett 1988, 232(1):221-226. 232. Nalin CM, McCarty RE, Role of a disulfide bond in the gamma subunit in activation of the ATPase of chloroplast coupling factor 1. J Biol Chem 1984, 259(11):7275-7280. 233. Pan W, Ko YH, Pedersen PL, Delta subunit of rat liver mitochondrial ATP synthase: molecular description and novel insights into the nature of its Biochemistry

PAGE 252

231 234. Walker JE, Fearnley IM, Gay NJ, Gibson BW, Northrop FD, Powell SJ, Runswick MJ, Saraste M, Tybulewicz VL, Primary structure and subunit stoichiometry of F1-ATPase from bovine mitochondria. J Mol Biol 1985, 184(4):677-701. 235. Ovchinnikov Yu A, Modyanov NN, Grinkevich VA, Aldanova NA, Kostetsky PV, Trubetskaya OE, Hundal T, Ernster L, Oligomycin sensitivity-conferring protein (OSCP) of beef heart mitochondria. Internal sequence homology and structural relationship with other proteins. FEBS Lett 1984, 175(1):109-112. 236. Ovchinnikov YA, Modyanov NN, Grinkevich VA, Aldanova NA, Trubetskaya OE, Nazimov IV, Hundal T, Ernster L, Amino acid sequence of the oligomycin sensitivity-conferring protein (OSCP) of beef-heart mitochondria and its homology with the delta-subunit of the F1-ATPase of Escherichia coli. FEBS Lett 1984, 166(1):19-22. 237. Uh M, Jones D, Mueller DM, The gene coding for the yeast oligomycin sensitivity-conferring protein. J Biol Chem 1990, 265(31):19047-19052. 238. Van Walraven HS, Lutter R, Walker JE, Organization and sequences of genes for the subunits of ATP synthase in the thermophilic cyanobacterium Synechococcus 6716. Biochem J 1993, 294 ( Pt 1):239-251. 239. Devenish RJ, Papakonstantinou T, Galanis M, Law RH, Linnane AW, Nagley P, Structure/function analysis of yeast mitochondrial ATP synthase subunit 8. Ann N Y Acad Sci 1992, 671:403-414. 240. Walker JE, Runswick MJ, Poulter L, ATP synthase from bovine mitochondria. The characterization and sequence analysis of two membrane-associated sub-units and of the corresponding cDNAs. J Mol Biol 1987, 197(1):89-100. 241. Walker JE, Lutter R, Dupuis A, Runswick MJ, Identification of the subunits of F1F0-ATPase from bovine heart mitochondria. Biochemistry 1991, 30(22):5369-5378. 242. Torok K, Joshi S, Cross-linking of bovine mitochondrial H+-ATPase by copper--o-phenanthroline. Interaction of the oligomycin-sensitivity-conferring protein with a 24-kDa protein. Eur J Biochem 1985, 153(1):155-159. 243. Houstek J, Kopecky J, Zanotti F, Guerrieri F, Jirillo E, Capozza G, Papa S, Topological and functional characterization of the F0I subunit of the membrane moiety of the mitochondrial H+-ATP synthase. Eur J Biochem 1988, 173(1):1-8. 244. Kimura T, Nakamura K, Kajiura H, Hattori H, Nelson N, Asahi T, Correspondence of minor subunits of plant mitochondrial F1 ATPase to F1F0ATPase subunits of other organisms. J Biol Chem 1989, 264(6):3183-3186.

PAGE 253

232 coupling of F1 to F0. J Biol Chem 1994, 269(10):7532-7537. 245. Manson MD, Tedesco P, Berg HC, Harold FM, Van der Drift C, A protonmotive force drives bacterial flagella. Proc Natl Acad Sci U S A 1977, 74(7):3060-3064. 246. Boyer PD, The binding change mechanism for ATP synthase--some probabilities and possibilities. Biochim Biophys Acta 1993, 1140(3):215-250. 247. Yasuda R, Noji H, Kinosita K, Jr., Yoshida M, F1-ATPase is a highly efficient molecular motor that rotates with discrete 120 degree steps. Cell 1998, 93(7):1117-1124. 248. Ariga T, Masaike T, Noji H, Yoshida M, Stepping rotation of F(1)-ATPase with one, two, or three altered catalytic sites that bind ATP only slowly. J Biol Chem 2002, 277(28):24870-24874. 249. Yasuda R, Noji H, Yoshida M, Kinosita K, Jr., Itoh H, Resolution of distinct rotational substeps by submillisecond kinetic analysis of F1-ATPase. Nature 2001, 410(6831):898-904. 250. Nagle JF, Tristram-Nagle S, Hydrogen bonded chain mechanisms for proton conduction and proton pumping. J Membr Biol 1983, 74(1):1-14. 251. Deckers-Hebestreit G, Altendorf K, The Fo complex of the proton-translocating F-type ATPase of Escherichia coli. J Exp Biol 1992, 172:451-459. 252. Dimroth P, Sodium ion transport decarboxylases and other aspects of sodium ion cycling in bacteria. Microbiol Rev 1987, 51(3):320-340. 253. Fillingame RH, Coupling H+ transport and ATP synthesis in F1F0-ATP synthases: glimpses of interacting parts in a dynamic molecular machine. J Exp Biol 1997, 200(Pt 2):217-224. 254. Junge W, ATP synthase and other motor proteins. Proc Natl Acad Sci U S A 1999, 96(9):4735-4737. 255. Vik SB, Antonio BJ, A mechanism of proton translocation by F1F0 ATP synthases suggested by double mutants of the a subunit. J Biol Chem 1994, 269(48):30364-30369. 256. Inesi G, Kirtley MR, Structural features of cation transport ATPases. J Bioenerg Biomembr 1992, 24(3):271-283. 257. Mosher ME, White LK, Hermolin J, Fillingame RH, H+-ATPase of Escherichia coli. An uncE mutation impairing coupling between F1 and Fo but not Fo-mediated H+ translocation. J Biol Chem 1985, 260(8):4807-4814. 258. Fraga D, Hermolin J, Oldenburg M, Miller MJ, Fillingame RH, Arginine 41 of subunit c of Escherichia coli H(+)-ATP synthase is essential in binding and

PAGE 254

233 260(20):11207-11215. 259. Shin K, Nakamoto RK, Maeda M, Futai M, F0F1-ATPase gamma subunit mutations perturb the coupling between catalysis and transport. J Biol Chem 1992, 267(29):20835-20839. 260. Nakamoto RK, Maeda M, Futai M, The gamma subunit of the Escherichia coli ATP synthase. Mutations in the carboxyl-terminal region restore energy coupling to the amino-terminal mutant gamma Met-23-->Lys. J Biol Chem 1993, 268(2):867-872. 261. Nakamoto RK, al-Shawi MK, Futai M, The ATP synthase gamma subunit. Suppressor mutagenesis reveals three helical regions involved in energy coupling. J Biol Chem 1995, 270(23):14042-14046. 262. Dunn SD, Futai M, Reconstitution of a functional coupling factor from the isolated subunits of Escherichia coli F1 ATPase. J Biol Chem 1980, 255(1):113-118. 263. Boyer PD, A perspective of the binding change mechanism for ATP synthesis. Faseb J 1989, 3(10):2164-2178. 264. Futai M, Kanazawa H, Structure and function of proton-translocating adenosine triphosphatase (F0F1): biochemical and molecular biological approaches. Microbiol Rev 1983, 47(3):285-312. 265. Gibson F, Downie JA, Cox GB, Radik J, Mu-induced polarity in the unc operon of Escherichia coli. J Bacteriol 1978, 134(3):728-736. 266. Nielsen J, Jorgensen BB, van Meyenburg KV, Hansen FG, The promoters of the atp operon of Escherichia coli K12. Mol Gen Genet 1984, 193(1):64-71. 267. Kasimoglu E, Park SJ, Malek J, Tseng CP, Gunsalus RP, Transcriptional regulation of the proton-translocating ATPase (atpIBEFHAGDC) operon of Escherichia coli: control by cell growth rate. J Bacteriol 1996, 178(19):5563-5567. 268. McCarthy JE, Expression of the unc genes in Escherichia coli. J Bioenerg Biomembr 1988, 20(1):19-39. 269. Brusilow WS, Porter AC, Simoni RD, Cloning and expression of uncI, the first gene of the unc operon of Escherichia coli. J Bacteriol 1983, 155(3):1265-1270. 270. Brusilow WS, Assembly of the Escherichia coli F1F0 ATPase, a large multimeric membrane-bound enzyme. Mol Microbiol 1993, 9(3):419-424. 271. Aris JP, Klionsky DJ, Simoni RD, The F0 subunits of the Escherichia coli F1F0-ATP synthase are sufficient to form a functional proton pore. J Biol Chem 1985,

PAGE 255

234 203 Pt 1:19-28. 272. Cox GB, Jans, D., Fimmel, A., Gibson, F., Hatch, L., The role of the b subunit in the integrated assembly of the F1F0 ATP synthase. Biochim Biophys Acta 1984, 768:201-208. 273. Klionsky DJ, Brusilow WS, Simoni RD, In vivo evidence for the role of the epsilon subunit as an inhibitor of the proton-translocating ATPase of Escherichia coli. J Bacteriol 1984, 160(3):1055-1060. 274. Xiong H, Vik SB, Alanine-scanning mutagenesis of the epsilon subunit of the F1-F0 ATP synthase from Escherichia coli reveals two classes of mutants. J Biol Chem 1995, 270(40):23300-23304. 275. Xiong H, Vik SB, Construction and plasmid-borne complementation of strains lacking the epsilon subunit of the Escherichia coli F1F0 ATP synthase. J Bacteriol 1995, 177(3):851-853. 276. Xiong H, Zhang D, Vik SB, Subunit epsilon of the Escherichia coli ATP synthase: novel insights into structure and function by analysis of thirteen mutant forms. Biochemistry 1998, 37(46):16423-16429. 277. Pati S, Brusilow WS, Deckers-Hebestreit G, Altendorf K, Assembly of the F0 proton channel of the Escherichia coli F1F0 ATPase: low proton conductance of reconstituted Fo sectors synthesized and assembled in the absence of F1. Biochemistry 1991, 30(19):4710-4714. 278. Pati S, Brusilow WS, The roles of the alpha and gamma subunits in proton conduction through the Fo sector of the proton-translocating ATPase of Escherichia coli. J Biol Chem 1989, 264(5):2640-2644. 279. Monticello RA, Angov E, Brusilow WS, Effects of inducing expression of cloned genes for the F0 proton channel of the Escherichia coli F1F0 ATPase. J Bacteriol 1992, 174(10):3370-3376. 280. Grabar TB, Cain BD, Genetic complementation between mutant b subunits in F1F0 ATP synthase. J Biol Chem 2004. 281. Bulygin VV, Duncan TM, Cross RL, Rotation of the epsilon subunit during catalysis by Escherichia coli FOF1-ATP synthase. J Biol Chem 1998, 273(48):31765-31769. 282. Turina P, Structural changes during ATP hydrolysis activity of the ATP synthase from Escherichia coli as revealed by fluorescent probes. J Bioenerg Biomembr 2000, 32(4):373-381. 283. Altendorf K, Stalz W, Greie J, Deckers-Hebestreit G, Structure and function of the F(o) complex of the ATP synthase from Escherichia coli. J Exp Biol 2000,

PAGE 256

235 284. Deckers-Hebestreit G, Altendorf K, Accessibility of F0 subunits from Escherichia coli ATP synthase. A study with subunit specific antisera. Eur J Biochem 1986, 161(1):225-231. 285. Deckers-Hebestreit G, Simoni RD, Altendorf K, Influence of subunit-specific antibodies on the activity of the F0 complex of the ATP synthase of Escherichia coli. I. Effects of subunit b-specific polyclonal antibodies. J Biol Chem 1992, 267(17):12364-12369. 286. Markwell MA, Haas SM, Bieber LL, Tolbert NE, A modification of the Lowry procedure to simplify protein determination in membrane and lipoprotein samples. Anal Biochem 1978, 87(1):206-210. 287. Tamarappoo BK, Handlogten ME, Laine RO, Serrano MA, Dugan J, Kilberg MS, Identification of the protein responsible for hepatic system N amino acid transport activity. J Biol Chem 1992, 267(4):2370-2374. 288. McLachlin DT, Bestard JA, Dunn SD, The b and delta subunits of the Escherichia coli ATP synthase interact via residues in their C-terminal regions. J Biol Chem 1998, 273(24):15162-15168. 289. Smith PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, Fujimoto EK, Goeke NM, Olson BJ, Klenk DC, Measurement of protein using bicinchoninic acid. Anal Biochem 1985, 150(1):76-85. 290. Boyer PD, The ATP synthase--a splendid molecular machine. Annu Rev Biochem 1997, 66:717-749. 291. Capaldi RA, Aggeler R, Mechanism of the F(1)F(0)-type ATP synthase, a biological rotary motor. Trends Biochem Sci 2002, 27(3):154-160. 292. Greie JC, Deckers-Hebestreit G, Altendorf K, Subunit organization of the stator part of the F0 complex from Escherichia coli ATP synthase. J Bioenerg Biomembr 2000, 32(4):357-364. 293. Ziegler M, Xiao R, Penefsky HS, Close proximity of Cys64 and Cys140 in the delta subunit of Escherichia coli F1-ATPase. J Biol Chem 1994, 269(6):4233-4239. 294. Vogel PD, Insights into ATP synthase structure and function using affinity and site-specific spin labeling. J Bioenerg Biomembr 2000, 32(4):413-421. 295. Berzborn RJ, Klein-Hitpass L, Otto J, Schunemann S, Oworah-Nkruma R, Meyer HE, The "additional subunit" CF0II of the photosynthetic ATP-synthase and the thylakoid polypeptide, binding ferredoxin NADP reductase: are they different? Z Naturforsch [C] 1990, 45(7-8):772-784.

PAGE 257

236 296. Lill H, Steinemann D, Nelson N, Mutagenesis of the b'-subunit of Synechocystis sp. PCC 6803 ATP-synthase. Biochim Biophys Acta 1994, 1184(2-3):284-290. 297. Gardner JL, Cain BD, The a subunit ala-217 --> arg substitution affects catalytic activity of F1F0 ATP synthase. Arch Biochem Biophys 2000, 380(1):201-207.

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BIOGRAPHICAL SKETCH Tammy Weng Bohannon Grabar, daughter of William and Chu Shia Bohannon, was born in Altus, Oklahoma, on February 6, 1977. She lived in Okinawa, Japan, Patrick Air Force Base, Florida, Selfridge Air National Guard Base, Michigan, and Orange Park, Florida, before settling with her family in Melbourne, Florida. In her youth she participated in a variety of extracurricular activities and sports. She graduated in the top 1% of her class from Eau Gallie High School in June 1995. The following August she attended the University of Florida where she majored in microbiology. She received a Bachelor of Science degree with honors from the College of Agriculture in May 1999. During the summer of 1999 Tammy joined Dr. Brian Cains laboratory in the Department of Biochemistry and Molecular Biology at the University of Florida to gain her first experiences in scientific research. The following Fall semester she was accepted to the University of Florida graduate program in the Department of Biochemistry and Molecular Biology. She joined the laboratory of Dr. Brian Cain in February 2000 where she began the work described in this dissertation. During her graduate career Tammy married her high school sweetheart and boyfriend of 11 years, Charles Raymond Grabar, Jr., on May 11, 2002. Their daughter, Kaia Mae Grabar, was born on August 16, 2003. 237


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Title: Contributions of the Individual b Subunits to the Function of the Peripheral Stalk of F1F0 ATP Synthase
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Permanent Link: http://ufdc.ufl.edu/UFE0006623/00001

Material Information

Title: Contributions of the Individual b Subunits to the Function of the Peripheral Stalk of F1F0 ATP Synthase
Physical Description: Mixed Material
Copyright Date: 2008

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Source Institution: University of Florida
Holding Location: University of Florida
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CONTRIBUTIONS OF THE INDIVIDUAL b SUBUNITS TO THE FUNCTION OF
THE PERIPHERAL STALK OF F1Fo ATP SYNTHASE













By

TAMMY WENG BOHANNON GRABAR


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


2004
































Copyright 2004

by

Tammy Weng Bohannon Grabar


































This document is dedicated to my husband, Chuck, and my daughter, Kaia.
















ACKNOWLEDGMENTS

The work illustrated in this dissertation and my growth as a scientist could not have

been accomplished without the guidance, encouragement and support of several people

on both the professional and personal levels. The first person I would like to thank is my

mentor, Dr. Brian Cain. He allowed me to join his laboratory when I was fresh out of

college, even though I had no real experiences in a scientific lab. He exhibited extreme

patience while teaching me everything from how to hold and operate a pipette to pouring

agarose gels to cloning my own plasmids. His evident passion and excitement about

science opened my eyes to a whole new world of opportunity. Prior to joining his

laboratory, I had never even dreamed of joining graduate school and pursuing a PhD;

therefore, I feel extreme gratitude and consider myself very fortunate to have j oined his

lab. Once I joined the lab, Dr. Cain allowed me the freedom to make my own initial

scientific and experimental decisions, which was an excellent teaching method for me,

but he was always there for guidance and support whenever it was needed. I would also

like to thank him for being so involved in his lab. On any given day, I knew I could have

his undivided attention if I needed to consult with him. Over the years Dr. Cain has spent

a tremendous amount of time teaching me to think critically about scientific experiments,

how to communicate my data and ideas to others, how to give a professional scientific

presentation, and how to write scientific papers, and for those countless hours I thank

him.









I would also like to thank every person on my committee: Dr. Linda Bloom, Dr.

Art Edison and Dr. Dan Purich from my department, and Dr. Julie Maupin-Furlow from

the Microbiology and Cell Sciences Department. I have known Dr. Maupin-Furlow the

longest. She taught one of the most challenging courses I took as an undergraduate.

When an unexpected death occurred in my family, she was kind enough to allow me to

postpone an exam without questioning my motive, which was very unusual for most of

my professors while I was an undergraduate. Thanks to her, that was the only class in

which my grade was not affected that semester. I would also like to thank her for her

continual support in helping me get into graduate school and then subsequently taking the

time to hike across campus to join my committee meetings. I would like to thank Dr.

Purich for teaching me, in the middle of a physical biochemistry class, that sometimes we

have to take some time off to go outside and get some fresh air. That was always an

important lesson when endless hours in the lab led to careless mistakes. I also enjoyed

his sometimes unusual stories and adventures that he had to share with me when I was

spending entire days in the biochemistry library studying for exams. I would like to

thank Dr. Edison for his constant support and encouragement. He has always been the

first person who publicly and very kindly commended me after my journal club

presentation. I believe his kind words of support through the years helped me to gain the

courage I needed to believe in myself to really deliver a good presentation. I would also

like to thank him for his eagerness to understand every aspect about my proj ect. And

last, but not least, I would like to thank Dr. Bloom. As a woman in my department and a

new mother with a career in academia, she has been a wonderful role model. She has

always had kind words and smiles to bestow on me. I would also like to thank her for









assisting me during my committee meetings when discussions of fluorescence started to

go over my head.

I would also like to take the time to thank everyone that I had the pleasure of

working with in the lab. These are the people I spent countless hours with during the

course of the day and held many scientific and personal conversations with, and I am

happy to be able to call them my friends. I could not have spent the last five years with a

better group of people. Drs. Tammy Otto and Debra Zies were members of the lab when

I first j oined and were the ones who taught me the ways of the lab. Dr. Michelle Gumz

joined the graduate program and subsequently joined Dr. Cain's lab the same time as I. I

would like to thank Tammy, Debbie and Michelle for their scientific and personal support

as well as sharing with me memories of pool barbeques, wedding showers and baby

showers. Dr. Deepa Bhatt joined the lab as a postdoc during my graduate career. Her

friendship and scientific guidance have been very valuable to me. I would like to thank

her for giving me fantastic advice on all of my oral presentations and reviewing all of my

papers.

And finally, I would like to thank my family for all the encouragement, love and

support they have unwaveringly offered over the years. I would like to thank my dad for

always finding the positive in everything that was negative and always encouraging me to

overcome the many obstacles that graduate school hurled towards me. He never lost faith

in me, even when I was ready to give up. I would like to thank my mom for her

tremendous support as well. She spent a lot of time and energy stocking my refrigerator

and freezer full of meals when I found that I did not have the time to care for myself. She

has spent many weeks and weekends at my home since my daughter was born so that I










could have extra time to work on my dissertation. And last, but not least, I would like to

thank my husband for his constant emotional support and belief in me. He has been with

me through the thick and thin of graduate school and never once complained of my

emotional torment when things were not going my way. I could not have accomplished

this without him.





















TABLE OF CONTENTS

Page


ACKNOWLEDGMENT S .............. .................... iv


LI ST OF T ABLE S ........._..... .......... ............... xii..


LIST OF FIGURES ........._..... ..............xiii..._._. ......


ABBREVIATIONS .............. .................... xvi


AB S TRAC T ......_ ................. ............_........x


CHAPTER


1 BACKGROUND AND SIGNIFICANCE ................. ...............1.........._....


Introducti on ............... .... ... ....._..._ .. ... ......._ ..........1
Structure and Function of F1Fo ATP Synthase .............. ...............3.....
The Catalytic Core ................. ...............6........._. ....
The of hexamer ........._... ......___ ...............8....

The y subunit ................. ...............12................
The Rotor Stalk ................. ...............12........... ....

The y subunit ................. ...............14................
The a subunit ........._.. ........... ...............16....
The ring of c subunits ................. ....___ ....___ ...........2
T he Stator Stalk ................. ...............25..............
The a subunit ........._.. ........... ...............27....
The 6 subunit ........._.. ........... ...............33....
The b subunit ........._..._.._ ...._._. ...............37....
Subunit Equivalence ........._..._.._ ...............51.._.._._ .....
F1Fo ATP Synthase Mechanism............... ...............5
Proton Translocation: Driving Rotation .............. ...............57....
Coupling ................. ......... ......... .........6
Catalysis: The Binding Change Mechanism .............. ...............62....
Genetic Expression and Assembly .............. ...............63....
Summary ........._... ...... ._ ._ ...............65....










V111















2 INTEGRATION OF UNEQUAL LENGTH b SUBUNITS INTO F1Fo ATP
SYNTHASE .............. ...............67....


Introducti on ................. ...............67.................
Materials and Methods .............. ...............69....
M material s ................. ...............69.......... .....
Strains and Media ................. ...............70........... ....
Recombinant DNA Techniques............... ...............7
Mutagenesis and Strain Construction ................. ...............75................
Crude Preparative Procedures .............. ...............78....
Determination of Protein Concentration ................. ..............................80
Ni-Resin Purification..................... .... ........8

Assays of F1Fo ATP Synthase Activity .............. ...............82....
Immunoblot Analysis .............. ...............85....
R e sults............... ... .. .. .. ......_ .......... .............8

HA-Epitope Tagged b Subunits................. ................8
Construction and growth characteristics of mutants .............. ...................88
Effects of epitope tags ................ .. .......... .. ...............90.....
Expression of different b subunits in the same cell ........._._............_._.....91
Ni-Resin Purification............... ..............9

V5-Epitope Tagged b Subunits .............. ..........................9
Construction and growth characteristics of mutants .............. ...................97
Effects of epitope tags ................. ...............100...............
Detections of heterodimers. .................... .......... ......_ .............0
Formation of mixed length b subunits in F1Fo ATP synthase ..........................107
Discussion ............ .... __ ...............111..



3 GENETIC COMPLEMENT ATION BETWEEN MUTANT b SUBUNITS IN F1Fo
ATP SY NTHASE ................. ...............114......... ......


Introducti on ................. ...............114................
Materials and Methods ................. ...............116...............
M materials ................ ...............116................
Strains and Media ................. ...............117................
Recombinant DNA Techniques ................. ...............117................
Mutagenesis and Strain Construction ................. ................. ......... ...11
Preparative Procedures ................ ...............120................
Immunoblot Analysis .................... .... ..............12
Assays of F1Fo ATP Synthase Activity .............. ...............121....
Re sults................. .. .... ......... ..... ........ .. ..................2
Construction and Growth Characteristics of Mutants ................. ................. .121
Heterodimer Formation of barg36 Defective Subunits with bwt .......................... 123
Heterodimer formation of bAl534end-his Complemented with bwt-vs ................... 128
Heterodimer Formation of b+124-130-his Complemented with bwt-vs .................... 131












Mutual Complementation ...._._.... .........__........._ ............13
Discussion .........._...._ ......... ...............139.....



4 DEVELOPMENT OF CYSTEINE CHEMICAL MODIFICATIONS OF ALTERED
b SUB UNIT S ................. ................. 143........ .....


Introducti on ................. ...............143................
Materials and Methods .............. ...............146....
M materials ................ ...............146................
Strains and M edia ................. ...............146................
Recombinant DNA Techniques............... ..............14
Mutagenesis and Strain Construction ................. ............. ......... .......148
Crude Preparative Procedures .............. ...............152....
Assays of F1Fo ATP Synthase Activity .............. ...............152....
Immunoblot Analysis .............. ...............153....
Re sults................. .. .... ......... ..... ........ .. ..................5
Construction and Growth Characteristics of Mutants ................. ................. .153
Effects of Cysteine Mutations .............. .....................157
Discussion ................. ...............158................



5 MUTAGENISIS OF THE AMINO AND CARBOXYL TERMINI OF THE b
SUBUNIT INT F1Fo ATP SYNTHASE .............. ...............161....


Introducti on ................. ...............161................
M materials and M ethods .............. ...............165....
M materials ................ ...............165................
Strains and M edia ................. ...............165................
Recombinant DNA Techniques............... ..............16
Mutagenesis and Strain Construction ................. ............. ......... .......16
Crude Preparative Procedures .............. ...............168....
Assays of F1Fo ATP Synthase Activity .............. ...............169....
R e sults............... .. .............. ...............169......
Amino Terminal Mutations ............... .. .. .... ..... .. ............... 169....
Construction and growth characteristics of mutants .............. ................. 169
Effects of amino terminal mutations .............. .....................171
Carboxyl Terminal Mutations ............... ........ ..............17
Construction and growth characteristics of mutants .............. ................. 173
Effects of carboxyl terminal mutation............... ................175
Discussion ................. ...............176................



6 CONCLUSIONS AND FUTURE DIRECTIONS .............. .....................8


C onclusions............... .. .. ..... .. .. ... ...... .................18

Integration of Unequal Length b Subunits into F1Fo ATP Synthase ................. 181











Genetic Complementation between Mutant b Subunits in F1Fo ATP
synthase .............. ... ..... ._ _. .........._.... ... ..... .. .. .. ........ 8
Development of Cysteine Chemical Modifications of Altered b Subunits.......185
Mutagenesis of the Amino and Carboxyl Termini of the b subunit in F1Fo
ATP Synthase ................. ...............186................
Future Directions ................. .. ...... ....... ...............188......
Complementing Mutant b Subunits .................. .... .. ......... .. ...... ._._............8
Function of F1Fo ATP Synthase Incorporated with b Subunit Heterodimers....190
Positions of the Individual b Subunits in F1Fo ATP Synthase...........................190
Length of the Peripheral Stalk in F1Fo ATP Synthase Complexes
Incorporated with Shortened and Lengthened b Subunits ................... ..........191
Other Implications .............. ...............195....



APPENDIX

A MUTAGENIC OLIGONULCEOTIDES .............. ...............202....

B DEVELOPING PROTOCOL FOR PURIFYING F1Fo ATP SYNTHASE .............206

Purification of Enzyme Complexes Incorporated with b Subunit Heterodimers .....206
Culture ............ _...... .. ...............206...
Di sruption of Bacteria .............. ...............207....
Ni-Resin Purification................ .............20
V5-Epitope limmunoprecipitation............... ..........1
Detection of Purified Enzyme Complexes ........._.._ ..... ._ ...............2 10
Assays of F1Fo ATP Synthase Activity .........._.._ ......... .............. ....21 1


LIST OF REFERENCES .........._._ .. ...._.. ...............212.....

BIOGRAPHICAL SKETCH .............. ...............237....

















LIST OF TABLES


Table pg

1-1. F1Fo ATP synthase subunit equivalency............... ..............5

2-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity of
mutants expressing epitope tagged uncF(b) genes............... ...............90.

3-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity
of mutants expressing epitope tagged uncF(b) genes ................. .........___.......122

4-1. Description of uncF(b) cysteine mutations .............. ...............149....

4-2. Description of the unc operon cysteine mutations ............. ....................15

4-3. Aerobic growth properties and membrane-associated ATP hydrolysis activity
of mutants expressing cysteine. ................ .............. ......... ........ .....156

5-1. Aerobic growth properties and membrane-associated ATP hydrolysis activity
of mutants expressing uncF(b) mutations at the amino terminus. ................... ....... 170

5-2. Aerobic growth properties and membrane-associated ATP hydrolysis activity
of mutants expressing uncF(b) insertions or deletions throughout the b subunit ..175

6-1. Preliminary data of coexpressed mutant b subunits. ............. .....................190

A-1. Oligonucleotide sequences. ............. ...............203....

A-2. Oligonucleotide description. .............. ...............204....


















LIST OF FIGURES


Figure pg

1-1. Timeline of developing views of F1Fo ATP synthase............... ...............5

1-2. Space-filling structural model of Escherichia coli F1Fo ATP synthase............._.._. .....7

1-3. Structure of the a subunit of E coli F1Fo ATP synthase............... ...............19

1-4. Controversial models of the a subunit topology ....._.._.. .... ... .__. ........_.......29

1-5. Amino acid sequence of the E. coli F1Fo ATP synthase b subunit............_..._... ..........3 8

1-6. Gross structure of the E. coli F1Fo ATP synthase and the domains of the b
subunit ........._..._... ...............40.._.._.. .....

1-7. Model for F1Fo ATP synthase peripheral stalk orientation dependent upon the
direction of rotation during ATP synthesis or hydrolysis ..........._.._ ........._..__....47

1-8. Speculative models for the b-like subunits ........._..._... ........._..._...55......_. .

1-9. Model of proton translocation and torque generation in Fo........ .............. .59

1-10. The binding change mechanism .............. ...............63....

2-1. Oligonucleotides for epitope tags and mutagenesis of uncF(b) ................ ...............74

2-2. Construction of the single transcript expression system .............. ....................7

2-3. Histidine and HA-epitope-tagged b subunit expression system ............... ..............89

2-4. Western blot analysis s of hi stidine and HA-epitope tagged b subunits .....................92

2-5. Investigation of detergent solubilization of F1Fo ATP synthase complexes .............94

2-6. Ni-resin purification of F 1Fo, expressing different length b subunits, treated with
the cross-linker BS3 ........._.__ ..... ._._ ...............95...

2-7. Ni-resin purification of histidine and HA-epitope tagged F1Fo treated with the
cross-linker BS3 ................ ...............96........... ....

2-8. Histidine and V5-epitope-tagged b subunit expression system ........._...... ..............98











2-9. ATP-driven energization of membrane vesicles prepared from uncF(b) gene
m utants. ............. ...............101....

2-10. NADH-driven acidification of membrane vesicles prepared from uncF(b)
m utants ................. ...............102.............


2-11. Ni-resin purification ofF FoF ATP synthase treated with the cross-linker BS3.....106

2-12. Ni-resin purification ofF Fo ATP synthase expressing unequal length b
subunits ........._... ...... ..... ...............108....

2-13. Quantitation of b subunit heterodimeric F1Fo ................. ............................110

2-14. Interactions of b subunits of unequal lengths ...._.__... .... .._._.. ........_........112

3-1. Oligonucleotides for epitope tags and C-terminal truncation of uncF(b) ................1 19

3-2. Ni-resin purification ofF FoF ATP synthase incorporated with barg36 Subunit
mutations ....._. ................ ................. 124....

3-3. ATP-driven energization of membrane vesicles prepared from uncF(b) arg36
gene mutants ........... ..... .._ ...............127..

3-4. Ni-resin purification ofF FoF ATP synthase containing a b subunit carboxyl-
terminal truncation .............. ............... 129...

3-5. ATP-driven energization of membrane vesicles incorporated with F,Fo ATP
synthase containing a b subunit carboxyl-terminal truncation................ .............13

3-6. Ni-resin purification of membranes incorporated with b+124-130-his subunit
mutati on ................. ...............132................

3-7. ATP-driven energization of membrane vesicles incorporated with a defective
b+124-130 Subunit mutation ................. ...............133................

3-8. Ni-resin purification ofF FoF ATP synthase incorporated with complementing
defective b subunits ........._._. ._......_.. ...............136....

3-9. ATP-driven energization of membrane vesicles incorporated with F Fo ATP
synthase containing complementing defective b subunits .............. ...................137

3-10. Interactions of defective b subunit with wild type b subunits found in intact
F1Fo ATP synthase complexes .............. ...............140....

3-1 1. Model of F1Fo ATP synthase incorporated with complementing defective b
subunits .............. ...............142....











4-1i. Model of E. coli F 1Fo ATP synthase ................. ...............144...........

4-2. Oligonucleotides for cysteine mutagenesis of the unc operon .............. .... ........._...150

4-3. Expression plasmid of cysteine mutants ................. ...............155........... ..

4-4. Western blot analysis of cysteine mutant b subunits of differing length................. 158

4-5. Model of F1Fo ATP synthase with cysteine substitutions in the b and 6 subunits ..159

5-1 Amino acid sequence and domains of the E. coli b subunit ................. ........_._.....162

5-2. Oligonucleotides for mutagenesis at the amino and carboxyl termini in the unc
operon ................ ...............167.............

5-3. ATP-driven energization of membrane vesicles prepared from b subunit
membrane domain mutants .............. ...............172....

5-4. Amino acid insertion and deletion analysis of the E. coli b subunit ................... ..... 174

5-5. Mutations constructed throughout the b subunit ........._._. ......___ ................177

6-1. Design of FRET experiments to measure the peripheral stalk ..........._..._ ...............193

6-2. Model of rotation inhibition due to a fusion protein on the a subunit ................... .. 195

6-3. Sequence alignments of subunits b and b from various species with the b
subunit of E coli .............. ...............200....

B-1. Diagram of purification procedures for homogeneous heterodimeric bvs/bhis
F1Fo ATP synthase complexes .............. ...............208....















ABBREVIATIONS

ACMA, 9 -ami no-6 -chl oro-2 -methoxy acri di ne

ADP, adenosine-5' -diphosphate

ala, alanine

AO, tegamineoxide WS-35

Ap, ampicillin

Apr, ampicillin resistant

asn, asparagine

ADP, adenosine-5' -diphosphate

ATP, adenosine-5' -triphosphate

b+7-his, SOVen amino acid insertion in the b subunit with a 6X histidine epitope tag at the

amino terminus

ba7-vs, seven amino acid deletion in the b subunit with a V5 epitope tag at the carboxyl

terminus

bser84 cys, Substitution of a cysteine for serine at amino acid position 84 in the b subunit

P-ME, P-mercaptoethanol

bp, base pair

BS3, bis(3 -sulfo-N-hydroxysuccinimide ester)

BCA, bicinchoninic acid

BSA, bovine serum albumin

Cm, chloramphenicol










Cmr, chloramphenicol resistance

cys, cystemne

DACM, N-(7-dimethylamino-4-methyl-coumarinyl)-maemd

DCCD, dicyclohexylcarbodiimide

Su2z, the second co-helix in the epsilon (s) subunit

ECD, 1-ethyl-3 [3 -dimethylamino]propyl carbodiimide

ECL, enhanced electrochemiluminescence

EDTA, ethylenediaminetetraacetic acid

FPLC, fast polynucleotide liquid chromatography

FRET, fluorescence resonance energy transfer

g, gravitational force

gln, glutamine

glu, glutamate

GFP, green fluorescent protein
HA, peptide epitope of hemagglutinin protein of human influenza virus

ICBR, Interdisciplinary Center for Biotechnology Research

IPTG, isopropyl-1 -thio-P-D-galactopyranoside

kb, kilobase

kD, kilodalton

LB, Luria Bertani medium

LBG, Luria Bertani media supplemented with 0.2% glucose

LDAO, lauryldimethylamine oxide

LSB, Laemmli sample buffer

mG, milligram










mL, milliliter

MOPS, 3- [N-morpholino]propanesulfonic acid

NADH, P-nicotineamide adenine dinucleotide, reduced form

NFDM, nonfat dry milk

Ni-CAM, high capacity nickel chelate affinity matrix

NMR, nuclear magnetic resonance spectroscopy

PAGE, polyacrylamide gel electrophoresis

PBS, phosphate-buffered saline

PBST, phosphate-buffered saline supplemented with 0.1% tween20

PCR, polymerase chain reaction

Pi, inorganic phosphate

P/O, number of ATP's made per 2 e- transferred to oxygen

PVDF, polyvinylidene fluoride

rms, root mean square

ser, serine

SDS, sodium dodecyl sulfate lauryll sulfate)

TID, 3 -(trifluoromethyl)-3 -(m- [125I]i Odophenyl)di azirine

TBS, tris-buffered saline

thr, threonine

TTBS, tris-buffered saline supplemented with 0. 1% tween20

TD, taurodeoxycholate

TE, tris[hydoxymethyl]aminomethane, ethylenediaminetetraacetic acid buffer, pH 8.0

Thl, tri s[hy doxym ethyl iami nom ethane, magne sium sulfate buffer, pH 7.5


XV111










Tm, melting temperature of double stranded DNA

Tris, tris[hydoxymethyl]aminomethane

Gig, microgram

CIL, microliter

V5, epitope found in the P and V proteins of the paramyxovirus, SV5

v/v, volume/volume

wt, wild type

w/v, weight/volume
















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

CONTRIBUTIONS OF THE INDIVIDUAL b SUBUNITS TO THE FUNCTION OF
THE PERIPHERAL STALK OF F1Fo ATP SYNTHASE

By

Tammy Weng Bohannon Grabar

August 2004

Chair: Brian D. Cain
Major Department: Biochemistry and Molecular Biology

The universal molecule of biological energetic is adenosine triphosphate (ATP),

and the enzyme involved in providing the maj ority of cellular ATP is F1Fo ATP synthase.

Enzymes in this family utilize the electrochemical gradient of protons across membranes

to synthesize ATP from ADP and inorganic phosphate in a coupled reaction. The

cytoplasmic Fl and the membrane-bound Fo sectors are linked by two stalk structures, the

rotor stalk and the peripheral stalk. Proton conduction through the Fo sector drives the

rotation of the rotor stalk within the catalytic core, which is held steadfast by the

peripheral stalk. In Escherichia coli, the 6 subunit of F1 and a parallel homodimer of

identical b subunits constitute the peripheral stalk of F1Fo ATP synthase. Work

accomplished in this dissertation indicates that the bacterial enzyme does not require two

identical b subunits to form the dimer. Two different length b subunits, with a size

difference of at least 14 amino acids, were capable of forming the b dimer of an intact

F1Fo ATP synthase complex. Also, in work presented in this dissertation, a defective









mutation in one region of the b subunit was overcome by dimer formation with a second

b subunit that contained defective mutation in a different region but had a wild-type

sequence in the region of the former defective b subunit. This mutual complementation

between fully defective b subunits indicated that each of the two b subunits makes a

unique contribution to the functions of the peripheral stalk, such that one mutant b

subunit is making up for what the other is lacking. Interestingly, the equivalent of the

bacterial b subunit in plants exists as two genetically different subunits, and the mammal

counterpart exists as at least four subunits. This work suggests that the individual

functions of the b subunits may be reflected in the fact that higher organisms evolved to

encode multiple b-type subunits.















CHAPTER 1
BACKGROUND AND SIGNIFICANCE

Introduction

The premiere of Peter Mitchell's chemiosmotic theory in 1961 eventually resulted

in the major breakthrough of the characterization of F1Fo ATP synthases. Basically, his

theory stated that protons are pumped across energy transducing membranes, thereby

creating an electrochemical gradient of protons (1). This proton gradient, also known as

the proton-motive force, consists of two components: i) a chemical component due to the

concentration gradient of protons and ii) an electrical component, or membrane potential,

due to the positive charge of the protons (H ). As a result, one side of the membrane is

more positive than the other. The potential energy of this gradient can then be transduced

to chemical energy or utilized to perform work when the protons diffuse back across the

membrane from the higher to the lower potential (2). The protons can diffuse across the

membrane through specific transmembrane proton conductors, which can synthesize

adenosine 5'-triphosphate (ATP) or co-transport solutes, and in the case of bacteria drive

flagellar rotation.

The ability to consume nutrients and convert them to energy is required of all living

organisms, from microscopic bacteria to plants to humans. The universal molecule of

biological energetic is ATP, and in almost all organisms, the central enzyme involved in

providing the maj ority of cellular ATP is F1Fo ATP synthase (3-6). F1Fo ATP synthases

are responsible for the production of ATP in the final step of processes called oxidative

phosphorylation and photophosphorylation. They provide the bulk of cellular energy in









the maj ority of eukaryotes and prokaryotes. The synthesis of ATP occurs at a rate of

about 100 s- which maintains a concentration of about 3 mM ATP in Escherichia coli

and greater concentrations in mitochondria and chloroplasts with no noticeable product

inhibition (3). In eukaryotes, they are located in the inner mitochondrial membrane, or in

the thylakoid membrane of chloroplasts. In most bacteria, F1Fo ATP synthase is located

in the cytoplasmic membrane. Enzymes in this family utilize the electrochemical

gradient of protons across these membranes in order to synthesize ATP from ADP and

inorganic phosphate (Pi) in a coupled reaction. In bacteria, the reaction of ATP synthases

can be reversed if the situation of a dissipated electrochemical proton gradient arises. In

this case, ATP derived from glycolysis can be hydrolyzed in order to pump protons

across the membrane, creating a membrane potential. The membrane potential can then

be utilized to drive other cellular processes such as the extrusion of sodium ions, nutrient

uptake and flagellar rotation.

An explosion of research concerning F1Fo ATP synthase has occurred during the

past few decades. In particular, a great deal of knowledge of the enzyme has been solved

only in the past decade. A plethora of relatively recent reviews concerning every aspect

of F1Fo ATP synthase can be found in the special editions of Journal ofBioenergetics

and Biomembranes (volume 32, 2000) and Biochimica et Biophysica Acta (volume 1458,

2000) as well as reviews authored by Noji and Yoshida (2001), Senior et al. (2002),

Capaldi et al. (2002) and Weber and Senior (2004). This chapter will provide detailed

explanations of what is currently known about F1Fo ATP synthase including the

mechanism of the enzyme as a whole as well as structure and functions of individual

subunits, equivalence of the bacterial enzyme to its eukaryotic equivalents and genetic










expression and assembly. The research presented in this dissertation primarily concerns

the b subunit of the Escherichia coli (E. coli) F1Fo ATP synthase. Hence the b subunit

will be discussed extensively later in this chapter.

Structure and Function of F1Fo ATP Synthase

The structure and function of F1Fo ATP synthases are remarkably similar from

bacteria to humans. In E. coli, the simplest form of the enzyme, F1Fo ATP synthase is a

complex enzyme composed of twenty-two polypeptides of eight different types with the

stoichiometry of u3 38ysab2C10 (Figure 1-2) (6, 7). The deduced molecular size is

approximately 530 kDa. The structure of F1Fo ATP synthase in chloroplasts is very

similar with the exception that there are two isoforms of the b subunit. On the other

hand, the mitochondrial enzyme is more complex, including an extra 7-9 small subunits

which are thought to have roles in enzyme regulation (8-10). Discussion of F1Fo ATP

synthase is commonly divided into two portions, Fl and Fo. The Fl portion of the enzyme

is composed of the cytoplasmic subunits, a3 3ysS, and is responsible for the synthesis of

ATP. The Fo portion consists of the membrane-bound subunits, ab2C10, and is responsible

for the translocation of protons through the membrane. New insights concerning the

functions and the intersubunit contacts have refined the way F1Fo ATP synthase is

perceived, dividing discussions of the enzyme into the catalytic core, the rotor (central)

stalk and the stator (peripheral) stalk (3, 11-14). The catalytic core consists primarily of

the u3 37 subunits, the rotor stalk consists of the ysclo subunits and the stator stalk

consists of the b26 Subunits.

Over the years, F1Fo ATP synthase has received worldwide recognition as the

tiniest rotary motor known to mankind (15, 16). Protons passing through the enzyme










complex drive the rotation of the rotor ysclo subunits at about 100 Hz. This rotation,

which is absolutely essential for the machinery of the enzyme, transmits energy over a

distance greater than 100 A+ by providing the means by which conformational changes in

the Fl catalytic core, u3 3, take place for the synthesis of ATP (3, 4).

Structural studies of F1Fo ATP synthase commenced in the early 1960's and persist

to this day in pursuit of a complete high-resolution structure. Negative staining

procedures in the early 1960's initially revealed the traditional tripartite features of the

enzyme complex from sub-mitochondrial particles, consisting of what was referred to as

the headpiece, stalk and basepiece (Figure 1-1) (17). Ten years later, the first electron

micrograph of a detergent-solubilized F1Fo ATP synthase was published, confirming the

existing idea of a tripartite molecule (18). Appreciably, electron microscopy (EM) in

combination with other biochemical data of isolated Fl exposed a hexagonal arrangement

of alternating subunits with a seventh mass found in the center of the array (Figure 1-1)

(7, 19). Based on this premise it was first suggested that Fl consisted of an alternating

hexagonal array of three a and three P subunits with the y, 6 and a situated centrally (7).

The idea did not gain favorable recognition for some twenty years until verified by x-ray

crystallography (20, 21). Continuous improvements in EM technology led to numerous

publications of various ATP synthases which defined the average overall dimensions of

about 190 A+ from top to bottom and about 37 A+ assigned to the stalk structure (22-25).

Using a combination of traditional biochemical, molecular biological and immunological

techniques along with EM, many important discoveries were made that led to what we

now understand of F1Fo ATP synthase. The first direct evidence for rotation of the stalk

appeared in 1990 (24), but was not followed by the visualization of the peripheral stalk,









Trip artite
features


Evidence of
rotatio n


First X-ray
crystallography
of rat F, at
9A


1974


1982


1990


Peripheral
stalk


---+


1994


1999


Figure 1-1. Timeline of developing views of F1Fo ATP synthase. Electron microscopy
and biochemical analysis from the early 1970's through the 1980's allowed
visualization of the classical tripartite features of F1Fo ATP synthase
consisting of what was referred to as the headpiece, stalk and basepiece.
Furthermore, the arrangement of the F1 subunits were first proposed in 1974,
though it did not gain favor until high resolution structure was obtained
twenty years later. The first direct evidence for rotational catalysis appeared
in 1990. Improved EM techniques showed the existence of a peripheral stalk,
assigned the role of the "stator" to hold the F1 sector in place against the
proposed rotation, and a "cap" in the late 1990's. In 1994, the first high-
resolution structure (2.8 A+) of F1 appeared, consisting of the OC3 3 hexamer
with partial structure of the y subunit. Currently there is no high resolution
structure for the entire F1Fo ATP synthase complex though many of the
subunits have been solved individually or in part by model polypeptides.


Proposecl
F, moiety


1970's









First high-
resolution
structure of
bovine F


at 2.8 A


1998


2004









assigned the task of the "stator" to hold Fl in place against the rotation of the centrally

located stalk, until many years later (26-29).

Today there is still no high-resolution structure of the entire F1Fo ATP synthase

enzyme complex from any organism. X-ray crystallographic and NMR data of partial

complex structures from rat, bovine, yeast and E. coli or model polypeptides deduced

from nucleotide sequences have accumulated over the past twenty years to allow for a

composite structural model with both high- and low-resolution structures (Figure 1-2).

Currently, complete high-resolution structures are available for the a, P, y, E and c

subunits and partial structures for the 6 and b subunit. There is currently no high-

resolution data for the membrane-integral a subunit.

The Catalytic Core

The first high-resolution structure, resolved to 2.85 A+, consisted of a3 3Y Of Fl

prepared from bovine heart under inhibited conditions and in the absence of Pi and

substoichiometric amounts of ADP (20). This major breakthrough was shortly followed

by a 2.80 A+ Fl isolated from rat liver in the absence of the physiological cation, Mg2+

(21). The arrangement of the subunits in the two structures obtained were exceptionally

similar and confirmed Catteral and Pedersen's proposal made two decades prior by

showing the three a and three P subunits arranged alternatively with the amino and

carboxyl-termini of the y subunit, each forming an a-helix, extending up through the

center of the hexamer (Figure 1-2) The only difference in the structures, which was the

occupancy of the three nucleotide binding sites located at the up interfaces, was likely

due to the difference in preparation conditions (7) or crystal quality. In the former,















SF1





rotor
stalk


Figure 1-2. Space-filling structural model ofEscherichia coli F1Fo ATP synthase. The
model is based on a composite of high and low resolution structures taken
from E. coli, yeast and bovine F1Fo ATP synthases. F1Fo subunits included in
the model were u3 38ysab2C10. The subunits were color coded as follows: a,
red; p, green; y, cyan; s, orange; 6, yellow; b, blue; a, brighter yellow; c,
darker blue. The direction of the arrow indicates the direction of rotation of
ysclo during ATP synthesis. The yellow and blue cylinders represent the a
subunit and portions of the b subunit that currently have no high-resolution
structures from any species.


catallytic
b_ core


peripheral
stalk









one of the catalytic sites was empty and in the latter, all three active sites were occupied

with nucleotides (20, 21). Since the occupancy state of the three catalytic sites has been

of considerable debate, a more accurate depiction of the Fl moiety, which would give

some insight about the mechanism of ATP synthesis, must be attained from crystals

obtained under physiological conditions. A more accurate depiction of the structures and

roles of each individual subunits of the Fl ATPase follows.

The up hexamer

Homology. The a subunit of the E. coli F1Fo ATP synthase, product of the uncA

gene, is the largest subunit consisting of 513 amino acids with a deduced molecular

weight of 55,3 13 Da. The P subunit, a product of the uncD gene, is a 459 amino acid

subunit, with a molecular weight of 50,325 Da. Based on the primary sequences, the a

and p subunits of E coli Fl have the most obvious homologies in the chlororplast and

mitochondrial enzymes (30). The highest conserved subunit from the E. coli F1Fo ATP

synthase is the p subunit with approximately 70% homology with the chloroplast and

mitochondria equivalents (31). The a subunits exhibit roughly 50% homology (31). A

total of 6 nucleotide binding sites are housed at the up interfaces, three catalytic

contributed primarily by the P subunit and three noncatalytic housed primarily by the a

subunit (32, 33). The nucleotide binding regions have sequence homologies with other

proteins that bind nucleotide or phosphate, including secA protein, N-ethylmaleimide

sensitive fusion protein, herpes simplex virus UL15, Ca2+-ATPase, H /K+ ATPase and

Na /K+ ATPase (34-37). Furthermore, the nucleotide binding motif, GXXXXGKT/S,

known as the Walker A motif, which was first identified in the a and P sequences ofF1,

has been found to be conserved in the high-resolution structures of other proteins









including p21'", adenylate kinase, RecA, elongation factor Tu, and transducin-a (20, 38-

42).

Tertiary structure. The first high-resolution structure of bovine Fl resolved at the

atomic level (2.8 A+) was solved by Walker's group a decade ago (20). It was found to be

a flattened sphere approximately 80 A+ high and 100 A+ wide with the three a and three P

subunits arranged as a hexamer of alternating subunits around a centrally located 90 A~

long a-helix formed by the y subunit. A dimple 15 A+ deep is located at the top of F1.

The amino-terminal regions of the a and P subunits were once thought to be in close

proximity to the membrane due to labeling experiments (43). Contrary to this early data,

the crystallographic data placed the amino-terminal regions on the top of the u3 3

hexamer over 100 A+ away from the lipid bilayer.

The folds of the a and P subunits were found to be nearly identical. They each

consisted of a six-stranded P-barrel at the amino terminus (al9-95, P9-82), ai central a-p

domain containing the nucleotide-binding site (u96-379, P83-363) and a bundle of seven and

six helices at the carboxyl termini of the a and P subunits, respectively (u380-510, P364-474)

(20). The nucleotide binding domain consisted of a nine stranded P-sheet with nine

associated a-helices, of which the a-carbons of the seven P-strands and the seven

associated helices can be superimposed onto the RecA protein ATP binding site with an

rms separation of 1.9 A+ (20).

The three catalytic sites were located at the interfaces of the u3 3 hexamer. In the

original crystal structure, now commonly referred to as the reference structure, two of the

three sites were occupied by nucleotide, containing MgADP ("(PDP Site") and MgAMP-

PN~P ("(PTP Site"). The third site was empty and designated "(PE." The PDP and PTP









subunits were in similar, closed conformations whereas the PE adopted an open

conformation, differing from the other two by a large hinge motion of the carboxyl

terminal domain of greater that 20 A+. Subsequently, several high-resolution structures of

crystals obtained under various nucleotide conditions gave the same overall structure of

bovine Fl with two nucleotides bound ("two nucleotide structures") (44-49). The PTP Site

was found to occasionally contain a diphosphate nucleotide, establishing that there is no

requirement for PTP to be occupied by a nucleotide triphoshate to produce a

conformational change (48). A more recent structure of bovine Fl solved by the Walker

group at 2.0 A+ showed all three catalytic sites bound by nucleotide (50). Both the PTP

and PDP Sites contained MgADP, adopting the closed conformation, whereas the site

corresponding to the PE Site in previous structures contained MgADP+Pi and adopted a

half-closed conformation. It is thought that the PDP Site is actually the catalytic site. The

structure of rat liver Fl was solved (2.8 A+) in the presence of physiological concentrations

of nucleotides but in the absence of the physiological cation, Mg2+. In this structure, all

three nucleotide binding sites adopted strikingly similar conformations, analogous to the

PDP and PTP Of the previously reported structure of the bovine Fl. This structure had no

indication of the open conformation and showed the presence of nucleotide in all three

sites (21). The structure of an OE3s 3COmplex from a thermophilic bacterium was solved

in the absence of nucleotides and exhibited all three P subunits in the open conformation,

suggesting there is a correlation between the open conformation and the absence of

nucleotide (5 1). A low resolution crystal structure (4.4 A+) of the E. coli Fl has been

obtained by Capaldi's group, in which the catalytic sites are thought to be very similar to

that of the bovine structure; however, the occupancy state of the nucleotide binding sites









was unclear (52). The frequent reports of "two nucleotide" structures have bewildered

scientists due to the vast body of biochemical data from numerous laboratories, using a

variety of techniques, which establish indisputably that all three catalytic sites are readily

filled with nucleotide (32, 53). It is possible that the enzyme preferentially crystallizes in

a ground state intermediate which may occur after the release of product, leaving one site

empty and opened (54). Nevertheless, the accumulating structural data may be indicative

of several intermediary steps that may form during the synthesis of ATP. A detailed

account of the mechanism of ATP synthesis follows later in this chapter.

The crystal structure does offer some insight of the chemical mechanism of ATP

synthesis (20). In the P subunit, 4.4 A+ from the terminal phosphate of the bound

nucleotide triphosphate, there is clearly a density for a water molecule hydrogen bonded

to the carboxylate of PglU1SS. This carboxylate is positioned to allow an inline

nucleophilic attack of the water molecule on the terminal phosphate. The guanidinium of

a neighboring residue, aarg373, iS thought to help stabilize the negative charge on the

terminal phosphate during the transition state (20). This same arrangement can be found

in the catalytic site of transducin-a (42). The crystal structure also provides some insight

as to why the nucleotide binding sites in the a subunit are noncatalytic. There is no

spacial equivalent of the carboxylate of PglU1SS in the a subunit. The spatial equivalence

in the a subunit is filled by a agln208, with the side chain pointed away from the terminal

phosphate (20). The binding of the adenine to the noncatalytic site of the a subunit is

highly specific, unlike the P subunit nucleotide binding site, which can accommodate

GTP, ITP as well as ATP (55, 56). This specificity is due to several hydrogen bonds as

well as the presence of the Ptyr368 ClOse to the 2-position of the adenine ring in the a










subunit, while in the P subunit the adenine is in contact with a hydrophobic interface

(20). Though the binding sites in the a subunits are highly specific, the roles remain

obscure.

The y subunit

The y subunit plays an important role in the catalytic core. Interactions between the

amino- and carboxyl-terminal a-helices and the u3~ 3Subunits are responsible for the

conformational changes that result in ATP catalysis. The y subunit is a fundamental part

of the rotor stalk.

The Rotor Stalk

Two narrow stalks, a centrally located stalk and a peripheral stalk, have been

observed to link the catalytic core of F1 and the membrane-bound proton translocating Fo

with about of 40-45 A+ in between (27). The central stalk came into view three decades

ago via EM and has since become widely referred to as the rotor stalk. The rotor stalk

consists of the y and a subunits. The bottom of the rotor stalk is connected firmly to the

Fo ring of c subunits located in the membrane and the top extends 90 A+ within the u3 3

hexamer of F1 where it forms crucial interactions with both the a and P subunits (57-59).

F1Fo ATP synthase is an extraordinary enzyme due to its ability to couple potential

energy, obtained from proton translocation through Fo in the membrane, to the synthesis

of chemical energy, over 100 A+ away in Fl, by a rapid rotation of subunits. Although

predicted by Boyer in the 1970's, evidence of rotation did not appear until the early

1990's. The X-ray crystal structure solved by Walker's group suggested that the y

subunit was the rotating subunit by suggesting it could distribute itself to all three P

subunits as opposed to just one (20). Consistent with this idea was inhibition of the Fl









complex by crosslinking the y subunit to one of the a or P subunits (60, 61) and recovery

after photobleaching experiments (62, 63). More convincing evidence was provided

when Duncan et al. crosslinked the y subunit to an unlabeled P subunit by disulfide bond

and then mixed the y-P complex with 35S-labeled P subunit (along with the a, 6 and a

subunits). When the disulfide bond was broken and ATP was added, the y subunit was

observed to switch from labeled to unlabeled P subunit (64). Finally, direct evidence was

achieved in single molecule experiments by attachment of a fluorescent actin filament to

the y subunit and observance of unidirectional rotation of the actin filament upon addition

of ATP (15). Direct observation of the rotating y subunit was soon followed by

observance of the rotation of the s and c subunits at the same speed and direction,

indicating that these three subunits rotate in synchrony, forming the central rotary

machinery of the enzyme complex (65-67). Until very recently, rotation has only been

observed in the direction of ATP hydrolysis. Direct evidence for the synthesis of ATP by

Fl has been shown by attaching a magnetic bead to the y subunit ofF1 fixed to a glass

surface and the rotating the bead, in the appropriate direction, using electrical magnets

(68).

The first structural information obtained for the a subunit ofE coli was

accomplished by nuclear magnetic resonance (NMR) studies (69) and is good agreement

with the crystal structure solved at 2.3 A (70). In all previous crystal structures of the

rotor stalk, the portion of the rotor stalk' s y subunit protruding from the Fl ap hexamer

and the a subunit were disordered. Recently, the structure of the bovine homologs of the

rotor stalk y and a subunits has been solved and refined to 2.4 A+ (48). The structures of

the E. coli y and a subunits are remarkably similar with that from bovine Fl. When









comparing the structures of the y and a rotor stalk obtained under different conditions or

from different sources, in combination with an overwhelming amount of biochemical and

immunological evidence, it is clear that the domains of the two subunits undergo maj or

shifts in position, which reflects its fundamental role in the synthesis of ATP (20, 44, 48,

71-80).

The y subunit

In E. coli, the y subunit is the third largest subunit of F1Fo ATP synthase, encoded

by the uncG gene as a 286 residue polypeptide with a deduced molecular weight of

3 1,563 Da. It plays an essential role in coupling proton transport to the synthesis of ATP.

The first visualization of a portion of the y subunit was a bovine Fl partial structure

solved in combination along with the u3 3 hexamer revealing three a-helices (20). The

Y209-272 (TOSidues 223-286 in the E. coli sequence) carboxyl terminus formed a long (90 A+)

a-helix extending from the stalk structure seen by EM to about 15 A+ from the top of the

hexamer. The bottom half of this helix formed a left-handed anti-parallel coiled coil with

a shorter a-helix composed of the amino-terminal residues Yl-45 (20). The two helices

protruded about 30 A+ from the bottom of F1. An approximately 200 kink in the latter

helix was produced by ypro40 and a similar but less pronounced kink was induced by Yleu217

in the former (48). A third, much smaller a-helix, composed of y73-90 (TOSidues 83-99 in

the E. coli sequence) was inclined at about 45 degrees from the larger helices and located

directly under the Fl hexamer.

More recently, the complete structure of the bovine rotor stalk has been solved to

2.4 A+ (48). The overall length of the stalk, from the carboxyl terminus of the y subunit to

the very bottom where it contacts the ring of c subunits, was 1 14 A+. The portion that









protrudes from the u3 3 hexamer, i.e. the part seen in electron micrographs, was 47 A~

long and 54 A+ wide at its largest cross-section. A completely new a/0 domain,

consisting of a five-strand P-sheet (1-5) and six a-helices (a-f), was identified in the

complete structure of the y subunit. Helices a and f extended into the u3 3 hexamer to

form the antiparallel coiled coil discussed previously. Strands 1-3 along with helices b

and c formed a Rossman fold that forms extensive interactions with the a subunit as well

as the ring of c subunits in the membrane (discussed below). This fold was linked to a P-

hairpin, formed by strands 4 and 5, by helix d. Overall, this a/0 domain had a globular,

oval shape with the dimensions 5 1 A+ wide by 41 A+ high. The positioning of the a/p

domain at the base of the rotor stalk may provide stability to the structure of the rotor

stalk during rotational catalysis (48).

A low-resolution crystal structure of the E. coli F1Fo ATP synthase y subunit was

solved to 4.4 A (52). Upon comparison with the high-resolution bovine structure

obtained one year later, a few differences were observed (48). Helices a and f (see

above) were extended by and extra 12 and 20 residues, respectively. Four additional

putative a-helices, designated B and D-F, were found in the E. coli structure and had

little agreement with the bovine y subunit structure. E. coli helix B runs parallel with the

bovine P-strand 5 and may correspond to it. Helix D has no apparent equivalent in the

bovine model and helices E and F appear to overlap with regions in the bovine 6

(bacterial s, see Table 1-1) and a (no equivalent in bacteria) subunits. The remainder of

the E. coli structure appeared similar to the bovine structure.

The crystal structure displays a strikingly asymmetrical Fl due to differences in the

domains of the a and p subunits and the interactions formed with the single y subunit










(20). The obvious asymmetric positioning of the coiled coil of the y subunit is a key

feature to the mechanics of the binding change mechanism of F1Fo ATP synthase. Its

large carboxyl terminus a-helix passes through a hydrophobic sleeve formed by six

proline-rich loops of the a and P subunits, undoubtingly resulting in the conformational

changes occurring in the catalytic sites (20). In the PE Subunit (see above), several

hydrogen bonds are formed with the y subunit, which forms a "catch", resulting in

conformational changes. Specifically, residues Yarg254 and ygln255 in the carboxyl terminal

helix form hydrogen bongs with PE-asp317, PE-thr318 and PE-asp319. Also, a second "catch" is

formed between the carboxyl terminal domain of the PT Subunit and the short helix of the

y subunit. Hydrogen bonds formed between Yylss, Yys,90 and Yalnso with PT-asp394 and PT-

glu398. This sequence of the P subunit, DELSEED (P394- 400), iS a portion of the binding

site of amphipathic cationic inhibitors and putatively the ATPase inhibitor protein (81-

83). Recently, mutations of residues involved in the "catch" loops were shown in inhibit

ATP hydrolysis activity by the soluble F1-ATPase (84). Structural information suggests

the two antiparallel coiled coil a-helices of the y subunit may unwind during rotational

catalysis and the a subunit rotates around the Fl axis while undertaking a net translation

of about 23 A+ (85). It is likely that these gross changes observed in the structures

revealed individual functional states of the enzyme complex during catalysis.

The E subunit

The a subunit ofE. coli F1Fo ATP synthase consists of 138 amino acids with a

molecular weight of 15, 068 Da and is encoded by the uncC gene. The a subunit has

several putative functions in the F1Fo ATP synthase complex including structural,

inhibitory and coupling roles. Structurally, the binding of F1 to Fo has long been known









to require the presence of the a subunit, which had implicated it as part of stalk structure

(86). In isolated Fl, and in isolated F1Fo to a lesser extent, it has been shown to inhibit

ATP hydrolysis activity (87-89). The removal of a from isolated Fl resulted in up to a

10-fold increase in ATP hydrolysis activity. Furthermore, a truncated version of the E

subunit lost all inhibitory functions but still promoted binding of F1 to Fo, hence the

inhibitory feature has been assigned to the extreme carboxyl terminus (90). It was

speculated that it acts as an inhibitor by reducing the rate at which the product is released

from the catalytic site (91). Also, the a subunit, as part of the rotor stalk, plays a role in

the coupling of proton translocation to the catalytic site. The a subunit' s diversity of

functions is supported by the findings that it produces several points of interactions with

the oc (61, 74, 92), P (61, 88, 89, 93, 94) and y (74, 95-97) subunits of F1 and the c (98,

99) subunits of Fo.

An innovative set of experiments conducted by members of the Dunn laboratory

made use of different sized fluorescent proteins, ranging from the 12 kDa cytochrome

b562 prOtein to the 30 kDa flavodoxin reductase protein with a 20 residue linker (100).

The proteins were fused to the carboxyl-terminus of the a subunit. Since the a subunit is

part of the rotor stalk, according to the concept of rotational catalysis, the fusion of a

large protein at this site should sterically hinder rotation due to the presence of the

peripheral stalk. Cells expressing the smaller cytochrome b562 prOtein fused to the E

subunit grew on minimal media, indicating a functional F1Fo ATP synthase complex.

However, cells expressing the larger flavodoxin reductase protein fused to the a subunit,

though found in an intact enzyme, failed to grow. These results provided the first

evidence, in vivo, supporting rotational catalysis.










High-resolution structural data of the E. cobi s subunit was first solved by NMR

followed by the X-ray crystal structures of the isolated E and completed ye subunits (69,

70, 101). The isolated a subunit consisted of an 84 residue amino-terminal P-sandwich

domain and a 48 residue carboxyl-terminal helix-turn-helix domain in which the two a-

helices formed an antiparallel hairpin (70). The P-sandwich consisted of two five-

stranded P-sheets folded as a rigid, flattened P-barrel. The structure of the isolated a

subunit from E. cobi was very similar to the Fl complex isolated from bovine, which

included the u3 378 subunits (48, 70). Superimposition of 127 of the amino acid Ca

resulted in an rms deviation of 1.6 A+. On the other hand, in the E. cobi y-s complex,

resolved to 2. 1 A+, the a subunit assumed a strikingly different conformation, in which the

two a-helices of the antiparallel hairpin at the carboxyl-terminus are wide apart and

wrapped around the y subunit (Figure 1-3A) (101). Subsequently, both conformations of

the a subunit have been trapped in E. cobi F1Fo ATP synthase by crosslinking

experiments, confirming the existence of both in an intact enzyme complex (102).

Furthermore, Capaldi's group observed that when the carboxyl-terminal helices assume

the hairpin conformation, bringing them closest to the Fo sector, ATP hydrolysis was

activated. Still, the enzyme was fully coupled in the direction of either hydrolysis or

synthesis. In contrast, when the two helices were open, assuming a position closer to the

Fl sector, ATP hydrolysis was inhibited and the enzyme functioned only in the direction

















~L~ 11
~3

EOL1


PDELSEED


E P-sandwich


Figure 1-3. Structure of the a subunit of E coli F1Fo ATP synthase. The residue numbers
and subunit labels are color coded to match the subunits it represents. The E
subunit has been suggested to undergo large conformational changes during
catalysis from an overwhelming amount of biochemical data. Two different
structures have been obtained for the a subunit, confirming the previous data.
A) Superimposition of the Ca trace of the E structure obtained from isolated a
subunit (u-helices shown in red and the P-sandwich shown in blue) with the
structure obtained from the y-s complex (yellow). B) Composite structural
model. Rotor stalk is based on the crystal structure obtained from the
y-s complex. The PDELSEED, Ecu2 and the s P-sandwich are indicated to show
the close proximity of PDELSEED and Su2 aS well as the relative distance
between the Su2z and the s P-sandwich.


134









of ATP synthesis. This conformational switch of the a subunit was therefore suggested to

play a key role as a selective inhibitor of ATP hydrolysis and directional regulator of

rotational catalysis by acting as a ratchet (102).

Movement of the two E a-helices was consistent with other observations. Changes

in the a subunit conformation due to nucleotide occupancy in the catalytic sites has been

observed in tryptic proteolysis experiments (89). Cysteine replacements in the carboxyl-

terminal a-helix (Su2) TOSulted in crosslinks with the a and P subunit (61, 103). More

importantly, treatment with a zero-length crosslinker, 1-ethyl-3 [3-dimethylamino]propyl

carbodiimide (EDC), resulted in a high yield of crosslinks between the a subunit and the

DELSEED (P380- P386) TegiOn of the P subunit (PDELSEED) following ATP hydrolysis in

the catalytic sites, but these interactions are disrupted upon the subsequent binding of

ATP. Also, in a composite structure of F1Fo ATP synthase incorporated with the E. coli

e-y complex as solved by Rodgers and Wilce, the s P-sandwich was at least 10 A+ away

from the PDELSEED TegiOn (Figure 1-3B) (101). The carboxyl-terminal Su2 prOduces

several points of interactions with the a, p and y as well as points of interactions with its

own P-sandwich domain a (61, 74, 88, 89, 92-99). In order for the Su2 to interact with

the PDELSEED, Ecul and the a subunit P-sandwich domain, it is clear from the structure that

the a subunit would be required to undergo large movements during the catalytic cycle.

In E. coli, F1Fo ATP synthase can act in two functional directions. In the case of a

dissipated electrochemical gradient, the F1Fo complex acts primarily as an ATPase in

order to pump protons across the membrane to provide a gradient to drive various ion

transport activities in the cell. Under severe conditions where cellular ATP levels are

exceedingly low the enzyme acts predominantly in the direction of ATP synthesis.









Therefore, one can imagine that the ability to selectively turn off ATP hydrolysis, while

preserving ATP synthesis function, may be important for E. coli. In mitochondria the

ability to control the F1Fo ATP synthase complex is essential. It is believed to act

exclusively in the direction of ATP synthesis and is strictly regulated (104).

The ring of c subunits

The c subunit is one of the three membrane-bound Fo subunits of F1Fo ATP

synthase. Ten copies of the c subunit form a ring in the membrane that plays a crucial

role in both proton translocation and rotation of the rotor stalk (105). It is the smallest

subunit of the F1Fo ATP synthase enzyme complex with 79 amino acids and a molecular

weight of 8,256 Da and it is encoded by the uncE gene.

Structure and topology. Early biochemical, genetic and immunological data had

suggested the structure of the c subunit to be that of a helical hairpin with two lipophilic

co-helices (amino acids 1-41 and 50-79) separated by a hydrophilic loop (amino acids 42-

49). Both of the putative transmembrane helices in the regions of Cleu4-leul9 and cphe53-phe76

were vulnerable to chemical modification by the nonpolar photoreactive reagent 3-

(trifluoromethyl)-3 -(m- [125I]i Odophenyl)di azirine (TID), whi ch i s a hydrophobi c carb ene

generator that is believed to react from the nonpolar region of the lipid bilayer, indicating

that these regions were in fact in the hydrophobic phase of the bilayer (106). The loop

region of the hairpin is substantially more polar and antibodies against it were shown to

bind to F1-stripped inverted membrane vesicles suggesting that it resides in the

cytoplasmic of the cell (107, 108). Both of the transmembrane helices are devoid of

charged amino acids with the noteworthy exception of Casp61 in the center of the second

helix, which undergoes a protonation and deprotonation cycle during proton translocation









(discussed in "F1Fo ATP Synthase Mechanism" below) Dicyclohexylcarbodiimide

(DCCD) reacts specifically with casp61, blocking proton translocation, and this reaction is

blocked by the mutationS Cala24-ser Of Cile28-thr, Suggesting that the c subunit is folded in

such a way so that the asp61 of the second helix is in close vicinity to residues 24 and 28

of the first helix (109, 110). This model was further supported by the ability to move the

critical aspartate from residue 61 in the second helix to residue 24 in the first helix

without disruption of enzyme function (1 11). Also, only one subunit in the ring of 10 c

subunits need be modified by DCCD to inhibit activity, indicating that each one of the c

subunits is consecutively involved in proton translocation (112, 113). Furthermore,

modification of the c subunit by DCCD trapped the configuration of the a subunit

(discussed above), providing evidence for a connection between the c and a subunits (89).

Intersubunit contacts made by the c subunit are evident from mutational data and

gives some insight to the topology. Mutations constructed in the polar loop region can

disrupt the binding of F1 to Fo (114-117). Three conserved amino acids, carg41-Cgln42-Cpro43,

lie at the apex of the polar loop region and are predicted to interact with the F1 s subunit

(98, 116). F1Fo ATP synthase complexes with the uncoupling mutation, Cgln42-glu, were

found to be recoupled with a second site suppressor mutation in the a subunit of F1,

Sglu31-gly, val, or cys (98) and was shortly followed by the observance of disulfide bridge

formation between the c subunit and the s and y subunits (99, 118). Also, switching the

essential aspartate from residue 61 in the second helix of the c subunit to residue 24 of

the first helix (discussed above) resulted in a functional F1Fo ATP synthase complex

though the cells were not as healthy compared to cells containing a wild-type enzyme

complex (111). Eighteen third-site suppressor mutants were found that helped to









optimize this cala24-asp,asp61-gly defect, with only five laying on the c subunit and 13 in the

a subunit, all near the aarg210 TOSidue, which is required for proton translocation (further

discussed below) (119, 120). Early models of the organization of Fo suggested that the

ring of c subunits were situated on the periphery, surrounding the centrally located a and

b subunits, which rotated in the center of the ring (121). This model was proved wrong

by high-resolution NMR data (122) and cross-linking experiments (123, 124) which

indicated that the oligomer of c subunits are closely packed with a lipid filled core less

than 25 A+ wide. The individual c subunits are packed front-to-back such that the second

helix of each is situated towards the exterior and the first helix is located on the interior,

which renders the casp61 exposed to the lipid environment. The uncommonly high pKa

(7. 1) of the casp61 carboxyl side chain is likely due to this hydrophobic environment (125).

Furthermore, scanning force and cryoelectron microscopy demonstrated that Fo is

asymmetrically arranged in the membrane (27, 126, 127). For these reasons, the a and b

subunits are thought to be situated to the periphery of the ring of c subunits.

High resolution structures of membrane-bound proteins were nonexistent for many

decades past structural determination of soluble proteins and still prove difficult to this

day due to their highly hydrophobic nature. The membrane intrinsic c subunit of E coli,

which was solved by NMR in an organic solvent (chloroform-methanol-water) in the

1990's, was one of the earliest high-resolution structures of a transmembrane helical

protein (122, 128-130). Notable, the c subunit could be reconstituted from the organic

solvent mixture with complete preservation of function; therefore, it was clearly not

irreversibly denatured (113). As predicted two decades prior, the c subunit folds as a

hairpin of two extended co-helices with the casp61 Of the second helix packed less than 5 A~









from the cala24 and cile28 Of the first helix (122). With the exception of Casp61 in the second

helix, both helices consist entirely of nonpolar amino acids. The first helix is greatly

enriched in glycines and alanines, which led to a smaller diameter. The a-helical

structure of the second helix is interrupted around casp61 due to disrupted hydrogen bonds

around cpro64 which cause the angle of the helical packing to change direction from there

to the carboxyl terminus (122). A recent study, using parallax analysis of fluorescence

quenching, the proton binding site casp61 WAS found to be deeply embedded in the

membrane at about 1.8 A+ from the center of the bilayer (13 1).

Stoichiometry. The stoichiometry of c subunits would be valuable in determining

the number of protons transported per ATP synthesized and will directly relate to the P/O

ratio of oxidative phosphorylation. However, the number of c subunits in Fo had been a

matter of controversy for many years. The number of c subunits in an F1Fo ATP synthase

complex was suggested to be between 9 and 14, but whether this number fluctuated based

on the species or environmental conditions or whether it was a fixed number were the

two prevailing arguments until just a few years prior. Based on a related family of

vacuolar (V-type) ATPases, in which the proposed subunit c had evolved into a fused

dimer of four transmembrane helices with a single proton-transporting glutamate in the

center of the fourth helix, Fillingame et al. set out to genetically fuse the E. coli c subunit

by introducing a flexible loop of similar length (123). The generated c-c dimers and c-c-c

trimers resulted in functional enzyme complexes. In combination with crosslinking

studies and normalization to the a/p content of the membranes, the favored stoichiometry

was fixed to 12 c subunits per F1Fo ATP synthase complex (132). More recently, the

experiment was revised to include only trimers and tetramers of the c subunit (105).









Partial activity was observed in complexes incorporated with eight (c4 C4) Of Hine C (C3

+ c3) Subunits and crosslinked products of more than 10 c subunits were observed but did

not purify in intact enzyme complexes. Crosslinking showed that the preferred

stoichiometry of c subunits in intact E. coli F1Fo ATP synthase was c4 C3 C3, Or 10 c

subunits. This number is consistent with the clo oligomer found in the yeast crystal

structure of a yeast F1Fo ATP synthase consisting of a3 3y8C10 TOSOlVed to 3.9 A+ (58).

However, the preferred number may still vary in different species. The archaebacteria

M\~ethanococcus janna;scjii ATP synthase has a natural c subunit trimer and therefore

cannot incorporate the E. coli equivalent of clo in the membrane (133). With 10 c

subunits present in the membrane 3.3 protons are required per ATP synthesized, which

was compatible with the early experimentally determined ratio of 3 H /ATP estimated

from E. coli whole cells (134). This value also indicates a P/O value of 2.3 from NADH-

linked substrates and 1.4 for succinate, also compatible with the predicted values of 3 and

2, respectively (135).

The Stator Stalk

The b and 6 subunits were once believed to form the central, rotating stalk of F1Fo

ATP synthase. However, high resolution crystallographic data refuted this idea in the

mid 1990's (20). The stator stalk did not come into view until improved EM technology

observed a peripheral stalk in the late 1990's (26-29) and the visualization of the 6

subunit, as a "cap" structure atop F1Fo ATP synthase, soon followed (27-29). As its name

implies, the role of the stator is to hold the u3 3 hexamer in place against the rotation of

the y subunit during rotational catalysis. Based on chemical crosslinking data, it is

currently believed that the a subunit resides to the periphery of the ring of c subunits with









the membrane-spanning domain of the b dimer situated to one side of the a subunit where

it is in close proximity to both the a and c subunits. The b dimer extends out of the

membrane and in a highly elongated conformation reaches to near the top of F1, making

contacts with the a and p subunits along the way and the 6 subunit at its extreme

carboxyl terminus.

The stator stalk consists of the b subunit of Fo and the 6 subunit of F1. Although

the primary function of the a subunit of Fo is considered to be a role in proton

translocation along with the clo subunits, it nevertheless plays the part as a stator and will

be discussed in this section. Structurally, the a subunit plays a role in both the formation

of a dynamic interface with the ring of c subunits as well as the formation of a secure

complex with the b dimer. Pursuit of a high-resolution structure for the a subunit remains

a challenge to this day. The high-resolution structure of the region of the a subunit that

forms the interface with the clo subunits is eagerly anticipated since it appears to be the

crucial region for proton translocation.

In regard to the stator stalk, partial structural information has been obtained for the

E. coli b subunit membrane spanning domain and the 6 subunit amino terminus by NMR

studies (136, 137). X-ray crystallography has solved the structure of a model polypeptide

based on the dimerization domain of the b subunit (138). The binding of F1 to the

membrane-bound Fo requires both the 6 and a subunits, suggesting that each of the

subunits are involved with the stalk structures of F1Fo ATP synthase (87, 139, 140). In

fact, the 6 subunit forms an integral part of the peripheral stalk and the a subunit

functions as part of the central stalk (discussed below). The 6 subunit of F1 has been

visualized seated at the very top of the Fl u3 3 hexamer by EM (11). However, recent










evidence has suggested that the 6 subunit may actually be positioned slightly to the side

of F1 in association with only a single oc subunit (Figure 1-2) (100, 141-143). The b

subunit is the primary focus of this dissertation and will be discussed at length later in

this chapter.

The asubunit

The E. coli a subunit is a large, extremely hydrophobic protein encoded by the

uncB gene and consists of 271 amino acids with a molecular weight of 30,276 Da. All

enzymes of the F1Fo ATP synthase family contain an a subunit homolog with strong

primary sequence homology even among evolutionarily diverse species (144). The most

highly conserved region resides in the carboxyl-terminal one-third end, amino acid

residues al90-263. Notably, in the region that is involved in proton translocation, there is a

remarkable conservation of the amino acid residues aleu207, aarg210, aleu211, asn214 and agln252

and an evident conservation of aglu219 and ahis245 at the homologous positions in all a

subunits from different species (144). The aarg210 iS the most strictly conserved among all

species and does not tolerate substitution with any other amino acid (further discussed

below) (145-147).

As mentioned above, there is no high-resolution structural data for the a subunit.

Also, contradicting models exist concerning the number of transmembrane helices as well

as the orientation in the membrane. Difficulty in studying the structure of the a subunit

arises from its extreme hydrophobicity and the necessity to include the denaturant,

trichloroacetate, in purification procedures. This is compounded by the fact that it cannot

be expressed at high levels in E. coli and is not found in the membrane without the

presence of both the b and c subunits (148-150). Furthermore, the a subunit is known to









be a substrate of the protease FtsH, which will rapidly degrade the subunit if it is not in

its native state (151). It was readily labeled with TID, which is a hydrophobic carbene

generator that is believed to react from the nonpolar region of the lipid bilayer, but its

solubility properties made it unsuitable for analysis as was done with the c and b subunit

(106). Consequently, the amino acids in contact with the lipid phase of the bilayer were

not identified. Due to difficulties in obtaining high-resolution structural data, much of

what is known of the a subunit arises from mutational studies.

Topology. Hydropathy analyses indicated five definite membrane-spanning

regions and one putative membrane span (121, 140, 152, 153). Much of what is known

of the a subunit structure and has come from the analysis of cysteine mutagenesis.

Greater than 50 cysteine substitutions, which resulted in a functional F1Fo ATP synthase,

were used in two kinds of experiments (154-158). Various maleimide derivatives were

used to search for the surface-accessible regions (154, 155). And double cysteine

mutations were used to search for disulfide formation between a-a and a-c (153). The

results supported the model in which the a subunit spans the membrane five times and the

fourth span, which includes aarg210 iS in contact with the second transmembrane ot-helix of

the c subunit (Figure 1-4A). Additionally, residues that were originally thought to be

located in the cytoplasm were not labeled, indicating that the six-membrane span model

was incorrect (132, 159).

The location of the amino-terminus of the a subunit has also been very

controversial. A substantial amount of evidence indicates that the carboxyl terminus



















66 10R


N




XXI


22T 238]


Periplasm



Membrane



Cytoplasm


127 138


257


BN


Ill 142


228 241


P~riplasm

0333330
t
Membrane

0333330


4~ 1 154


I


70 97


166 206


~C Cytoplasm


261


Figure 1-4. Controversial models of the a subunit topology. There is no high-resolution
structural data for the a subunit. Mutagenesis, crosslinking and
immunological experiments were used to study the topology. Roman
numerals indicate the number of transmembrane helices. Small numbers
indicate the relative position of the amino acid residue. Several crosslinking
reactions were observed between the fourth helix of the a subunit (IV*) and
the second helix of the c subunit in double cysteine mutants. Contradicting
models exist for the topology of the a subunit. A) Model with five-
transmembrane helices and the amino terminus residing in the periplasm. B)
Model with six transmembrane helices and the amino terminus located in the
cytoplasm.


19 206









resides in the cytoplasm (154, 155, 160). This observation, in combination with the five-

transmembrane helices, indicates that the amino-terminus should reside in the

periplasmic space. Polyclonal antibodies against a peptide model of the extreme

carboxyl-terminus as well as antibodies against epitope tags constructed at the carboxyl

terminus of the a subunit revealed this region to be located in the cytoplasm (160).

Moreover, cysteine substitutions at a266 Of G277 were highly reactive on the cytoplasmic

side of the membrane (154, 155). The orientation of the amino and carboxyl-termini was

studied by gene fusion proteins and peptide-directed antibodies, revealing a cytoplasmic

location of both termini (161, 162). Insertion of epitope tags at various positions also

confirmed the cytoplasmic local of both termini, arguing in favor of the controversial six-

transmembrane model of the a subunit (Figure 1-4B) (160). In the five-transmembrane

model a stretch of about 37 amino acids at the amino-terminus resides in the periplasm

with only one transmembrane helix, approximately up to residue a66, preSent (Figure 1-

4A) (154-156, 158). In the six-transmembrane model the amino-terminus resides in the

cytoplasm with two transmembrane helices present before the first cytoplasmic loop,

which range from approximately residues a33-49 and a54-70 (Figure 1-4B) (160).

A series of a subunit amino-terminal truncations and internal deletions were

constructed and the F1Fo ATP synthase function was tested by growth on a succinate

minimal media. Assembly of intact complexes was tested by membrane-associated

ATPase activity and the presence of the a subunit was analyzed by immunoblot analysis

(163). Four sections were found to be particularly interesting. The first 33 residues at the

amino terminus were shown to be necessary for the insertion of the a subunit into the

membrane. Two internal deletions, from residues a91-99 and al63-177, TOSulted in functional










enzyme complexes, indicating that these regions were not important for function. A

fourth deletion, from residues al20-124 WAS concluded to be important for function, but not

assemble because high levels of a subunit were found in the membrane, but the enzyme

was not functional.

The importance of the carboxyl-terminus was also analyzed by constructing a series

of early termination codons (164). Sequence alignment of the a subunit demonstrates

that many bacterial homologues contain glutamate and histidine residues at the extreme

carboxyl-terminus (glu-glu-his in E. coli). However, truncation of the Einal four residues

had no effect, and truncation of the Einal nine residues were tolerated at 250C, suggesting

that the extreme carboxyl terminus of the a subunit did not significantly contribute to

proton conduction or functional interactions with other subunits.

Proton translocation. The first indication that the a subunit was directly involved

in proton translocation appeared nearly two decades ago when mutations constructed in

the a subunit (aser206-leu and ahis245-tyr) WeTO found to affect Fo-mediated proton pumping

without influencing F1Fo ATP synthase assembly (165). Since then, not including the

cysteine mutations described above, more than 75 missense mutations have been

constructed and analyzed in or near the conserved regions of the a subunit to chart the

amino acids involved in proton translocation. In general, mutation of a conserved amino

acid residue impaired Fo-mediated proton translocation, but the severity of the defects

varied (166).

The only F1Fo ATP synthase a subunit residue that is strictly conserved amongst all

species, from bacteria to humans, and cannot endure any amino acid substitution, whether

basic, acidic or nonpolar, was aarg210 (145-147). Mutations at this site abolished both









ATP-driven proton pumping and passive Fo-mediated proton translocation. Growth on

succinate minimal media indicated no ATP synthesis by the mutants. The observed

effects were shown not to be due to failure of F1Fo ATP synthase to assemble because

treatment with the detergent lauryldimethylamine oxide (LDAO) released Fl from the

prepared membranes and revealed abundant ATP hydrolysis activity. The presence of

assembled F1Fo ATP synthase complexes incorporated with an aarg210 mutant was later

directly confirmed by Dr. James Gardner (167). Substitution with an alanine allowed

passive Fo-mediated proton translocation indicating that the proton channel was intact

and suggested that the aarg210 iS not obligatorily protonated or deprotonated during proton

conduction (168). A second site suppressor mutation, agln252 arg, which partially

compensated for the aarg210 gln mutation, was identified, and suggested to be in close

proximity to each other with residence on the transmembrane helix 5 and 4, respectively,

in the five-transmembrane model (121). The a210 TOSidue is thought to have a direct role

in proton translocation. The orientation of the a subunit' s fourth transmembrane helix

had been determined relative to the orientation of the c subunit' s second transmembrane

co-helix by crosslinking double cysteine mutants (157). Crosslinking data has positioned

a214 in ClOse proximity to c62 and c65, and a211 ClOse to c69 (157). This places the putative

fourth helix of the a subunit in contact with the second helix of the c subunit. Models

have the a210 TOSidue positioned near the center of the fourth helix at a level in the lipid

bilayer very close to the essential casp61 TOSidue (14). Whether a210 iS directly

protonated/deprotonated or controls protonation of the casp61 TOSidue remain unanswered

(169). Insight from a high resolution structure of an intact Fo is greatly desired and

would provide extremely valuable answers to many of the unsolved questions.










Single mutations at residues a218, a219 Of G245 WeTO Shown to have a considerable

impact on Fo-mediated proton conduction (144, 146, 170). When comparing amino acid

sequences of various mitochondria, chloroplast and bacteria, there appears to be an

instance of evolutionary covariation with these three amino acids (144). This suggests

that when a mutation occurred in one of the three residues, it was accompanied by a

second mutation to compensate for any loss in activity. This would cause the two

residues to pass through evolution as a hereditary unit. Based on this observation, double

mutants were constructed in the E. coli a subunit to imitate other lines of evolution (144,

171). Every double mutant studied resulted in functional F1Fo ATP synthase complexes

with considerably more activity than any of the single residue mutants. Due to the

functional relationship, it is possible that these three amino acids are in close proximity to

each other.

A few other strongly conserved amino acid residues located on the fourth and fifth

transmembrane helices are worth mentioning. Residues aasp214 and agln252 were both

strongly conserve but found nonessential, with the effects of mutations at these residues

varied widely (146, 170, 172, 173). Models of the a subunit have these residues lining a

water-filled proton channel. Recently, the aqueous accessibility of residues along

transmembrane helices 2 and 5 has been shown to extend to both sides of the membrane

(174). Also, a mutation at residue 217, aala217,ar,, blocked proton conduction and

inhibited F1Fo ATP synthase activity (167).

The 8 subunit

The E. coli 6 subunit is one of the F1 subunits. It is discussed here because it is an

essential part of the F1Fo ATP synthase stator stalk. The simplest stator stalks occur in









nonphotosynthetic prokaryotes and consist of a dimer of the Fo b subunits (discussed in

the following section) and a single Fl 8 subunit. The 6 subunit displays a very low level

of conservation across various species. It is a globular protein encoded by the uncH gene

that consists of 177 residues with a molecular weight of 19,332 Da. It plays essential

roles in both the binding of Fl to Fo as well as coupling of the catalytic activities of Fl

and Fo (139, 175-180). Circular dichroism (CD) spectroscopy and sedimentation analysis

studies performed on the 6 subunit suggested a highly helical and elongated conformation

(139).

Structure of the 8 subunit. A partial high-resolution structure of the 6 subunit has

been solved by NMR (137). During the purification procedure, a truncated form of the 6

subunit was produced by a bacterial protease. It was revealed to be the amino-terminal

134 amino acids (81-134) by mass spectroscopy and N-terminal sequencing. The same

sized 6 subunit fragment was often seen in Fl preparations and could be produced in

isolated E. coli Fl by treatment with trypsin without liberating the 61-134 frOm the Fl

complex (89). Furthermore, purified 81-134 Stably binds to 6-free Fl preparations. The

high affinity of the 61-134 Subunit for Fl indicated that the conformation of the 6 fragment

was preserved during the purification procedure. NMR was performed on both the 61-134

fragment and the intact 8 subunit; however, the quality of data for the intact subunit was

not sufficient enough for structural analysis due to its propensity to aggregate at high

concentrations. To date, the carboxyl-terminal 43 amino acid residues (6135-177) Of the 6

subunit is the only portion of the Fl sector not known at the atomic level.

The amino-terminal 105 residues of the 6 subunit formed a dense globular domain,

while the region from residues 106-134 was mostly disordered with the exception of one










a-helix (137). The amino-terminal domain, 61-105, consisted of a six a-helix bundle with

the dimensions 45 x 20 x 30 A+. Helices 1 (64-20) and 2 (624-38), and helices 5 (670-81) and 6

(688-104) Organized into V-shapes that intercalated to form a core. Helices 3 (641-47) and 4

(653-64) were packed compactly against this four-helix core. Following this globularly

packed domain there was a loop region followed by a seventh a-helix (6118-129).

Comparison of the structural data for the intact 8 subunit against that of the 61-134

fragment illustrated the same structure for residues 1-104, but the spectral shift of

residues 105-134 was very different. It was possible that the carboxyl-terminal 42

residues missing from the 61-134 fragment affects this region of the 6 subunit.

8 subunit topology. Taken together with biochemical and immunological data, the

structure revealed by NMR revealed that the 6 subunit consists of two domains, an amino

terminal domain, 61-104, and a carboxyl-terminal domain, 6105-177. Under oxidizing

conditions, two native cysteines present in the amino- and carboxyl-terminal domains of

the intact 8 subunit, Scys64 and Scysl40, TOSpectively, formed a disulfide bond. Furthermore,

NMR data indicated some NOE's between the carboxyl-terminal a-helix and the amino-

terminal domain. The data indicated that there is probably a close interaction between

the amino- and carboxyl-terminal domains of the intact 8 subunit.

Proteinase accessibility and immunological analysis were used to examine the

topology of the 6 subunit (89). The 6 subunit was susceptible to trypsin digestion at the

carboxyl-terminal 20 residues in isolated Fl, but not in intact F1Fo ATP synthase,

indicating a protection of the amino-terminal region by Fl. Deletion analysis of the

carboxyl-terminal region also implied the importance of the 6 subunit in binding Fl to Fo.









Taken together, these observations suggested that the amino-terminal domain is

predominantly involved in the binding of the 6 subunit to Fl and the carboxyl-terminal

domain is involved in binding to Fo.

The location of the 6 subunit has had a history of being very controversial. Prior to

the high resolution structure obtained by Abrahams et al. (1994), the b and 6 subunits

were expected to form part of the central stalk of the F1Fo ATP synthase enzyme, which

is now known to consist of the y and a subunits (20). Due to the dimensions, it seemed

unlikely that the b and y subunits could fit as part of the central stalk, which implied that

they must form a separate connection between Fl and Fo. Improving EM technology did

not allow visualization of the second stalk structure at the periphery of the F1Fo ATP

synthase complex until many years later (27, 28). Prior to visualization by EM, several

early crosslinking studies had been reported in the quest to find the location of the

6 subunit binding on Fl, finding it to be on the a subunit (89, 181-184). Notably,

crosslinking the 6 subunit to the a subunit did not have a great impact on F1Fo ATP

synthase function, as would be expected if the 6 subunit formed part of the stator stalk

(185). High-resolution structure of the Fl u3 3 hexamer with a partial structure of the y

subunit had revealed a dimple in the top of the hexamer approximately 15 A+ deep that

was adj acent to the core space where the amino- and carboxyl-terminal a-helices of the y

subunit resided (20). EM studies had revealed a "cap" structure at the very top of F1 in

both E. coli and mitochondrial complexes (27, 29, 186). It was thus believed that the 6

subunit resided in the dimple of F1 as the cap seen in the EM structures (187). This

possibility was refuted when Prescott et al. demonstrated the ability to stably incorporate

the green fluorescent protein (GFP), via varying length peptide linkers (0, 4 or 27 amino









acids), to the carboxyl-terminus of the y subunit without interrupting function of the

enzyme complex. GFP forms a rigid, stable structure with the dimensions 24 A+ wide and

48 A+ high (188). This study indicated that the putative cap structure could not possibly

occupy the entire dimple atop Fl. More recent evidence has suggested that the 6 subunit

may actually be positioned slightly to the side of F1 in association with only a single a

subunit (Figure 1-2) (100, 141, 142, 189).

The bsubunit

The b subunit is required for the normal assembly and function of F1Fo ATP

synthase (190). The E. coli F1Fo ATP synthase has two identical b subunits, which form

a homodimer, that are the product of a single gene (Figure 1-2). It is an elongated

amphipathic polypeptide that crosses the membrane one time at its amino-terminus and

has an extensive hydrophilic carboxyl-terminal domain. This pattern is characteristic of

b-type subunits of ATP synthases, although the mitochondrial b has two consecutive

membrane-spanning segments at the amino-terminus (191). Most ATP synthase b-type

subunits consist of between 150 and 170 amino acid residues. The E. coli b subunit,

encoded by the uncF gene, consists of 156 amino acid residues and has a deduced

molecular weight of 17, 265 Da (Figure 1-5).

Domains and Structure. Currently there is no high-resolution structure of the

entire b subunit. Several factors probably contribute to the difficulty of structural

analysis. The b dimer is a thin, highly extended, mostly a-helical structure, its

dimerization is comparatively weak and reversible (192), and it displays evidence of

flexibility (193-195). This has led to alternative low-resolution approaches to study the

structure of the b dimer such as circular dichroism (CD) spectroscopy, deletion analysis,










74=, 75= 78 227= 228 a,
t 200 bend a a
++ A* A* A f 40
MNLNATILGQ AIAFVLFVLF CMKYVWVPPLM AAlEKRQKEl
Mnembrane-Spanning Tether



At* .. X 8
ADGLASAERA HKDLDLAKAS AT











QVAILAVA GAEKllERSV DEAANSDIVD KLVAEL

F,-Binding






Figure 1-5. Amino acid sequence of the E. coli F1Fo ATP synthase b subunit. The E. coli
b subunit is a 156 residue amphipathic polypeptide. The amino acid sequence
and the four domains are shown. The transmembrane domain (bl-22), tether
domain (b24-60), dimerization domain (b63-122) and the 6-binding domain (bl23-
156) are Shown in blue, orange, green and red, respectively. The large purple
stars indicate residues capable of forming high yields of b-b crosslinks upon
cysteine substitution. The smaller purple stars indicate residues found to form
low-yields of crosslinks. The arrows indicate positions crosslinked to other
subunits of ATP synthase. High-resolution structures based on model
polypeptides consisting of bl-34 and b62-122 (underlined residues) have been
solved by NMR and crystallography, respectively. NMR analysis of residues
1-34 has revealed a a-helical structure with a rigid 200 bend at positions 23-
26. X-ray crystallography revealed a highly a-helical structure with modeled
into a right-handed coiled coil.










analytical ultracentrifugation, chemical crosslinking, and the analysis of tendencies for

disulfide bond formation. CD spectroscopy analysis has predicted the secondary

structure of the b subunit to be approximately 80% ot-helical with about 14% p-turn

conformation (196).

Although there is no high-resolution structure of the intact Fo sector, an abundance

of evidence suggests the necessity of the b subunit to exist in the dimeric state. The

hydrophilic region of b, consisting of residues b24-156 (alSo known as bsol), has been

expressed and shown to form highly extended dimers capable of binding to F1-ATPase in

solution (197). Sedimentation equilibrium ultracentrifugation gives a molecular weight

value of about 30,000 Da for bsoi, consistent with a dimer of two 15,000 Da bsol

monomers (13). The existence of the dimeric state of the b subunits was confirmed by

covalently cross-linking the two b subunits in the complex and verifying the activity of

the enzyme (198). Furthermore, the ability of b to bind to Fl was discovered to be

directly proportional to the ability of b to form dimers, suggesting the necessity of the b

dimer formation before the binding of Fl to the complex (199).

The dimerization of the b subunit has been shown to be relatively weak and

reversible. The monomeric and dimeric forms of bsol were shown to exist in a dynamic

equilibrium and the dimer was converted to the monomeric state at 400C (192). This

same melting characteristic was observed with CD spectroscopy (200). Furthermore, the

similar traits were observed in photosynthetic organisms, which encode two different b-

type subunits, b and b '(13). When the cytoplasmic regions of the b and b subunitss from

the cyanobacterium Synechocystis were expressed individually, the polypeptides were

found to only exist in the monomeric state. However, when they were mixed together,








the formation of the dimers was observed by chemical crosslinking and sedimentation
equilibrium ultracentrifugation (13). Also, the dimers were observed to melt at 400C as
was the case with the E. coli b subunit. The striking similarity between the E. coli b
subunit and the photosynthetic organism b and b subunits indicate that the former is a
good model in which to study the b subunit.
Cross-linking and deletion analysis has led to the development of a four-domain
model of the E. coli F1Fo ATP synthase b subunit (Figures 1-5 and 1-6) (12). Amino acid




FDB 1 adinn


Dim eriz ation
Dom ain



Tether
Dom ain

M~embrane
Dom ain


I


Figure 1-6. Gross structure of the E. coli F1Fo ATP synthase and the domains of the b
subunit. The b subunit domains were described by Dunn et al. (12) The
membrane-spanning domain roughly corresponds to amino acid residues 1-22,
the tether domain is approximately residues 24-60, the dimerization domain is
considered to be residues 60-122, and the F1-binding domain is roughly
residues 123-156.









residues bl-22 COrresponds to the hydrophilic membrane spanning domain. Residues b24-60

roughly corresponds to the tether domain, which is the portion of the peripheral stalk

often observed in electron micrographs. The dimerization domain, approximately b60-122,

is required for the dimerization of the two b subunits. And finally, the Fl binding

domain, roughly amino acid residues bl23-156, iS required for the binding of F1 to Fo.

The amino-terminal membrane spanning domain, bl-22, forms a single

transmembrane span while the large remainder is a polar hydrophilic domain which

extends above the cytoplasmic leaflet of the lipid bilayer and reaches towards the top of

Fl (Figure 1-6) (13, 191). A wealth of evidence has suggested this proposed topology for

the b subunit. The amino terminal region was uniformly vulnerable to chemical

modification by the a nonpolar photoreactive reagent, TID, which is a hydrophobic

carbene generator that is believed to react from the nonpolar region of the lipid bilayer,

indicating that this region was in fact in the hydrophobic phase of the bilayer (106). This

observation was consistent with other labeling procedures including the labeling of boys21

by hydrophobic nitrenes (201) or the hydrophobic maleimide N-(7-dimethylamino-4-

methyl-coumarinyl)-maleimide (DACM) (148). Modification of boys21 interfered with

intersubunit interactions within Fo. Furthermore, reconstitution of the Fo subunits upon

labeling the b subunit with DACM resulted in reduced proton translocation as well as Fl

binding affinities (148). TID failed to label the region basn2-gln10, indicating that the first

few residues at the amino-terminus protrude into the periplasm (106)

Despite attempts by several laboratories, there is presently no high-resolution

structure of the entire b subunit. Therefore, model polypeptides have been constructed in

order to elucidate the structure of the b subunit by domain. A model polypeptide










comparable to residues bl-34, which contains the membrane-spanning domain, dissolved

in a 4:4: 1 v/v mixture of chloroform/methanol/water previously used for solving the

structure of subunit c, has been solved by NMR (136). The data revealed an a-helical

monomeric structure with a 20o bend at residues 23-26 (KYVW) (Figure 1-5). The

hydrophobic residues, b4-22, formed an a-helix, followed by the 200 bend, and then

resumed with a-helical structure from residues b27-34. The bend was proposed to be

positioned as the b subunit exits the membrane. A series of cysteine substitutions

resulted in a high yield of crosslinks formed at residues b2, b6, and blo (Figure 1-5). A

lower yield of crosslinks were observed to form at residues b3, bs, b9 and bll (Figure 1-5).

No crosslinks were observed when cysteines were substituted for residues 12-21. The

observance of continuous crosslinks between residues b6-11 WeTO Suggested to indicate a

dynamic interaction between the contacting faces of the two b subunits in this region

(132). These observations led to a dimeric model in which the extreme amino-termini of

the b subunits crossed each other in close proximity at an angle of about 350 in the region

of residues b4-11, and then the two b subunits angled apart as they traverse the membrane

towards the cytoplasmic side (Figure 1-5) (136). The region of the 200 bend, b23-26, WAS

suggested to change the direction of the second a-helix, b27-33, Such that it would extend

into the cytoplasm at an angle perpendicular to the plan of the membrane. This model

was confirmed by a systematic mutational analysis of the membrane-spanning domain

performed by Hardy et al. (202).

The tether domain of the b subunit, roughly b24-60, iS the least defined part of the

subunit domains from a structural point of view. It corresponds to the portion of the

peripheral stalk often seen in electron micrographs and is called "tether" simply because









it is the section of the b subunit that links the more defined membrane-spanning and

dimerization domains Figure 1-6). There is no high resolution structure for this region of

the b subunit. The NMR structure of bl-34, described above extended slightly within this

domain, revealing an a-helix at least up through residue b34 .(136) Also, a heptad repeat,

extending from just outside of the membrane and continuing without interruption up to

residue bala79 Suggests the structure to be a coiled coil (197, 203). Though crosslinking

studies showed that the tether domains of the two b subunits are in parallel and in close

proximity, this domain contributes little to the stability of the dimerization of the b

subunits (192).

Deletion constructs analyzed by sedimentation equilibrium experiments suggested

that a form of the b subunit truncated in each end, b53-122, WAS capable of forming dimers

with an efficiency close to the complete cytoplasmic domain, b24-156, indicating that the

most pertinent intersubunit contacts of the b subunit was located within this central

region, referred to as the dimerization domain (192). More recently, the dimerization

domain has been refined to residues b63-122; however, the amino-terminal boundary is

likely to decrease even further due to the observation that deletion of residues b54-64 Of

b65-75 TOSulted in intact and functional F1Fo ATP synthase complexes (Figure 1-6) (194).

A crystal structure of a monomeric dimerization domain, based on a model

polypeptide consisting of residues b62-122, has recently been solved and refined to 1.55 A+

(138). Based on an undecad repeat and crosslinking data, Dunn and coworkers have

constructed a model in which the two a-helices of the b62-122 TegiOn formed a coiled-coil

with a right-handed superhelical twist. A number of previous studies supported this

coiled-coil arrangement. First, the shape of the b53-122 pOlypeptide was consistent with a









coiled-coil of similar length as determined from its frictional coefficient (1.60) in an

ultracentrifuge and from NMR relaxation parameters (192). Secondly, small-angle X-ray

scattering by b52-122 in Solution specified a maximum length to be about 95 A+, consistent

with the expected coiled-coil length (13). Thirdly, CD spectroscopy indicated that this

polypeptide was 100% ot-helical and the similar intensities of the minima suggested the

helices to be arranged in a coiled-coil (204). Fourthly, cysteine substitution and

crosslinking studies suggested a periodicity consistent with a parallel coiled-coil (Figure

1-5) (204). Finally, b subunit sequence analysis of E coli and other prokaryotes revealed

a conservation of an undecad pattern, which is a distinctive characteristic of a right-

handed coiled coil. Mutation of amino acid residue barg-83, which interrupts the undecad

repeat, markedly stabilized the dimer, as expected for the proposed two-stranded, right-

handed coiled-coil structure.

The carboxyl-terminal F1-binding domain, bl24-156, alSo referred to as the 6-binding

domain, has a more globular conformation and is required for the binding of F1 to Fo

(205, 206). Work accomplished by Futai and coworkers two decades ago revealed that

truncation of the extreme carboxyl-terminus of the F1-binding domain by only a few

amino acids resulted in assembly defects in F1Fo ATP synthase (206). Subsequently,

work performed by Dunn and coworkers demonstrated that the final two to four amino

acids of the b subunit were necessary for binding the 6 subunit of Fl (205). An addition

of a cysteine at the carboxyl-terminus was chemically crosslinked to a cysteine

introduced at Siss.

A close association of the two b subunits in the F1-binding domain was indicated

by crosslinks formed between cysteines individually substituted at positions bl24, bl25,









bl26, bl27, bl28, bl29, bl30, bl31, bl32, bl39, bl44, bl46, Or bl56 (Figure 1-5) (198, 200, 207).

Hydrodynamic evidence favors a folded structure for this domain of the b subunit as

opposed to the highly elongated structure of the remainder of the b subunit. Also, several

studies have shown that either a balal28-glu mutation, deletion of the last four residues, or

cold temperature dramatically decreased the sedimentation coefficient, by 23%,

suggesting that the F1-binding domain underwent a conformational change from a

globular structure to a less folded more extended conformation (192, 205, 208). The

mutation, balal28-glu, may have caused an electrostatic repulsion that would cause the two

b subunits to push apart. The carboxyl-terminal residues may form an amphipathic helix,

so the deletion would have disrupted essential interactions. And cold temperatures have

been shown to weaken hydrophobic interactions in proteins, suggesting the importance of

the hydrophobic amino acids in the folding of this domain (13). Dunn et al. suggested

that these observations implied that the carboxyl-terminus of the b subunit has a weakly

folded structure in which the hydrophobic amino acids are arranged to impart structural

stability and create hydrophobic patches on the surface (13). The folded conformation

appears to be required for the exposed hydrophobic patches to interact with Fl.

Mutagenesis. Several mutant searches and site-directed mutagenesis studies have

been performed in the membrane-spanning domain of the b subunit. However, only a

single mutation, bgly9,asp, lOcated near the periplasmic side of the lipid bilayer, resulted in

a defective proton pore in an intact F1Fo ATP synthase complex (209). Second site

suppressors of this mutation have been found in the a(apro240-ala Of Opro240-leu) and c

(cala62-ser) Subunits that partially repaired the defect, indicating an interaction between the

b subunit and both the a and c subunits (210, 211). The membrane-spanning domain









contains one charged residue, blys23, but mutations generated at this site did not influence

proton translocation, suggesting that this domain of the b subunit did not have a direct

function in proton conduction, although it was required for the maintenance of a

functional Fo complex (212). A systematic mutational study of the membrane-spanning

domain conducted by Hardy et al. was described above and a triple mutant, bN2A,T6A,Q10A,

is described in Chapter 5 of this dissertation (202).

A couple notable characteristics can be attributed to the tether region of the b

subunit. Relatively large deletions and insertions of up to 11 and 14 amino acids,

respectively, were accommodated in this region and the altered b subunits still assembled

into fully functional F1Fo ATP synthase complexes (193, 194). In fact, decreases

observed in enzyme activity paralleled the decrease of b subunit found in the membrane,

suggesting that the alterations affected assembly of the enzyme, but not the function.

Assuming co-helical structure, an 11 amino acid deletion would shorten the b subunit

tether region by approximately 16.5 A+ and a duplication of 14 amino acids would

increase the length by about 21 A+. This implies that the b subunit is highly flexible and

the altered b subunits may compensate for the lost or gained distance via that flexibility.

The fact that the peripheral stalk must extend from within the membrane to up near the

top of Fl suggests that some part of the stalk must be flexible enough to stretch or

straighten in the shortened b subunits, or in the case of the lengthened b subunits, bend to

take up the slack. Prior to these observations, the b subunit was often viewed as a rigid,

rod-like structural feature during rotational catalysis. Also, though the b subunit is the

least conserved subunit of F1Fo ATP synthase, gaps are rarely found in sequence

alignments of numerous organisms (213, 214). Therefore, the ability to manipulate the


















































Figure 1-7. Model for F1Fo ATP synthase peripheral stalk orientation dependent upon the
direction of rotation during ATP synthesis or hydrolysis. The subunits are
color coded as follows: ot, light blue; P, grey; y, dark blue; E, green; 6, orange;
a, yellow; b, red; c, cyan. The panel on the left indicates the orientation of the
b2 dimer if the enzyme is actively synthesizing ATP. The panel on the right
represents the position of the b2 dimer during ATP hydrolysis. The arrows
indicate the direction of rotation of the rotor stalk subunits (ysclo). The red
cylinders indicate regions of the b subunits for which there is no high-
resolution structure.









length of the peripheral stalk was an unexpected surprise to the field. The length of the

wild-type b subunit is probably the optimum length for assembly of F1Fo ATP synthase.

The apparent flexibility has been proposed to help alleviate torsional strain brought about

by rotational catalysis (193). Another hypothesis concerning the flexibility of the tether

domain is that this region of the b2 dimer serves as a hinge, allowing reorientation of the

stator depending on the direction of rotation as the enzyme carries out ATP synthesis or

hydrolysis (Figure 1-7) (195, 202).

Another important feature of the tether domain is the evolutionarily conserved

barg36, that has been implicated in a structural role influencing proton conduction through

Fo. Mutational studies at this amino acid residue led to numerous defects from failure to

assemble or function to uncoupling phenotypes (215). Amino acid substitutions, barg26-ile

or barg26-glu, TOSulted in assembled F1Fo ATP synthase complexes that displayed defects

in Fo-mediated proton translocation or a disruption of coupling activity, respectively.

Substitution with a cysteine at this residue led to a crosslink product with the a subunit of

Fo (216, 217). The close proximity of this conserved residue to the a subunit indicates

that it may play a role in aligning the proton exit channel.

Protein-protein interactions between the two b subunit dimerization domains have

been shown to be essential for forming the peripheral stalk (13). Mutations at a

conserved residue, bala79, TOSulted in maj or F1Fo ATP synthase assembly defects (203,

218). The bala79 mutations were modeled in the bsol polypeptide to investigate the affects

of the mutations. The model polypeptides were shown to retain co-helical structure, but

chemical crosslinking and sedimentation experiments suggested that the bala79 mutants

were incapable of forming dimers (199).









In the F1-binding domain, a mutation was found, balal28-asp, that had little effect on

the dimerization of the b subunits but led to an assembly defect of F1Fo ATP synthase

(208). However, the mutant was found to have a reduced tendency to interact with Fl and

sedimentation equilibrium ultracentrifugation experiments revealed a 12% decrease in the

sedimentation coefficient, indicating a structural perturbation (discussed above). The

studies suggested that the balal28 TOSidue was not important in b subunit dimerization, but

it had an important structural role in the F1-binding domain.

Intersubunit interactions. The formation of disulfide bonds between cysteine

residues introduced in the membrane-spanning domain of the b subunit and the a subunit

as well as second site suppressors of the bgly9,asp found in the a subunit (discussed above)

strongly suggests an interaction between the two stator subunits (209-211). The bgly9-asp

mutation was also partially suppressed by a mutation in the c subunit (discussed above),

but it is not known whether it is due to a direct interaction between the b and c subunits

or if the suppression is mediated through the a subunit.

The b subunit has also been shown to interact with the ot, P and 6 subunits of F1

(198, 216). The interaction of the Fo b subunit with the Fl 8 subunit has been well

documented (200, 205, 219, 220). The interaction is mediated by the carboxyl-termini of

both subunits and is essential for the binding of F1 and Fo. The critical role of the 6

subunit mediating the interaction between the b subunit and the bulk of F1 has been

demonstrated by the inability of 6-depleted Fl to bind to Fo (219). Crosslinking the b and

6 subunits via introduced cysteines did not affect F1Fo ATP synthase activity, which was

consistent with the proposed role of the b28 peripheral stalk as a stator and demonstrated

that the b-6 interaction need not be dynamic (221). Though it is believed that the binding










of Fl to Fo heavily relies on the b-6 interaction, evidence of other subunit contacts may

also influence binding. Furthermore, the binding of b to 8 was shown to be relatively

weak by analytical ultracentrifugation, indicating that other subunit contacts may

contribute to binding (220). Many crosslink formations were found when cysteines were

introduced into the b and a or P subunits (Figure 1-5). A cysteine introduced to the

carboxyl-terminus of the b subunit has been shown to crosslink to acvs90 (198). Also,

cysteines positioned at b92 Or bl09 formed crosslinked products with the a subunit or both

the a and p subunits, respectively (216). These results confirm that the b subunit is

proximal to the u3 3 hexamer, but a direct interaction has not been confirmed.

Stator stalk function. The necessity of a stator in F1Fo ATP synthase was

recognized after the realization that rotation was a fundamental feature of ATP catalysis.

The current view of the peripheral stalk is primarily that of the stator which forms a

connection the a subunit and the u3 3 hexamer, holding these subunits in place against

the rotation of the rotor subunits, cloys. The idea of a flexible stator stalk has led to other

proposed features of the b subunit. The apparent flexibility of the b subunit has been

suggested to transiently store energy during rotational catalysis which could be

potentially be expended to force the conformational change that allows the release of

ATP(222). Another model describes the flexibility of the tether region as a hinge that

could reorient the b subunit when switching between ATP synthesis and hydrolysis

(discussed above) (Figure 1-7) (202). However, there is no direct evidence of whether

the b subunit is actually acting in a flexible manner. Finally, the b subunit has recently

been suggested to influence the nucleotide binding sites in the P subunits (223). In these

experiments, a spin label was incorporated at residue P331. Upon addition of bsoi, the










spectrum of this spin label was observed to change in a way that implied that the catalytic

sites were in a more open conformation. These results indicate that the current view of

the stator stalk as a structural feature may soon be revised such that the b subunit has a

more direct role during rotational catalysis.

Subunit Equivalence

The E. coli ATP synthase complex has, by far, the simplest architecture of all the

F1Fo ATP synthase enzyme complexes. It is composed of twenty-two polypeptides of

eight different types with the stoichiometry u3 3y~sab2010 (Figure 1-2, Table 1-1) (112,

224). The u3 3YiS Subunits comprise the Fl sector and the ab2C10 Subunits comprise the

Fo sector. In the E. coli enzyme, all eight subunits are necessary for the function of F1Fo

ATP synthase (225, 226). Chloroplasts also have a relatively simple architecture with the

exception that they have nine different subunits (227) due to the fact that the two b

subunits are products from two different genes and are not identical (Figure 1-8). In

contrast, the F1Fo ATP synthase from mammalian mitochondria is composed of at least

thirty-one polypeptides of sixteen different types with the Fl stoichiometry of u3 3Y68

and a much more complex Fo consisting of a, b2, C10-14, d, e,J; g, (F6)2, A6L, OSCP, IFI

(7, 228-230). Yeast mitochondria F1Fo ATP synthase has an extra three subunits

compared to the mammalian enzyme, stflp, i, and k.

The F1 sector. In the Fl sectors, homologues of the E. coli F1Fo ATP synthase

have been identified for the a, P and y subunits, based on amino acid sequence

homology, in the chloroplast and mitochondrial enzymes (30). Based on the primary

sequences, the highest conserved subunit from the E. coli F1Fo ATP synthase is the P

subunit with approximately 70% homology with the chloroplast and mitochondria










Table 1-1. F1Fo ATP synthase subunit equivalency
Mitocl-ondria
Bacteria Chloroplast Yeast Bovine Function
a a a a catalytic site
P P P p catalytic site
y y y rotor
6 6 OSCP OSCP stator
a E 8 8 rotor
-~ E stabilization?
a a (or IV) a (or 6) a poton channel, stator
b b (or I)b (or 4) b stator
Sb '(or II) 9 -stator
c c (or III) c (or 8) c poton channel, rotor
-~~ d'a stator?
8 A6L stator?
e e ?


h F6 stator
inh1p IF1 inhibitor
Sstf~ ?

k -?

equivalents (31). The a subunits exhibit roughly 50% homology (31). The nucleotide

binding regions of these two subunits also have sequence homologies with other proteins

that bind nucleotide or phosphate, including the E. coli secA protein, N-ethylmaleimide

sensitive fusion protein, herpes simplex virus UL15, Ca2+-ATPase, H /K+ ATPase and

Na /K+ ATPase (34-37). Furthermore, the nucleotide binding motif, GXXXXGKT/S,

which was first identified in the a and P sequences ofF1, has been found to be conserved

in the high-resolution structures of other proteins including p21,as, adenylate kinase,

RecA, elongation factor Tu, and transducin-a (20, 38-42).

Interestingly, the y subunit in the chloroplast F1Fo ATP synthase complex contains

an insert of about 35 amino acids that is not present in the mitochondrial or









nonphotosynthetic eubacteria (231). This loop contained two cysteine residues that were

found to be reduced in the active enzyme complex during photosynthesis and oxidized to

a disulfide bond in the inactive enzyme while in the dark (232). The E. coli s subunit is

unfortunately known as the 6 subunit in the mitochondrial enzyme (233) and also shares

primary sequence homology with the mitochondrial IF1 inhibitor protein. The E. coli s

and bovine 6 subunits share 60% sequence identity, which had suggested that they are

functionally equivalent (191, 234). The availability of high resolution structures for these

subunits has revealed that they are strikingly similar. Superimposition yields a 1.64 A+

rms deviation (48). The bacterial a subunit has been suggested to be an inhibitor of ATP

hydrolysis, undergoing large, ratchet-like conformational changes to selectively switch

off ATP hydrolysis (102). In mitochondria, this inhibitor action of the bacterial a subunit

is ascribed to the IF1 protein of the Fo sector. It was suggested that the bacterial a subunit

was separated into the two polypeptides in the mitochondrial enzyme complex, E and IF1.

Finally, the bacterial and chloroplast 8 subunits ofF1 share a significant sequence

homology (234). The 6 subunit's equivalent in the mitochondrial F1Fo ATP synthase is

known as oligomycin-sensitivity-conferring protein (OSCP) (23 5-23 7). The carboxyl-

terminal region of the mitochondrial b subunit was been demonstrated to bind to the

OSCP subunit (E. coli 6 subunit) through subunit interactions (Collinson et al., 1994) and

chemical crosslinking analysis (Soubannier et al., 1999).

The Fo sector. The Fo sectors of the F1Fo ATP synthase family is by far more

diverse than the Fl sector with an additional eight different subunit types in mammalian

and an extra ten different subunits in yeast mitochondria (Table 1-1). The E. coli a and c

subunits are respectively equivalent to the chloroplast IV and III subunits and the









mitochondrial ATPase-6 and ATPase-8 subunits. In every case, both subunits are

hydrophobic proteins required for proton translocation (120, 140). Unlike the a and c

subunits, the E. coli b subunit does not have any obvious homologues in chloroplasts or

cyanobacteria F1Fo ATP synthases. However, both of them have two distinct subunits

with similar hydrophobic and hydrophilic residue distribution (238). These subunits are

referred to as subunits b and b 'in cyanobacteria and subunits I and II in chloroplasts. It

is believed that only one of each of these subunits is present in the F1Fo ATP synthase

complex, incorporating into the enzyme as a b-b heterodimer as opposed to the b subunit

homodimer present in E. coli. No obvious homologue of the E. coli b subunit has been

found in the mitochondrial F1Fo ATP synthase, even upon analysis of sequence, function

or three-dimensional structure (239). However, hydropathy plot analysis does indicate

that the mammalian mitochondrial b subunit that may be analogous to the E. coli b

subunit (240, 241). Proteolysis studies and crosslinking data supported the location of

the mitochondria b subunit at a position analogous to the E. coli b subunit (242, 243). At

least three other subunits may play the role of the E. coli b dimer including subunit 8 (or

A6L), d' and F6. The mitochondrial b subunit is believed to have two a-helical

transmembrane spans at its amino-terminus arranged in an antiparallel configuration.

The extreme amino-terminus of the mitochondrial b subunit is thought to begin on the

cytoplasmic side of the membrane, traverses the membrane to the periplasmic leaflet of

the membrane as an a-helix, then turn back and traverses the membrane again as it exits

the membrane in the cytoplasm and reaches towards the top of F1 (Figure 1-8). The

remainder of what would be equivalent to the E. coli b dimer is highly speculative,

though the overall shape and characteristics of the b8dF6 subunits favor this explanation








(Figure 1-8). In this model, the mitochondrial subunit 8 contributes to a third membrane

spanning region, and a combination of subunit d and subunit F6 forms the hydrophilic
domain.

The Fo subunit known as the a subunit in mitochondrial F1Fo ATP synthase has no

counterpart in the bacterial or chloroplast enzymes. It is a small polypeptide (50 amino

acids) folded into a helix-loop-helix. It is believed to play a role in the stabilization of the

central stalk and its absence in the bacterial and chloroplast enzymes may explain why


b2


Domains




F1 binding



Dimerization


Tether

Membrane


Bacteria


Mitochondri on


Figure 1-8. Speculative models for the b-like subunits. Shown are models for the
bacterial, chloroplast and mitochondrial b subunits. The membrane-spanning
regions are indicated by the black lines. An abundance of evidence supports
the parallel arrangement of the bacterial b2 homodimer and the chloroplast bb '
heterodimer. The mitochondrial analogue of the b subunit is believed to
consist of up to four polypeptides. The model shown for the mitochondrial
b8dF6 Structure is highly speculative.


bb'


b8dF,


Chloroplast










the bacterial a subunit of the Fl sector (equivalent to the 6 subunit in mitochondria) easily

dissociates from the Fl complex whereas this has not been observed for the mitochondrial

enzyme. Mitochondrial Fo consists of several additional subunits not found in the E. coli

or chloroplast enzymes including the e, fand g subunits as well as an extra three in yeast,

stflp, j, and k subunits.

E. coli F1Fo ATP synthase as a model. Initial studies of the F1Fo ATP synthase

complex were achieved with enzymes isolated from mitochondria or chloroplast.

Although the bacterial, chloroplast and mitochondrial enzymes differ in oligomeric

complexity, the enzymes show acceptable overall structural resemblance and primary

sequence homology that it is widely accepted that the mechanism of action is the same in

all organisms (240, 244). Therefore, studies using the bacterial model became widely

accepted since it was more versatile and offered a large range of research that could not

be readily undertaken with the more complex organisms. Other advantages include ease

of genetic techniques, the ability of bacteria to grow via glycolysis, which allowed

characterization of defective F1Fo ATP synthases, and ease of large-scale purification

procedures due to a practically unlimited supply of bacteria.

F1Fo ATP Synthase Mechanism

The overall function of F1Fo ATP synthase can be divided into three distinct parts:

proton conduction, coupling, and catalysis. The a and c subunits of Fo are responsible for

the translocation of protons through the membrane. The Fl y and a subunits of the rotor

stalk are responsible for coupling the energy acquired from the proton gradient to the Fl

catalytic sites. And the three catalytic sites located at the interfaces of the three oc and P









subunits are responsible for the synthesis of ATP or sometimes, as in the case of bacteria,

ATP hydrolysis. All three functions must be tightly integrated for the production of ATP.

Proton Translocation: Driving Rotation

The demonstration that the electrochemical gradient of protons drives the rotation

of bacterial flagella (245) in combination with Peter Mitchell's chemiosmotic theory (2)

began the search for evidence of rotation in F1Fo ATP synthesis. At the same time, a

model for proton transport was suggested by Cox et al. (212) and Boyer developed his

ideas for the binding change mechanism (discussed below) (246). But an indication of

rotational catalysis was not evident until the high-resolution crystal structure of F1

became available (20). This was followed a few years later by the first direct observation

of rotation when Noji et al. fixed the top of F1 to a glass coverslip and attached a

fluorescently labeled actin filament to the y subunit (15). Upon addition of ATP, rotation

of the actin filament was observed under an optical microscope at 0.2-10 revolutions per

second. At low concentrations of ATP (<600 nM), the actin filaments were observed to

rotate in a step-wise manner at 1200 intervals, which reflects the three catalytic sites in

the F1 003 3 hexamer (247). Experiments, in which two phenylalanine residues in the

nucleotide binding pocket were mutated to reduce the binding affinity of ATP, indicated

that the binding and hydrolysis of ATP is initially accompanied by a 900 substep

followed by a 300 substep attributed to product release (248, 249). These observations

were consistent with the two proposed "catches" observed between the P and y subunits

in the high resolution structure (see "The y subunit:" above) (20). The observance of the

rotation of the s and c subunits at the same speed and direction, indicating that these three

subunits rotate in synchrony, forming the central rotary machinery of the enzyme










complex (65-67). The concept of rotational catalysis with the rotation of the Yscio

subunits relative to the OC3 3 hexamer is now well accepted.

Several models have been proposed for proton translocation. One of the earliest

models suggested a series of side chains spanning the lipid bilayer formed a "proton

wire", involving amino acid residues casp61, aarg210, aglu219 and ahis245, 18 Which the protons

"hop" from one side chain to another until it passes through the membrane (212, 226,

250). Other models include a water-filled "proton channel" model, formed by the

charged residues of the a and c subunits, in which hydronium ions (H30 ) pass through

the lipid bilayer (251) and a "proton carrier" model, in which a proton binds on the

exterior of the membrane followed by a conformational change that brings the complex

through the membrane and releases the proton on the outer surface (252, 253).

The current prevailing model suggests that protons, in the form of H30 enter a

half channel created by the a subunit (Figure 1-9) (254, 255). The H30' is then believed

to protonate one of the clo subunits at residue casp61, which is positioned near the center of

the lipid bilayer (131). Besides forming the proposed half channel, the a subunit is

thought to play another crucial role in the protonation of the casp61. The pKa of the casp61

carboxyl side chain is uncommonly high, which is likely due to its lipid environment

(125). The essential aarg210 TOSidue is thought to facilitate a pKa shift of the casp61

carboxyl side chain to a lower pKa form during proton translocation (14, 132). Either the

protonation of the carboxylate, or possibly the release of a proton from a previously

protonated casp61 into an exit channel housed by the a subunit, somehow promotes the

generation of torque (Figure 1-9). The torque produced by proton translocation is

believed to drive the rotation of the ring of clo subunits relative to the a and b subunits











during rotational catalysis. Translocation of three to four protons generates enough


torque energy to rotate the clo ring by 1200 and results in the synthesis of a single

molecule of ATP.



~'-;
r


H' 10O
H'


H* H'


H' H'


Figure 1-9. Model of proton translocation and torque generation in Fo. Subunits included
are y (green), E (orange), a (yellow), b2 (purple), clo (blue). Protons (red) are
traveling in the direction of ATP synthesis. Protons are believed to travel
through a half channel housed in the a subunit as hydronium, H30 An
essential residue located near the center of the lipid bilayer in the c subunit,
casp61, iS thought to be protonated as another proton exits through another
"exit" half channel located on the cytoplasmic side of the membrane. The
protonation/deprotonation drives the rotation of the clo ring. The model was
drawn from schemes proposed by Junge (254) and Vik and Antonio (255).









Coupling

In F1Fo ATP synthase, the mechanism of energy coupling requires both the rotor

stalk and the peripheral stalk. Rotation of the ring of clo subunits consequently results in

the rotation of the entire rotor stalk, which is essential in coupling the energy obtained

from proton translocation to the synthesis of ATP in the catalytic sites of the u3 3

hexamer located over 100 A+ away. The role of the peripheral stalk is to hold the hexamer

in place while the amino- and carboxyl-terminal a-helices of the y subunit rotates within.

In a fully functional and coupled enzyme, the kinetics of proton translocation and ATP

synthesis are linked so that one cannot proceed without the other, and vise versa (256).

Mutational studies suggested that the polar loop of the clo subunits was involved in

the coupling function. In F1Fo ATP synthases incorporated with the cgln42 glu Subunit

mutant, Fl was found to bind normally to the Fo mutant, but the passive leakage of

protons through this complex was not prevented as in must be in coupled enzyme

complexes (257). Also, ATP was hydrolyzed normally by this mutant, but hydrolysis

was not coupled to active proton translocation. A similar uncoupled phenotype was

found in complexes incorporated with a carg41 lys mutant (258). Another mutation at the

same site, carg41 his, WaS found to prevent the binding of F1 to Fo. Thus, this loop region

in the c subunit appears to play essential roles in both the binding of F1 and the coupling

of proton translocation to ATP synthesis. Second site suppressor mutations to the

cgln42 glu mutation were found in the a subunit, specifically, Sglu31 gly,val,1ys, that recouped

proton translocation and ATP synthesis (98). Crosslinking studies of cysteine double

mutants found cross-linked products between Scys31 and ccys40, Ccys42 and ccys43 (99).

Moreover, a functional contribution of the y subunit in energy coupling was demonstrated










by y subunit mutants that uncoupled proton transport and ATP hydrolysis (259). Second

site suppressor mutations were found in other regions of the y subunit (260, 261).

Cysteine substitutions formed crosslinks between the y subunit, cys205o, and ccys40, Ccys42

and ccys43 (118). Crosslinking studies also provided evidence for an interaction between y

and p (75, 77). In addition, the a subunit has been cross-linked to both the Fl and Fo

subunits, via introduced cysteines, indicating that it spans the entire length of the stalk

with the y subunit (60, 99). Combined, these observations had suggested that the

coupling mechanism occurred by direct interactions of the c subunit loop regions and the

y and a rotor stalk of F1, which convey the proton gradient energy to the catalytic sites,

probably by direct interactions.

The crystal structure later confirmed these interactions (20). The crystal structure

displayed a strikingly asymmetrical Fl due to differences in the domains of the a and P

subunits and the interactions formed with the single y subunit (20). The obvious

asymmetric positioning of the coiled coil of the y subunit is a key feature to the

mechanics of the binding change mechanism (discussed below) of F1Fo ATP synthase.

Its large carboxyl terminus a-helix passes through a hydrophobic sleeve formed by six

proline-rich loops of the a and P subunits, presumably resulting in the conformational

changes occurring in the catalytic sites (20). In the PE Subunit (see above), several

hydrogen bonds are formed with the y subunit which forms a "catch", resulting in

conformational changes. Specifically, residues yarg254 and ygln255 in the carboxyl terminal

helix form hydrogen bongs with PE-asp317, PE-thr318 and PE-asp319. Also, a second "catch" is

formed between the carboxyl terminal domain of the PT Subunit and the short helix of the










y subunit. Hydrogen bonds form between ylsss, Yys,90 and Yalaso within the PDELSEED

region, PT-asp394 and PT-glu398. Structural information suggests the two antiparallel coiled

coil a-helices of the y subunit may unwind during rotational catalysis and the a subunit

rotates around the Fl axis while undertaking a net translation of about 23 A+ (85). It is

likely that these gross changes observed in the structures revealed individual functional

states of the enzyme complex during catalysis.

Catalysis: The Binding Change Mechanism

F1Fo ATP synthases house three catalytic sites, located at the u3 3 interfaces, with

the predominate sites positioned in the P subunit and some contributions made by the a

subunits (20). The minimal complex capable of normal ATP hydrolysis activity is the

a3 37 complex (262). Boyer predicted that ATP synthesis requires a chronological

involvement of the three catalytic sites, each of which changes its binding affinity for the

substrates and products as it continues through a cyclical mechanism, referred to as the

"binding change mechanism" (246, 263). The mechanism of ATP synthesis, in terms of

the u3, P3 and Y subunits and the substrates, ADP and Pi, is described here (Figure 1-10).

The principles of the binding change mechanism have become the most commonly

used model for recounting the means of ATP synthesis by F1Fo ATP synthase. The three

distinct catalytic sites were described as the tight (T) site containing ATP, an empty open

(0) site, and the loose (L) site illustrated with ADP and Pi bound (Figure 1-10).

According to Boyer, catalysis starts with the binding of ADP and Pi at the open site. The

energy input from proton translocation drives the rotation of the rotor stalk, which

ultimately results in the conformational changes responsible for ATP synthesis such that

the tight site is converted to an open site, the open site assumes a loose site conformation,









and the loose sites becomes a tight sight. ATP is formed in the new tight site and the

molecule of ATP that was found in the original tight site is released and the binding

change mechanism starts fresh (Figure 1-10). Boyer's mechanism included three

proposals: i) only one site is actively synthesizing ATP at any given moment, ii) the

reaction occurs reversibly at this site, and iii) energy input is required to bind the

substrates, ADP and Pi, into the catalytic sites and to release the synthesized ATP, but not

for the actual reaction to occur. Strong evidence for this model has come from the crystal

structure of Fl (20) and the observance of rotation via an attached fluorescent actin

filament (discussed previously) (15, 65-67).



ATP Hydrolys s












ATP~ Syn~thesis



Figure 1-10. The binding change mechanism. This is a simplified model of a more
detailed enzymatic mechanism described by Weber and Senior (189) in which
all three catalytic sites are transiently filled with nucleotide during ATP
synthesis. Subunits included are oc (red), P (green) and y (pink).

Genetic Expression and Assembly

The E. coli F1Fo ATP synthase is encoded in the 7 kb unc operon, which was

cloned and sequenced in its entirety (224, 264). The genes encoded are called the uncB,










unZcE, uncF, uncH, uncA, uncG, uncD and uncC coding for the a, c, b, 6, a, y, P and a

subunits, respectively. A single copy of each gene is transcribed into a single

polycysternic mRNA transcript, from which multiple polypeptides can be translated

(265-267). The synthesis of the correct number of subunits, resulting in the

stoichiometry of u3 38ysab2C10, iS thought to occur by translational regulation (264, 268).

The efficiency at which the individual subunits are synthesized were observed to be

variable and roughly corresponded to the stoichiometry of the intact enzyme complex

(269, 270).

Far less is understood about the assembly of the complex F1Fo ATP synthase

enzyme compared to the structural and mechanistic studies. Some have proposed that no

particular pathway is necessary based on the observation that the complex can be

dissembled into individual subunits and then reconstituted in vitro (113, 148, 271). On

the contrary, some believe that the assembly follows an integrated pathway in vivo (272).

The idea of an assembly pathway appealed to many since it would prevent a newly

assembled Fl sectors from freely hydrolyzing cellular ATP in the cytoplasm, and newly

assembled Fo sectors from acting as open proton pores in the membrane. If assembly

were a random event, the potential to create isolated Fl and Fo sectors would exist. Some

evidence supporting the integrated pathway does exist. The a subunit is thought to

function as an inhibitor of ATP hydrolysis activity, undertaking large conformational

changes to allow the enzyme to switch from ATP synthesis to ATP hydrolysis under

conditions of low ATP or low proton gradient, respectively (102, 177, 273-276). Its

inhibitory action may possibly act to inhibit free ATPase activity in the partially

assembled state. Some have speculated that the binding of the Fl subunits to Fo may









influence the opening of the Fo proton channel in vivo (176, 277-279). The a subunit was

observed to be absent from membranes of cells lacking the b or c subunits (150).

Furthermore, work accomplished in our laboratory has showed that the b subunit

monomer does not integrate into the membrane has no affinity for Fl, indicating that the

formation of the b dimer in the membrane is an early event in F1Fo ATP synthase

assembly (199, 203, 218).

Summary

There is now ample evidence indicating that F1Fo ATP synthases are composed of

three functional parts, the catalytic core, the rotor stalk and the stator stalk. In the E. coli

enzyme complex, the catalytic core consists of the u3 3y Subunits and functions as an

ATP synthase or an ATPase. The rotor stalk consists of the ysclo subunits and couples

the energy of proton conduction to the synthesis of ATP by rotating within the u3 3

hexamer. Finally, the stator stalk consists of the Sab2 Subunits and remains in a fixed

position, anchored to the membrane by the a and b2 Subunits and to the u3 3 hexamer via

interactions made by the 6 subunit. Much of what was known of F1Fo ATP synthase has

been irrefutably confirmed by high resolution structures of partial complexes or model

polypeptides. To date, a high resolution structure of the complete enzyme, or at least the

complete Fo is eagerly anticipated.

Since the visualization of the peripheral stalk less than a decade ago a plethora of

data characterizing the b2 homodimer has emerged. The observation that the b2

homodimer was likely not a rigid structure, and possibly more of an elastic structure,

created the foundation for the work described in the following chapters. The work

illustrated in Chapters 2, 3, 4, 5 and 6 of this dissertation will characterize the role of the










peripheral stalk' s dimer of identical b subunits in the E. coli F1Fo ATP synthase by using

a combination of site-directed mutagenesis and biochemical methods. Chapter 2

demonstrates the ability of the E. coli b subunit to form heterodimers and the capability

of F1Fo ATP synthase complex to tolerate the incorporation of two different length b

subunits with a size difference of at least 14 amino acids (195). Chapter 3 demonstrates

the formation of b heterodimers including at least one and up to two defective b subunits

and documents indisputable evidence that F1Fo ATP synthases incorporated with b

subunit heterodimers are functional (280). Furthermore, the work accomplished in the

chapter indicates, for the first time, that each of the two b subunits makes a unique

contribution to the functions of the peripheral stalk, such that one mutant b subunit is

making up for what the other is lacking. Chapter 4 describes cysteine chemical

modifications constructed in the 6 subunit and shortened, lengthened and wild-type

length b subunits. The unc operon expression plasmids generated in this study will be

used in future fluorescent labeling experiments. Chapter 5 documents mutagenesis

experiments conducted on the extreme amino- and carboxyl termini of the b subunit

(202) (Bhatt et al., manuscript in preparation, 2004). Finally, Chapter 6 summarizes the

conclusions of this study and suggests the future directions that the work described in this

dissertation has offered.















CHAPTER 2
INTEGRATION OF UNEQUAL LENGTH b SUBUNITS INTO F1Fo ATP SYNTHASE

Introduction

F1Fo ATP synthases provide the bulk of cellular energy production in both

eukaryotes and prokaryotes (3, 5, 6). Enzymes in this family utilize the electrochemical

gradient of protons across membranes in order to synthesize ATP from ADP and

inorganic phosphate in a coupled reaction (16). In Escherichia coli, F,Fo ATP synthase is

a complex enzyme composed of approximately twenty-two polypeptides with the

stoichiometry of u3 3y~sab2C10 (6, 7). The F, portion is composed of the subunits

a3 3yis and is responsible for enzymatic catalysis. The Fo portion of the enzyme consists

of the ab2C10 Subunits and is responsible for the translocation of protons through the

membrane.

Electron microscopy has shown that the Fl and Fo sectors are linked by two slender

stalk structures (11). During ATP synthesis proton translocation drives the rotation of the

central stalk, which consists of subunits ys, within the u3 3 hexamer held stationary by

the peripheral stalk. This rotation propagates the conformational changes in the active

sites located at the up interfaces driving catalytic activity (3, 15, 62, 65, 281, 282). The 6

subunit of Fl and a dimer of two identical b subunits from Fo comprise the peripheral

stalk acting as the stator. The 6 subunit has been visualized seated at the top of the Fl

a3 3 hexamer (187). However, recent evidence has suggested that the 6 subunit may be

positioned slightly to the side of F1 in association with a single a subunit (100, 141-143).









The C-terminal region of the 6 subunit is in direct contact with the extreme C-terminal

end of the b dimer (3, 185, 200, 219, 220). The b subunit dimer constitutes the majority

of the peripheral stalk stretching from within the membrane to near the top of F1 (12).

Dimerization of the b subunits is required for the normal assembly and function of

F1Fo ATP synthase (199). The two b subunits are believed to exist in parallel as an

extended structure spanning from the periplasmic side of the membrane to near the top of

F1. Each has a N-terminal transmembrane domain, a tether domain extending from the

surface of the membrane to the bottom of F1, a dimerization domain and a 6-binding

domain (13). The ability of b to bind to F1 was proportional to the ability of b to form

dimers, suggesting the necessity of the b dimer formation before the binding of F1 to the

complex (199). Presently, there is no high-resolution structure of the entire b subunit. A

model polypeptide of the first 34 residues of the N-terminus has been solved by NMR,

revealing a hydrophobic membrane-spanning a-helix (136). A crystal structure of a

monomeric dimerization domain, consisting of residues 62-122, has been solved and

refined to 1.55 A (138). Dunn and coworkers have constructed a model in which the two

a-helices of the b62-122 TegiOn form a right-handed coiled coil. Much of the structural

information on the b dimer has been gleaned from classical biochemical approaches such

as CD-spectroscopy, crosslinking and sedimentation experiments (30, 166, 192, 196, 197,

203, 283). These studies revealed that the overall structure of the b subunit dimer is a

highly extended conformation with approximately 80% a-helix.

Previous studies have shown that b subunits with deletions of up to eleven amino

acids and insertions of up to fourteen amino acids, corresponding to approximately 16 A~

and 21 A+, respectively, formed functional FiFo complexes (193, 194). When b subunits










with either a seven amino acid deletion or an insertion, b or b 7, respectively, were

incorporated into the F Fo ATP synthase complex, the properties of the enzymes were

essentially wild type. These observations suggested that the role of the b dimer is more

of a flexible structural feature. However, it was not known whether this flexibility

extended to the dimerization of two b subunits of unequal lengths and their incorporation

into an enzyme complex. In the present study an experimental system was developed to

allow expression of two different b subunit genes and determine whether the differing b

subunits were assembled into an F1Fo ATP synthase complex. Here, we demonstrate that

the F1Fo ATP synthase complex can tolerate b subunits with a size difference of at least

14 amino acids.

Materials and Methods

Materials

Molecular biology enzymes and mutagenic oligonucleotides were obtained from

Invitrogen (Carlsbad, CA), Life Technologies, Inc. (Grand Island, NY), New England

Biolabs (Beverly, MA) and Stratagene (La Jolla, CA). Reagents were obtained from

Sigma (St. Louis, MO), BioRad Laboratories (Hercules, CA) and Fisher Scientific

(Pittsburgh, PA). Plasmid purification kits were acquired from Qiagen Inc. (Valencia,

CA). The anti-rabbit immunoglobulin horseradish peroxidase-linked whole antibody

(from donkey), anti-mouse immunoglobulin horseradish peroxidase-linked whole

antibody (from sheep), Hybond ECL Nitrocellulose membrane,

electrochemiluminescence Western blotting reagents and high performance

chemiluminescence film were purchased from Amersham Biosciences (Piscataway, NJ).

Polyclonal antibodies against SDS-denatured b subunit (284, 285) were generously









provided by Dr. Karlheinz Altendorf (Universitait Osnabriick, Osnabriick, Germany).

Mouse monoclonal antibodies against the peptide epitope of hemagglutinin protein of

human influenza virus (HA epitope tag) were purchased from Roche Molecular

Biochemicals (Indianapolis, IN). Monoclonal antibodies against the epitope found in the

P and V proteins of the paramyxovirus, SV5 (V5 epitope tag) were purchased from

Invitrogen .

Strains and Media

The bacterial strains used to create the epitope tagged b subunits include the wild

type b subunit expression plasmid, pKAM14, and plasmids used to express b subunits

shortened or lengthened by 7 amino acids, pAUL3 and pAUL19, respectively, and have

been described previously (193, 194, 203). The plasmids encoding the different uncF(b)

genes were used to compliment E. coli strain KM2 (Ab) carrying a chromosomal deletion

of the gene (218). All strains were streaked onto plates containing Minimal A media

supplemented with succinate (0.2% w/v), to determine enzyme viability. Cells harvested

for membrane preparation were grown in Luria Bertani media supplemented with glucose

(0.2% w/v) (LBG). Isopropyl-1 -thio-P-D-galactopyranoside (IPTG)(40 Clg/ml),

ampicillin (Ap) (100Clg/ml), and chloramphenicol (Cm) (25 Clg/ml) were added to media

as needed. All cultures were incubated at 37oC for the appropriate duration.

Recombinant DNA Techniques

Plasmid purification. Plasmid DNA was purified with the Qiagen Mini-Prep and

Maxi-Prep kits according to the protocols provided from the manufacturer. Mini-preps

required 4 mL (high copy plasmid) or 6 mL (low copy plasmid) of an overnight bacterial

culture carrying the desired plasmid. Maxi-preps required a 500 mL culture grown










overnight. Final elution volumes for the mini-prep kit was 30 or 50 C1L, for low copy and

high copy plasmids, respectively, and 200-250 C1L for the maxi-prep kit. Final

concentrations of 0.5 and 1.0 Clg/CIl plasmid were routinely obtained with the Qiagen mini

and maxi-prep kits.

Digestions, ligations, and transformations. Restriction endonuclease digestions,

ligations, and transformations were performed according to the recommendations of the

manufacturers (New England Biolabs, Stratagene and Invitrogen). For analytical

purposes, restriction endonuclease digestions were normally prepared in a total volume of

20 CIL, including plasmid DNA, enzyme, buffer, ddH20 and occasionally BSA, and then

incubated for an hour at the temperature specified by the manufacturer. Ligations

required two purified double-stranded DNA fragments, a vector and an insert, of known

length and concentration (ng/C1L). DNA fragments were routinely separated in a 0.8 %

agarose gel by electrophoresis and the appropriate sized fragment was excised and

purified using a Qiagen, Inc. QIAquick Gel Extraction kit. The vector fragment

contained the antibiotic resistance gene and the origin of replication. The insert typically

contained the desired gene or a site specific mutation. Two control reactions and two

ligation reactions were set up. The typical reaction was set up in 20-40 CIL and included

vector, insert, ATP, T4 DNA ligase buffer, T4 DNA ligase and ddH20. The first control

reaction was a control for uncut plasmids, containing no insert and no ligase, and the

second, containing no insert, was a control for the vector' s ability to ligate with itself.

The femptomolar concentration (fmol/CIL) was determined from the known size and

measured concentration. Two ligation reactions were then set up, the first had 1 part

vector to 3 parts insert and the second was 1:10. Typically 5-15 fmol of vector was used.









The four reactions were incubated for 5 minutes at room temperature if T4 High

Concentration DNA ligase was used or overnight at 16 oC if T4 DNA ligase was used and

then transformed into competent E. coli. Transformations were performed in one of three

different E. coli strains. DH~a competent cells and XL10-Gold Ultracompetent cells

were competent bacteria purchased from Invitrogen and Stratagene. These bacteria were

used for purposes of plasmid preparations or to transform with a mutagenesis reaction

such as Quikchange or ligations. The DH~a and XL10-Gold bacteria were stored at -80

oC. Basically, 25 CIL pre-aliquoted cells were thawed on ice and 1-5 CIL plasmid DNA

was added and mixed by gentle stirring with the pipette tip. The bacteria were incubated

on ice for 30 minutes, heat shocked at 42 oC for 45 seconds, and then incubated on ice for

2 minutes. 1 mL LBG was added and then incubated at 37 oC taped to a roller drum. The

cells were harvested by centrifugation for 1 minute, the supernatant was discarded, the

bacteria were resuspended in the remaining media, spread onto a LBG plate

supplemented with the appropriate antibiotic and incubated at 37 oC overnight. XL10-

Gold cells required treatment with P-mercaptoethanol (P-ME) before transformation to

increase the efficiency. 2 C1L of the provided P-ME mix was added to 45 CIL pre-

aliquoted cells and incubated on ice for 10 minutes prior to transformation with gentle

swirling every 2 minutes. The transformation proceeded as described above. KM2 (Ab)

was a strain of E coli generated by a previous lab member (218) and maintained in our

laboratory. This strain was used for purposes of plasmid expression and the study of the

b subunit of F1Fo ATP synthase. Before transformation, KM2 had to be made competent.

Sterile technique was important since KM2 cells cannot be selected for by an antibiotic.

All reagents and supplies were pre-chilled unless otherwise noted. A culture of KM2










cells was grown overnight at 37 oC in 5 mL LBG. The overnight culture (100 CIL) was

inoculated into 10 mL pre-warmed (37 oC) LBG and allowed to grow for 2-4 hours. The

fresh culture was poured into sterile polypropylene tubes and incubated on ice for 10

minutes. The bacteria were then harvested by centrifugation (6,000 rpm in a ss-34 rotor)

for 10 minutes. The supernatant was discarded and the tubes were inverted on a

Kimwipe for 1 minute to allow excess LBG to drain. The bacteria pellet was

resuspended in 10 mL cold, sterile 50 mM CaCl2 and incubated on ice for 45-60 minutes.

The bacteria were harvested as described above, resuspended in 1 mL of the 50 mM

CaCl2, and stored at 4 oC until use. Generally, the competent KM2 cells could be used

after two hours on ice but were most efficient for transformation at 24 hours and expired

at 72 hours. For transformation, 100-200 CIL of cells were used as previously described.

Site-directed mutagenesis. Site-directed mutagenesis was performed either by

means of a Stratagene Quikchange XL kit or by ligation-mediated mutagenesis.

Oligonucleotides containing the desired mutation(s) were designed to anneal to the same

sequence on opposite strands of the plasmid (sense and antisense primers) (Appendix A)

(Figure 2-1). When possible, a silent mutation was encoded to add or delete a restriction

endonuclease recognition sequence to allow for easy screening of the mutation. Primers

were optimally designed by ensuring the mutation was in the middle of the sequence, a

cytosine (C) or guanine (G) flanked both ends of the sequence, and the melting

temperature (Tm) was greater than or equal to 78 oC. The Tm was calculated as

Tm=81.5+0.41 (%GC)-675/N-%mismatch, where N was the primer length (bases). When

introducing insertions or deletions, "%mismatch" was dropped from the formula.










Mutagenic Primers (sense strand)

A
histidine tag Sphl H H H
5' CTAAATAGAAGCATGCTGCTGTGCACCACCAC
HHH
CACCACCACAATC-TTAACGCAcACAATCCTCGGC 3'


Ndel
HA tag Y P Y D V
5' CTAAATAGAGGCATTGTGCTATGTACCCATATGAC GTG
P D Y A
C CGGAC TACGCGAcATC-TTAcACGCAAICAATCCTCGGC 3'



V5 tag SacI G K P I P N P
5' CGTGGATAAAC-TTGTCGCTGAGCTC GGTAAAC CGATCC CGAAC CCG
L L G L D S
CTGCTGGGTCTGGACTCTAC CTAAGGAGGGAGGGGCTGATGTCT 3'

B

+ Sphl site Sphl
5' CTGTAAGGAGGGAGGGGCATGC GTCTGAATTTTA-TTACG 3'



-Sphl site +ATG H H H
5' GGAGGGAGG GGCTGATGCACCACCAC 3'

Figure 2-1. Oligonucleotides for epitope tags and mutagenesis of uncF(b). Shown are the
sense strands of the mutagenic primer pairs. Green and red codons specify
translation start and stop sites, respectively. A) Mutagenic primers encoding
the sequence of the desired epitope tag insertion along with a silent mutation
that introduced a new endonuclease recognition sequence. Epitope tags were
inserted as described in the Materials and Methods. The oligonucleotides
sequences specifying the histidine, HA or V5 epitope (bold blue) tag are
labeled with the corresponding amino acid. The Sphl, Ndel and SacI
restriction sites (underlined) were added along with the histidine, HA and V5
tags, respectively, to facilitate screening. B) Oligonucleotides designed to
construct the one-plasmid expression system.









Primers were not fast polynucleotide liquid chromatography (FPLC) or polyacrylamide

gel electrophoresis (PAGE) purified as called for in the protocol. The reaction mixture

consisted of 5 CIL Stratagene's 10X Pfu buffer, 3 CIL QuikSolution, 50 ng wild type

plasmid, 125 ng each of the sense and antisense oligonucleotides, 1 C1L of 25 mM

dNTP's, and 1 unit Pfu turbo polymerase. The total volume was brought up to 50 C1L

with ddH20 and PCR was performed in a Perkin Elmer GeneAmp PCR System 2400

thermocycler according to the following cycling parameters: 950C 1 minute presoak; 18

cycles of 950C 50 seconds, 600C 50 seconds, 680C 1 minute per kb of plasmid length;

40C 7 min. Upon completion of the thermocycling, 1 C1L DpnI restriction endonuclease

was added to the reactions in order to digest the methylated (nonmutated) plasmid DNA.

The plasmids carrying the desired mutation were then transformed into competent DH500

cells, purchased from Life Technologies, and grown on LBG plates supplemented with

the appropriate antibiotic. Plasmids carrying the desired mutation(s) were screened for

by restriction endonuclease analysis and then the nucleotide sequences were directly

determined by automated sequencing in the core facility of the University of Florida

Interdisciplinary Center for Biotechnology Research (ICBR).

Mutagenesis and Strain Construction

Plasmids pKAM14 (b, Apr) (203), pAUL3 (ba7, Apr) (194) or pAUL19 (b 7, Apr)

(193) were used to construct the epitope tagged b subunits. Epitope tags were inserted

into each of the plasmids using the Stratagene Quikchange kit. A histidine epitope tag

was inserted at the N-terminus by mutagenesis between the first and second codons of the

uncF(b) gene to express bhis or b+7-his (Appendix A) (Figure 2-1). Plasmids pTAM37

(bhis) and pTAM3 5 (b+7-his) were created by digesting both of the recombinant histidine-









tagged b subunit plasmids with PstI and Ndel and subsequently lighting the genes into a

plasmid conferring the chloramphenicol resistance gene and the pACYC 184 origin of

replication (Table 2-1). Likewise, an HA epitope tag was inserted at the N-terminus by

mutagenesis between the first and second codons of the uncF(b) gene to generate

plasmids pTAM36 (bHA) Or pTAM34 (bA7-HA). A V5 epitope tag was added to the C-

terminus by site-directed mutagenesis before the termination codon of the uncF(b) gene

to express bvS or ba7~vs from plasmids pTAM46 and pTAM47 (Figure 2-1). The

recombinant HA-tagged and V5-tagged b subunit plasmids included the ampicillin

resistance gene and the pUC18 origin of replication. Unique restriction enzyme sites

Sphl, Ndel and SacI were constructed near the histidine, HA and V5 epitope tag

sequence, respectively, for an initial detection of the insertions, and then the nucleotide

sequence was subsequently confirmed by automated sequencing in the ICBR core

facility. Additionally, a set of plasmids was designed in order to express two different-

tagged b subunits from a single transcript (Figure 2-2). As an example, a unique

restriction enzyme site, Sphl, was created in conjunction with the histidine tag between

the Shine Dalgarno sequence and the first codon of uncF(b) to create pTAM3 5 (Figure 2-

2A). In a separate site-directed mutagenesis reaction, an Sphl site was added to the

pTAM34 plasmid downstream of the HA-tagged b subunit and behind another favorable

Shine Dalgarno sequence (Figure 2-2B). The two plasmids were digested with Sphl and

BstEII restriction endonucleases. The 3.2 kb vector fragment from pTAM34 and the 623

bp insert fragment from pTAM3 5 were isolated and then excised from a 0.8 % (w/v)

agarose gel and purified with a QIAquick gel extraction kit. The vector and insert

fragments were then allowed to ligate overnight at 16 oC. It was then crucial to mutate an









A ori B
Cm' L 4
Sph| HA tag


pTAM34
pTAM35 t
6.9 kb I~H) SD~td~h
(b+7-hi SD Ofl (b-y+AdSh





Diplest
Sph|



Vector Insert
3 1 kb Ligate 623 bp


C HA tag
rp SD b,


pTA M40
3.7 kb SD
Sph|
(bH -~ _HA & +7-his





Figure 2-2. Construction of the single transcript expression system. A) A unique
restriction enzyme site, Sphl, was created in conjunction with the histidine tag
between the Shine Dalgarno sequence and the first codon of uncF(b) to create
pTAM3. B) An Sphl site was added to pTAM34 downstream of the HA-
tagged b subunit and behind another favorable Shine Dalgarno sequence. The
two plasmids were digested with Sphl and BstEII. The 3.2 kb vector fragment
from pTAM34 and the 623 bp insert fragment from pTAM3 5 were ligated.
Additional mutagenesis (see Materials and Methods) resulted in C) a 3.7 kb
plasmid, pTAM40, which expressed bA7-HA and b+7-his from the same promoter
and included the ampicillin resistance gene and the pUC 18 origin of
replication. In similar constructions, plasmids pTAM41 (bwt-HA and b+7-his)
and pTAM42 (bwt-HA and bwt-his) were created.









intrinsic added start codon, which is in the Sphl recognition sequence, to prevent a

missense mutation. The mutagenic oligonucleotide was designed to accomplish three

tasks in one reaction: 1) mutate the "ATG" found in the Sphl recognition sequence, 2)

add a new Shine Dalgarno sequence in a favorable position from the true start codon, and

3) mutate the "GTG" start codon to a more favorable "ATG" start site (Figure 2-1). The

resulting 3.7 kb plasmid, pTAM40, expressed bA7-HA and b+7-his from the same promoter

and included the ampicillin resistance gene and the pUC 18 origin of replication (Figure

2-2C). In similar constructions, plasmids pTAM41 (bwt-HA and b+7-his) and pTAM42 (bwt-

HA and bwt-his) were created (Table 2-1). Throughout this dissertation, the insertion or

deletion and the epitope tag are indicated after the plasmid name for clarity, for example

plasmid pTAM35 (b+7-his). Each plasmid and the control plasmids pKAM14 (b) and

pBR322 were expressed in the E. coli cell line KM2 (Ab) for study, so that the only b

subunits in the cells were the product of the plasmid genes.

The two plasmid expression system successfully allowed expression of various

combinations of histidine tagged and V5-tagged b subunits in the same cell (Figure 2-7).

Appropriate antibiotics were added to the growth medium, and in the case of the

coexpressed plasmids, both ampicillin and chloramphenicol were added to select for cells

expressing both plasmids.

Crude Preparative Procedures

Inverted membrane vesicles from KM2(Ab) strains expressing the desired b subunit

epitope tagged F1Fo ATP synthase complex were prepared for activity assays, Ni-resin

purification and Western Blot analysis. Unless otherwise noted, all reagents, rotors and

materials were kept at 4 oC. The membrane preparations were prepared by inoculating a









10 mL starter culture, grown overnight, into a 2 L Erlenmeyer flask containing 500 mL

LBG, supplemented with the appropriate antibiotic, ampicillin (Ap) and/or

chloramphenicol (Cm). Similarly, 1 mL of a starter culture was inoculated into a nephalo

flask containing 50 mL LBG (Ap and/or Cm). The bacteria were grown at 37 oC in a

New Brunswick Scientific incubator shaker (220 rpm) and the turbidity was monitored

using a Klett-Summerson photoelectric colorimeter. IPTG (40 CIM) was added when the

turbidity reached 75 Klett units and the cells were collected when the turbidity reached

150 Klett units. The bacteria were harvested by centrifugation for 10 minutes at 8,000Xg

in a Sorvall GSA rotor. The pellets were rinsed once with TM buffer (50 mM tris-HC1,

10 mM MgSO4, pH 7.5) and then resuspended in a final volume of 10 mL TM buffer.

DNasel (10 mg/mL) was added to a final concentration of 10 Clg/mL and the bacteria

were broken by passing through a French Pressure Cell one time at 14,000 psi. Cellular

debris and unbroken cells were removed by centrifuging twice at 10,000Xg for 10

minutes. Membranes were then collected by ultracentrifugation at 150,000Xg in a

Beckman 70.1 Ti rotor for 1.5 hours. The membrane pellets were rinsed once with TM

buffer and then resuspended in TM buffer to a final volume of 2 mL using a 2 mL

Wheaton tissue grinder. For the purposes of Western blot analysis or Ni-resin

purification, one ultracentrifugation step sufficed. However, activity analysis required an

additional ultracentrifugation step in order to remove nonspecifically bound ATPases. In

this case, the membrane pellets were resuspended in a final volume of 10 mL using a 10

mL Wheaton tissue grinder and the ultracentrifugation step was duplicated.