Binding of the Nonnucleoside Reverse Transcriptase Inhibitor

Efavirenz to HIV-1 Reverse Transcriptase Monomers and Dimers

by

Valerie Ann Braz

Submitted in partial fulfillment of the requirements

for the degree of Doctor of Philosophy

Thesis Advisor: Mary D. Barkley

Department of Chemistry

CASE WESTERN RESERVE UNIVERSITY

January 2010

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

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candidate for the ______degree *.

(signed)______(chair of the committee)

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*We also certify that written approval has been obtained for any proprietary material contained therein.

TABLE OF CONTENTS

Table of Contents 1

List of Tables 4

List of Schemes 5

List of Figures 6

Abstract 8

Acknowledgements 10

List of Abbreviations 11

Chapter 1: Introduction

1.1 HIV and AIDS 15

1.2 HIV 16

1.2.1 HIV Retrovirus and Dynamics 16

1.2.2 HIV Structure and Biology 18

1.2.3 The HIV Lifecycle 24

1.2.4 HIV-1 Genetic Diversity 27

1.3 HIV Reverse Transcription 28

1.4 HIV RT 31

1.4.1 HIV RT Inhibitors 33

1.5 Research Techniques 39

1.5.1 Equilibrium Dialysis 39

1.5.2 Fluorescence Spectroscopy 41

1.5.3 Mass Spectrometry 44

1 1.6 Research Objectives 50

1.7 References 55

Chapter 2: Efavirenz Binding to HIV-1 Reverse Transcriptase Monomers and Dimers

2.1 Abstract 65

2.2 Introduction 66

2.3 Experimental Procedures 70

2.4 Results 74

2.5 Discussion 98

2.6 Appendix 106

2.7 Acknowledgements 109

2.8 References 110

Chapter 3: The Efavirenz Binding Site in HIV-1 Reverse Transcriptase Monomers

3.1 Abstract 118

3.2 Introduction 119

3.3 Experimental Procedures 124

3.4 Results 126

3.5 Discussion 127

3.6 References 143

Chapter 4: Separation of Protein Oligomers by Blue Native Gel Electrophoresis

4.1 Abstract 148

4.2 Introduction, Results, Discussion 149

4.3 Acknowledgements 155

4.4 References 157

2 Chapter 5: Conclusions and Future Directions

5.1 Conclusions and Future Directions 159

5.2 References 163

Appendix: Equilibrium Dialysis Data 164

3 LIST OF TABLES

Table 1-1. Function of Accessory/Auxiliary Proteins 22

Table 2-1 Equilibrium Dialysis Dissociation Constants 80

Table 2-2 Kinetic Parameters of Efavirenz Binding 91

Table 2-3 Quantum Yields of Efavirenz Binding 95

Table 3-S1 Sequence and Residue Number of Analyzed Peptides 141

4 LIST OF SCHEMES

Scheme 1-1. Thermodynamic Linkage of NNRTI Binding and Dimerization 51

Scheme 2-1 Thermodynamic Linkage of Efavirenz Binding and Dimerization 68

Scheme 2-2 Mechanisms of Slow Binding Inhibitors 86

5 LIST OF FIGURES

Figure 1-1 The HIV-1 Virion 17

Figure 1-2 Organization of the HIV-1 Genome 19

Figure 1-3 Steps of Viral Infection 25

Figure 1-4 Model of Reverse transcription 29

Figure 1-5 Structure of HIV-1 RT 32

Figure 1-6 Structure of NRTIs 34

Figure 1-7 Structure of HIV-1 RT─NNRTI Complex 37

Figure 1-8 Structure of NNRTIs 38

Figure 1-9 Principle of Equilibrium Dialysis 40

Figure 1-10 The Jablonski Diagram 43

Figure 1-11 Standard tandem mass spectrometry experiment (MS2) 47

Figure 1-12 Schematic of Hydrogen/Deuterium Exchange 49

Figure 1-13 Schematic of RT Species in Solution 54

Figure 2-1 Equilibrium dialysis data for wild-type p51 and p51W401A 77

Figure 2-2 Structure of HIV-1 RT complexed with efavirenz (1FK9) 82

Figure 2-3 Association of efavirenz to p66/p51 and p51 84

Figure 2-4 Progress curves for p51W401A binding efavirenz 88

Figure 2-5 Dependence of kobs on efavirenz concentration 90

Figure 2-6 Dissociation of p51W401A─ and p66W401A─EFV complexes 93

Figure 2-7 BN-PAGE of p66 ± NNRTI 98

Figure 2-8 Structures of HIV-1 RT (1DLO) and (1FK9) 104

Figure 3-1 Crystal structure of unliganded HIV-1 RT (1DLO) 121

6 Figure 3-2 Percent exchange of peptides in p66W401A monomer ± efavirenz. 127

Figure 3-3 Difference in number of deuteria exchanged in bound and unbound

p66 and p51 129

Figure 3-4 Mass spectra of peptide 232–246 in p51W401A and p51W401A─EFV 131

Figure 3-5 FT-ICR mass spectra of p66W401A, p51W401A, and p51W401A─EFV 132

Figure 3-6 Five peptides stabilized in monomer─EFV complexes 138

Figure 3-S1 Percent exchange of peptides in p51W401A ± efavirenz 142

Figure 4-1 Migration of HIV-1 p66 and p51 by blue native electrophoresis 151

Figure 4-2 Migration of proteins by native agarose gel electrophoresis 153

7 Binding of the Nonnucleoside Reverse Transcriptase Inhibitor

Efavirenz to HIV-1 Reverse Transcriptase Monomers and Dimers

ABSTRACT

By

Valerie Ann Braz

Human immunodeficiency virus type 1 (HIV-1) reverse transcriptase (RT) was

the first target of antiretroviral therapy in the treatment of AIDS. RT converts single-

stranded viral RNA into double-stranded proviral DNA. The enzyme has two activities,

DNA polymerase and RNase H. The biologically relevant form is a heterodimer

composed of two subunits, p66 and p51. The subunits are products of the same gene and

have identical N-terminal amino acid sequences; p51 lacks the C-terminal RNase H

domain. In solution RT exists as a complex equilibrium mixture of p66/p51 heterodimer,

p66/p66 and p51/p51 homodimers, and p66 and p51 monomers. Two classes of inhibitors

have been developed and approved for clinical use, NRTIs and NNRTIs. NNRTIs are highly effective and relatively noncytotoxic small, amphiphilic, noncompetitive inhibitors that nestle into a hydrophobic pocket ~10 Å away from the polymerase active site in the p66 subunit of RT. NNRTIs also have diverse effects on RT subunit dimerization; some enhance dimerization and others weaken dimerization. Efavirenz is an NNRTI capable of affecting several steps in HIV-1 reverse transcription and replication. This work reports two novel functions of the inhibitor: (1) efavirenz, and presumably also other NNRTIs, binds to monomeric forms of RT and (2) efavirenz is a slow binding inhibitor of

8 heterodimer and monomers. In addition, the binding site on p66 and p51 monomers was identified. Five techniques were used to characterize the interaction of efavirenz to all species of RT; equilibrium dialysis, tryptophan fluorescence, native gel electrophoresis, and hydrogen-deuterium exchange and Fourier transform ion cyclotron resonance mass spectrometry. Although the biological significance of efavirenz binding to monomeric forms of RT is not known, this work suggests that monomeric forms of RT may be potential targets for HIV-1 therapeutics.

9 ACKNOWLEDGEMENTS

I begin by thanking my parents, Shirley and Charles Braz, for their love and countless support. To my best friend and sister, Darla, for helping me realize that I should go with the flow and things will (eventually) work out. And to my brother, Julian, for listening and teaching me to work “smarter, not harder”.

I thank my advisor, Dr. Mary D. Barkley, for her guidance and encouragement.

She has taught me many invaluable lessons of life and science. I must also thank the members of my thesis committee, Dr. James Burgess, Dr. Anthony Pearson, Dr. Clemens

Burda, and Dr. Patrick Wintrode.

To the members of the Barkley laboratory, Kathyrn Howard, Carl Venezia, and

Brendan Meany, I greatly appreciate your friendship and persistent knowledge. This is truly a group of wonderful thinkers with unconditional patience and kindness.

Finally, I would like to thank my better half for laughing on the good days and enduring the bad. Your support and encouragement has lasted before and after my time away.

“Facts are stubborn things;

and whatever may be our wishes, our inclinations,

or the dictates of our passions, they cannot alter the state of facts and evidence.”

-John Adams

10 LIST OF ABBREVIATIONS

A absorbance AIDS acquired immunodeficiency syndrome ANS 1-anilino-8-naphthalenesulfonate AZT zidovudine BBNH N-(4-tert-butylbenzoyl)-2-hydroxy-1-naphthaldehyde hydrazone BBSH (4-tert-butylbenzoyl)-2-hydroxy-1-salicylyl hydrazone BN-AGE blue native agarose gel electrophoresis BN-PAGE blue native polyacrylamide gel electrophoresis c concentration CA capsid CTS central termination signal d path length DCM dichloromethane delaviridine 1-[3-[(1-methylethyl)amino]-2-pyridinyl]-4-[[5- [(methylsulfonyl)amino]-1H-indol-2-y1]carbonyl]-piperazine DMF Dimethyl formamide dNTP deoxynucleotide triphosphate DNA deoxyribonucleic acid DTT dithiothreitol ε molar extinction coefficient E. coli Escherichia coli EDTA ethylenediaminetetraacetic acid efavirenz (4S)-6-chloro-4-cyclopropylethynyl-1,4-dihydro-4- trifluoromethyl-2H-3,1-benzoxazin-2-one EFV efavirenz Env envelope etravirine 4-[[6-amino-5-bromo-2-[(4-cyanophenyl)amino]-4- pyrimidinyl]oxy]-3,5-dimethylbenzonitrile E1 Cooperative unfolding/correlated exchange kinetics E2 uncorrelated exchange kinetics

11 Φ fluorescence quantum yield F fluorescence intensity f fractional intensity FDA U.S. Food and Drug Administration FT-ICR Fourier transform ion cyclotron resonance Gag group-specific antigen gp41 transmembrane envelope glycoprotein gp120 surface envelope glycoprotein gp160 envelope precursor glycoprotein G–250 Coomassie G–250 HIV human immunodeficiency virus HIV-1 human immunodeficiency virus type 1 HXMS hydrogen deuterium exchange mass spectrometry

I fluorescence intensity decay IN integrase ITC isothermal titration calorimetry kass association rate constant kdiss dissociation rate constant

kobs observed rate constant

Ka equilibrium association constant

Kd equilibrium dissociation constant l path length LC Langerhans cells LTR long terminal repeat M molar mass MA matrix mRNA messenger ribonucleic acid MS mass spectrometry m/z mass to charge ratio n refractive index NATA N-acetyltryptophanamide

12 NC nucleocapsid Nef negative regulatory factor nevirapine 11-cyclopropyl-5,11-dihydro-4-methyl-6h-dipyrido(3,2-b:2',3'- e)(1,4)diazepin-6-one NRTI nucleoside reverse transcriptase inhibitor NNRTI nonnucleoside reverse transcriptase inhibitor NTA nitrilotriacetic acid NVP nevirapine p51W401A W401A, tryptophan to alanine site mutation on p51 p66W401A W401A, tryptophan to alanine site mutation on p66 p51L234A L234A, leucine to alanine site mutation on p51 PBS primer binding site PMSF phenylmethylsulfonyl fluoride Pol polymerase PPT polypurine tract PR protease 2 χr reduced chi-square r radial distance

r0 reference position R gas constant Rev regulator of virion RNA ribonucleic acid RT reverse transcriptase SDS/PAGE sodium dodecyl sulfate/polyacrylamide gel electrophoresis SPR surface Plasmon resonance T temperature Tat trans-activator of transcription TBE tris-borate EDTA buffer TCEP tris(2-carboxyethyl)phosphine Tris tris(hydroxymethyl)aminomethane tRNA transfer RNA

13 TSAOe3T 1-{spiro[4-amino-2,2-dioxo-1,2-oxathiole-5,3'- [2',5'-bis-O-(tert-

butyldimethylsilyl)--D-ribofuranosyl]]}-3-ethylthymine UNAIDS Joint United Nations Programme on HIV/AIDS UV ultraviolet Vif Viral infectivity factor Vpr viral protein R Vpu viral protein U WHO World Health Organization wt wild-type

14 Chapter 1

1.1 HIV and AIDS

Over a quarter of a century has passed since the human immunodeficiency virus

(HIV), the causative agent of acquired immunodeficiency syndrome (AIDS), was first isolated (1). Although it has been an up-hill battle against the virus, many important scientific achievements have been made along the way. As tools and technology advance, researchers continue to gain a better grasp of the virus’s biochemistry, immuno- and molecular biology, genetics, and pathogenesis. In total, 25 therapeutics encompassing 6 different classes of drugs have been developed that significantly prolong the lives of those afflicted (2). However, we are still incapable of curing the disease. This is highlighted by the failure to fully understand the virus and is also in part due to the exceptional genetic flexibility of HIV which results in resistance to chemotherapeutic agents. Because of this, the worldwide incidence of HIV infection increases and many currently infected still suffer.

As reported by the Joint United Nations Programme on HIV/AIDS (UNAIDS) in the “2008 Report on the Global AIDS Epidemic”, it is estimated that approximately 25 million people worldwide have already died from AIDS and there are currently over 33 million people living with HIV/AIDS. Not surprisingly, HIV has become the most infectious disease resulting in human death. Because of this, the epidemic has sparked a global awareness of health disparities and has spurred political, financial, and technological action.

15 1.2 HIV

1.2.1 HIV Retrovirus and Dynamics

HIV-1 is a member of lentivirus class of retroviruses. In specific, lentiviruses use viral-mediated killing or impairing of cells which characterizes them as slow progressive viruses (3). In general, retroviruses are a large group of enveloped RNA viruses that employ reverse transcription of genomic RNA to create double-stranded (ds) DNA using the host’s machinery. This ds DNA then becomes integrated into the host’s cellular genome. Common to all retrovirus are structural, replicative, and envelope proteins used during the process of infection. These are found in three coding domains termed gag, pol, and env, respectively (4). A figure of the HIV-1 virion is shown in Figure 1-1.

16 Figure 1-1. The mature HIV-1 Virion. Credit: US National Institutes of Health.

17 1.2.2 HIV-1 Structure and Biology

1.2.2a The HIV Virion

The virion is encapsulated by a viral envelop. The envelope is a lipid-bilayer studded with glycosylated envelope (Env) proteins. There are two Env proteins, a transmembrane (TM, gp41) and a surface (SU, gp120). Inside the virion are structural and replicative proteins. Structural proteins are encoded in the gag gene and house the

Gag polyprotein precursor (Pr55gag). Replicative proteins are located in the pol gene and code for enzymes essential for viral replication, protease (PR), reverse transcriptase (RT), and integrase (IN). Additionally there are six auxiliary genes, tat, rev, nef, vif, vpr, and vpu (5–7). The organization of the HIV-1 genome is shown in Figure 1-2.

18 Figure 1-2. Organization of the HIV-1 genome.

19 1.2.2b Env Gene

The env gene produces gp160, a glycoprotein that is cleaved into envelope

proteins SU (gp120) and TM (gp41). Although these proteins remain associated in the

lipid bilayer, they have two different functions. SU, which remains on the exterior surface

of the bilayer, aids in adsorption of the virus on host cells (8). It is also important for

receptor recognition, viral attachment, and entry (9, 10). TM is a transmembrane protein

that noncovalently links SU to the surface and aids in cellular fusion (11-13).

1.2.2c Gag Gene

Gag precursor protein (Pr55gag) is essential for viral assembly. It is cleaved into

matrix (MA), capsid (CA), nucleocapsid (NC), and three smaller proteins (p6, p2, and p1). MA is located under the viral membrane and forms the viral matrix. It has two functions; targeting Pr55gag to the viral membrane and aids in translocating the viral

preintegration complex into the nucleus (5, 14, 15). CA is a hydrophobic protein that also

has two functions by enclosing the viral particle and the NC complex (6, 16). NC is a

small, basic protein that is essential to the viral life cycle. The roles of the smaller p6, p2,

and p1 proteins are not clearly understood. However, p6 is known to play a part in viral

budding and incorporation of the regulatory protein Vpr (17).

1.2.2d Pol Gene

Pol encoded proteins are essential for viral replication. The Pol polyprotein is

cleaved into PR, RT, and IN. PR is an aspartic protease that autocatalytically cleaves

itself from a Gag/Pol fusion protein to form a symmetric dimer and is required for

processing and maturation of viral proteins (18–21).

20 RT is responsible for viral replication. It is proteolytically cleaved from Gag/Pol fusion

protein by PR. The biologically active form is a heterodimer consisting of a 66 kDa (p66) and a 51 kDa (p51) subunit (22). The heterodimer has three catalytic activities (1) RNA- dependent DNA polymerase activity, (2) DNA-dependent DNA polymerase activity, and

(3) RNase H activity (23, 24). IN is a 32 kDa protein that incorporates proviral DNA, synthesized by RT, into the DNA of the host’s genome (25). The virus then uses the host’s own machinery for future viral production (26).

1.2.2e Accessory/Auxiliary Proteins

Accessory proteins necessary to HIV-1 are Tat, Rev, Nef, Vpr, Vif and Vpu (27).

These six proteins can be separated into the time of their appearance after infection. Tat,

Rev, and Nef are present early, while Vpr, Vif, and Vpu are present at late stages of the viral life cycle (28). Table 1-1 shows the major functions associated with these proteins

(29).

21 Table 1-1. Function of Accessory/Auxiliary Proteins.

Protein Function

Tat Promotes transcription of viral RNA, induces apoptosis, and inhibits siRNA formation by DICER Rev Exports un-spliced viral RNA from nucleus, and effects stability and translation of viral RNA Nef Modulates cellular receptors, enhances viral activity, interferes with host signal transduction, and regulates cholesterol trafficking Vpr Transports pre-integration complex into the nucleus, interferes with host cell cycle propagation, induces apoptosis, and transactivates HIV-LTR and host genes Vif Stimulates reverse transcription, and counteracts host anti- virus mechanisms Vpu Degrades CD4, and promotes virion defense

22

In short, the transactivator protein, Tat, is a transcriptional activator responsible

for promoting viral RNA transcriptional initiation and elongation. It is also a positive feedback protein that upregulates synthesis of itself and other viral proteins (27). An

important note is that viral particles that contain tat-negative mutants are not capable of

replicating. Tat is also a regulator of the protein REV. REV regulates virion production

through viral RNA splicing and transport of unspliced viral RNA from the nucleus to the cytoplasm. Due to REV’s ability to regulate splicing, it serves as a molecular switch from early to late stages of viral gene expression, causing expression of viral structural proteins required for assembly of the virion. Nef, the negative regulation factor, is a weak regulator of viral transcription and may act as a modifier in the expression of cellular and viral genes. Little is known on the exact function of Vpr, but it has been shown to have multiple effects during viral replication. Vif, virion infectivity factor, promotes viral particle infection. Vpu promotes assembly of Gag proteins, which is required for efficient viral budding. It also aids in processing of viral envelope proteins (27).

23 1.2.3 HIV-1 Life cycle

The HIV viral life cycle depicts how a single viral particle infiltrates a cell,

Langerhans cells and/or CD4+ T cells, and uses the host’s machinery to produce new HIV viral particles. The ultimate result is gradual depletion of helper T-cells. Viral infection is associated with entrance of multiple viral particles, and can be broken down into 10 steps: (1) absorption, (2) uncoating, (3) reverse transcription, (4) transport into the nucleus and subsequent integration, (5) transcription/splicing, (6) RNA transport, (7) translation, (8) assembly, (9) budding, and (10) maturation. This is shown in Figure 1-3.

24 Figure 1-3. Steps of Viral Infection. Credit: Vanderbilt University Medical School

(illustration by Dominic Doyle).

25 The initial process of absorption occurs with the virus binding to the cell membrane of the target cell. Viral SU binds to target cell receptors CD4 and CXCR or

CCR5 (30-33). Next, with the help of gp41 (TM) the viral and host membrane fuse, leading to the release of the core viral particle (containing RT, PR, IN, TM, SU, Vpr, Vif and tRNALys3) into the cytoplasm of the host (31).

Once in the host cell the virus begins reverse transcription. Double-stranded viral

DNA is produced from a tRNALys3 primer and single stranded genomic RNA (34). After ds viral DNA is made, a preintegration complex forms allowing transport into the nucleus. IN then cleaves the host’s DNA and integrates the newly transcribed ds viral

DNA (15, 35). After integrated, the host’s machinery is then used for subsequent production of viral particles via normal cellular transcription and translation of its genome, in particular the Tat and Rev proteins (6, 7). Tat and Rev proteins are responsible for regulating HIV-1 protein expression. Production of the Tat protein increases cellular transcription and therefore the production of HIV proteins (36, 37).

Production of Rev directs the splicing of the RNA transcripts, and once high enough levels of Rev are translated these proteins assist in transport of mRNAs into the cytoplasm where they are translated (7, 28, 38).

Maturation of the virus, that is released after budding from the cell membrane, occurs in the viral particle once the Gag polyprotein is cleaved from virally encoded PR.

Gag is housed in the Gag-Pol precursor polyprotein that is brought into the virion (39).

First, dimerization of the Gag-Pol polyproteins must occur. Subsequently, dimerization of the large polyprotein precursor results in dimerization of the aspartic protease, PR. The active PR dimer then processes Gag-Pol into its functional components. These steps are

26 necessary for viral maturation and subsequent infection. Cleavage of Gag from Gag-Pol

induces internal structural organization in the virion. This process is a requirement for

efficient viral infection.

1.2.4 HIV-1 Genetic Diversity

Currently there are two forms of HIV: HIV-1 and HIV-2. However, HIV-1 is the

most prevalent and virulent form of the virus. Unfortunately, there is also a high degree of diversity in HIV-1. This is achieved by high mutation rates, genetic recombination,

and rapid viral production (40). An important player that causes diversity is the viral

enzyme RT. RT transcribes viral RNA into DNA. However, as the DNA is being

transcribed from the viral RNA template, the RNase H domain of the enzyme

simultaneously destroys the RNA template. This prevents any proofreading during the

transcription process and facilitates viral diversity from the high error rates after

transcription occurs. The inability to proofread leads to an error rate of approximately 1 in 4,000 nucleotides (41, 42). It is estimated that the concentration of RT inside the virion

is roughly 106 and approximately 106 reverse transcription events occur each day. If, for instance, one RT corresponds to one reverse transcription event, and if one point mutation occurs during reverse transcription, then almost every point mutation would be produced on a daily basis. This would lead to many variants within the HIV-1 population and it can be postulated that within an infected individual multiple molecular variants may be present. However, not all mutations lead to enzymatically active viral proteins.

Productive mutations, which create variation in the viral genome but do not impair enzymatic function, are the causative agent of antiviral drug resistance.

27 1.3 HIV Reverse Transcription

Once the core viral proteins enter the cytoplasm of the cell, reverse transcription begins. Reverse transcription is the process of converting genomic viral RNA into ds

DNA (34). A schematic for this process is shown in Figure 1-4 (43).

28 Figure 1-4. Model of Reverse Transcription

29

RT begins by elongating a partially unwound tRNA primer that is hybridized to

the primer binding site (PBS) in the 3'–long terminal repeat (LTR) of genomic RNA to initiate minus-strand DNA synthesis. The replication primer in HIV is a tRNALys3.

Synthesis proceeds to the 5' end of the genome. This generates the minus-strand strong- stop DNA. As RT reaches the end of the template, the RNase H domain degrades the rest of the RNA strand in the RNA/DNA duplex. Next there is a strand transfer in which the newly synthesized DNA is transferred to the 3' end of the genome. This strand transfer is guided by repeat, R, sequences of the LTRs present at both ends of the RNA. Minus- strand DNA synthesis occurs accompanied by RNase H–mediated incomplete degradation of the template, leaving behind a purine rich RNA called the polypurine tract, PPT (44). Plus-strand DNA synthesis begins primarily at the PPT and continues copying minus-strand DNA to its 5' end. RNase H then removes the tRNALys3 primer facilitating a second strand transfer event. During the second strand transfer, complementary PBS segments of the plus- and minus-strand DNA anneal. Lastly, completion of plus- and minus-strand DNA synthesis occurs ending at the central termination signal, CTS, with each strand serving as a template for the other (45, 46).

30 1.4 HIV RT

RT has three different enzymatic functions: (1) RNA-dependent DNA polymerase

activity, (2) DNA-dependent DNA polymerase, and (3) RNase H activity (47). The

biologically active form of HIV RT is the heterodimer (48). Additionally, the individual

subunits can form homodimers. The subunits are products of the same gene and have

identical N-terminal amino acid sequences; p51 lacks the C-terminal RNase H domain

(49-51). The p66 subunit in the heterodimer has both polymerase and RNase H active

sites (52). The polymerase domain contains four subdomains: fingers, palm, thumb, and

connection. The structure of the heterodimer (Figure 1-5) is asymmetric, and the four

subdomains common to each subunit are in different orientations relative to one another

(53). In general, the p66 subunit is described as having an “open” conformation while the p51 is described as having a “closed” conformation (54). Although much is known about

the functions of RT, little is known about the individual subunits. Heterodimeric RT is

not the only form present in solution. RT exists as an equilibrium between the

heterodimer (p66/p51), two homodimers (p66/p66, and p51/p51), and monomers (p66

and p51). Both homodimers have DNA polymerase activity (55) and the p66/p66

homodimer also has RNase H activity (51). The monomeric subunits are devoid of

enzymatic activity (50, 51). Hence, due to its essential role in the HIV lifecycle, RT is a

major target of antiretroviral chemotherapeutic strategies (56).

31 Figure 1-5. Structure of HIV-1 RT (1DLO). The p66 and p51 subunits of RT contain four common subdomains in the polymerase domain: fingers (blue), palm (red), thumb

(green), and connection (orange). The p66 also has an RNase H subdomain (magenta).

32 1.4.1 HIV RT Inhibitors

Over a decade has been spent developing antiviral agents for HIV-1 RT.

Currently there are two classes of inhibitors targeting RT, nucleoside RT inhibitors

(NRTIs) and nonnucleoside RT inhibitors (NNRTIs). NRTIs are dideoxy nucleoside triphosphate (ddNTPs) analogs that try to prevent DNA chain elongation. These also act as competitive inhibitors of RT. NNRTIs are highly specific noncompetitive inhibitors that bind in a pocket near the catalytic site of RT. The mechanisms of action of NNRTIs are still unclear.

1.4.1a NRTIs

NRTIs are essentially precursor compounds that mimic natural nucleosides and require phosphorylation by cellular enzymes into an active triphosphate moiety (57). A feature common to all NRTIs is the lack of a 3' hydroxyl (OH) on the sugar group of the inhibitor. This missing OH prevents nucleophilic attack of an incoming nucleotide ensuring termination of DNA synthesis. Currently there are eight NRTIs approved for treatment of HIV-1. These compounds comprise five subgroups that are classified by the type of sugar moiety (58): Group 1, 3' OH group is replaced by a hydrogen; Group 2, chemical substitution of the 3' OH group; Group 3, contain an oxathiolane ring, with a sulfur atom at the 3' position of the sugar ring; Group 4, contain a double bond between the 2' and 3' positions of the sugar moiety; and Group 5, these are acyclic and lack a sugar ring. All types of NRTIs are shown in Figure 1-6 with examples representing each group.

33 Figure 1-6. Structure of NRTIs.

34 35 1.4.1b NNRTIs

NNRTIs are a diverse class of small molecules that bind in a hydrophobic pocket

~ 10 Å away from the polymerase active site of RT, shown in Figure 1-7 (59). Currently

there are four clinically used NNRTIs, nevirapine, delaviridine, efavirenz, and etravirine

Figure 1-8. These drugs are also highly effective and relatively noncytotoxic (60).

NNRTIs have been found to affect both early and late stages of the HIV-1 replication cycle by multiple mechanisms (61-63). This includes inhibition of DNA polymerase activity, enhancing polymerase-dependent RNase H activity (3'-DNA directed), and partially inhibiting polymerase-independent RNase H activity (5'-RNA directed). They also inhibit plus-strand initiation by affecting the ability of RT to bind the RNA polypurine tract (PPT). During late stages of HIV-1 replication, NNRTIs enhance processing and homodimerization of a 90 kDa Pol polyprotein in a yeast two-hybrid system. NNRTIs have also been shown to increase intracellular processing of Gag and

Gag-Pol precursor polyproteins in HIV-1-transfected cells (62). They have also been shown to have diverse effects on RT dimerization; efavirenz (EFV) and nevirapine

(NVP) enhance subunit interactions (64, 65), delaviridine has little or no effect (64), and

TSAOe3T, BBNH and BBSH weaken subunit interactions (66, 67).

36 Figure 1-7. Structure of HIV-1 RT─NNRTI Complex (1FK9). The subdomains of p66 and p51 subunits of RT are color coded: fingers (blue), palm (red), thumb (green), and connection (orange). The p66 also has an RNase H subdomain (magenta), efavirenz

(cyan) shown using van der Waals radii.

37 Figure 1-8. Structure of clinically used NNRTIs

Br O N

H2N N Etravirine (TMC-125) Intelence™

HN N

38 1.5 Research Techniques

1.5.1 Equilibrium Dialysis.

Equilibrium dialysis is an effective tool for studying the interactions of small molecules with proteins (68). An important feature of this technique is that the results provide the true nature of the interaction because it is conducted under equilibrium conditions. The principle of this technique is to partition the free ligand from the protein─ligand complex by allowing the ligand to dialyze through a semi-permeable membrane until the concentration of the unbound ligand is equal on both sides of the membrane (Shown in Figure 1-9).

39 Figure 1-9. Principle of Equilibrium Dialysis. Adapted from NestGroup Inc.

40

In a typical equilibrium dialysis experiment there are two buffered solutions

separated by a membrane that only allows diffusion of a low molecular mass ligand. The

protein is placed in compartment 1 and compartment 2 contains only the dialysate buffer.

At the start of the reaction, the ligand, whose molecular weight is lower than the

molecular weight cut off of the membrane, is added to compartment 2. The ligand freely

diffuses until equilibrium is reached in both compartments. The bound ligand is

calculated from

[Ibound] = [Iin] – [Iout] (1)

where [Iin] is the total concentration of free and bound ligand in one compartment and

[Iout] is the concentration of free ligand in the other compartment. These experiments

were conducted using a [14C] radiolabeled NNRTI, efavirenz. After each experiment,

aliquots from both compartments were removed and counted using a Beckman Coulter

LS6500 Multi purpose scintillation counter. The data from the scintillation counter were

converted to molarity using the specific activity of the NNRTI, calculated by Moravek

Biochemicals after synthesis, and fit to mathematical models in the Dialfit program.

Details of the Dialfit program are provided in Chapter 2, Section 2.7.

1.5.2 Fluorescence Spectroscopy.

Fluorescence spectroscopy is a widely used technique in biochemistry, biophysics and material sciences. Fluorescence is governed by several parameters; fluorescence intensity, emission, quantum yield, lifetime, and polarization (69). In essence, a photon of light excites a susceptible molecule on the femtosecond timescale (10–15 s). After the

absoption of light energy, the electron at the higher energy state cannot be sustained and

41 there is a transitional relaxation from excited singlet states to different vibrational levels of the ground states. This occurs on a slower, measureable timescale of picoseconds (10–

12 s). The energy decay from the excited state emits a longer wavelength photon as the

molecule returns to the ground state. The overall process occurs on the timescale of

nanoseconds (10–9 s). This process termed fluorescence is shown in Figure 1-10. The

interaction between light and matter are measured by steady-state and time-resolved

fluorescence experiments. Here, only steady-state fluorescence is described as it pertains

to the research conducted.

42 Figure 1-10. A Jablonski diagram. Credit: Molecular Expressions™

43

Steady-state fluorescence experiments are performed under conditions of constant

illumination. The intensity of emitted light is measured as a function of wavelength. The

fluorescence quantum yield (Ф) is the ratio of photons absorbed to photons emitted and

gauges the efficiency of fluorescence emission during relaxation,

absorbed photons # photons absorbed kr Ф =  (2) em itted photons # photons emitted  kk nrr where kr is the cumulative rate constant of all radiative processes and knr is the

cumulative rate constant of all non-radiative processes. The value of Ф is determined

experimentally from comparison of a known standard by

2 F As n Ф = Фs (3) 2 s F A ns

where Фs is the quantum yield of the standard, and F, A, n, Fs, As, and ns, are the integrated fluorescence intensity, absorbance, and refractive index of the sample and standard respectively.

Advances in fluorescence allow the unique properties of fluorescent molecules to be detected with extraordinary sensitivity and selectivity. This is especially useful for proteins. There are three aromatic amino acids that can be used to examine conformational changes in proteins; tryptophan, tyrosine, and phenylalanine. The most influential is tryptophan. Tryptophan is exquisitely sensitive to environmental changes from nearby residues in the protein that result from structural rearrangements as well as externally added quenchers into the system. Fortunately, RT contains 37 tryptophan residues, 19 in the p66 subunit and 18 in the p51 subunit. These well distributed

44 tryptophan residues were used to report conformational changes that occur due to drug

binding.

1.5.3 Mass Spectrometry.

Mass spectrometry (MS) is an incredibly useful tool for determining elemental

composition of molecules and for studying protein structural dynamics (70). Unlike other spectroscopic techniques, detection of compounds can be accomplished with very minute quantities (10-12–10-15 M), depending on the type of ionization method and mass analyzer

used. A mass spectrometer measures the masses of individual molecules that have been

electrically charged and converted into ions, i.e. the mass-to-charge ratio (m/z).

The instrument primarily has two main components, an ion source and a mass

analyzer which governs the sensitivity, the accuracy, and resolution of the results. Two

types of ion sources are used to generate gaseous ions, electrospray ionization (ESI) and

matrix-assisted laser desorption/ionization (MALDI). Both techniques are forms of soft

ionization and produce positively charged ion droplets after application of high voltage

(ESI) or irradiation of UV-laser pulses (MALDI). Formation of gaseous ions is

subsequently directed to a mass analyzer for m/z measurement. For purposes of this

research ESI-MS is further elaborated.

To evaluate the changes in solution structure that ligand binding has on monomers

of RT both tandem mass spectrometry and hydrogen/deuterium exchange mass

spectrometry experiments were conducted. In a tandem mass spectrometry experiment

m/z measurements of peptide ions and fragmented ions were monitored. These types of experiments are also referred to as MSn, where n represents the number of m/z

measurements performed by the mass analyzer. For example, in a MS2 experiment the

45 mass analyzer first records the m/z of the peptide ion. Next the peptide ion is bombarded with inert helium gas to fragment the ion. This is termed collisionally induced dissociation (CID) and produces fragmentation patterns unique to that peptide. This process is shown in Figure 1-11.

46 Figure 1-11. Standard tandem mass spectrometry experiment (MS2)

47 In HXMS experiments, dilution into deuterium buffer is used to label backbone

amide hydrogen atoms. During the course of an experiment various levels of labeling are

witnessed as a function of incubation time. The backbone amide hydrogen atoms exchange when the backbone is exposed to the deuterium buffer. The labeling is due to structural fluctuations that naturally occur in solution or after the addition of ligands to the reaction. Deuterium uptake is a function of regional flexibility and solvent accessibility of the protein. In a typical HXMS experiment, the protein is incubated at 5

°C in deuterium buffer and quenched after different time intervals by dropping the pH of the solution to ~ 2.3. Performing the reaction at low temperatures prevents rapid back exchange with hydrogen atoms. Region-specific deuterium uptake is monitored by digestion with porcine pepsin to generate peptic fragments of protein. The peptic

fragments are then separated by high-performance liquid chromatography (HPLC) and

the amount of deuterium uptake of each peptide is determined by the mass analyzer.

Addition of each deuterium atom to the peptide increases the mass by one as compared to

the non-deuterium labeled sample. Percent deuterium uptake is given by the following

equation

 mm D  %0  100 (4)  mm %0%100

where m is the mass of the deuterated peptide, m0% is the mass of a non-deuterated

peptide, and m100% is the mass of a fully deuterated peptide. The fully deuterated peptide

is prepared by incubating the protein in a high concentration of denaturant-D2O buffer, such as guanidinium or urea to completely unfold the protein. After denaturation in deuterated buffer, the sample is similarly digested with a protease and analyzed. An example of this is shown in Figure 1-12.

48 Figure 1-12. Schematic of Hydrogen/Deuterium Exchange

49 1.6 Research Objectives

The goal of this research project is to determine the stoichiometry and binding constants of the NNRTI efavirenz to monomers and homodimers of RT. In addition, the structural changes that occur due to drug binding to monomers are also examined. Our hypothesis proposes a thermodynamic linkage between NNRTI binding and subunit dimerization (Scheme 1-1).

50 Scheme 1-1. Thermodynamic Linkage of NNRTI Binding and Dimerization

2I

p66/p51 p66/p51 I + I Kd(1)

Kd(4) Kd(2) 2I p66+ p51 p66 I + p51 I K (3) d

51 The thermodynamic circle proposes a link between the effect NNRTIs have on

dimerization and their affinity for binding monomers and dimers of RT. Therefore, if an

NNRTI enhances dimerization then the inhibitor binds more tightly to the dimer.

Conversely, if the NNRTI weakens dimerization then the inhibitor binds more tightly to

the monomer. By examining protein─ligand interactions we can achieve a better understanding of the inhibition mechanism. Additionally, we can open the possibility of using RT monomers as new potential drug targets.

Current literature reports of NNRTIs indicate that these compounds have diverse

effects on RT subunit dimerization; efavirenz (EFV) and nevirapine (NVP) are shown to

enhance subunit interactions, delavirdine (DLV) has little or no effect, and TSAOe3T,

BBNH and BBSH decrease subunit stability (64–67, 71). Our initial aim was to

determine: (1) if NNRTIs are capable of binding monomers, (2) equilibrium monomer

binding constants, and (3) the binding stoichiometry. All three were achieved through

equilibrium dialysis, intrinsic tryptophan fluorescence, and native gel electrophoresis.

Our second aim was to determine the changes that occur in the solution structure of the

monomers upon drug binding. This was accomplished by HXMS.

The research described here is a completely new addition to the knowledge base

of RT, and that in itself is a completely rewarding triumph. RT is an incredibly

interesting protein that is difficult to work with under in vitro conditions. This project is

based upon tremendous efforts by Dr. Mary D. Barkley’s research group to understand

this complicated protein. Previous analytical ultracentrifugation experiments were critical

for conducting experiments in this project. In solution, the protein can exist as five

species, three dimeric, and two monomeric as shown in Figure 1-11. Based on the

52 group’s findings, experiments were set up accordingly. Results corresponding to aim one are described in Chapter 2, whereas solution structural studies are described in Chapter 3.

53 Figure 1-13. Schematic of RT Species in Solution

54

References

1. Barre-Sinoussi, F., Cherman, J. C., Rey, F., Nugeyre, M. T., Chamaret, S., Gruest,

J., Dauguet, C., Axler-Blin, C., Vezinet-Brun, F., Rouzioux, C., Rozenbaum,

W., and Montagnier, L. (1983) Isolation of a T-lymphoptrohpic retrovirus from a

patient at risk for acquired immunodeficiency syndrome (AIDS). Science 220,

868–871.

2. Shafer, R. W., and Schapiro, J. M. (2008) HIV-1 drug resistance mutations: an

updated framework for the second decade of HAART. AIDS REV. 10, 67–84.

3. Weiss, R. A. (1996) Retrovirus classification and cell interactions. J. Antimicrob.

Chemother. 37, 1–11.

4. Levy, J. A., and Coffin, J. M. (1992) "Structure and Classification of

Retroviruses". The Retroviridae (1st ed.). New York: Plenum Press, 26–34.

5. Gelderblom, H. R., Hausmann, E. H. S., Ozel, M., Pauli, G., and Koch, M. A.

(1987) Fine structure of human immunodeficiency virus (HIV) and

immunolocalization of structural proteins. Virology 156, 171–176.

6. Haseltine, W. A. (1991) Molecular biology of human immunodefiency virus type

1. FASEB J. 5, 2349–2360.

7. Cullen, B., R. (1991) Regulation of HIV-1 gene expression. FASEB J. 5, 2361–

2368.

8. McDougal, J. J., Kenedy, M. S., Seigh, J. M., Cort, S. P., Mawla, A., and

Nicholson, J. K. A. (1986) Binding of HTLV-III/LAV to T4+ cells by a complex

of the 100K viral protein and the T4 molecule. Science 231, 382–385.

55 9. Cann, A. J., Churcher, M. J., Boyd, M., O’Brien, W., Zhao, J. Q., Zack, J. A., and

Chen, I. S. Y. (1992) The region of the envelop gene of human immunodeficiency

virus type 1 responsible for determination of cell tropism. J. Virol. 66, 305–309.

10. Kato, K., Sato, H., and Takebe, Y. (1999) Role of naturally occurring basic amino

acid substitutions in the human immunodeficiency virus type 1 subtype E

envelope V3 loop on the viral coreceptor usage and cell tropism. J. Virol. 73,

5520–5526.

11. Koito, A., Harrowe, G., Levy, J. A., and Cheng-Mayer, C. (1994) Functional role

of the V1/V2 region of human immunodeficiency virus type 1 envelope

glycoprotein gp120 in infection of primary macrophages and soluble CD4

neutralization. J. Virol. 68, 2253–2259.

12. Gallaher, W. R., Ball, J. M., Garry, R. F., Griffen, M. C., and Montelaro, R. C.

(1989) A general model for the transmembrane proteins of HIV and other

retroviruses. AIDS Res. Hum. Retroviruses 5, 431–440.

13. Kowalski, M., Potz, J., Bsiripour, L., Dorfman, T., Goh, W. C., Terwillger, E.,

Dayton, A., Rosen, C., Haseltime, W., and Sodroski, J. (1987) Functional regions

of the envelope glycoprotein of human immunodeficiency virus type 1. Science

237, 1351–1355.

14. Spearman, P., Wang, J. J., Vander Heyden, N., and Ratner, L. (1994)

Identification of human immunodeficiency virus type 1 Gag protein domains

essential to membrane binding and particle assembly. J. Virol. 68, 3232–3242.

15. Bukrinsky, M. I., Sharova, N., Dempsey, M. P., Stanwick, T. L., Bukrinskaya, A.

G., Haggerty, S., and Stevenson, M. (1992) Active nuclear import of human

56 16. Ganser, B. K., Li, S., Klishko, V. Y., Finch, J. T., and Sundquist, W. I. (1999)

Assembly and analysis of conical models for the HIV-1 core. Science 283, 80–83.

17. Göttlinger, H. G. (2001) The HIV-1 assembly machine. AIDS 15, S13–S20.

18. Graves, M. C., Lim, J. J., Heimer, E. P., and Kramer, R. A. (1988) An 11-kDa

form of human immunodeficiency virus protease expressed in Escherichia coli is

sufficient for enzyme activity. Proc. Natl. Acad. Sci. 85, 2449–2453.

19. Mous, J., Heimer, E. P., and LeGrice, S. F. J. (1988) Processing protease and

reverse transcriptase from human immunodeficiency virus type 1 polyprotein in

Escherichia coli. J. Virol. 62, 1433–1436.

20. Freed, E. O., and Martin, M. A. (2001) HIVs and their replication, in Fields

Virology, 4th ed., Lippincott, Williams, and Wilkins, Philadelphia.

21. Debouck, C., Gorniak, J. G., Strickler, J. E., Meek, T. D., Metcalf, B. W., and

Rosenberg, M. (1987) Human immunodeficiency virus protease expressed in

Escherichia coli exhibits autoprocessing and specific maturation of the gag

precursor. Proc. Natl. Acad. Sci. 84, 8903–8906.

22. von der Helm, K. (1996) Retroviral proteases: structure, function and inhibition

from a non-anticipated viral enzyme to the target of a most promising HIV

therapy. Biol. Chem. 377, 765–774.

23. Gilboa, E., Mitra, S. W., Goff, S. P., and Baltimore, D. (1979) A detailed model

of reverse transcription and tests for crucial aspects. Cell 18, 93–100.

57 24. Goff, S. P. (1990) Retroviral reverse transcriptase: synthesis. Structure, and

function. J. Acquired Immune Defic. Syndr. 3, 817–831.

25. Katz, R. A., and Skalka, A. M (1994) The retroviral enzymes. Annu. Rev.

Biochem. 63, 133–173.

26. Bushman, F. D. and Cragie, R. (1992) Integration of human immunodeficiency

virus DNA: adduct interference analysis of required DNA sites. Proc. Natl. Acad.

Sci. 89, 3458–3462.

27. Haseltine, W. A. (1988) Replication and pathogenesis of the AIDS virus. J.

Acquired Immune Defic. Syndr. 1, 217–240.

28. Cullen, B. R. (1992) Mechanism of action of regulatory proteins encoded by

complex retroviruses. Microbiology Rev. 56, 375–394.

29. Li, L., Li, H. I., Pauza, C. D., Burkinsky, M., and Zhao, R. Y. (2005) Roles of

HIV-1 auxiliary proteins in viral pathogenesis and host-pathogen interaction. Cell

Res. 15, 923–934.

30. Maddon, P. J., Dalgeish, A. J., McDougal, J. S., Clapham, P. R., Weiss, R. A.,

and Axel, R. (1986) The T4 gene encodes the AIDS virus receptor and is

expressed in the immune system and the brain. Cell 78, 333–348.

31. Stein, B. S., Gowda, S. D., Lifson, J. D., Penhallow, R. C., Bensch, K. G., and

Engelman, E. G. (1987) pH-independent HIV entry into CD4-positive T cells via

virus envelope fusion to the plasma membrane. Cell 49, 659–668.

32. Wu, L., Gerard, N. P., Wyatt, R., Choe, H., Parolin, C., Ruffing, N., Borsetti, A.,

Cardoso, A., Desjardin, E., Newman, W., Gerard, C., and Sodroski, J. (1996)

58 33. Trkola, A., Dragic, T., Arthos, J., Binley, J. M., Olson, W. C., Chen-Meyer, C.,

Robinson, J., Maddon, P. J., and Moore, J. P. (1996) CD4-dependent, antibody-

sensitive interaction between HIV-1 and it co-receptor CCR-5. Nature 384, 184–

187.

34. Baltimore, D. (1970) RNA-dependent DNA polymerase in virions of RNA

tumour viruses. Nature 226, 1209–1211.

35. Vink, C., and Plasterk, R. H. (1993) The human immunodeficiency virus

integrase protein. Trends Genet. 9, 433–438.

36. Kim, J. B., Yamaguchi, Y., Wada, T., Handa, H., and Sharp, P. A. (1999) Tat-SF1

protein associates with RAP30 and human SPT5 proteins. Mol. Cell. Biol. 19,

5960–5968.

37. Weeks, K. M., Ampe, C., Schultz, S. C., Steitz, T. A., and Crothers, D. M. (1990)

Fragments of the HIV-1 Tat protein specifically bind TAR RNA, Science 249,

1281–1285.

38. Cullen, B. R. (1998) HIV-1 auxiliary proteins: making connections in a dying

cell. Cell 93, 685–692.

39. Bukrinskaya, A. G. (2004) HIV-1 assembly and maturation. Arch. Vir. 149,

10637–1082.

40. Ratner, L. (1993) HIV life cycle and genetic approaches. Perspect. Drug. Disc.

Design 1, 3–22.

59 41. Preston, B. D., Poiesz, B. J., and Loeb, L. A. (1988) Fidelity of HIV-1 reverse

transcriptase. Science 242, 1168–1171.

42. Mansky, L. M., and Temin, H. M. (1995) Lower in vivo mutation rate of human

immunodeficiency virus type 1 than predicted from the fidelity of purified reverse

transcriptase. J. Virol. 69, 5087–5094.

43. Rausch, J. W., and Le Grice, S. F. J. (2004) ‘Binding, bending, and bonding’:

polypurine tract-primed initiation of plus-strand synthesis in human

immunodeficiency virus. Int. J. Biochem. Cell Biol. 36, 1752–1766.

44. Sorge, J., and Hughes, S. H. (1982) Polypurine tract adjacent to the U3 region of

Rous sarcoma virus genome provides a cis-acting function. J. Virol. 43, 482–488.

45. Huber, H. E., McCoy, J. M., Seehra, J. S., and Richardson, C. C. (1989) Human

immunodeficiency virus 1 reverse transcriptase: template binding, processivity,

strand displacement synthesis and template switching. J. Biol. Chem. 264, 4669–

4678.

46. Charneau, P., Mirambeau, G., Roux, P., Paulous, S., Buc, H., and Clavel, F.

(1994) HIV-1 reverse transcription. A termination step at the center of the

genome. J. Mol. Biol. 241, 651–662.

47. Patel, P. H., Jacobo-Molina, A., Ding, J., Tantillo, C., Clark, A. D., Raag, Jr., R.,

Nanni, R. G., Hughes, S. H., and Arnold, E. (1995) Insights into DNA

polymerization mechanisms from structure and function analysis of HIV-1

reverse transcriptase. Biochemistry 34, 5351–5363.

48. di Marzo Veronese, F., Copeland, T. D., DeVico, A. L., Rahman, R., Oroszlan, S.,

Gallo, R. C., and Sarngadharan, M. G. (1986) Characterization of highly

60 49. Hizi, A., McGill, C., and Hughes, S. H. (1988) Expression of soluble,

enzymatically active, human immunodeficiency virus reverse transcriptase in

Escherichia coli and analysis of mutants, Proc. Natl. Acad. Sci. U.S.A. 85, 1218–

1222.

50. Restle, T., Muller, B., and Goody, R. S. (1990) Dimerization of human

immunodeficiency virus type 1 reverse transcriptase, J. Biol. Chem. 265, 8986–

8988.

51. Restle, T., Muller, B., Goody, R. S. (1992) RNase H activity of HIV reverse

transcriptase is confined exclusively to the dimeric forms, FEBS Lett. 300, 97–

100.

52. Le Grice, S. F. J., Naas, T., Wohlgensinger, B., and Schatz, O. (1991) Subunit-

selective mutagenesis indicates minimal polymerase activity in heterodimer-

associated p51 HIV-1 reverse transcriptase, EMBO J. 10, 3905–3911.

53. Wang, J., Smerdon, S. J., Jager, J., Kohlstaedt, L. A., Rice, P. A., Friedman, J. M.,

and Steitz, T. A. (1994) Structural basis of asymmetry in the human

immunodeficiency virus type 1 reverse transcriptase heterodimer. Proc. Natl.

Acad. Sci. U.S.A. 91, 7242–7246.

54. Jacobo-Molina, A., Ding, J., Nanni, R. G., Clark, Jr., A. D., Lu, X., Tantillo, C.,

Williams, R. L., Kamer, G., Ferris, A. L., Clark, P., and et al. (1993) Crystal

structure of human immunodeficiency virus type 1 reverse transcriptase

61 55. Bavand, M. R., Wagner, R., and Richmond, T. J. (1993) HIV-1 reverse

transcriptase: polymerization properties of the p51 homodimer compared to the

p66/p51 heterodimer. Biochemistry 32, 10543–10552.

56. Tsai, C. H., Lee, P. Y., Stollar, V., and Li, M. L. (2006) Antiviral therapy

targeting viral polymerase, Curr. Pharm. Des. 12, 1339–1355.

57. Furman, P. A., Fyfe, J. A., St. Clair, M. H., and et al. (1986) Phosphorylation of

3’-azido-3’ deoxythymidine and selective interactions of the 5’-triphosphate with

human immunodeficiency virus reverse transcriptase. Proc. Natl. Acad. Sci.

U.S.A. 83, 8333–8337.

58. Menendez-Arias, L. (2008) Mechanisms of resistence to nucleoside analogue

inhibitors of HIV-1 reverse transcriptase. Vir. Res. 134, 124–146.

59. Kohlstaedt, L. A., Wang, J., Friedman, J. M., Rice, P. A., and Steitz, T. A. (1992)

Crystal structure at 3.5 Å resolution of HIV-1 reverse transcriptase complexed

with an inhibitor, Science 256, 1783–1790.

60. De Clercq, E. (1998) The role of non-nucleoside reverse transcriptase inhibitors

(NNRTIs) in the therapy of HIV-1 infection, Antiviral Res. 38, 153–179.

61. Sluis-Cremer, N., and Tachedjian, G. (2008) Mechanisms of inhibition of HIV

replication by non-nucleoside reverse transcriptase inhibitors, Virus Res. 134,

147–156.

62. Grobler, J. A., Dornadula, G., Rice, M. R., Simcoe, A. L., Hazuda, D. J., and

Miller, M. D. (2007) HIV-1 reverse transcriptase plus-strand initiation exhibits

62 63. Figueiredo, A., Moore, K. L., Mak, J., Sluis-Cremer, N., de Bethune, M.-P., and

Tachedjian, G. (2006) Potent nonnucleoside reverse transcriptase inhibitors target

HIV-1 Gag-Pol. PLoS Pathog. 2, 1051–1059.

64. Tachedjian, G., Orlova, M., Sarafianos, S. G., Arnold, E., and Goff, S. P. (2001)

Nonnucleoside reverse transcriptase inhibitors are chemical enhancers of

dimerization of the HIV type 1 reverse transcriptase, Proc. Natl. Acad. Sci. U.S.A.

98, 7188–7193.

65. Venezia, C. F., Howard, K. J., Ignatov, M. E., Holladay, L. A., and Barkley, M.D.

(2006) Effects of efavirenz binding on the subunit equilibria of HIV-1 reverse

transcriptase, Biochemistry 45, 2779–2789.

66. Sluis-Cremer, N., Arion, D., and Parniak, M. A. (2002) Destabilization of the

HIV-1 reverse transcriptase dimer upon interaction with N-acyl hydrazone

inhibitors, Mol. Pharmacol. 62, 398–405.

67. Sluis-Cremer, N., Dmitrienko, G. I., Balzarini, J., Camarasa, M.-J., and Parniak,

M. A. (2000) Human immunodeficiency virus type 1 reverse transcriptase dimer

destablilization by 1-spiro[4-amino-2,2-dioxo-1,2-oxathiole-5,3-[2,5-

bis-O-(tert-butyldimethylsilyl)-β-D-ribofuranosyl]]-3-ethylthymine,

Biochemistry 39, 1427–1433.

68. The Nest Group (2002) Guide to Equilibrium Dialysis, Harvard Biosciences Inc,

Southborough, MA.

63 69. Lakowicz, J. R. (2006) Principles of Flourescence Spectroscopy, 3rd ed., Springer,

New York.

70. Kinter, M., and Sherman, N. (2000) Protein Sequencing and Identification Using

Tandem Mass Spectrometry, John Wiley & Sons Inc., New York, NY, USA.

71. Tachedjian, G., Moore, K. L., Goff, S. P., Sluis-Cremer, N. (2005) Efavirenz

enhances the proteolytic processing of an HIV-1 pol polyprotein precursor and

reverse transcriptase homodimer formation, FEBS Letters 579, 379-384.

64 Chapter 2: Efavirenz Binding to HIV-1 Reverse Transcriptase Monomers and Dimers

2.1 Abstract

Efavirenz (EFV) is a nonnucleoside reverse transcriptase inhibitor (NNRTI) of

HIV-1 reverse transcriptase (RT) used for the treatment of AIDS. RT is a heterodimer composed of p66 and p51 subunits; p51 is produced from p66 by C-terminal truncation by HIV protease. The monomers can form p66/p66 and p51/p51 homodimers as well as

p66/p51 heterodimer. Dimerization and efavirenz binding are coupled processes. In the

crystal structure of the p66/p51─EFV complex, the drug is bound to the p66 subunit. The binding of efavirenz to wild-type and dimerization-defective RT proteins was studied by equilibrium dialysis, tryptophan fluorescence and native gel electrophoresis. A 1:1 binding stoichiometry was determined for both monomers and homodimers. Equilibrium dissociation constants are ~2.5 µM for both p66─ and p51─EFV complexes, 250 nM for p66/p66─EFV complex, and 7 nM for p51/p51─EFV complex. An equilibrium dissociation constant of 92 nM for p66/p51─EFV complex was calculated from the thermodynamic linkage between dimerization and inhibitor binding. Binding and unbinding kinetics monitored by fluorescence were slow. Progress curve analyses revealed a one-step, direct binding mechanism with association rate constants k1 ~13.5

–1 –1 –4 –1 M s for monomers and heterodimer and dissociation rate constants k–1 ~1  10 s for monomers. A conformational selection mechanism is proposed to account for the slow association rate. These results show that efavirenz is a slow, tight-binding inhibitor capable of binding all forms of RT and suggest that the NNRTI binding site in monomers and dimers is similar.

65 2.2 Introduction

HIV-1 RT converts single-stranded viral RNA into double-stranded proviral

DNA. The enzyme has two activities, DNA polymerase and RNase H. The biologically relevant form is a heterodimer composed of two subunits, p66 and p51 (1). The subunits

are products of the same gene and have identical N-terminal amino acid sequences; p51

lacks the C-terminal RNase H domain (2-4). The individual subunits can also form

homodimers. The p66 subunit in the heterodimer has both polymerase and RNase H

active sites (5). The monomeric subunits are devoid of enzymatic activity (3, 4). Due to

its essential role in the HIV lifecycle, RT is a major target of antiretroviral drugs (6). Two

classes of inhibitors have been developed and approved for clinical use, NRTIs and

NNRTIs. The NNRTIs are highly effective and relatively noncytotoxic (7). These small,

amphiphilic, noncompetitive inhibitors nestle into a hydrophobic pocket ~ 10 Å away

from the polymerase active site in the p66 subunit of RT (8, 9). NNRTIs primarily interfere with reverse transcription, but they also affect late stages of HIV replication in

Gag-Pol polyprotein processing (10-12).

NNRTIs have diverse effects on RT subunit dimerization. Efavirenz (EFV) and

nevirapine (NVP) enhance subunit interactions (13, 14), delavirdine has little or no effect

(13), and TSAOe3T, BBNH and BBSH weaken subunit interactions (15, 16). The evidence for these results derives from multiple techniques including yeast two-hybrid, pull-down assays, urea induced dissociation, size exclusion chromatography, and sedimentation equilibrium studies. To explain the contrasting effects of NNRTI binding on RT, our group previously proposed a thermodynamic cycle (14). In Scheme 2-1, P

66 denotes p66 or p51 monomer, P/P is p66/p51 heterodimer, p66/p66 homodimer, or p51/p51 homodimer, and I is NNRTI.

67 Scheme 2-1: Thermodynamic Linkage of NNRTI Binding and Subunit Dimerization

I

P/P P/P I Kd(1)

Kd(4) Kd(2) I

P+P PPI + K (3) d

68 The thermodynamic linkage between NNRTI binding and RT subunit dimerization makes

the following predictions: (1) NNRTIs bind to both monomeric and dimeric forms of RT.

The crystal structures of RT—NNRTI complexes show one drug bound per heterodimer

(8, 9). In solution, the stoichiometry of drug binding to dimer is not known. (2) NNRTIs

that enhance dimerization bind more tightly to dimers. Conversely, NNRTIs that weaken

dimerization bind more tightly to monomers. Identifying and quantifying the various

protein-ligand interactions is essential for thorough understanding of the inhibition

mechanism of NNRTIs. The previous thermodynamic cycle (Scheme 1 in ref 14) makes the additional prediction that low concentrations of inhibitor will promote dimerization if

Kd(1) < Kd(3). However, eventually Le Châtelier's principle will shift the equilibrium

towards the formation of P─I at high concentrations of inhibitor.

Previous sedimentation equilibrium studies showed that efavirenz enhances the

formation of p66/p51, p66/p66, and p51/p51 by 25-, 50-, and 600-fold (14). Here we

measure the binding of efavirenz to p66 and p51 monomers in wild-type and dimerization

defective mutant RTs and determine the binding stoichiometry of monomers and

homodimers. Binding stoichiometry and equilibrium dissociation constants for drug

binding to dimer and monomer, Kd(1) and Kd(3), were determined by equilibrium

dialysis. The kinetics of drug binding to monomers and heterodimer were monitored by

intrinsic protein fluorescence. Finally, the binding of [14C] efavirenz to p66 monomer and

p66/p66 homodimer was visualized by Blue Native gel electrophoresis.

69 2.3 Experimental Procedures

Materials. Efavirenz was obtained from the NIH AIDS Research and Reference

Reagent Program (Germantown, MD). [14C] efavirenz (Specific Activity: 52 mCi/mmol)

was purchased from Vitrax (Placentia, CA). Dialysis tubing was purchased from

Spectrum Labs (Rancho Dominguez, CA). Rapid Equilibrium Dialysis (RED) Device and

TCEP were purchased from Pierce (Rockford, IL). Econo-Safe scintillation fluid was

purchased from Atlantic Nuclear Corporation (Canton, MA). Oligodeoxynucleotide

primers, 5% Coomassie blue G-250 sample additive and the NativePAGE Novex Bis-Tris

gel system were purchased from Invitrogen Corp. (Carlsbad, CA). EZ-Run Protein Gel

Staining solution was purchased from Fisher Scientific (Fair Lawn, NJ). Biochemical

reagents were purchased from Roche Applied Science (Indianapolis, IN). Other

chemicals were from Sigma Chemicals (St. Louis, MO). RT buffer D is 0.05 M Tris (pH

7.0), 25 mM NaCl, 1 mM EDTA, and 10% (v/v) glycerol.

Protein Preparation. HIV-1 RT proteins with N-terminal hexahistadine

extensions were expressed in Escherichia coli M15 strains containing plasmid p6H RT

for p66, p6H RT51 for p51, or p6H RT-PR for p66/p51 heterodimer and purified by Ni–

NTA, S-Sepharose, and DEAE chromatography as previously described (14, 17). Protein concentration is determined from absorbance at 280 nm and is expressed in monomer units (14, 18). Protein stock solutions prior to use were dialyzed overnight into RT buffer

D containing 1 mM TCEP.

Dimerization defective RT proteins were prepared from plasmids p6H RT and p6H RT51 containing the W401A mutation (19). The W401A mutation was introduced

by one round of mutagenesis using the QuickChange site-directed mutagenesis kit

70 (Stratagene, La Jolla, CA). The oligonucleotide primer sequences were: forward, 5’-

GGGAAACAGCGTGGCCAGAGTATTGGCAAGCCACCTG-3’; reverse, 5’-

CAGGTGGCTTGCCAATACTCTGTCCACGCTGTTTCCC-3’. All mutations were confirmed by DNA sequencing at Agencourt Bioscience (Beverly, MA).

Equilibrium Dialysis. Equilibrium dialysis experiments were conducted using 1.5 mL RNase/DNase free amber microcentrifuge tubes and 4-mm dialysis tubing with 3,500 molecular weight cut off or RED devices. A 1 mM stock solution of [14C] efavirenz in

DMF was prepared. A 250 μL aliquot of RT solution was loaded into dialysis tubing or one chamber of the RED device. RT concentrations were 0.1–10 µM p51, 1–10 µM

51W401A, 2–4 µM p51L234A, 0.4–5 µM p66, and 0.8–7.5 µM p66W401A. RT buffer D containing 1 mM TCEP and 0.2–20 µM [14C] efavirenz was used as dialysate buffer. For microcentrifuge tubes, the dialysis bag and 1 mL of dialysate buffer were placed in the tube and the tube was capped. For RED devices, 0.4 mL of dialysate buffer was placed in the other chamber. The samples were set up in triplicate, secured to a benchtop rotator and dialyzed at 4 ºC. Wild-type RT proteins were dialyzed for up to 5 days; W401A mutant proteins were dialyzed for 30 h. Equilibration of efavirenz across the membrane occured by 20 h.

Efavirenz binding was quantified by counting three 50 μL aliquots of the inside protein solution and outside dialysate solutions in 5 mL of scintillation fluid using a

Beckman Coulter LS6500 Multi purpose scintillation counter. A buffer blank and 50 μL aliquots of the initial dialysate solution were also counted. Bound ligand concentration was calculated from:

[Ibound] = [Iin] – [Iout] (1)

71 where [Iin] is the total concentration of free and bound efavirenz inside the dialysis

tubing or RED chamber, and [Iout] = [I] is the concentration of free efavirenz in the

outside dialysate. Scintillation counting data were converted to molarity and fit to

mathematical models in the Dialfit program as described in the Appendix. The value of

Ka(4) was fixed in the data analysis using ln Ka = 8.3 for p51/p51 homodimer and ln Ka =

12.4 for p66/p66 homodimer (14).

Isothermal Titration Calorimetry. ITC experiments were performed on a Microcal

VP-ITC microcalorimeter. Wild-type p51 solutions (1.5 and 3.0 μM) were titrated with efavirenz (200 μM) in RT buffer D containing 3% DMF at 5 ºC. Prior to the reaction p51 was dialyzed into RT buffer D containing 3% DMF to eliminate any solvent effects.

Aliquots of 5, 10, and 15 μL of the efavirenz solution were added over 60 min to a final concentration 40 μM. The amounts of heat released after each addition of efavirenz into the p51 solution and the buffer blank were identical, indicating that (1) the binding event is too slow to measure by this technique or (2) ΔΗ = 0.

Fluorescence. Absorbance was measured on a Cary 3E UV–vis

spectrophotometer at 5 °C. Fluorescence was measured on a PC1 photon counting

spectrofluorometer (ISS, Champaign, IL) in ratio mode under magic angle conditions

using 4-nm excitation and 16-nm emission bandwidths at 5 °C. The sample compartment

was flushed with nitrogen to prevent condensation. Samples were placed in 45 μL quartz cells with 3-mm path length (Starna Cells, Inc., Atascadero, CA). Absorbance at 280 nm was < 0.3 to avoid inner filter effects. Fluorescence quantum yields Φ were measured at

295-nm excitation wavelength relative to NATA in water with Φ = 0.23 at 5 °C. The

72 quantum yield of NATA at 5 °C was determined relative to tryptophan in water at 295-

nm excitation wavelength, 25 °C, with Φ = 0.14 (20).

Association and dissociation kinetics of RT proteins and efavirenz were

monitored by fluorescence using Vinci 1.6.SP7 software (ISS, Champaign, IL). Intrinsic

tryptophan fluorescence was measured at 295-nm excitation wavelength, 340-nm

emission wavelength using NATA in water as reference. Slow kinetic intensity data were collected from samples and NATA every 30 s (signal averaged over 5 s) for 4–5 h, then every 5 min (signal averaged over 10 s) for 27 h. Fluorescence intensity F = Is / Ir was calculated from the ratio of sample intensity Is to reference intensity Ir to correct for

instrumental drift.

Association reactions were started by adding 2 μL of a diluted efavirenz stock

solution (250 mM in DMF) to 80 μL of 2.5 μM p66W401A or p51W401A, 4.5 μM wild-type

p51, or 20 μM p66/p51 (85% dimer). The solution was mixed in the cell for 5 s and immediately placed in the fluorometer. Final efavirenz concentrations were 5–40 μM.

Dissociation reactions were started by 100-fold dilution of a 20 μM solution of p66W401A or p51W401A complexed with 35 μM efavirenz. The change in intrinsic tryptophan

fluorescence due to binding or unbinding of efavirenz was fit to a single exponential

function.

(F(t) – F0) / (F∞ – F0) = C (1 – exp[– kobs t]) (2a)

(F(t) – F∞) / (F0 – F∞) = C1exp[– kdiss t] + C2 (2b)

where F(t) is intensity at time t, F0 is intensity at t = 0, F∞ is intensity of the last time

point, and C is a constant.

73 Native Gel Electrophoresis. BN-PAGE was carried out using the Novex Bis-Tris gel system as described previously (21). A 5–10 μL aliquot of 2 μM p66W401A and 0.8–5

μM p66 in the absence or presence of NNRTI was mixed with 0.3 μL Coomassie G-250 sample additive, 2.5 μL NativePAGE Sample Buffer, and water to a final volume of 15

μL. Gels were stained with EZ-Run Protein Gel Staining solution and destained in water.

For gels containing [14C] efavirenz, p66 was incubated for 2 h or 1 week and subjected to

BN-PAGE. Gels were imaged by a PhosphorImager (Amersham Biosciences,

Piscataway, NJ), viewed with ImageQuant software, and then stained in EZ-Run Protein

Gel Staining solution and destained in water.

74 2.4 Results

Equilibrium Dialysis. Binding of efavirenz to p66 and p51 is coupled to formation

of homo- and heterodimers (Scheme 1). Dimerization constants in the absence and

presence of NNRTI are characterized by Kd(4) and Kd(2), while inhibitor dissociation constants of dimer and monomer complexes are Kd(1) and Kd(3). Dimerization constants

for p66/p66 and p51/p51 homodimers in the absence and presence of efavirenz were

previously determined by sedimentation equilibrium (14). Equilibrium dialysis was used

to determine inhibitor dissociation constants Kd(1) and Kd(3).

Equilibrium binding experiments were initially set up with p51, because the

dimerization constants in the absence and presence of efavirenz, Kd(4) = 230 μM for

p51/p51 and Kd(2) = 0.37 μM for p51/p51─I, provide access to both monomer and

homodimer. The first binding experiments used 10 µM p51 (7.5% homodimer) and 20

µM [14C] efavirenz. Dialysis was terminated and samples were analyzed at 30 h and at 3,

5 and 7 days. After 30 h the ratio of efavirenz to p51 was ~0.84:1, indicating a binding stiochiometry of either one inhibitor per p51 monomer or two inhibitors per p51/p51

homodimer. The ratio of efavirenz to p51 decreased to 0.68:1 after 3 days, 0.52:1 after 5

days, and 0.49:1 after 7 days. A ratio of one efavirenz per p51/p51 homodimer is

consistent with the stoichiometry in the crystal structure of p66/p51─EFV complex (9).

Due to the slow dimerization all experiments examining efavirenz binding to dimeric species were allowed to bind for 5 days. To confirm that the 30 h dialysis with wild-type p51 represents efavirenz binding to monomer, equilibrium dialysis experiments were also performed using dimerization defective RT proteins. Two dimerization defective mutations reported in the literature are L234A (22, 23), and W401A (19). L234A is a

75 primer grip mutation while W401A is a mutation in the tryptophan repeat motif of the

connection subdomain. The presence of either of these mutations in the p66 or p51

subunit of the heterodimer result in dimerization deficiency, the mutation in p66 having

the most detrimental effect. Equilibrium dialysis experiments set up with 3–6 µM of p51L234A and 5–12 µM of [14C] efavirenz failed to detect any bound efavirenz. Thus

L234A mutation prevents not only dimerization but also efavirenz binding. This is not

surprising given that L234 is a contact residue in the NNRTI binding pocket.

Inhibitor dissociation constants Kd(1) and Kd(3) were determined by

simultaneously varying protein and efavirenz concentrations in equilibrium dialysis

experiments. The data sets for multiple concentrations of protein and efavirenz were

analyzed with the Dialfit program (Appendix). The ln Ka(4) value, where Ka(4) is the equilibrium association constant of p51/p51 or p66/p66 homodimers, is set as a constant

(14) and ln Ka values for inhibitor binding to monomers and dimers are allowed to float.

The data sets for wild-type RT proteins equilibrated for 5 days with efavirenz were fit to the coupled equilibria in Scheme 1. Weighted least square fits were performed until the fits converged. Figure 1A shows the efavirenz binding data for wild-type p51 together

with the fit to eqs A1a – A1c for the coupled equilibria. The log of [Ibound] is plotted for

clarity; [Ibound] in μM was used in the data analysis. To illustrate the range of efavirenz and protein concentrations used in the experiments, the residuals [Ibound]exp –

[Ibound]calc are plotted versus total protein concentration (inset). Figure 1B shows the

efavirenz binding data for p51W401A and the fit to eq Ala for a simple binding equilibrium.

76 Figure 2-1. Equilibrium dialysis data for p51. (A) Wild-type p51 equilibrated with efavirenz for 5 days: (○) experimental values [Ibound]exp, and (●) calculated values

W401A [Ibound]calc from Dialfit using eqs A1a–Alc. (B) p51 equilibrated with efavirenz for

30 h: (○) [Ibound]exp, and (●) [Ibound]calc from Dialfit using eq A1a. Insets show

residuals ([Ibound]exp – [Ibound]calc) versus total protein concentration [Protein]Tot in each measurement.

-5.0 A

-5.5

-6.0 M

 -6.5 0.8 0.6

0.4

-7.0 M 0.2 

0.0 -7.5 -0.2

log 10 [Ibound], -0.4

-8.0 Residuals, -0.6

-0.8 -8.5 0246810 [Protein]Tot , M -9.0 0246810 [I], M

77 -5.0 B -5.2

-5.4

-5.6 2.5 M  -5.8 2.0

1.5 -6.0 M

 1.0

[Ibound], -6.2 10 0.5 log -6.4 0.0 Residuals, -6.6 -0.5

-1.0 -6.8 0246810 [Protein]Tot , M -7.0 0246810 [I], M

78 Table 2-1 shows the results of the global analyses for wild-type and dimerization

defective RT proteins. The dissociation constants Kd(1) and Kd(3) for efavirenz binding

to dimers and monomers were calculated from the average ln Ka values with Kd = 1/Ka.

The inhibitor dissociation constant Kd(1) of wild-type homodimer complexes is about

100-fold tighter for p51/p51─EFV than p66/p66─EFV: Kd(p51/p51─I) = 7 nM compared

to Kd(p66/p66─I) = 250 nM. The inhibitor dissociation constants Kd(3) of wild-type and

dimerization defective monomer complexes are much weaker, in the µM range. The

Kd(3) values for wild-type p66 and p51 measured after equilibration with efavirenz for 5

days are inaccurate. At high protein concentrations the free monomer concentration is too

low to detect, and at low protein concentrations binding of efavirenz to monomer is too

weak to detect. The data set for wild-type p51 equilibrated for 30 h with efavirenz and

analyzed neglecting the dimerization reaction provides a more reliable value for

Kd(p51─I) = 1.7 µM, because at 30 h wild-type p51 is ~70% monomers. The data sets for

the dimerization defective mutants were also fit neglecting the dimerization reaction. The

W401A W401A inhibitor dissociation constants Kd(3) for p51 and p66 monomers are

approximately equal: Kd(p51─I) = 2.4 µM and Kd(p66─I) = 2.7 µM. The similarity

between wild-type and W401A values obtained for Kd(3) suggest that the W401A

substitution does not alter the NNRTI binding site.

79

Table 2-1.

Table 2-1: Equilibrium Dialysis Dissociation Constantsa

protein Dialysis lnKa(1) Kd (1), µM lnKa(3) Kd (3), µM

time

p51 30 h 17.3 ± 0.6 0.030 13.26 ± 0.6 1.7

(0.017 – 0.056) (0.95 – 3.2)

p51 5 days 18.9 ± 0.8 0.0068 10.5 ± 1.0 28

(0.0028 – 0.014) (10.1 – 75.0)

p51W401A 30 h 12.93 ± 0.6 2.5

(1.3 – 4.4)

p66 5 days 15.2 ± 0.5 0.25 10.9 ± 1.2 19

(0.15 – 0.41) (5.6 – 61)

p66W401A 30 h 12.83 ± 0.3 2.7

(2.0 – 3.6)

a In 0.05 M Tris, pH 6.5, 25 mM NaCl, 1 mM EDTA, 1 mM TCEP, and 10% glycerol

at 5 ºC. Data sets collected for each protein and dialysis time are analyzed with the

Dialfit program (see Appendix). Errors are 95% confidence intervals.

80 Change in Intrinsic Fluorescence Upon Binding Efavirenz. RT contains multiple tryptophan residues, 19 in p66 and 18 in p51 (Figure 2-2).

81 Figure 2-2 Structure of HIV-1 RT complexed with efavirenz (1FK9): p66 (purple), p51

(green), efavirenz (cyan), and tryptophans (red).

82 Tryptophan fluorescence is exquisitely sensitive to the local electrostatic environment of

the indole chromophore (24, 25). Changes in RT fluorescence associated with

dimerization and NNRTI binding have been reported (26, 27). The fluorescence changes due to dimerization were attributed to the tryptophan repeat motif in the connection subdomain spanning residues 398–414. The NNRTI-binding pocket region contains - sheet β12–β13–β14, which has two tryptophans W229 and W239; W229 is in the loop between β-strands 12 and 13 and W239 is in β-strand 14. These two tryptophans may report conformational changes upon inhibitor binding.

The kinetics of inhibitor binding to RT proteins were monitored by fluorescence.

Figure 2-3 plots fluorescence versus time for efavirenz binding to p51 monomer and p66/p51 heterodimer.

83 Figure 2-3. Association of efavirenz to (─) p66/p51 and (─) p51 monitored by tryptophan fluorescence at 5 ºC. λex = 295 nm, λem = 340 nm. 20 µM p66/p51, 83%

heterodimer diluted to 2 µM prior to adding efavirenz; 4.5 µM p51, 97% monomer.

[EFV]:[protein] 2:1.

84 In order to measure heterodimer fluorescence, a 20 μM solution containing 83% dimer was diluted 10-fold and efavirenz was added immediately to start the kinetics experiment before dissociation of the dimer occurs (t1/2 = 2 days; 28) The overall intensity change for the heterodimer is about half that of the monomer, consistent with efavirenz only affecting tryptophan residues in the p66 subunit.

Progress Curve Analysis of Efavirenz Binding. Two kinetic mechanisms have been used to account for the slow binding of inhibitors to enzymes (Scheme 2-2; 29, 30).

85 Scheme 2-2. Mechanisms of Slow Binding Inhibitors

86 Mechanism A depicts direct, reversible binding of inhibitor I to enzyme E, where the

association and dissociation rate constants k1 and k–1 are inherently slow. Mechanism B

depicts an induced-fit model with fast equilibration of inhibitor and enzyme to form an

intermediate complex EI, followed by slow isomerization of EI complex to form the final

complex EI*. To discriminate between the two mechanisms, the observed rate constant

kobs from the progress curve of the enzyme reaction, is determined as a function of

inhibitor concentration. A plot of kobs versus inhibitor concentration is linear for

Mechanism A and hyperbolic for Mechanism B.

Progress curves were measured for p66W401A, p51W401A, and p66/p51 at multiple concentrations of efavirenz. Figure 2-4 shows the set of curves for p51W401A. The solid

lines are the fits to eq 2a. Similar curves were obtained for p66W401A and p66/p51.

87 Figure 2-4 Progress curves for p51W401A binding to efavirenz monitored by tryptophan

W401A fluorescence at 5 ºC. λex = 295 nm, λem = 340 nm. 2.5 µM p51 ; (─) 5 µM, (─) 8

µM, (─) 11 µM, and (─) 14 µM efavirenz. Data acquired at 30 s intervals for first 4–5 h,

then at 5 min intervals. Data were fit to eq 2a to obtain kobs.

88 Figure 2-5 shows the plots of kobs versus inhibitor concentration. The linear fits are consistent with Mechanism A, where

kobs = k–1 + k1[I] (3)

The values of the rate constants k1 and k–1 calculated from the slopes and intercepts of

Figure 5 are given in Table 2-2. All three proteins have similar association rate constants

–1 –1 k1 of 13.5 M s . Additionally, the dissociation rate constants k–1 were about 5.9–8.1 

–5 –1 10 s , corresponding to t1/2 ~2.7 h. Having defined the binding modality of efavirenz,

app the values of Kd(3) were calculated from the ratio of k–1/k1.

89 W401A Figure 2-5. Dependence of kobs on efavirenz concentration (●) p51 , (▲) p66/p51, and (■) p66W401A. Data from duplicate progress curves at each inhibitor concentration were averaged; solid lines are fits to eq 3.

3.0

2.8

2.6

2.4 -1

s 2.2 - 4

2.0 10 1.8 obs k 1.6

1.4

1.2

468101214 EFV, µM

90 Table 2-2.

Table 2-2: Kinetic Parameters of Efavirenz Bindinga

b –1 –1 c –5 –1 app, d e –5 –1 Protein k1 , M s k–1 , 10 s Kd(3) , µM k –1(diss) , 10 s

p51W401A 13.7 ± 0.7 5.9 ± 0.7 4.2 ± 0.5 9.1 ± 0.2

6.6 ± 0.5 f

p66W401A 13.4 ± 0.6 8.1 ± 0.8 6.0 ± 0.3 8.9 ± 0.2

6.6 ± 0.3 f

p66/p51 13.3 ± 0.9 6.7 ± 0.9 5.0 ± 0.3 nd a In 0.05 M Tris, pH 6.5, 25 mM NaCl, 1 mM EDTA, 1mM TCEP, 10% glycerol, and varying concentrations of EFV at 5 ºC. b Slope and error from Figure 4. c Intercept and

d e error in intercept from Figure 4. Kd(3) = intercept/slope = k–1/ k1. Dissociation

f kinetics. Error is range of average values for two experiments. Kd(3) = k–1(diss)/k1.

91 The kinetics of dissociation of p66W401A─ and p51W401A─EFV complexes were measured under essentially irreversible conditions, so that at equilibrium < 3% of monomer─EFV complex is present (Figure 2-6).

92 Figure 2-6. Dissociation of (─) p51W401A─EFV and (─) p66W401A─EFV complexes monitored by tryptophan fluorescence 5 ºC. λex = 295 nm, λem = 340 nm. Red curves are

fits to eq 2b. Data acquired at 30 s intervals for first 4–5 h, then at 5 min intervals.

1.0

) 0.8 0 F

- 0.6 inf F (

/

) 0.4 0 - F )

0.2 t ( (F 0.0

02468101214 Time, h

93 –5 –1 Dissociation rate constants k–1(diss) of 8.9–9.1 ± 0.2  10 s , or t1/2 = 2.1 h, for both

monomers were determined from fitting the curve to eq 2b. The k–1(diss) values from

kinetics measurements are close to the k–1 values determined from the plots of kobs versus

[I] (Table 2-2). The equilibrium dissociation constants Kd(3) calculated from the ratio of the rate constants k–1(diss)/k1 for monomer binding are 2.5-fold higher than the value

determined by equilibrium dialysis.

Fluorescence Quantum Yields. Fluorescence quantum yields of dimerization

defective monomers and wild-type heterodimer were measured in the absence and

presence of efavirenz. To measure the extent of quenching, most of the protein must be

bound to efavirenz. The quantum yields of monomer─EFV complexes were measured on

solutions containing 1 µM monomer and 30 µM efavirenz equilibrated for 30 h at 5 °C,

giving 94% monomer─EFV complex. The quantum yields of p66W401A and p51 W401A monomers are the same within error (Table 2-3).

94 Table 2-3.

Table 2-3: Quantum Yields of Efavirenz Binding

Protein Φ

p51W401A 0.14 ± 0.01

+ efavirenz 0.05 ± 0.02

p66W401A 0.15 ± 0.01

+ efavirenz 0.05 ± 0.02

p66/p51 0.11 ± 0.01

+ efavirenz 0.07 ± 0.01

NATA 0.23

In 0.05 M Tris, pH 6.5, 25 mM NaCl, 1 mM

EDTA, 1 mM TCEP, and 10% glycerol at 5 ºC, λex

= 295 nm. Errors are standard deviations of 4

experiments.

95 Efavirenz binding decreases the quantum yield of both monomers by a factor of 3. In

order to measure the quantum yield of the heterodimer, a 20 μM solution containing 83%

dimer was diluted 50-fold and scanned immediately as above. The quantum yield of the

heterodimer is approximately 20% lower than that of the monomers. The quantum yield

of p66/p51─EFV complex was measured on solutions containing 20 µM p66/p51 and 40

µM efavirenz equilibrated for 1 wk at 5 °C prior to dilution. Because efavirenz enhances

dimerization 25-fold and binds more tightly to dimer than monomer, this solution

contains 98% p66/p51─EFV complex. Efavirenz binding to heterodimer only quenches

the fluorescence by a factor of 1.6.

Native Gel Electrophoresis of NNRTI Binding. BN-PAGE has been used to

monitor dimerization of RT proteins in the absence and presence of efavirenz (21). The

slow dissociation rate of efavirenz (t1/2 ~2 h) makes it possible to visualize binding of

[14C] efavirenz to monomer on gels (Figure 2-7 A and B). The p66W401A and wild-type

p66 were incubated with a 0.7:1.0 ratio of [14C] efavirenz to protein. The wild-type p66 concentration was 5 µM or approximately 53% homodimer. Lane 1 shows [14C] efavirenz

binding to p66W401A monomer. Lane 2 shows [14C] efavirenz binding to the mixture of

wild-type p66 monomer and p66/p66 homodimer. Lastly, Lane 3 shows enhancement of

dimerization by efavirenz after equilibration of wild-type p66 with [14C] and excess cold

efavirenz for one wk, giving 91% p66/p66─EFV complex. Thus BN-PAGE supports the conclusions from equilibrium dialysis that efavirenz binds RT monomers as well as homodimers.

Nevirapine has been reported to have disparate effects on RT dimerization. Yeast-

two hybrid experiments indicate small enhancement of dimerization, whereas urea

96 denaturation studies find no effect (16, 23). BN-PAGE was performed using p66 incubated with excess efavirenz or nevirapine for 1 wk. Figure 2-7B shows that both

NNRTIs enhance dimerization with efavirenz having the greater effect. These results are consistent with the findings by yeast-two hybrid.

97 Figure 2-7. Blue native polyacrylamide gel electrophoresis of p66 in the absence and

presence of NNRTIs. (A) Monomer and homodimer binding to [14C] EFV; (lane 1) 2 µM

p66W401A + [14C] EFV incubated 2 h, (lane 2) 5 µM p66 + [14C] EFV incubated 2 h and

(lane 3) 5 µM p66 + [14C] EFV for 1 wk (lane 3). (B) wild-type p66 incubated in the

absence and presence of excess NNRTI for 1 wk; (lane 1) Native Markers, (lane 2) 0.8

µM p66, (lane 3) 3 µM p66 + EFV, (lane 4) 3 µM p66 + NVP.

98 2.5 Discussion

Although RT has been extensively studied for almost two decades, new functions

of this enigmatic enzyme continue to be discovered. This chapter reports two novel

functions: (1) efavirenz, and presumably also other NNRTIs, binds to monomeric forms

of RT and (2) efavirenz is a slow binding inhibitor of heterodimer and monomers. The

biological significance of monomer binding is presently unknown. NNRTIs have been

found to affect both early and late stages of the HIV-1 replication cycle by multiple

mechanisms (31, 32). Efavirenz interacts at the level of reverse transcription by inhibiting

DNA polymerase activity, enhancing polymerase-dependent RNase H activity (3'-DNA directed) and partially inhibiting polymerase-independent RNase H activity (5'-RNA

directed). It also inhibits plus-strand initiation by affecting the ability of RT to bind the

RNA polypurine tract. During late stages of HIV-1 replication, efavirenz enhances processing and homodimerization of a 90 kDa Pol polyprotein in a yeast two-hybrid system and increases intracellular processing of Gag and Gag-Pol precursor polyproteins in HIV-1-transfected cells (11). By increasing the processing of these polyproteins,

efavirenz lowers viral production by reducing the amount of the full constructs that would become incorporated into a budding particle. Essential to the above processes is

defining the binding properties of the species, whether monomer or dimer, to which

efavirenz binds. Drug design requires an immense understanding of the target. This study

suggests that monomeric forms of RT may be potential targets for HIV-1 therapeutics. It

also sparks development of high throughput screening assays based on p66 and p51

monomers to evaluate binding of new drugs to wild-type and drug resistance mutant RTs.

99 The two crystal structures of RT─EFV complexes show 1:1 binding stoichiometry (9, 33). Currently no crystal structures are available for homodimers or monomers of RT. Equilibrium dialysis indicated a 1:1 stoichiometry for p66/p66─ and

p51/p51─EFV complexes. A 1:1 binding stoichiometry for monomer─EFV complexes

was also obtained by equilibrium dialysis for wild-type p51 and dimerization-deficient

p66W401A and p51W401A, albeit with a lower affinity than the homodimers (Table 2-1). The

278 apparent free energies of efavirenz binding to homodimers at 5 °C, ΔG = –RTlnKa, are

–35.1 kJ/mol for p66/p66─I and –43.6 kJ/mol for p51/p51─I. The more favorable binding energy of the p51/p51 homodimer may be attributed to better contacts between efavirenz and the protein or more facile formation of the binding pocket in a dimer lacking 2 RNase H domains. All monomers have similar efavirenz binding energies,

ΔG278 ~ –30 kJ/mol, which is 5–12 kJ/mol less favorable than binding to homodimers.

The dissociation constants Kd(1) and Kd(3) from equilibrium dialysis (Table 2-1)

and Kd(2) and Kd(4) from previous sedimentation equilibrium experiments, allow us to

complete the thermodynamic linkage of NNRTI binding and subunit dimerization

proposed for RT (14). In the closed cycle of Scheme 1, ∆G = 0. Substituting ∆G278 gives

–RT ln Ka(2) – RT ln Ka(3) = –RT ln Ka(1) – RT ln Ka(4) (4)

In the case of p66, the left side of eq 4 sums to –68 ± 6 kJ/mol and the right side to –64 ±

4 kJ/mol. In the case of p51, the left side of eq 4 totals –64 ± 4 kJ/mol and the right side –

63 ± 6 kJ/mol. These results for homodimers confirm the hypothesis that NNRTI binding

is coupled to subunit dimerization. We can then calculate the dissociation constant of the p66/p51─EFV complex Kd(1) from the cycle in Scheme 1, where Kd(2) and Kd(4) are the

dissociation constants of the heterodimer in the presence and absence of EFV (14), and

100 Kd(3) is the dissociation constant of p66─ or p51─EFV complexes. Rearranging eq 4

gives RT lnKa(1) = 38 ± 6 kJ/mol or Kd(1) = 92 nM ± 5 nM. Most studies of efavirenz

binding to RT have employed polymerase activity assays, carried out in the presence of

template/primer and dNTP (12, 34, 35). Maga et al. (35) reported a dissociation constant

of 150 nM for free RT─EFV complex extracted from enzymatic data. Geitmann et al.

(36) measured binding of several NNRTIs to immobilized wild-type and drug resistance

mutant RTs by SPR at 25 °C in buffer containing 0.005% surfactant and 3% (v/v)

DMSO. An overall dissociation constant of 45 nM was obtained for efavirenz binding to

wild-type RT.

The binding kinetics monitored by tryptophan fluorescence establish that

efavirenz is a slow, tight binding inhibitor of RT. Dissociation rate constants k–1(diss) = 9

–5 –1  10 s , or t1/2 = 2.2 h, were obtained for monomer─EFV complexes. Association rate

–1 –1 constants k1 ~ 13.5 M s were obtained for a reversible, direct binding reaction of

efavirenz to both monomers and heterodimer (Scheme 2, Mechanism A). This suggests

that formation of the NNRTI binding pocket occurs by an analogous process for

monomeric and dimeric species of RT, despite the 25-fold difference in efavirenz binding

affinity. Slow binding inhibitors described in the literature follow either direct binding or

conformational change inhibition models (Scheme 2). For example, small azasugar

inhibitors of β-glucosidase and yeast isomaltase bind by the direct Mechanism A with

association rate constants ranging from 23 M–1 s–1 to 7.3  104 M–1 s–1 (37). These inhibitors also have slow dissociation rate constants of 0.16–6.7  10–2 s–1. By contrast,

peptide α-ketoacid analogues are slow binding inhibitors of hepatitis C virus NS3

protease that bind by the induced-fit Mechanism B (3). These inhibitors undergo rapid

101 equilibration with the enzyme in the first step of EI complex formation with k1 = 6.5 

7 –1 –1 –1 10 M s and k–1 = 0.2 s . The subsequent step is a slow isomerization to EI* with k2 =

–3 –1 –5 –1 1.7–7.5  10 s and k–2 = 0.57–1.8  10 s or t1/2 = 11–48 h.

A few previous reports noted slow onset of inhibition by NNRTIs. For example,

5–20 min pre-incubation periods of RT with NNRTIs were required to witness inhibition of polymerase and RNase H activity (31, 35, 39, 40). The slow association rate constants

reported here (Table 2-2) could be caused by a conformational selection step involving exclusive binding of a lowly populated conformer of either protein or inhibitor. The chemical structure of efavirenz is provided in Figure 2-2. The benzoxazinone ring system is rigid with free rotation of the cyclopropyl ethynyl group (41). Efavirenz is in the same

position in the binding pocket in both crystal structures of wild-type RT─EFV complex.

However, the cyclopropyl ethynyl group is rotated ~100º in the drug resistance mutant

K103N RT─EFV complex relative to the position in the wild-type structures (9). The rigidity of the efavirenz core together with the ability of the binding pocket to accommodate different orientations of the cyclopropyl ethynyl group excludes conformational selection of the inhibitor as the culprit.

A selected-fit model has been proposed in which a conformational pre- equilibration of the protein precedes inhibitor binding (36, 42). In this model for the slow

binding, efavirenz would bind preferentially to a less populated conformational state of

RT proteins. Productive collisions occurring between inhibitor and this conformational state would induce a slow shift in the conformational equilibrium favoring formation of the binding pocket and subsequently the EI complex. A less likely alternative would be severe orientation effects resulting in unproductive collisions of E and I that slow

102 formation of EI complex in the direct binding Mechanism A. A selected-fit model gave the best fit to the SPR data for wild-type RT and efavirenz with an overall association

4 –1 –1 rate constant kon = 5.5  10 M s . This kon value is 3 orders of magnitude faster than our association rate constant k1 for efavirenz binding to p66/p51. The dissociation rate

–3 –1 constant koff ≈ 2.3  10 s is about 20-fold faster than the k–1 value obtained from progress curve analysis. A probable explanation for the faster binding kinetics is the

different solution conditions used in the SPR assay. Osmolytes such as detergents,

organic solvents, and salts affect protein solution structure and interactions (43, 44).

The NNRTI binding pocket is absent from structures of RT and RT─substrate complexes (45, 46). The polymerase domain is composed of four subdomains: fingers,

palm, thumb, and connection. In the asymmetric structure of the heterodimer the

subdomains are in different orientations in the p66 and p51 subunits. The positions of

these subdomains in each subunit in the absence of efavirenz, is shown in Figure 2-8

(upper). In RT–EFV complexes, the binding pocket is in the palm of the p66 subunit in

RT─NNRTI complexes (Figure 2-8 (lower), 9).

103 Figure 2-8. Structures of HIV-1 (upper) RT (1DLO) and (lower) RT─EFV complex

(1FK9). Polymerase domains: fingers (blue), palm (red), thumb (green), and connection

(orange) subdomains; RNase H domain (magenta). Efavirenz (yellow) and contact residues (grey) are shown using the van der Waals radii.

104 Efavirenz contacts 14 residues in the NNRTI binding pocket of the p66 subunit of the heterodimer: L100, K101, K103, V106, V179, Y181, Y188, G190, F227, W229, L234,

H235, P236 and Y318. The ligand-protein contacts were derived with LPC software (47).

The efavirenz contact residues are highlighted in the two subunits to illustrate the relative locations. Although the contacts residuals are clustered in the p51 subunit efavirenz does not bind at this site. It can be hypothesized that the p66/p66 and p51/p51 homodimers probably also have asymmetric structures because they both have polymerase activity (3).

Additionally they probably have similar NNRTI binding pockets in the subunit that binds the inhibitor, because p66 and p51 have identical amino acid sequences and similar folding patterns of the polymerase subdomains. Given that p66 and p51 monomers are capable of forming a competent NNRTI binding pocket, presumably the polymerase domain of both monomers must adopt a conformation analogous to that of the p66 subunit in the heterodimer.

105 2.6 Appendix

Dialfit: Mathematical Models and Data Analysis Leslie A Holladay§

There are two different ways in which the model equations may be written: Case

A and Case B.

Case A. For Case A the following equilibrium constants are defined as:

P + I = PI Ka(3) = [PI] / ([P] [I]) [PI] = Ka(3) [P] [I] (A1a)

2 2 P + P = PP Ka(4) = [PP] / [P] [PP] = Ka(4) [P] (A1b)

PP + I = PPI Ka(1) = [PPI] / ([PP] [I]) [PPI] = Ka(1) [PP] [I] (A1c)

The equilibrium dialysis experiment involves computing the total molar

concentration of inhibitor bound to both monomer and dimer [Ibound], by subtracting the concentration of free inhibitor outside the dialysis bag [Iout] = [I] from the total concentration of inhibitor inside the bag [Iin] as in eq 1. For the data analysis, the observable variable we wish to model is [Ibound]. The total concentration of protein

[P]tot inside the bag is presumed to be known to higher precision than that of the free and

bound inhibitor concentrations. [Ibound] is the observable to be predicted knowing [I]

and [P]tot along with the current estimates for Ka(1), Ka(3), and Ka(4). Dialfit is

applicable to any experiment that provides data for [Ibound] and [I], knowing [P]tot.

To compute [Ibound], the concentration of free protein monomer [P] must first be

computed. The conservation of mass equation is

[P]tot = [P] + [PI] + 2 [PP] + 2 [PPI] (A2)

Rearranging and substituting terms with the equilibrium constants from eqs A1 results in

a quadratic in [P],

a [P]2 + b [P] + c = 0 (A3a)

106 where

a = 2 Ka(4) + 2 Ka(1) Ka(4) [I] (A3b)

b = 1 + Ka(3) [I] (A3c)

c = – [P]tot (A3d)

The only physical meaningful root of eq (A3a) is the positive one. The fitting function may be written as

2 [Ibound] = Ka(3) [P] [I] + Ka(1) Ka(4) [P] [I] (A4)

The values of [Ibound] determined over a wide range of total inhibitor and total protein

concentrations are globally fitted to eq A4. Note that Ka(1) and Ka(4) cannot be separated and thus one value must be a fixed parameter.

Case B. For Case B two equilibrium constants are defined in eqs (A1a) and

(A1b); the third equilibrium constant is defined as

PI + P = PPI Ka(2) = [PPI] / ([PI] [P]) [PPI] = Ka(2) [PI] [P](A5)

Here too [Ibound] is the observable to be predicted knowing [I] and [P]tot, but with the

current estimates for Ka(2), Ka(3), and Ka(4). The conservation of mass eq (A2) and

quadratic eqs (A3a), (A3c), and (A3d) are the same as in Case A; eq (A3b) becomes

a = 2 Ka(4) + 2 Ka(2) Ka(3) [I] (A6)

Again, the physically meaningful root of eq (A3a) is the positive one. The fitting function

is

2 [Ibound] = Ka(3)[P] [I] + Ka(2) Ka(3)[P] [I] (A7)

107 Note that Cases A and B are mathematically equivalent from the relationship Ka(1) Ka(4)

= Ka(2) Ka(3). The two parameters Ka(2) and Ka(3) are not separable, and thus the value

of Ka(2) must be fixed.

Weighted Least Squares and Parameter Standard Errors. The global data sets

have a very wide range of values for [Ibound] and [I]. For radioactive counts, the relative standard deviation is equal to √N/N, where N is the number of counts (48). The relative

variance is equal to 1/N. [Ibound] is computed from the difference in counts inside and

outside the bag. Define the number of counts inside the bag as Ni and the number of

counts outside the bag as No. Then the relative variance of the difference (Ni – No) = (Ni

+ No) / (NiNo). In the situation in which the values for [Ibound] and [I] vary over several

orders of magnitude, it is essential to use weighted least squares because the variance will

also vary over a wide range (49). The weight Wj of any (Ni – No)j value is the reciprocal

of the variance, Wj = (NiNo) / (Ni +No). The actual weights used are normalized so that

the j Wj = 1 to cause the returned residual error in the fit to be correct. It is clear from in

Figure 1 (insets) that the errors in [Ibound] are very heteroscedastic, The errors in the

fitted variable vary a lot with respect to the independent variable. Only if the errors are

homoscedastic is unweighted least squares appropriate.

Standard errors for the parameter values were computed using the “balanced

bootstrap” with 100 trials (50, 51). If any of the individual bootstrap trials was more than three standard deviations away from the parameter estimate that trial was deleted as an outlier and the standard deviation recomputed. The 95% confidence intervals are computed using the standard deviation from the estimated parameter value.

108 2.7 Acknowledgements

The authors are grateful to Ms. Kathryn J. Howard for suggesting the gel

experiment with [14C] efavirenz and for advice about site-directed mutagenesis and protein purification. We also thank Dr. Clare Woodward for suggesting that NNRTIs bind to monomers and Drs. Vernon Anderson and Jonathan Karn for helpful discussions.

109 2.8 References

1. di Marzo Veronese, F., Copeland, T. D., DeVico, A. L., Rahman, R., Oroszlan, S.,

Gallo, R. C., and Sarngadharan, M. G. (1986) Characterization of highly

immunogenic p66/p51 as the reverse transcriptase of HTLV-III/LAV, Science

231, 1289–1291.

2. Hizi, A., McGill, C., and Hughes, S. H. (1988) Expression of soluble,

enzymatically active, human immunodeficiency virus reverse transcriptase in

Escherichia coli and analysis of mutants, Proc. Natl. Acad. Sci. U.S.A. 85, 1218–

1222.

3. Restle, T., Muller, B., and Goody, R. S. (1990) Dimerization of human

immunodeficiency virus type 1 reverse transcriptase, J. Biol. Chem. 265, 8986–

8988.

4. Restle, T., Muller, B., Goody, R. S. (1992) RNase H activity of HIV reverse

transcriptase is confined exclusively to the dimeric forms, FEBS Lett. 300, 97–

100.

5. Le Grice, S. F. J., Naas, T., Wohlgensinger, B., and Schatz, O. (1991) Subunit-

selective mutagenesis indicates minimal polymerase activity in heterodimer-

associated p51 HIV-1 reverse transcriptase, EMBO J. 10, 3905–3911.

6. Tsai, C. H., Lee, P. Y., Stollar, V., and Li, M. L. (2006) Antiviral therapy

targeting viral polymerase, Curr. Pharm. Des. 12, 1339–1355.

7. De Clercq, E. (1998) The role of non-nucleoside reverse transcriptase inhibitors

(NNRTIs) in the therapy of HIV-1 infection, Antiviral Res. 38, 153–179.

110 8. Kohlstaedt, L. A., Wang, J., Friedman, J. M., Rice, P. A., and Steitz, T. A. (1992)

Crystal structure at 3.5 Å resolution of HIV-1 reverse transcriptase complexed

with an inhibitor, Science 256, 1783–1790.

9. Ren, J., Milton, J., Weaver, K. L., Short, S. A., Stuart, D. I., and Stammers, D. K.

(2000) Structural basis for the resilience of efavirenz (DMP-266) to drug

resistance mutations in HIV-1 reverse transcriptase, Structure 8, 1089–1094.

10. Merluzzi, V. J., Hargrave, K. D., Labadia, M., Grozinger, K., Skoog, M., Wu, J.

C., Shih, C.-K., Eckner, K., Hattox, S., Adams, J., Rosenthal, A. S., Faanes, R.,

Eckner, R. J., Koup, R. A., and Sutton, J. L. (1990) Inhibition of HIV-1

replication by a nonnucleoside reverse transcriptase inhibitor, Science 250, 1411–

1413.

11. Figueiredo, A., Moore, K. L., Mak, J., Sluis-Cremer, N., de Bethune, M.-P., and

Tachedjian, G. (2006) Potent nonnucleoside reverse transcriptase inhibitors target

HIV-1 Gag-Pol. PLoS Pathog. 2, 1051–1059.

12. Young, S. D., Britcher, S. F., Tran, L. O., Payne, L. S., Lumma, W. C., Lyle, T.

A., Huff, J. R., Anderson, P. S., Olsen, D. B., Carroll, S. S., Pettibone, D. J.,

O’Brien, J. A., Ball, R. G., Balani, S. K., Lin, J. H., Chen, I.-W., Schleif, S. A.,

Sardana, V. V., Long, W. J., Byrnes, V. W., and Emini, E. A. (1995) L-743,726

(DMP-266): A novel, highly potent nonnucleoside inhibitor of the human

immunodeficiency virus type 1 reverse transcriptase, Antimicrob. Agents

Chemother. 39, 602–2605.

13. Tachedjian, G., Orlova, M., Sarafianos, S. G., Arnold, E., and Goff, S. P. (2001)

Nonnucleoside reverse transcriptase inhibitors are chemical enhancers of

111 dimerization of the HIV type 1 reverse transcriptase, Proc. Natl. Acad. Sci. U.S.A.

98, 7188–7193.

14. Venezia, C. F., Howard, K. J., Ignatov, M. E., Holladay, L. A., and Barkley, M.D.

(2006) Effects of efavirenz binding on the subunit equilibria of HIV-1 reverse

transcriptase, Biochemistry 45, 2779–2789.

15. Sluis-Cremer, N., Arion, D., and Parniak, M. A. (2002) Destabilization of the

HIV-1 reverse transcriptase dimer upon interaction with N-acyl hydrazone

inhibitors, Mol. Pharmacol. 62, 398–405.

16. Sluis-Cremer, N., Dmitrienko, G. I., Balzarini, J., Camarasa, M.-J., and Parniak,

M. A. (2000) Human immunodeficiency virus type 1 reverse transcriptase dimer

destablilization by 1-spiro[4-amino-2,2-dioxo-1,2-oxathiole-5,3-[2,5-

bis-O-(tert-butyldimethylsilyl)-β-D-ribofuranosyl]]-3-ethylthymine,

Biochemistry 39, 1427–1433.

17. Le Grice, S. F. J., Cameron, C. E., and Benkovic, S. J. (1995) Purification and

characterization of human immunodeficiency virus type 1 reverse transcriptase.

Methods Enzymol. 262, 130–144.

18. Ignatov, M. E., Berdis, A. J., Le Grice, S. F. J., and Barkley, M. D., (2005)

Attenuation of DNA replication by HIV-1 reverse transcriptase near the central

termination sequence. Biochemistry 44, 5346–5356.

19. Tachedjian, G., Aronson, H.-E., de los Santos, M., Seehra, J., McCoy, J. M., and

Goff, S. P. (2003) Role of residues in the tryptophan repeat motif for HIV-1

reverse transcriptase dimerization, J. Mol. Biol. 326, 381–396.

112 20. Chen, R. F. (1967) Fluorescence quantum yields of tryptophan and tyrosine, Anal.

Lett. 1, 35–42.

21. Braz, V. A., and Howard, K. J. (2009) Separation of protein oligomers by blue

native gel electrophoresis, Anal. Biochem. 388, 170–172.

22. Ghosh, M., Jacques, P. S., Rodgers, D. W., Ottman, M., Darlix, J. L., and Le

Grice, S. F. J. (1996) Alterations to the primer grip of p66 HIV-1 reverse

transcriptase and their consequences for template-primer utilization, Biochemistry

35, 8553–8562.

23. Tachedjian, G., Aronson H.-E. G., Goff, S. P. (2000) Analysis of mutations and

suppressors affecting interactions between subunits of the HIV type 1 reverse

transcriptase, Proc. Natl. Acad. Sci. U.S.A. 97, 6334–6339.

24. Vivian, J. T., and Callis, P. R. (2001) Mechanisms of tryptophan fluorescence

shifts in proteins, Biophys. J. 80, 2093–2109.

25. Callis, P. R., Petrenko, A., Muino, P. L., and Tusell, J. R. (2007) Ab initio

prediction of tryptophan fluorescence quenching by protein electric field enabled

electron transfer, J. Phys. Chem. B 111, 10335–10339.

26. Divita, G., Restle, T., and Goody, R. S. (1993) Characterization of the

dimerization process of HIV-1 reverse transcriptase heterodimer using intrinsic

protein fluorescence, FEBS Lett. 324, 153–158.

27. Barnard, J., Borkow, G., and Parniak, M. A. (1997) The thiocarboxanilide

nonnucleoside UC781 is a tight-binding inhibitor of HIV-1 reverse transcriptase,

Biochemistry 36, 7786–7792.

113 28. Venezia, C. F., Meany, B. J., Braz, V. A., and Barkley, M. D. (2009) Kinetics of

association and dissociation of HIV-1 reverse transciptase subunits, Biochemistry

in press.

29. Copeland, R. A. (2005) Evaluation of Enzyme Inhibitors in Drug Discovery, John

Wiley & Sons, New Jersey.

30. Szedlacsek, S. E., and Duggleby, R. G. (1995) Kinetics of slow and tight-binding

inhibitors, Methods Enzymol. 249, 144–180.

31. Sluis-Cremer, N., and Tachedjian, G. (2008) Mechanisms of inhibition of HIV

replication by non-nucleoside reverse transcriptase inhibitors, Virus Res. 134,

147–156.

32. Grobler, J. A., Dornadula, G., Rice, M. R., Simcoe, A. L., Hazuda, D. J., and

Miller, M. D. (2007) HIV-1 reverse transcriptase plus-strand initiation exhibits

preferential sensitivity to non-nucleoside reverse transcriptase inhibitors in vitro,

J. Biol. Chem. 282, 8005–8010.

33. Lindberg, J., Sigurdsson, S., Löwgren, S., Andersson, H. O., Sahlberg, C.,

Noréen, R., Fridborg, K., Zhang, H., and Unge, T. (2002) Structural basis for the

inhibitory efficacy of efavirenz (DMP-266), MSC194 and PNU142721 towards

the HIV-1 RT K103N mutant, Eur. J. Biochem. 269, 1670–1677.

34. Xia, Q., Radzio, J., Anderson, K. S., and Sluis-Cremer, N. (2007) Probing

nonnucleoside inhibitor-induced active site distortion in HIV-1 reverse

transcriptase by transient kinetics analysis, Protein Sci. 16, 1728–1737.

35. Maga, G., Ubiali, D., Salvetti, R., Pregnolato, M., and Spadari, S. (2000)

Selective interaction of the human immunodeficiency virus type 1 reverse

114 transcriptase nonnucleoside inhibitor efavirenz and its thio-substituted analog

with different enzyme-sustrate complexes, Antimicrob. Agents Chemother. 44,

1186–1194.

36. Geitmann, M., Unge, T., and Danielson, H. (2006) Interaction kinetic

characterization of HIV-1 reverse transcriptase non-nucleoside inhibitor

resistance, J. Med. Chem. 49, 2375–2387.

37. Lohse, A., Hardlei, T., Jensen, A., Plesner, I. W., and Bols, M. (2000)

Investigation of the slow inhibition of almond β-glucosidase and yeast isomaltase

by 1-azasugar inhibitors: evidence for the ‘direct binding’ model, Biochem. J.

349, 211–215.

38. Narjes, F., Brunetti, M., Colarusso, S., Gerlach, B., Koch, U., Biasiol, G., Fattori,

D., De Francesco, R., Matassa, V. G., and Steinkühler, C. (2000) α-Ketoacids are

potent slow binding inhibitors of the hepatitis C virus NS3 protease, Biochemistry

39, 1849–1861.

39. Borkow, G., Fletcher, R. S., Barnard, J., Arion, D., Motakis, D., Dmitrienko, G.

I., and Parniak, M. A. (1997) Inhibition of the ribonuclease H and DNA

polymerase activities of HIV-1 reverse transcriptase by N-(4-tert-butylbenzoyl)-2-

hydroxy-1-napthaldehyde hydrazone, Biochemistry 36, 3179–3185.

40. Spence, R. A., Kati, W. M., Anderson, K. S., and Johnson, K. A. (1995)

Mechanism of inhibition of HIV-1 reverse transcriptase by nonnucleoside

inhibitors, Science 267, 988–993.

115 41. Wang, D-.E., Rizzo, R. C., Tirado-Rives, J., and Jorgensen, W. L. (2001)

Antiviral drug design: computational analyses of the effects of the L100I mutation

for HIV-RT on the binding of NNRTIs, Bioorg. Med. Chem. Lett 11, 2799–2802.

42. Weikl, T. R., and von Deuster, C. (2009) Selected-fit versus induced-fit protein

binding: kinetic differences and mutational analysis, Proteins 75, 104–110.

43. Pegram, L. M., and Record, M. T. (2008) Thermodynamic origin of Hofmeister

ion effects, J. Phys. Chem. B 112, 9428–9436.

44. Rösgen, J., Pettitt, B. M., and Bolen, D. W. (2005) Protein folding, stability, and

solvation structure in osmolyte solutions, Biophys. J. 89, 2988–2997.

45. Hsiou, Y., Ding, J., Das, K., Clark, Jr. A. D., Hughes, S. H., and Arnold, E.

(1996) Structure of unliganded HIV-1 reverse transcriptase at 2.7 Å resolution:

implications of conformational changes for polymerization and inhibition

mechanism, Structure 4, 853–860.

46. Huang, H., Chopra, R., Verdine, G. L., and Harrison, S. C. (1998) Structure of a

covalently trapped catalytic complex of HIV-1 reverse transcriptase: implications

for drug resistance, Science 282, 1669–1675.

47. Sobolev, V., Sorokine, A., Prilusky, J., Abola, E. E., and Edelman, M. (1999)

Automated analysis of interatomic contacts in proteins, Bioinformatics 15, 327–

332

48. Willard, H. H., Merritt, Jr. L. L., and Dean, J. A. (1965) Instrumental Methods of

Analysis, D. Van Nostrand Company, New Jersey, 256.

49. Draper, N. R., and Smith, H. (1966) Applied Regression Analysis, Wiley & Sons,

New York, 77–81.

116 50. Efron, B., and Tibshrirani, R. (1986) Bootstrap methods for standards errors,

confidence intervals, and other measures of statistical accuracy, Stat. Sci. 1, 54–

77.

51. Hall, P. (1992) The Bootstrap and Edgeworth Expansion, p 293, Springer-Verlag,

NY.

117 Chapter 3: The Efavirenz Binding Site in HIV-1 Reverse Transcriptase Monomers

3.1 Abstract

Efavirenz (EFV) is a potent nonnucleoside reverse transcriptase inhibitor

(NNRTI) used in the treatment of AIDS. NNRTIs bind in a hydrophobic pocket located

in the p66 subunit of reverse transcriptase (RT), which is not present in crystal structures

of unliganded RT. These drugs have diverse effects on dimerization of the two subunits,

p66 and p51. Recent studies showed that p66 and p51 monomers bind efavirenz with

micromolar affinity (Braz VA, Holladay LA, and Barkley MD (2009) Biochemistry

submitted). The formation of monomer─EFV and RT─EFV complexes follows a slow, one-step, direct binding mechanism. The binding site on p66 and p51 monomers was identified by hydrogen-deuterium exchange mass spectrometry (HXMS). HXMS data reveal that five peptides, four of which contain efavirenz contact residues seen in the crystal structure of the RT─EFV complex, show reduced exchange in monomer─EFV complexes. Moreover, peptide 232–246 undergoes slow cooperative unfolding in the bound monomers, but at a much slower rate than observed in the p66 subunit of RT.

These results suggest that the efavirenz binding site on p66 and p51 monomers is similar to the NNRTI binding pocket in the p66 subunit of RT. FT-ICR mass spectra of intact monomers show two conformational populations in the absence of efavirenz and a single, more compact conformation in the presence of efavirenz. The population shift is consistent with a selected fit binding mechanism. This study provides valuable information about the NNRTI binding site in monomers and a potential screening tool for drug discovery.

118 3.2 Introduction

HIV-1 reverse transcriptase (RT) was the first target of antiretroviral therapy in the treatment of AIDS. Currently, nucleoside and nonnucleoside reverse transcriptase inhibitors are an essential component of highly active antiretroviral therapy (HAART).

The biologically active form of RT is an asymmetric heterodimer composed of 66 and 51 kDa subunits (1). The p66 subunit contains polymerase and RNase H domains. The p51 subunit is a proteolytic product of p66 without the C-terminal RNase H domain.

Polymerase subdomains in each subunit are fingers (residues 1–85, 118–155), palm

(residues 86–117, 156–236), thumb (residues 237–318) and connection (residues 319–

426) (2, 3). Crystal structures of nonnucleoside reverse transcriptase inhibitors (NNRTIs) bound to RT have identified amino acid contacts and conformational changes associated with inhibitor binding (4). The NNRTI binding pocket, which only exists in structures of

RT—NNRTI complexes, resides in the palm of the p66 subunit with an additional contact in the p66 thumb and in the fingers of the p51 subunit. Efavirenz (EFV) and other

NNRTIs are a class of small amphiphilic compounds that bind ~10 Å away from the polymerase active site (5, 6) and have diverse effects on RT subunit dimerization and polymerase activity (7-10).

In solution, RT is a reversible equilibrium mixture of monomers, homodimers, and heterodimer. There are no crystal structures of either monomers or homodimers. All dimers have enzymatic activity. NNRTI binding is coupled to dimerization. Efavirenz enhances dimerization of both homo- and heterodimers (11, 12) and processing of precursor polyprotein constructs (8). The monomers have folded conformations, but lack activity and do not bind nucleic acid substrates. However, recent equilibrium dialysis

119 experiments showed that the two monomers, p66 and p51, bind efavirenz with the same

micromolar affinity (Chapter 2). The binding stoichiometry is one efavirenz per monomer

and one efavirenz per homodimer. These results confirmed the previously proposed

thermodynamic linkage between NNRTI binding and subunit dimerization (12). Kinetics experiments using tryptophan fluorescence also showed that efavirenz is a slow binding inhibitor. The kinetics data indicate a one-step direct binding mechanism with binding

–1 –1 rate constant ka = 13.5 M s for p66 and p51 monomers as well as for RT. We attributed the slow binding kinetics to conformational selection, where efavirenz preferentially binds to a conformer present at low concentrations (14). Additional support for this hypothesis comes from surface plasmon resonance studies indicating that

NNRTIs bind to RT by a two-step mechanism consisting of a conformational equilibrium followed by complex formation (15).

In the structure of RT (Figure 3-1), the amino acid residues forming the consensus

NNRTI binding pocket are clustered in the two subunits.

120 Figure 3-1. Crystal structure of unliganded HIV-1 RT (1DLO). Four subdomains of the polymerase domain in p66 and p51 subunits: (blue) fingers, (pink) palm, (green) thumb, and (orange) connection; (grey) RNase H domain of p66 subunit; (black) EFV contact residues with side chains in p66 and p51 subunits.

121 In the p66 subunit, the efavirenz contact residues are located between the fingers and

thumb with side chains pointing into the pocket. In the p51 subunit, these contact residues

are located between the palm and thumb with more randomly oriented side chains. The

p51 subunit does not form a functional NNRTI binding pocket, as evident from the

crystal structures of RT—NNRTI complexes (4) and the binding stoichiometry of

homodimers (13). The above equilibrium and kinetics studies of efavirenz binding to

monomers raise intriguing questions about the binding site in the monomers. First, is the

structure of the bound monomer different from unbound monomer? If so, this population

shift is consistent with the proposed selected-fit binding mechanism (15). Second, do p66

and p51 monomers undergo similar conformational changes? The Kds for p66─ and

p51─EFV complexes are 2.7 and 2.5 μM, suggesting similar binding sites in p66 and p51. Third, do the monomers use the same residues as the heterodimer to bind efavirenz?

In the RT─EFV complex, efavirenz makes contacts with L100, K101, K103, V106,

V179, Y181, Y188, G190, F227, W229, L234, H235, and P236 in the palm and Y318 in

the thumb of the p66 subunit (16).

Here we use hydrogen/deuterium exchange mass spectrometry (HXMS) to

examine the solution conformation and dynamics of p66 and p51 monomers in the

presence of efavirenz. Analysis of the exchange kinetics of protein backbone amide

protons provides information on amide hydrogen bonding, flexibility, and local solvent

accessibility (17). Amide hydrogens that are located in elements of stable secondary

structure, α-helices and β-sheets, exchange slowly compared to amide hydrogens in

flexible regions and surface-exposed loops. Comparison of H/D exchange of the two

monomers with and without efavirenz reveals how inhibitor binding alters local

122 flexibility and solvent exposure. Additionally Fourier transform ion cyclotron resonance

mass spectrometry (FT-ICR MS) is used to examine intact unbound and efavirenz bound monomers.

123 3.3 Experimental Procedures

Materials. Efavirenz was obtained from the NIH AIDS Research and Reference Reagent

Program (Germantown, MD). D2O was purchased from Cambridge Isotope Laboratories

(Andover, MA). Biochemical reagents and chemicals were purchased from Roche

Applied Science (Indianapolis, IN) and Sigma Chemicals (St. Louis, MO) unless otherwise specified. RT buffer D is 0.05 M tris(hydroxymethyl)aminomethane (Tris,

RNase, DNase-free, pH 7.2), 25 mM NaCl, 1 mM ethylenediaminetetraacetic acid

(EDTA), and 10% (v/v) glycerol (molecular biology grade redistilled).

Purification of p66W401A and p51 W401A with N-terminal hexahistidine extensions

was performed as described (13). In brief, the dimerization defective mutant proteins

were expressed in Escherichia coli and purified by column chromatography; final protein

concentrations were determined by absorbance at 280 nm (12).

Peptide Mapping by Tandem Mass Spectrometry. Peptide mapping experiments were

carried out as previously described (32). Sequencing by tandem mass spectrometry was

carried out using a Finnigan™ LTQ quadrupole ion trap mass spectrometer

(ThermoElectron). Additional experiments were conducted on an LTQ-FT-ICR mass

spectrometer (ThermoElectron) to confirm peptide identification by exact mass.

W401A W401A HXMS. The p66 and p51 proteins (7.0 μg, 20 μM) in RT buffer D-H2O were diluted 10-fold into RT buffer D-D2O (pD 7.2) containing 5% glycerol and were

incubated for 5 to 5000 s at 5 ºC. For experiments in the presence of efavirenz, protein

samples were incubated with 40 μM efavirenz for 15 h prior to dilution into the D2O buffer containing 25 μM efavirenz. Exchange was quenched by 5-fold dilution into 100 mM NaH2PO4 (pH 2.4) at 5 ºC.

124 The deuterium-labeled protein was digested on ice with 5 μL of 1 mg/mL porcine pepsin in H2O for 5 min and analyzed by HPLC-MS as described elsewhere (33). The samples were loaded on a Vydac C18 reversed phase HPLC column, eluted by increasing acetonitrile concentration from 2% to 98%, and injected directly into the mass spectrometer. Deuterium levels for each peptide were corrected for back-exchange using

 mm 0 D   N (1)  mm 0100 where D is the number of amide hydrogens exchanged with deuterium, m is the centroid mass of the peptide at a given time point, m0 is the mass of the undeuterated peptide, m100 is the mass of the fully deuterated peptide, and N is the number of amide hydrogens.

FT-ICR MS. Nano-spray ion cyclotron resonance mass spectrometry was performed on a Bruker Daltonics APEX-QE equipped with a 7 Tesla magnet and Apollo 2 electrospray ionization source. The p66W401A and p51 W401A monomers (20 μM) were dialyzed overnight at 5 °C into 100 mM NH4OAc (pH 7.0) and then incubated in the absence or presence of 40 μM efavirenz for 15 h at 5 °C. Passive nano-ESI was accomplished using borosilicate tips pulled to a ~3 μm opening using a Sutter P-97 capillary puller. A platinum wire inserted into the solution-containing nano-ESI tip acted as a grounded electrode while a potential between –0.9 to –2.0 kV was applied to the inlet to the mass spectrometer. Spectra shown correspond to the transformation of 16 k data points digitized at a rate of 555.6 kHz.

125 3.4 Results

Conformational Changes in Monomer─EFV Complexes. The p66 and p51 monomers contain the dimerization defective W401A substitution (18) to ensure that p66 and p51 remain monomeric in the presence of efavirenz (13, 19). HXMS was used as described

(20). In short, HXMS was monitored at various times after dilution into deuterated buffer.

The exchange was quenched, the protein was digested with pepsin, and the fragments were analyzed by LC-MS. The peptic fragments provided ~80% sequence coverage for each monomer [supplemental information (SI) Table 3-S1]. Comparison of HXMS data for wild-type and W401A monomers confirmed that the mutation has no effect on the solution structure.

Figure 3-2 shows the full peptide maps of p66 in the absence and presence of efavirenz.

126 Figure 3-2. Percent exchange of peptides in p66W401A monomer in (upper) absence and

(lower) presence of efavirenz. Color of amino acid sequence indicates subdomains: (blue) fingers, (red) palm, (green) thumb, and (orange) connection; (magenta) RNase H domain;

(black) EFV contact residues. Colored bars below sequence from top to bottom give exchange at 10, 50, 100, 500, 1000, and 5000 s.

127 The peptide maps for p51 are shown in SI Figure 3-S1. In unbound monomers, most of the peptides show little exchange at 10 s, indicative of secondary structure or inaccessibility to solvent. The exceptions are peptides 210–231 in the palm, 232–246 in the junction between the palm and thumb, 417–425 in the connection, and 534–560 in the

RNase H domain; these peptides are either solvent-exposed loops or unfolded. In bound monomers, peptides 88–109 and 210–231 in the palm, 232–246 in the palm-thumb junction, 257–282 in the thumb, and 299–328 in the thumb-connection junction, are more rigid. Four of these five peptides contain NNRTI binding pocket residues (4), the exception being peptide 257–282 in the thumb. Two other peptides that contain binding pocket residues, 182–187 and 187-192, show very little exchange in either the absence or presence of efavirenz. Additionally, efavirenz has no effect on exchange in the RNase H domain.

The structural changes in the polymerase domain of the two bound monomers are similar. Figure 3-3 compares the difference in number of deuteria exchanged in the absence and presence of efavirenz for five peptides in p66 and p51 after 10, 100, and

1000 s in D2O.

128 Figure 3-3. Difference in number of deuteria exchanged in bound and unbound p66 and p51. Difference calculated by subtracting the exchange in unbound monomer from the exchange in the monomer─EFV complex. Differences are shown for p66W401A after (light blue) 10 s, (purple) 100 s, and (dark blue) 1000 s; for p51W401A after (yellow) 10 s,

(orange) 100 s, and (red) 1000 s.

129 A small decrease in exchange at 10 s and reduction in exchange at later times indicates stabilization of existing structure, whereas a large decrease in exchange at 10 s suggests formation of additional secondary structure or solvent exclusion. Therefore, the structure of peptides 88–109 and 257–282 are more rigid in bound monomers. On the other hand, peptides 210–231 and 299–328 have either increased secondary structure or some residues blocked by the inhibitor. Peptide 232–246 undergoes cooperative unfolding in the presence of efavirenz as described below.

Reversible Cooperative Unfolding in Efavirenz Binding Site. HXMS provides the ability to distinguish two types of hydrogen exchange kinetics, correlated exchange EX1 and uncorrelated exchange EX2 (21). EX1 kinetics results in a double isotopic envelope in the mass spectra. The two peaks with low and high mass-to-charge ratios (m/z) correspond to two states of the peptide, folded and unfolded. EX1 exchange kinetics is emblematic of slow reversible cooperative unfolding, which appears irreversible in the presence of excess D2O. The commonly observed EX2 kinetics shows a gradual shift of a single peak to higher average m/z. Figure 3-4 shows that the exchange kinetics of peptide

232–246 switch from EX2 kinetics in the absence of efavirenz to EX1 kinetics in the presence of the inhibitor.

130 Figure 3-4. Mass spectra of peptide 232–246 in (left) p51W401A and (right)

W401A p51 ─EFV complex after different incubations times in RT buffer D–D2O. High and low m/z peaks for p51W401A ─EFV complex fit to Gaussian distributions (dashed lines).

131 The EX1 mechanism is observed in both p66─ and p51─EFV complexes. In the absence of efavirenz, about 80% of the amide hydrogens exchange after 10 s in D2O, indicating that the peptide is largely unfolded. In the presence of efavirenz, two populations are clearly present at low and high m/z. For the concentrations used in the HXMS experiments, 94% and 90% of the monomer is bound to efavirenz at equilibrium before and after dilution into D2O (Materials and Methods; 13). Moreover, the t1/2 for unbinding of efavirenz is ~2.5 h (13). Thus, the double isotopic envelope is not an artifact due to unbound monomer. The low and high m/z peaks were fit to Gaussian distributions. The folded conformation is the major solution structure of peptide 232–246 in the bound monomers. Five and 10 s incubations in D2O produce little change in the ratio of folded to unfolded peptide. The shape of the double isotopic envelope remains relatively constant between 50 and 5000 s. By 2 h almost 50% of the peptide is still protected from exchange. After 4 h, most of the peptide has undergone cooperative unfolding/refolding, as evident from the shift to the high m/z peak.

The difference in centroid mass of the low and high m/z peaks of the doubly charged ion of peptide 232–246 corresponds to exchange of 5 amide hydrogens. In the crystal structures of RT and RT─EFV complex, this peptide comprises a loop and β strands 13 and 14 in the p66 subunit and is partially unstructured in the p51 subunit.

There are 6 amino acids in β strands 13 and 14. Additionally, the preceding peptide 210–

231 exchanges 8 fewer amide hydrogens in bound than in unbound monomers (Figure 3-

3). In the structure of RT─EFV complex this peptide has 9 amino acids forming secondary structures. The decrease in exchange in peptides 210–231 and 232–246 in

132 bound monomers is consistent with formation of structural elements similar to those of the p66 subunit of the RT─EFV complex.

Multiple Populations of Unbound Monomers. Generating intact proteins and their complexes in the gas phase is possible using electrospray ionization (ESI) (22, 23). The charge-state distribution of electrosprayed protein ions reflects the compactness of the protein in solution. Generally, unfolded proteins exhibit a relatively broad distribution centered around higher charge states in ESI mass spectra, while when folded the same proteins produce narrower distributions centered around lower charge states (24, 25).

Nanospray-ESI was used to produce intact multiply charged ions of p66 and p51 in the absence and presence of efavirenz. Figure 3-5 (upper and middle) presents the spectra of p66 and p51 obtained in the absence of efavirenz.

133 Figure 3-5. Nano-ESI mass spectra of (upper) p66W401A, (middle) p51W401A, and (lower) p51W401A─EFV complex. Deconvoluted masses for p66W401A are (▲) 66,210 Da, (●)

65,890 Da, and () 65,550 Da. For p51W401A in the absence and presence of EFV, the deconvoluted mass is 52,790 Da. The peaks marked with (*) correspond to a 47 kDa truncated protein.

134 The spectra of the monomers each show two distinct charge-state envelopes. For p66 monomer, the main distribution is centered around the 18+ charge state. For p51 monomer, a distribution centered around the 15+ charge state is the most intense, while there is a second distribution centered around the 20+ charge state. The presence of two charge-state envelopes in each spectrum indicates the existence of at least two different solution conformations of both monomers. The lower charge-state distributions, present at higher m/z, most likely correspond to relatively more folded structures of each monomer.

The ESI mass spectrum of p66 is complicated by the existence of multiple molecules or complexes of masses differing by several hundred Daltons, giving the appearance of splitting peaks. It is not clear whether these correspond to non-covalent adducts, which may be present in solution or formed during ESI, or cleavage products from the protein catalyzed by the ESI process. In contrast to the multiple species observed in the p66 ESI mass spectrum, peaks in the p51 mass spectrum correspond to a single mass.

While the p51─EFV complex was not directly detected, the effect of the drug on the protein mass spectrum is dramatic. In the presence of efavirenz, the higher charge- state distribution disappears from the ESI mass spectrum of p51 and only the lower charge-state distribution remains (Figure 3-5, lower). Moreover, the new charge-state distribution is centered one charge lower than the most intense charge state in the absence of efavirenz. This indicates that binding of efavirenz shifts the equilibrium of two unbound populations to one bound conformational state. Furthermore, the data suggest that the conformation of the p51─EFV complex is relatively compact. The ESI mass

135 spectrum of p66 with efavirenz did not contain peaks corresponding to either p66 or the p66─EFV complex. Instead, a signal corresponding to a 47 kDa species was observed.

The truncation of the protein may result from efavirenz rendering p66 more susceptible to cleavage during ESI or in the gas phase. The identity of the 47 kDa species is unclear; its mass does not correspond to a single cleavage of intact p66. This species is present in extremely low abundance in the mass spectrum of p66 in the absence of efavirenz (Figure

3-5, upper).

136 3.5 Discussion

Efavirenz is an NNRTI capable of affecting several steps in HIV-1 reverse transcription and replication (26, 27). We have recently shown that efavirenz also binds p66 and p51 monomers, a completely new function of NNRTIs for which the biological significance is unknown (13). Although equilibrium and kinetics studies defined the binding constants and binding mechanism for p66─ and p51─EFV complexes, the binding site on the monomers is not known. We used HXMS along with FT-ICR MS to identify the monomer binding sites as potential targets for drug design and to begin understanding the conformational selection process of NNRTI binding in both monomers and heterodimer.

The HXMS results localize the efavirenz binding site to the same five peptides in the polymerase domain of p66 and p51. Four of these peptides contain NNRTI binding pocket residues, implying that the efavirenz binding site in the monomers is similar to the binding site in the heterodimer. The two subunits of RT have different configurations in the crystal structure of the heterodimer. The p66 subunit is described as having an “open” conformation that contains the polymerase active site (28); the p51 subunit has a “closed” conformation that conceals the active site residues (29). In the absence of structures for the monomers, the five peptides that become more folded in the presence of efavirenz are mapped onto separate views of the p66 and p51 subunits from the crystal structure of the

RT─EFV complex (Figure 3-6).

137 Figure 3-6. Five peptides stabilized in monomer─EFV complexes mapped onto subunits of HIV-1 RT─EFV complex (1FK9): (left) p66 subunit and (right) p51 subunit from crystal structure of the heterodimer. Four subdomains of the polymerase domain: (blue) fingers, (pink) palm, (green) thumb, and (orange) connection; (grey) RNase H domain of p66 (grey); (black) five peptides in monomer─EFV complexes.

138 While identical peptides are affected in the two monomers, the secondary and tertiary structures of these peptides are quite different in the two subunits of the heterodimer. In the p66 subunit structure, the affected peptides are contiguous and concentrated in the vicinity of the NNRTI binding pocket. Together with the cooperative stabilization of β strands 13 and 14 that form the top of the NNRTI binding pocket (see below), these results strongly suggest that the conformation of the polymerase domain of bound monomers is similar to that of the p66 subunit in RT. This is supported by the fact that both homodimers have polymerase activity (30, 31), in which one subunit must have a catalytically active “open” conformation similar to the p66 subunit of the heterodimer.

Previous HXMS studies of RT found that peptide 232–246, located at the base of the thumb, undergoes EX1 exchange due to slow cooperative unfolding of β strands 13 and 14 with t1/2 ~6 s (20). No evidence for EX1 exchange or two conformationally distinct populations of this peptide is seen in the unbound monomers. In the presence of efavirenz, there are clearly two slowly interconverting populations. However, the interconversion rate is markedly slower than that of unliganded RT. These results suggest that efavirenz binds to the population with a more folded conformation. The enhanced local folding is accompanied by stabilization of the structure of the other four peptides in the palm and thumb. Examination of the amino acid sequence of these peptides shows multiple Lys, Arg, and His residues; five in 88–109, five in 210–231, two in 232–246, four in 257–282, and four in 299–328. Stabilization of the peptides may sequester these side chains as well as amide nitrogens or carbonyl oxygens, thereby changing the exposed surface area and reducing the ability to gain a charge during ESI. This is consistent with the shift in the ESI charge-state distributions to a more folded

139 conformation with lower charge state in the presence of efavirenz. Modulating the conformational stability of the binding pocket and thumb appears to be a key feature of

RT structure and function.

Acknowledgements

This work was supported by the National Institutes of Health (Grant GM071267). We are grateful to James M. Seckler for helpful discussions of HXMS data and to Matthew

Forbes for technical help in FT-ICR MS experiments.

140 Suplemental Information.

Table 3-S1. Sequence and Residue Number of Analyzed Peptides sequence residue no. KIKALVE 30–36 LVEICTE 34–40 MEKEGKISKIGPENPYNTPVF 41–61 NKRTQDF 81–87 WEVQLGIPHPAGLKKKKSVTVL 88–109 DVGAY 110–115 SVPLDEDF 117-124 FRKYTAFTIPSINNETPGIRYQY 124–146 NVLPQGWKGSPAIF 147–160 YMDDL 183–187 LYVGSD 187–192 LEIGQHRTKIEELRQHL 193–209 LRWGLTTPDKKHQKEPPFLWMG 210–231 YELHPDKWTVQPIVL 232–246 PEKDSWTVND 247–256 IQKLVGKLNWASQIYPGIKVRQLCKL 257–282 LRGTKALTEVIPLTEEAE 283–300 LELAENREILKEPVHGVYYDPSKDLIAE 299–328 IQKQGQGQWTYQ 329–340 IYQEPFKNLKTGKYARMRHAHTNDVKQLTE 341–370 AVKITTES 371–379 IVIWQKTPKFKLPIQKETWETA 380–401 VNTPPLVKL 426–439 WYQLEKEPIVGAETF 426–440 YVDGAASRETKL 427–438 LTNTTNQKTEL 469–479 EVNIVTDSQ 492–500 YALGIIQAQPDKSESEL 501–517 VNQIIEQLIKKEKVYL 518–533 AWVPAHKGIGGNEQVDKLVSAGIRKIL 534–560

141 Figure 3-S1: Percent exchange of peptides in p51W401A monomer in (upper) absence and

(lower) presence of efavirenz. Color of amino acid sequence indicates subdomains: (blue) fingers, (red) palm, (green) thumb, and (orange) connection; (black) EFV contact residues. Colored bars below sequence from top to bottom give exchange at 10, 50, 100,

500, 1000, and 5000 s.

142 3.6 References

1. Wang, J., et al. (1994) Structural basis of asymmetry in the human

immunodeficiency virus type 1 reverse transcriptase heterodimer. Proc. Natl.

Acad. Sci. USA 91, 7242–7246.

2. Sluis-Cremer, N., Temiz, A., and Bahar, I. (2004) Conformational changes in

HIV-1 reverse transcriptase induced by nonnucleoside reverse transriptase

inhibitor binding. Curr. HIV Res. 2, 323–332.

3. Kohlstaedt, L. A., Wang, J., Friedman, J. M., Rice, P. A., and Steitz, T. A. (1992)

Crystal structure at 3.5 A resolution of HIV-1 reverse transcriptase complexed

with an inhibitor. Science 256, 1783–1790.

4. Das, K., Lewis, P. J., Hughes, S. H., and Arnold, E. (2005) Crystallography and

design of anti-AIDS drugs: conformational flexibility and positional adaptability

are important in the design of non-nucleoside HIV-1 reverse transcriptase

inhibitors. Prog. Biophys. Mol. Biol. 88, 209–231.

5. Tsai, C. H., Lee, P. Y., Stollar, V., and Li, M. L. (2006) Antiviral therapy

targeting viral polymerase. Curr. Pharm. Des. 12, 1339–1355.

6. De Clercq, E. (1998) The role of non-nucleoside reverse transcriptase inhibitors

(NNRTIs) in the therapy of HIV-1 infection. Antiviral Res. 38, 153–179.

7. Spence, R. A., Kati, W. M., Anderson, K. S., and Johnson, K. A. (1995)

Mechanism of inhibition of HIV-1 reverse transcriptase by nonnucleoside

inhibitors. Science 267, 988–993.

8. Sluis-Cremer, N., Arion, D., Abram, M. E., and Parniak, M.A. (2004) Proteolytic

processing of an HIV-1 pol polyprotein precursor: insights into the mechanism of

143 9. Tachedjian, G., Moore, K. L., Goff, S.P., and Sluis-Cremer, N. (2005) Efavirenz

enhances the proteolytic processing of an HIV-1 pol polyprotein precursor and

reverse transcriptase homodimer formation. FEBS Lett. 579, 379–384.

10. Figueiredo, A., et al. (2006) Potent nonnucleoside reverse transcriptase inhibitors

target HIV-1 Gag-Pol. PLoS. Pathog. 2, 10511059.

11. Tachedjian, G., Orlova, M., Sarafianos, S. G., Arnold, E., and Goff, S. P. (2001)

Nonnucleoside reverse transcriptase inhibitors are chemical enhancers of

dimerization of the HIV type 1 reverse transcriptase. Proc. Natl. Acad. Sci. USA

98, 7188–7193.

12. Venezia, C. F., Howard, K. J., Ignatov, M. E., Holladay, L. A., and Barkley, M.

D. (2006) Effects of efavirenz binding on the subunit equilibria of HIV-1 reverse

transcriptase. Biochemistry 45, 2779–2789.

13. Braz, V. A., Holladay, L. A., and Barkley, M. D. (2009) Efavirenz binding to

HIV-1 reverse transcriptase monomers and dimers. submitted Biochemistry.

14. Weikl, T. R., and von Deuster, C. (2009) Selected-fit versus induced-fit protein

binding: kinetic differences and mutational analysis. Proteins 75, 104–110.

15. Geitmann, M., Unge, T., and Danielson, H. (2006) Interaction kinetic

characterization of HIV-1 reverse transcriptase non-nucleoside inhibitor

resistance. J. Med. Chem. 49, 2375–2387.

144 16. Sobolev, V., Sorokine, A., Prilusky, J., Abola, E. E., and Edelman, M. (1999)

Automated analysis of interatomic contacts in proteins. Bioinformatics 15, 327–

332.

17. Wales, T.E., Engen, J. R. (2006) Hydrogen exchange mass spectrometry for the

analysis of protein dynamics. Mass Spectrom. Rev. 25, 158–170.

18. Tachedjian, G., et al. (2003) Role of residues in the tryptophan repeat motif for

HIV-1 reverse transcriptase dimerization. J. Mol. Biol. 326, 381–396.

19. Braz, V. A., and Howard, K. J. (2009) Separation of protein oligomers by blue

native gel electrophoresis. Anal. Biochem. 388, 170–172.

20. Seckler, J. M., Howard, K. J., Barkley, M. D., and Wintrode, P. L. (2009)

Solution structural dynamics of HIV-1 reverse transcriptase heterodimer.

Biochemistry 48, 7646–7655.

21. Weis, D., Wales, T. E., Engen, J. R., Hotchko, M., and Ten Eyck, L. F. (2006)

Identification and characterization of EX1 kinetics in H/D exchange mass

spectrometry by peak analysis. J. Am. Soc. Mass. Spectrom. 17, 1498–1509.

22. Benesch, J. L., Ruotolo, B. T., Simmons, D.A., and Robinson, C. V. (2007)

Protein complexes in the gas phase: technology for structural genomics and

proteomics. Chem. Rev. 107, 3544–3567.

23. Fenn, J. B., Mann, M., Meng, C. K., Wong, S. F., and Whitehouse, C. M. (1989)

Electrospray ionization for mass spectrometry of large biomolecules. Science 246,

64–71.

24. Chowdhury, S. K., Katta, V., and Chait, B. T. (1990) Probing conformational

changes in proteins by mass spectrometry. J. Am. Chem. Soc. 112, 9012–9013.

145 25. Konermann, L., and Douglas, D. J. (1997) Acid-induced unfolding of cytochrome

c at different methanol concentrations: electrospray ionization mass spectrometry

specifically monitors changes in the tertiary structure. Biochemistry 36, 12296–

12302.

26. Sluis-Cremer, N., and Tachedjian, G. (2008) Mechanisms of inhibition of HIV

replication by non-nucleoside reverse transcriptase inhibitors. Virus Res. 134,

147–156.

27. Grobler, J. A., et al. (2007) HIV-1 reverse transcriptase plus-strand initiation

exhibits preferential sensitivity to non-nucleoside reverse transcriptase inhibitors

in vitro. J. Biol. Chem. 282, 8005–8010.

28. Jacobo-Molina, A., et al. (1993) Crystal structure of human immunodeficiency

virus type 1 reverse transcriptase complexed with double-stranded DNA at 3.0 Å

resolution shows bent DNA. Proc. Natl. Acad. Sci. USA 90, 6320–6324.

29. Le Grice, S. F. J., Naas, T., Wohlgensinger, B., and Schatz, O. (1991) Subunit-

selective mutagenesis indicates minimal polymerase activity in heterodimer-

associated p51 HIV-1 reverse transcriptase. EMBO J. 10, 3905–3911.

30. Bavand, M. R., Wagner, R., and Richmond, T. J. (1993) HIV-1 reverse

transcriptase: Polymerization properties of the p51 homodimer compared to the

p66/p51 heterodimer. Biochemistry 32, 10543–10552.

31. Restle, T., Muller, B., and Goody, R. S. (1990) Dimerization of human

immunodeficiency virus type 1 reverse transcriptase. J. Biol. Chem. 265, 8986–

8988.

146 32. Tsutsui, Y., Liu, L., Gershenson, A., and Wintrode, P. L. (2006) The

conformational dynamics of a metastable serpin studied by hydrogen exchange

and mass spectrometry. Biochemistry 34, 16337–16346.

33. Zhang, Z., and Smith, D. L. (1993) Determination of amide hydrogen exchange

by mass spectrometry: a new tool for protein structure elucidation. Protein Sci. 2,

522–531.

147 Chapter 4: Separation of Protein Oligomers by Blue Native Gel Electrophoresis

4.1 Abstract

Native gel electrophoresis is used as a tool to assess structural differences in proteins. This chapter presents an application to separate oligomeric forms of proteins, such as HIV-1 reverse transcriptase monomers and homodimers. Technical difficulties encountered with various native gel techniques and ways to circumvent them are described.

148 4.2 Introduction, Results, and Discussion

Human immunodeficiency virus type 1 reverse transcriptase (HIV-1 RT) is a multifunctional enzyme that catalyzes the conversion of genomic RNA into double- stranded proviral DNA. The enzyme is a heterodimer of 66 and 51 kDa subunits that can also form homodimers. The p51 subunit is derived by proteolytic cleavage of the C- terminal RNase H domain of p66. Analytical ultracentrifugation experiments determined

Kd values of 4.2 and 230 μM for homodimerization of p66 and p51, respectively (1). To evaluate dimerization over a range of conditions, we tried a commercially available kit for blue native polyacrylamide gel electrophoresis (BN–PAGE). Purified p66 behaved normally with one or two bands depending on solution conditions, but purified p51 produced a ladder of bands under conditions in which only monomer is present in solution. Therefore, we explored a variety of native gel electrophoresis techniques (2–6) and developed a modified protocol for blue native agarose gel electrophoresis (BN–AGE) that gave superior results for p51 as well as other soluble proteins.

A Novex Bis–Tris Gel System and NativeMark Protein Standards were purchased from Invitrogen (Carlsbad, CA, USA). SeaKem Gold Agarose was purchased from Lonza

(Rockland, ME, USA). Kaleidoscope Precision Plus Protein Standards were purchased from Bio-Rad (Hercules, CA, USA). EZ-Run Protein Gel Staining Solution was purchased from Fisher Scientific (Fair Lawn, NJ, USA). Bovine serum albumin (BSA) and T4 DNA ligase were purchased from Roche Diagnostics (Indianapolis, IN, USA).

Carbonic anhydrase, aprotinin, cytochrome c, and other biochemical reagents were purchased from Sigma Chemicals (St. Louis, MO, USA). HIV-1 p66 and p51 were purified as described previously (1).

149 BN–PAGE was carried out using the Novex Bis–Tris Gel System according to the manufacturer’s specifications. Precast NativePAGE Novex 4 to 16% (v/v) Bis–Tris gels were run with near-neutral pH at 90 V at 4 °C with stirring for 4.5 h. Protein samples (10

μL) were mixed with the sample buffer provided (2.5 μL) and 5% (w/v) Coomassie blue

G-250 (0.3 μL). Gels were stained in EZ-Run Protein Gel Staining Solution for 1 to 2 h and were destained in water. BN–AGE was carried out using a Bio-Rad mini-sub cell DT unit. A 3% (w/v) horizontal SeaKem Gold Agarose gel (10 cm  6 cm  5 mm) was prepared using native agarose gel buffer (NAGB: 25 mM Tris and 19.2 mM glycine at pH 7.0 or 8.5) (4). The horizontal gel was submerged in the apparatus containing NAGB, and electrophoresis was performed at room temperature at 40 V for 4.5 h. Protein samples (10 μL) were mixed with sample buffer (2.5 μL, NAGB containing 30% (w/v) glycerol) and 5% (w/v) Coomassie blue G-250 (0.3 μL, Novex Bis–Tris Gel System).

Gels were stained and destained using the same protocol as BN–PAGE.

The patterns of p66 and p51 generated in BN–PAGE are shown in Fig. 4-1. Fig.

4-1A shows BN–PAGE of p66 solutions containing monomeric and dimeric forms. Fig.

4-1B shows BN–PAGE of a p51 solution containing only monomers. Severe laddering of monomeric p51 was observed. To confirm that the multiple p51 bands were an artifact of

BN–PAGE, the bands from BN–PAGE were cut out, soaked, and subjected to sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE, not shown). The protein bands that originally migrated to different positions by BN–PAGE now migrated to positions identical to the p51 standard by SDS–PAGE. This confirms that protein degradation did not cause the ladder pattern.

150 Figure 4-1. Migration of HIV-1 p66 and p51 subunits by blue native electrophoresis. 5

µg of protein loaded per well; (% monomer, % homodimer) calculated from Kd values.

(A) BN-PAGE: Lane 1: Native Markers, Lane 2: 1 µM p66 (75% p66, 25% p66/p66),

Lane 3: 5 µM p66 (60% p66, 40% p66/p66). (B) BN-PAGE: Lane 1: Native Markers,

Lane 2: 5 µM p51 (96% p51, 4% p51/p51), Lane 3: 10 µM p51 (92% p51, 8% p51/p51).

(C) BN-AGE: Lane 1: 5 µM wt p51 (96% p51, 4% p51/p51), Lane 2: 5 µM wt p51 + efavirenz (46% p51, 54% p51/p51), Lane 3: 5 µM W401A p51 (100% p51), Lane 4: 5

µM W401A p51 + efavirenz (~95% p51, ~5% p51/p51), Lane 5: 5 µM (100% p51)

L234A p51, Lane 6: 5 µM (100% p51) L234A + efavirenz.

151 We tried several permutations of the native PAGE system. First, BN–PAGE was run in the absence of Coomassie blue G-250. p51 remained in the sample well and did not enter the gel (not shown). Second, detergents were added to ensure p51 solubility during BN–PAGE. Protein samples containing either 1% (w/v) n-dodecyl-β-D-maltoside or 1% (w/v) digitonin were prepared as described above. Third, voltage was reduced to eliminate the possibility of thermal denaturation during BN–PAGE. Gels were run at 60

V at 4 ºC with stirring for 8 h. Fourth, Bio-Rad Criterion Tris–HCl gels 8 to 16% (v/v)

(pH 8.5) were run at 100 V at 4 ºC with stirring for 2.5 h. Fifth, nongradient 16-cm native polyacrylamide gels were poured using the Protean II xi cell with a 3% (v/v) stack and

7.5, 10.0, or 12.5% (v/v) resolving gel. Gels were run using the Novex Bis–Tris buffer system at 90 V at 4 ºC with a circulating water system for 8 to 10 h. Neither omitting

Coomassie blue G-250, adding detergents, lowering the voltage, increasing the pH from

6.8 to 8.5, nor replacing the gradient with standard gels resolved the laddering. It is possible that an interaction of p51 with the polyacrylamide matrix contributes to the peculiar behavior despite identical amino acid sequences of p51 and the first 440 residues of p66. Therefore, agarose was evaluated as an alternative support medium (2, 3).

In the Invitrogen BN–PAGE kit and a published BN–AGE protocol, electrophoresis is carried out at near-neutral pH (2, 4–6). Fig. 4-2A and B show the migration of several proteins with different isoelectric points (pI) run on 3% (w/v) agarose gels in the presence and absence of Coomassie blue G-250 at pH 7.0.

152 Figure 4-2. Migration of proteins by native agarose gel electrophoresis. 5 µg of protein loaded per well. Lane 1: aprotinin (pI = 10.0─10.5), Lane 2: cytochrome c (pI =

10.0─10.5), Lane 3: carbonic anhydrase (pI = 6.6─7.2), Lane 4: BSA (pI = 4.7), Lane 5:

T4 DNA ligase (pI = 6.0─6.2), and Lane 6: p51 (pI = 8.7). (A) pH 7.0 with Coomassie blue G-250, (B) pH 7.0 without Coomassie blue G-250, (C) pH 8.5 with Coomassie blue

G-250, (D) pH 8.5 without Coomassie blue G-250. Gels in (A-C) destained 36 h, gel in

(D) destained 72 h.

153 As expected in the presence of Coomassie blue G-250, all proteins migrated toward the cathode (Fig. 4-2A). Aprotinin, cyctochrome c, BSA, and T4 DNA ligase migrated as single bands with approximately the same mobility, carbonic anhydrase migrated as three bands, and p51 migrated as a single band only a short distance from the well. In the absence of Coomassie blue G-250, aprotinin (pI = 10.0–10.5) migrated as a single band and cytochrome c (pI = 10.0–10.5) migrated as a diffuse band toward the anode, BSA (pI

= 4.7) and T4 DNA ligase (pI = 6.0–6.2) migrated as single bands toward the cathode, and carbonic anhydrase (pI = 6.6–7.2) and p51 (pI = 8.7) remained near the well (Fig. 4-

2B). Although p51 migrated as a single band on agarose gels at pH 7.0, the mobility was low in both the presence and absence of Coomassie blue G-250.

Native agarose gels have been run at pH 8.5 (4). In the presence of Coomassie blue G-250, aprotinin, carbonic anhydrase, BSA, T4 DNA ligase, and p51 all migrated as single bands, whereas cytochrome c migrated as a long diffuse band, toward the cathode

(Fig. 4-2C). In the absence of Coomassie blue G-250, the migration of the proteins was driven by their respective pI values; aprotinin and cytochrome c migrated toward the anode as single bands, whereas carbonic anhydrase, BSA, T4 DNA ligase, and p51 migrated toward the cathode as single bands (Fig. 4-2D).

Because p51 migrated cleanly by BN–AGE at pH 8.5, we tried to separate monomeric and dimeric p51 under these conditions. Fig. 4-1C shows the migration of wild-type (wt) and mutant p51 proteins in the presence of Coomassie blue G-250 as discrete bands representing monomer and homodimer. The mutant proteins W401A and

L234A are dimerization deficient (7, 8). Efavirenz is an inhibitor that enhances dimerization of RT subunits (1, 9). The wt and W401A p51 samples incubated with

154 efavirenz clearly show dimer formation as a result of drug binding, whereas L234A does not bind the drug (9).

The results presented here extend the use of blue native gel electrophoresis to the separation of oligomeric forms of proteins. Protein migration under native conditions is dependent on molecular mass, pI, buffer pH, and type and percentage of gel matrix. In the presence of Coomassie blue G-250, migration is also dependent on nonspecific binding of the dye by the protein to provide a net negative charge. Agarose acts as a superior solid phase for separation of monomeric and dimeric HIV p51 and may work well for other proteins. The availability of sieving agarose for high-resolution separations has allowed us to develop a novel protocol to study small proteins and protein–protein interactions. In conclusion, our experience dictates the use of due diligence in the choice of system and conditions used. The appropriate conditions may prove to be counterintuitive to theoretical expectations.

155 4.3 Acknowledgements

This work was supported by NIH grant GM071267. We would like to thank Dr.

Mary Barkley and Dr. Tsutomu Arakawa (Alliance Protein Laboratories, Thousand Oaks,

CA) for helpful discussions.

156 4.4 References

1. Venezia, C. F., Howard, K. J., Ignatov, M. E., Holladay, L. A., and Barkley, M.

D. (2006) Effects of efavirenz binding on the subunit equilibria of HIV-1 reverse

transcriptase, Biochemistry 45, 2779-2789.

2. Niepmann, M., and Zheng, J. (2006) Discontinuous native protein gel

electrophoresis, Electrophoresis 27, 3949-3951.

3. Kim, R., Yokota, H., and Kim, S.-H. (2000) Electrophresis of proteins and

protein-protein complexes in a native agarose gel, Anal. Biochem. 282, 147-149.

4. Henderson, N. S., Nijtmans, L. G. J., Lindsay, J. G., Lamantea, E., Zeviani, M.,

and Holt, I. J. (2000) Separation of intact pyruvate dehydrogenase complex using

blue native agarose gel electrophoresis, Electrophoresis 21, 2925-2931.

5. Schägger, H., Cramer, W. A., von Jagow, G. (1994) Analysis of molecular masses

and oligomeric states of protein complexes by blue native electrophoresis and

isolation of membrane complexes by two-dimensional native electrophoresis,

Anal. Biochem. 217, 220-230.

6. Swamy, M., Siegers, G. M., Minguet, S. M., Wollschield, B., and Schamel, W.

W. A. (2006) Blue native polyacrylamide gel electrophoresis (BN-PAGE) for the

identification and analysis of multiprotein complexes, Sci. STKE 345, 1-14.

7. Powell, M. D., Ghosh, M., Jacques, P. S., Howard, K. J., Le Grice, S. F. J., and

Levin, J. G. (1997) Alanine-scanning mutations in the “primer grip” of p66 HIV-1

reverse transcriptase result in selective loss of RNA priming activity, J. Biol.

Chem. 272, 13262-13269.

157 8. Tachedjian, G., Aronson, H.-E.G., de los Santos, M., Seehra, J., McCoy, J. M.,

and Goff, S. P. (2003) Role of residues in the tryptophan repeat motif for HIV-1

reverse transcriptase dimerization, J. Mol. Biol. 325, 381-396.

9. Tachedjian, G., Orlova, M., Sarafianos, S. G., Arnold, E., and Goff, S. P. (2001)

Nonnucleoside reverse transcriptase inhibitors are chemical enhancers of

dimerizarion of the HIV type 1 reverse transcriptase, Proc. Natl. Acad. Sci. USA,

98, 7188-7193.

158 Chapter 5

5.1 Conclusions and Future Directions

The results presented in Chapters 2 and 3 allow us to make the following conclusions: (1) monomers are capable of binding NNRTIs, (2) efavirenz is a slow and tight binding inhibitor, (3) two structural populations of the monomers exist in solution, and (4) binding of efavirenz selected one conformation of the protein over another.

The work described in Chapter 2 provides the first documented account that RT monomers are capable of binding NNRTIs. The importance of this is to further understand how NNRTIs affect subunit dimerization. This has further downstream implications on forming the biologically active heterodimer. Through the use of equilibrium dialysis, dissociation constants of each monomer and homodimer were measured in the absence and presence of efavirenz (Table 2-1). Based on these results, the previously proposed thermodynamic linkage (Scheme 1-1) of the effects of NNRTIs on RT dimerization was solved. The cycle was completed by the additional results for efavirenz binding to monomers and homodimers with ΔG = 0 around the closed path.

Additionally, by tryptophan fluorescence we are able to propose that the mechanism for efavirenz binding is simple and direct (Scheme 2-2, Mechanism A). However, due to the inability to detect a fast phase by steady-state fluorescence measurement, we propose that in addition to following a slow direct binding mechanism, NNRTIs bind preferentially to one solution structure of the protein. As such, in solution the monomers exist in two different structural populations; one which is unable to bind the ligand and another which is able to bind the ligand. Binding of efavirenz to one form shifts the equilibrium population. This accounts for the slow binding. Lastly, to demonstrate that efavirenz is

159 also a tight binding inhibitor, we used BN-PAGE. Electrophoresis was conducted on wt and p66W401A dimerization defective monomers over the course of 4–5 h. If unbinding of the inhibitor was fast, then dissociation of the ligand would occur during electrophorsis and no radiolabeled efavirenz would be detected upon completion of the run. This is not the case as shown in Figure 2-6. Chapter 2 describes, in depth, the binding mechanism and constants for efavirenz to monomers and dimers of RT.

Chapter 3 investigated the solution dynamics of RT monomers in the absence and presence of efavirenz. HXMS was used to report structural changes that occurred in different parts of the protein after binding of efavirenz. The changes in protein structure for p66 and p51 after various incubation times in D2O buffer are shown in Figures 3-2 and Figure S1, respectively. These studies also demonstrate that the monomers bind and form the NNRTI binding pocket in a similar manner to the heterodimer, making this a biologically relevant process. The data show that the peptides flanking the binding pocket are the most affected. It is also noted that there do not appear to be long range structural perturbations after binding since no changes in the RNase H subdomain of p66 are witnessed. Overall, the structure of the monomers in the presence of efavirenz becomes more rigid. This is additionally accounted for in production of intact gas phase ions of the p51W401A─EFV complex (Figure 3-5). The two distinct structural populations in the absence of efavirenz is reduced to one in the presence of efavirenz. There is also reduction in the m/z ratio in the p51W401A─EFV complex indicating that the structure has become more rigid and less solvent exposed (Figure 3-5).

Future directions for this project would be to determine the binding constants of other NNRTIs for monomers and homodimers of RT. Literature reports indicate that

160 NNRTIs have different effects on dimerization. Specifically, efavirenz and nevirapine have been shown to enhance dimerization (1); TSAOe3T, BBNH, and BBSH weaken dimerization (2, 3); and delaviridine has no effect (1). The proposed thermodynamic linkage provides a rational for this process and we can use equilibrium dialysis to discern whether NNRTIs that weaken dimerization bind more tightly to the monomers and vice versa. The only literature reports of determining binding constants of NNRTIs to wt and mutant RTs uses SPR. These studies were conducted in the presence of detergent and organic solvent.

There can be multiple forces driving conformational changes of proteins in solution. Many in vitro systems used to study biological molecules employ the use of various electrolytes and osmolytes. Measurements carried out on proteins and ligands in aqueous solutions containing osmolytes are consistent with the idea that these small molecules interact with the peptide backbone (4, 5). In addition to affecting the peptide backbone, osmolytes may also contribute the solvation of the ligand therefore driving the binding reaction forward. A common biological osmolyte, such as urea, could be studied to test the possibility of conformational selection for ligand binding to RT. A shift in the population of solution conformers due to urea would either increase or decrease the rate of ligand binding. Optimal urea concentrations that do not promote complete unfolding of polypeptide chains have been previously determined (2, 3). However, each protein reacts differently, and optimatization of urea concentration must be done. Initial studies should first be completed using CD spectroscopy. This would confirm the stability and degree of folded structure in the protein. Next would be to determine the effects of urea on the solution structure by HXMS. This would provide details of the structural rearrangements

161 that occur such as increase or decrease in subdomain flexibility. Lastly would be to determine the equilibrium binding constants in the presence of urea and an NNRTI by equilibrium dialysis. This data could then be used to design kinetics experiments using tryptophan fluorescence to further elucidate the binding mechanism via a one-step or two-step process.

Another interesting area to pursue would be to study the effects NNRTI mutations on the solution structure of RT for future drug design. These experiments would be accomplished through HXMS. Understanding the structural changes that occur due single amino acid substitutions would provide valuable insight for the future design of novel RT inhibitors. If a mutation results in enlargement of the pocket, advances should be made to increase the size of the NNRTI. If the NNRTI is too small to make contacts with residues in the pocket, then the barrier for retention would be low and the drug could easily diffuse out of the pocket. Conversely, if the mutation results in decreasing the size of the pocket, smaller inhibitors should be designed so that the entry barrier is reduced.

Drug design is a complex process that involves fully understanding the biological target and properties of organic molecules. Modification of functional groups that successfully interact with the target to elicit the desired response is essential in the development of chemotherapeutic agents.

162 5.2 References

1. Tachedjian, G., Orlova, M., Sarafianos, S. G., Arnold, E., and Goff, S. P. (2001)

Nonnucleoside reverse transcriptase inhibitors are chemical enhancers of

dimerization of the HIV type 1 reverse transcriptase, Proc Natl Acad Sci U.S.A.

98, 7188-7193.

2. Sluis-Cremer, N., Arion, D., and Parniak, M. A. (2002) Destabilization of the

HIV-1 reverse transcriptase dimer upon interaction with N-acyl hydrazone

inhibitors, Mol Pharmacol 62, 398-405.

3. Sluis-Cremer, N., Dmitrienko, G. I., Balzarini, J., Camarasa, M. J., and Parniak,

M. A. (2000) Human immunodeficiency virus type 1 reverse transcriptase dimer

destabilization by 1-[Spiro[4"-amino-2",2"-dioxo-1",2"-oxathiole-5",3'-[2', 5'-bis-

O-(tert-butyldimethylsilyl)-β-D-ribofuranosyl]]]-3-ethylthymine, Biochemistry

39, 1427-1433.

4. Auton, M., and Bolen, D. W. (2004) Additive transfer free energies of the peptide

backbone unit that are independent of the model compound and the choice of

concentration scale, Biochemistry 43, 1329-42.

5. Liu, Y., and Bolen, D. W. (1995) The peptide backbone plays a dominant role in

protein stabilization by naturally occurring osmolytes, Biochemistry 34, 12884-

91.

163 Appendix: Equilibrium Dialysis Raw Data

Table A1: wt p51 5 d [Ptotal] μM [Ibound] μM [Ifree] μM 0.1 0.0121 0.296 0.1 0.0188 0.300 0.4 0.032 0.443 0.4 0.049 0.446 0.48 0.071 0.241 0.48 0.053 0.262 0.48 0.052 0.264 0.51 0.09 0.311 0.51 0.112 0.338 0.56 0.26 0.71 0.56 0.19 0.81 1.0 0.24 0.81 1.0 0.27 3.05 1.15 0.141 1.06 1.15 0.151 1.99 1.15 0.309 0.712 1.15 0.384 0.745 1.26 0.35 1.04 1.26 0.38 1.05 1.26 0.37 2.69 2.0 0.32 1.2 2.0 0.59 1.24 1.5 0.321 1.01 1.5 0.268 1.014 1.5 0.46 2.49 2.6 0.75 3.72 2.6 0.7 3.59 3.6 0.169 2.28

164 3.6 0.172 4.66 10 4.11 1.35 10 4.29 1.67 10 3.9 6.47 10 4.08 6.62 10 5.44 10.57 10 4.98 11.28

Table A2: wt p51 30 h [Ptotal] μM [Ibound] μM [Ifree] μM 0.5 0.34 1.13 0.5 0.35 1.06 0.9 0.67 1.86 1.6 1.5 3.83 1.6 0.88 2.17 3.5 1.52 3.97 3.5 5.44 1.52 10 8.18 8.0 10 7.46 8.4 10 4.81 1.6 10 4.7 2.8 10 2.71 0.44 10 3.1 0.41

165

Table A3: W401A p51 30 h [Ptotal] μM [Ibound] μM [Ifree] μM 1.2 0.14 0.55 2.6 2.73 0.8 3 1.63 2.54 3.1 0.54 1.34 5.3 3.33 4.09 6 3.78 5.63 9.1 6.01 5.42 9.6 7.74 3.39 10 7.32 7.73

166

Table A4: wt p66 5 d [Ptotal] μM [Ibound] μM [Ifree] μM 0.4 0.09 0.74 0.45 0.12 1.4 0.45 0.13 2.22 0.55 0.12 0.7 0.82 0.43 2.03 0.82 0.38 2.05 0.86 0.26 1.12 0.96 0.37 0.89 1.0 0.33 0.25 1.0 0.33 0.19 1.5 0.96 2.38 1.5 0.56 3.66 1.5 0.58 3.55 2.0 0.59 0.99 3.9 1.2 3.19 4.0 0.96 3.34 5.0 1.74 4.73

167

Table A5: W401A p66 30 h [Ptotal] μM [Ibound] μM [Ifree] μM 0.79 0.11 2.53 0.83 0.15 2.63 1.1 1.34 2.48 1.5 0.55 5.46 2.29 0.95 5.36 2.74 1.01 4.95 4.49 2.72 5.99 5.99 3.75 9.67 7.34 5.85 11.41

168