STUDIES INTO THE ROLE OF SERINE/THREONINE RESIDUES IN HIV CORE STABILITY AND REPLICATION

SARAH MARY-ROSE CONNOR MARTIN

Bachelor of Science (Molecular Biology) (Honours)

Discipline of Microbiology and Immunology

School of Molecular & Biomedical Science

The Faculty of Science

The University of Adelaide

Adelaide, South Australia

HIV Research Laboratory

Infectious Diseases Laboratories

Institute of Medical and Veterinary Science

SA Pathology

Adelaide, South Australia

A thesis submitted to the University of Adelaide in fulfilment of the requirements for the degree of Doctor of Philosophy

August 2013 TABLE OF CONTENT ABSTRACT ...... VII

DECLARATION ...... IX

ACKNOWLEDGEMENTS ...... X

ABBREVIATIONS ...... XI

PRESENTATIONS ARISING ...... I

CHAPTER 1 - INTRODUCTION ...... 1

1.1 HIV AND AIDS ...... 1

1.1.1 History and identification ...... 1

1.1.2 Classification ...... 3

1.2 HIV-1 EPIDEMIOLOGY AND PATHOGENESIS ...... 4

1.2.1 Epidemiology ...... 4

1.2.2 Transmission ...... 5

1.2.3 Pathogenesis ...... 7

1.2.4 HIV-1 Prevention and Therapy ...... 10

1.3 HIV-1 VIRION ...... 13

1.3.1 HIV-1 virion ...... 13

1.3.2 HIV-1 ...... 14

1.4 HIV-1 REPLICATION CYCLE ...... 15

1.4.1 Overview of HIV-1 replication cycle ...... 15

1.4.2 Entry ...... 15

1.4.3 Core disassembly ...... 16

1.4.4 Reverse ...... 17

1.4.5 Nuclear translocation and integration...... 19

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1.4.6 Transcription, , assembly and budding...... 21

1.5 THE HIV-1 CORE ...... 23

1.5.1 CA protein ...... 23

1.5.2 The HIV-1 Core ...... 25

1.6 ASSEMBLY AND DISASSEMBLY OF THE HIV CORE ...... 28

1.6.1 Immature virion ...... 28

1.6.2 Core disassembly following viral entry ...... 30

1.6.3 Core stability and core yield ...... 31

1.6.4 Cellular factors, restriction factors and core disassembly ...... 33

1.6.5 Capsid (CA) and reverse transcription...... 38

1.6.6 Capsid (CA), nuclear import and integration ...... 40

1.7 PROTEIN PHOSPHORYLATION AND HIV ...... 44

1.7.1 Protein Phosphorylation ...... 44

1.7.2 Virion incorporated kinases ...... 44

1.7.3 Phosphorylation of HIV proteins ...... 46

1.7.4 CA phosphorylation & potential kinases ...... 49

1.8 AIMS AND SCOPE OF THESIS...... 51

CHAPTER 2 - MATERIALS AND METHODS ...... 53

2.1 MATERIALS ...... 53

2.1.1 culture ...... 53

2.1.2 Plasmid vectors ...... 54

2.1.3 Virus stocks ...... 54

2.1.4 Bacterial culture ...... 54

2.1.5 Common solutions ...... 54

2.1.6 Antibodies ...... 55

2.2 DNA CLONING AND EXTRACTION ...... 56

2.2.1 Restriction digestion ...... 56

2.2.2 Agarose gel electrophoresis ...... 56 ii

2.2.3 DNA extraction from agarose gels ...... 56

2.2.4 Ligation ...... 57

2.2.5 Transformation ...... 57

2.3 DNA EXTRACTION FROM CELLS ...... 57

2.3.1 Plasmid extraction and purification ...... 57

2.3.2 HIRT DNA extraction ...... 57

2.3.3 Spectrophotometry and DNA quantitation ...... 58

2.4 CONSTRUCTION OF CAPSID PHOSPHORYLATION MUTANTS ...... 58

2.4.1 PCR mutagenesis of the CA region ...... 58

2.4.2 Site-directed PCR mutagenesis of CA in pET-32a ...... 59

2.4.3 Overlap extension PCR mutagenesis ...... 59

2.4.4 PCR cycling parameters ...... 59

2.4.5 Subcloning mutant CA sequences to generate full length infectious pNL4-3 ...... 60

2.4.6 Screening CA mutant clones by restriction digest analysis ...... 60

2.5 CONFIRMATION OF MUTATIONS BY SEQUENCE ANALYSIS ...... 61

2.5.1 Sequencing primers...... 61

2.5.2 Sequencing PCR ...... 61

2.6 CELL TRANSFECTIONS AND VIRUS ISOLATION ...... 62

2.6.1 Transfection ...... 62

2.6.2 Recovery of virus for infections ...... 62

2.6.3 Concentration of virus preparations ...... 62

2.6.4 Pelleting virus preparations ...... 62

2.6.5 Harvesting transfected cell lysates ...... 63

2.7 CELL INFECTIONS ...... 63

2.7.1 Virus CA p24 measurement ...... 63

2.7.2 HuT-78 cell infection ...... 63

2.7.3 Single cycle infectivity assay ...... 64

2.7.4 Fate-of-capsid infection ...... 64

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2.8 REAL-TIME PCR ...... 65

2.9 PROTEIN ANALYSIS ...... 65

2.9.1 Protein quantitation assay ...... 65

2.9.2 Protein precipitation of sucrose gradient fractions ...... 66

2.9.3 Preparation of cell lysates from fate-of-capsid infections ...... 66

2.10 WESTERN BLOT ANALYSIS ...... 66

2.11 SUCROSE GRADIENT ULTRACENTRIFUGATION ...... 67

2.11.1 Sucrose solutions ...... 67

2.11.2 Preparation of gradients ...... 67

2.11.3 Collection of fractions ...... 67

2.11.4 Measuring refractive index ...... 68

CHAPTER 3 - CHARACTERISATION OF CAPSID MUTANT ...... 69

3.1 INTRODUCTION ...... 69

3.2 RESULTS ...... 71

3.2.1 Analysis of profiles of CA mutant viruses ...... 72

3.2.2 Some CA mutant viruses have altered replication competence and reverse transcription

ability……………………………………………………………………………………………………………………………………..74

3.3 DISCUSSION ...... 75

CHAPTER 4 - IN VITRO ANALYSIS OF CORE STABILITY AS A MEASURE OF CORE DISASSEMBLY ...... 79

4.1 INTRODUCTION ...... 79

4.2 RESULTS ...... 81

4.2.1 Methods to identify core structures...... 81

4.3 ANALYSIS OF DETERGENT TREATED VIRUS BY RATE ZONAL ULTRACENTRIFUGATION ...... 85

4.3.1 Increasing detergent concentration results in release of free CA protein from core

structures ...... 85

4.3.2 Development of conditions to remove the ...... 86

4.3.3 0.1% Triton-X-100 distinguishes between WT and hyperstable virus ...... 87

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4.3.4 WT and hypostable virus cores could not be distinguished using rate zonal

ultracentrifugation ...... 88

4.4 ANALYSIS OF CORE STABILITY OF CA MUTANTS ...... 89

4.4.1 CA mutants S41A, S109A, S146A, S149A, S178A and T188V are not hyperstable ...... 89

4.4.2 CA mutants exhibit WT (S41A, S146A, T188V) or hypostable (S109A, S149A, S178A)

cores………………………………………………………………………………………………………………………………………..90

4.5 DISCUSSION ...... 92

CHAPTER 5 - PERSISTENCE OF CA FOLLOWING INFECTION AS A MEASURE OF CORE DISASSEMBLY IN

THE INFECTED CELL ...... 98

5.1 INTRODUCTION ...... 98

5.2 RESULTS ...... 101

5.2.1 Optimisation of cell-free infection for fate-of-capsid assay ...... 101

5.2.2 Virus Stocks ...... 101

5.2.3 Enhancing the fate-of-capsid assay ...... 102

5.2.4 Protein Analysis of infected cell lysates ...... 103

5.3 ANALYSIS OF CA PERSISTENCE IN INFECTED CELLS ...... 104

5.3.1 CA is detected within infected target cells up to 6 hr post-infection...... 104

5.3.2 CA complexes change subtly 2-6 hr post infection ...... 105

5.3.3 CA persistence inside infected cells does not correlate with in vitro core stability ...... 106

5.4 DISCUSSION ...... 107

CHAPTER 6 - GENERAL DISCUSSION ...... 114

6.1 THE ROLE OF THE VIRAL CORE IN THE EARLY STAGES OF HIV REPLICATION ...... 114

6.2 REPLICATION OF HIV CA MUTANT VIRUSES ...... 115

6.3 HIV P24 CA AND CORE STABILITY ...... 119

6.4 HIV CORE DISASSEMBLY DURING INFECTION ...... 123

6.5 CONCLUDING REMARKS ...... 125

6.6 FUTURE WORK ...... 127

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APPENDIX 1...... 130

APPENDIX 2...... 131

APPENDIX 3...... 132

BIBLIOGRAPHY ...... 140

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ABSTRACT

Disassembly of the HIV viral core describes the rearrangement and release of capsid (CA) from the core following entry into the host cell. In this process, while the conical shaped core may be lost, some CA remains associated with the resulting reverse transcription (RTC) and preintegration complexes (PIC).

What triggers release of CA from the core is unknown. Cores from virus containing mutations in CA that show altered core stability, and release CA from the core at rates faster or slower than wild type (WT) virus demonstrate blocks in replication during reverse transcription and nuclear translocation. How the CA protein affects theses process is not understood, but intrinsic stability of the core is instrumental in regulating interactions with cellular factors.

Evidence suggests that core disassembly is critical for the early steps in HIV replication and it may regulate replication in a cell type dependent fashion.

Mutation of charged residues throughout CA results in viruses displaying altered core stability. Regulation of charge in the core, possibly by phosphorylation of CA, is one potential mechanism that may control core disassembly. Substitution of serine residues within CA illustrates five viruses, including three representing the major phospho-acceptor sites (S109,

S149 and S178) that show altered replication profiles.

To explore the role of these residues in core disassembly, the present study investigated the in vitro stability and the intracellular disassembly of the cores from these viruses. Chapter 3 describes the characterisation of viruses with mutations in CA at S41A, S109A, S146A,

S149A, S178A and T188V to analyse the effect of substitution at these sites on viral replication. Substitution at S109, S149, S178 and T188 reduced replication competence and altered the production of reverse transcription intermediates. S41A and S146A demonstrated altered reverse transcription, but did not result in blocks in replication.

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Chapter 4 describes modification of an assay to examine viral core stability. Using this assay,

CA mutant viruses (S109A, S149A and S178A) demonstrated reduced in vitro stability of the viral core in comparison with WT NL4-3 virus. Analysis of core disassembly following cell infection (Chapter 5) could not identify defects in core disassembly inside the cell, but suggested progressive changes occurred to viral complexes following infection.

The results in this thesis suggest that substitution in CA at S109, S149 and S178 alters in vitro core stability in these viruses, and may impact on core disassembly during HIV replication.

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DECLARATION

This work contains no material which has been accepted for the award of any other degree or diploma in any university or other tertiary institution to Sarah Martin and, to the best of my knowledge and belief, contains no material previously published or written by another person, except where due reference has been made in the text. I give consent to this copy of my thesis, when deposited in the University Library, being made available for loan and photocopying, subject to the provisions of the Copyright Act 1968. I also give permission for the digital version of my thesis to be made available on the web, via the University’s digital research repository, the Library catalogue, the Australasian Digital Theses Program (ADTP) and also through web search engines, unless permission has been granted by the University to restrict access for a period of time.

SARAH MARYROSE CONNOR MARTIN

August 2013

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ACKNOWLEDGEMENTS

Acknowledgements must go to my supervisors Dr Li Peng and Professor Chris Burrell, and Dr Adam Davis. Thank you for giving me the opportunity to carry out this project in your laboratory, and for your support and feedback throughout my extended PhD candidature. Many thanks must go to Dr Jill Carr for all your help and guidance shaping my thesis.

Thank you to all the members of the HIV laboratory. It has been a huge learning curve from Honours till PhD. Thank you to John Karlis for assistance with infection experiments. Also, thank you to Carl Coolen for advice on running sucrose gradients. Thank you to Adrian Purins for preparing my cells. Thank you Kelly Cheney and Satiya Wati, for so much advice on so many things.

Acknowledgement must go to Chris Bagley for help with protein modelling and images; and to Peter Zilm for access to 2D PAGE equipment. Many thanks to IMVS Photography, especially to Peter for scanning hundreds of western blot films for me.

Thank you to Mum and Dad for your love and encouragement.

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ABBREVIATIONS

Α Alpha DNA deoxyribonucleic acid A Alanine dNTP deoxynucleoside triphosphate Å angstrom(s) aa dsDNA double stranded DNA AIDS acquired immune deficiency dUTP deoxyuridine triphosphate syndrome E glutamic acid AK auto-activated protein kinase EDTA ethylenediaminetetraacetic acid AMV avian myeloblastosis virus EIAV equine infectious anaemia virus ARV AIDS-associated ELISA enzyme-linked immunosorbent ATP adenosine triphosphate assay AZT Azidothymidine EM electron microscopy Β Beta ER βME β-mercaptoethanol ERK2 extracellular-signal regulated kinase bp base pair(s) ERT endogenous reverse BSA bovine serum albumin transcription °C degrees Celsius FAK focal adhesion kinase CA, p24 Capsid FCS foetal calf serum cAMP cyclic AMP FDA Food and Drug Administration CaMV cauliflower mosaic virus FeLV feline leukaemia virus CCR5 chemokine (C-C motif) receptor FMDV foot and mouth disease virus 5 FPLC fast phase liquid CDC Centers for Disease Control and chromatography Prevention G gram(s) cDNA coding DNA x g g force CEL core envelope linkage G glycine CK-II casein kinase II g/mL grams per millilitre CO2 carbon dioxide GDP guanosine diphosphate CPK cytosolic protamine kinase GFP green fluorescent protein C-PKA cyclic AMP-dependent protein GTP guanosine triphosphate kinase HAART highly active anti-retroviral CTD carboxyl terminal domain therapy CTL cytotoxic T lymphocyte HBV CXCR4 chemokine (C-X-C motif) HCV virus receptor 4 HIV human immunodeficiency virus CypA cyclophilin A HLA-DR human leukocyte antigen DR D aspartic acid HPV human papilloma virus Da dalton(s) hr hour(s) DHBV duck hepatitis B virus HRP horse radish peroxidase DMEM Dulbecco's Modified Eagle's Hsp70 heat shock protein 70 medium DMSO dimethyl sulfoxide HSV virus

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HTLV human T-cell leukaemia virus N Asparagine ICTV International Committee on the NC Nucleocapsid Taxonomy of Viruses NDR nuclear Dbf2 related kinase IDAV immunodeficiency-associated negative replication factor virus NERT natural endogenous reverse IL Interleukin transcription IN NIH National Institute of Health K Lysine NMR nuclear magnetic resonance kb kilobase(s) NNRTI non-nucleoside reverse kDa kilodalton(s) transcriptase inhibitor L Leucine NPC nuclear pore complex LAV lymphadenopathy-associated NRTI nucleoside virus inhibitor LB Luria-Bertani broth nt Nucleotide Lck lymphocyte-specific protein NTD amino terminal domain tyrosine kinase OD optical density LTR long terminal repeat OH hydroxyl group M Methionine P Proline M molar (moles/litre) p.i. post infection MA Matrix PAGE polyacrylamide gel MAPK mitogen activated protein electrophoresis kinase PBS phosphate buffered saline MBPK myelin basic protein kinase PBS primer binding site MEK MAPK ERK2 kinase PCR polymerase chain reaction mg milligram(s) pI isoelectric point MgCl2 magnesium chloride PI protease inhibitor MHC major histocompatibility PIC preintegration complex complex PKC protein kinase C MHR major homology region PKR RNA dependent protein kinase min minute(s) PO4 Phosphate MIP macrophage inflammatory PPT polypurine tract protein PR protease inhibitor mL millilitre(s) PVA potato virus A MLV murine leukaemia virus PVDF polyvinylidene fluoride mm millimetre(s) pyk2 proline-rich tyrosine kinase 2 mM Millimolar Q Glutamine MMTV mouse mammary tumour virus R Arginine M- moloney murine leukaemia MuLV virus R repeat region MOI multiplicity of infection Rev regulator of virion expression mRNA messenger RNA RIPA radio immunoprecipitation MTOC microtubule-organising centre assay RNA ribonucleic acid MWCO molecular weight cut-off RNaseH ribonuclease H ng nanogram(s) rpm revolutions per minute nm nanometer(s)

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RRE Rev response element > greater than RT reverse transcriptase ≥ equal to or greater than RTC reverse transcription complex S serine < less than SDS sodium dodecyl sulfate sec second(s) SIV simian immunodeficiency virus ssDNA single stranded DNA T Threonine TAR trans-activating response region TAS temperature arrested state (of fusion) Tat trans-activator protein TCA trichloroacetic acid TEM transmission electron microscopy TNF tumour necrosis factor TRIM5α tripartite motif protein 5 alpha μg microgram(s) µL microlitre(s) U3 or U5 untranslated region (3' or 5') UNAIDS Joint United Nations Programme on AIDS UV Ultraviolet V Valine V volt(s) v/v volume per volume VAPK virus-associated protein kinase Vif virion infectivity factor VLP virus-like particles viral protein R Vpu viral protein U VSV-G vesicular stomatitis virus W Tryptophan WHO World Health Organisation WT wild type [3H]- tritiated thymidine dTTP

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PRESENTATIONS ARISING

Conference Presentations

Oral Presentations:

Oct 2006 Australian Centre for Hepatitis Virology & HIV Virology Interest Group 3rd National Scientific Workshop in Lorne, Victoria: HIV-1 capsid phosphorylation, capsid disassembly and reverse transcription

Poster Presentations:

Dec 2005 The 3rd Scientific Meeting of the Australian Virology Group at Cowes, Phillip Island, Victoria: Investigating a role for HIV-1 Capsid Phosphorylation in Virus Uncoating and Reverse Transcription (abstract #253)

July 2007 Annual Scientific Meeting of the Australian Society for Microbiology, in Adelaide, South Australia: HIV-1 Capsid Phosphorylation and Core Disassembly

Dec 2007 The 4th Scientific Meeting of the Australian Virology Group at Fraser Island, Queensland: Investigating the role of HIV-1 capsid phosphorylation in capsid disassembly and reverse transcription (abstract #143)

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CHAPTER 1 - Introduction

1.1 HIV and AIDS

1.1.1 History and identification

The first cases of acquired immunodeficiency syndrome (AIDS) were documented in 1981 in the United States (US) (1981). AIDS presented as an epidemic immunosuppressive disease.

Previously healthy individuals became predisposed to opportunistic infections and tumours such as Pneumocystis carinii pneumonia, cytomegalovirus and Kaposi’s sarcoma (Gottlieb et al., 1981; Siegal et al., 1981). Although epidemiological data was consistent with viral aetiology, it was some time before AIDS was attributed to the exogenous retrovirus human immunodeficiency virus type 1 (HIV-1) (Barre-Sinoussi et al., 1983; Gallo et al., 1987; Gallo et al., 1984; Levy et al., 1984). Prior to this, immunosuppressive properties of sperm, Epstein-

Barr virus, cytomegalovirus and even a fungus, were initially considered as potential causes of AIDS (Broder & Gallo, 1984).

In 1982 a retrovirus was proposed as the aetiological agent responsible for AIDS. Similarities suggested the retrovirus was potentially related to the human T-cell leukaemia virus (HTLV), which had only been recently identified at the time. Essex et al (1983) hinted that the most common cancer caused by feline leukaemia virus (FeLV) was a T–cell leukaemia, while the most common disease caused by this agent was an acquired T-cell immunodeficiency (Essex et al., 1983), characteristics echoed by the AIDS epidemic (Trainin et al., 1983).

HIV-1 shares several common features with HTLV-I and HTLV-II. They each possess a reverse transcriptase, infect mature T cells in vitro and form giant multi-nucleated cells called syncytia. Exogenous virus can be isolated from mature T cells, notably CD4+ T lymphocytes.

Additionally, these viruses share some nucleotide sequence homology. They have similarly sized major core proteins and possess cross reacting antigens.

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Early studies showed sera from many patients at risk of acquiring AIDS recognised HTLV-I- associated antigen (Essex et al., 1983). However, while most of these patients had antibodies against the cell membrane antigen (Essex et al., 1983), neither HTLV-I or II virions, nor antibodies against their structural core proteins were isolated from these individuals (Robert-Guroff et al., 1984). Cytopathic viruses similar to known members of the

HTLV family were detected in patients with AIDS as early as November 1982. However, many were not reported until 1984 due to difficulties propagating these viruses (Barre-

Sinoussi et al., 1983; Chermann et al., 1983; Gallo et al., 1984; Gallo et al., 1983; Gelmann et al., 1983; Levy et al., 1984). Further difficulties identifying the retrovirus responsible for

AIDS stemmed from the extreme genomic diversity displayed by these viruses. Between 1984 and 1987 no two HIV-1 virus isolates were identical (Barre-Sinoussi et al., 1983; Wain-

Hobson et al., 1985; Wong-Staal et al., 1985). Difficulties propagating virus due to its cytopathicity led to isolation of many genetically diverse viral (HIV-1) isolates by different laboratories, which were designated as different viruses; e.g. HTLV-III, human T-cell leukaemia virus III (Gallo, National Institute of Health (NIH), US); LAV, lymphadenopathy- associated virus; IDAV, immunodeficiency-associated virus (Montagnier, Pasteur Institute,

France); and ARV, AIDS-associated retrovirus (Levy, University of California School of

Medicine, US) (Gallo & Wong-Staal, 1985).

Françoise Barré-Sinoussi and Luc Montagnier at the Pasteur Institute in France reported isolation of from patients at risk of AIDS in 1983 (Barre-Sinoussi et al., 1983;

Chermann et al., 1983). Isolation of retroviruses was also reported in AIDS patients by Robert

Gallo at the NIH in the US (Gallo et al., 1983). Integrated HTLV proviral DNA sequences were also detected in patients with AIDS (Gelmann et al., 1983), and a correlation was made between patients with HIV-1 and AIDS in 1984 (Gallo et al., 1984; Markham et al., 1984).

Following much controversy, Barré-Sinoussi and Montagnier were awarded the Nobel Prize for Medicine for the discovery of HIV in 2008 (2008b).

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The production of the H9/HTLV-IIIB system (Popovic et al., 1984; Ratner et al., 1987), resulted in significant scientific headway. The permanent virus producing cell line allowed mass production of virus, and the production of the first specific reagents such as antibodies and nucleic acid probes. Enzyme-linked immunosorbent assays (ELISA) detected circulating antibodies against HIV-1 in 90% of AIDS patients (Sarngadharan et al., 1984). Shortly thereafter, western blot was successful in detecting antibodies against HIV-1 in 100% of patients (Safai et al., 1984).

In 1986 the virus responsible for AIDS was designated as the human immunodeficiency virus type 1 (HIV-1) by the International Committee on the Taxonomy of Viruses (ICTV) (Brown,

1986; Coffin et al., 1986). This follows the nomenclature convention identifying the species affected and a major pathogenic characteristic.

1.1.2 Classification

HIV-1 belongs to the retroviridae family of viruses, which primarily infect vertebrate hosts.

Retroviruses characteristically encode three regions; gag, and genes. HIV possesses two identical copies of a positive sense RNA genome, which is reverse transcribed by a virally encoded DNA polymerase into double stranded DNA (dsDNA), before integration into the host cell chromosome to form the provirus.

The retroviridae family is divided into two subfamilies: Orthoretrovirinae and

Spumaretrovirinae. These are further divided into genera, including both simple and complex retroviruses. Simple retroviruses encode only Gag, Pol and Env gene products, while complex retroviruses also encode regulatory or accessory proteins. There are three simple retrovirus genera: alpharetroviruses, betaretroviruses and gammaretroviruses; and four complex retroviruses: deltaretroviruses, epsilonretroviruses, lentiviruses and spumaviruses. All these virus genera fall under the Orthoretrovirinae subfamily, with the exception of spumaviruses which belong to the Spumaretrovirinae subfamily (2002).

HIV is subclassed as a lentivirus. These viruses cause slow progressive types of disease and are often characterised by long asymptomatic periods of infection before the onset of disease.

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There are two HIV strains, HIV-1 and HIV-2. HIV-1 is divided into three major phylogenetic groups: M (major), N (non-M or O) and O (outlier), all of which result in AIDS. Group M, which contains 95% of isolates globally, is further divided into 10 clades (A-J). Group O has fewer isolates, which are more restricted in their geographical location, while group N has very few isolates (Gao et al., 1999).

HIV-2 was first isolated from patients in West Africa in 1986 (Clavel et al., 1986). HIV-2 has remained confined to West Africa and is most prevalent in Guinea-Bissau. HIV-1 and HIV-2 share 40-60% structural and sequence homology. Unlike those infected with HIV-1, the majority of individuals with HIV-2 will not progress to disease. This contrast may provide useful insights for understanding the pathogenesis of HIV-1. All studies in this thesis use the

HIV-1 strains NL4-3 or HXB2.

1.2 HIV-1 Epidemiology and Pathogenesis

1.2.1 Epidemiology

In June 1981, AIDS was first brought to the attention of the international medical profession with the report of five previously healthy men who presented with severe immunodeficiency

(Gottlieb et al., 1981; Siegal et al., 1981). All individuals exhibited deficient cell-mediated immunity and reduced numbers of circulating CD4+ T cells. As a result, these men became susceptible to infections not normally seen in healthy individuals (Gottlieb et al., 1981; Siegal et al., 1981). By February 1983, 1000 cases of AIDS had been reported to the US Centers for

Disease Control and Prevention (CDC), and the numbers continued to rise rapidly (Jaffe et al., 1983).

By December 2011, there was an estimated 34 million people living with HIV/AIDS worldwide (Figure 1.1). This includes 16.7 million women and 3.3 million children under 15 years of age ((UNAIDS), 2012). 2.5 million new HIV infections occurred during the year of

2011 alone ((UNAIDS), 2012).

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Figure 1.1 Worldwide distribution of HIV In 2011 34 million people were estimated to be living with HIV. This image depicts the worldwide distribution of HIV in adults.

Adapted from Global report: UNAIDS report on the global AIDS epidemic 2010.

Sub-Saharan Africa remains the most affected region globally. It is home to greater than two thirds (69%) of HIV positive individuals (an estimated 23.5 million people). More than two- thirds (71%) of new infections (1.8 million) acquired during 2011 occurred in Sub-Saharan

Africa. Furthermore, 72% (1.3 million) of AIDS-related deaths occurred in Sub-Saharan

Africa in 2011 ((UNAIDS), 2012). In contrast to trends in other areas, the majority of infected individuals are women ((UNAIDS), 2010; 2012).

The latest estimates produced by The Joint United Nations Programme on HIV/AIDS

(UNAIDS) reported a decline in the numbers of new infections in Sub-Saharan Africa, South and South-East Asia, the Caribbean and Latin America ((UNAIDS), 2012). An increase in the number of infections was reported in East-Asia, Oceania, North America and Western

Europe, with marked increases in the Middle East, North Africa and Eastern Europe. The proportion of HIV infected men and women remained steady, with women making up 49% of infected individuals. 80% of all women living with HIV are situated in Sub-Saharan Africa

((UNAIDS), 2010).

In Australia between 1982 and December 2011, 32166 people were diagnosed with HIV infections, and 10839 were individuals diagnosed with AIDS. In the same time, 6846 individuals died from AIDS-related deaths (2012b). There were 1060 new diagnoses in the year to June 2012 (2012b). Trends show that the rate of new infection has declined in New

South Wales from 2002-2011, while an increase in the rate of infections was observed in

Victoria, South Australia, Tasmania, Northern Territory and Australian Capital Territory. The rates of new diagnoses in Queensland and Western Australia were stable over the same time frame. While HIV prevalence in Australia remains mainly in men who have unprotected sex with men, trends are changing. 60% of new HIV infections attributed to heterosexual contact during 2007-2011 were in people from high prevalence countries (2012a).

1.2.2 Transmission

Transmission of HIV occurs through infected bodily fluids, including blood, semen, vaginal secretions and breast milk (Oleske et al., 1983). The risk of infection is affected by the

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duration and frequency of contact, the infectious dose and the competence of the infected host’s immune system.

There are two general patterns in the incidence of HIV infection and AIDS. The distribution and trend of infection is related to social behaviour. Infection is widespread across the general population in Sub-Saharan Africa. In many other parts of the world it remains concentrated to

‘at risk’ populations: men who have unprotected sex with men, injecting drug users, sex workers and their sexual partners.

The first reported cases in the US illustrated transmission predominantly through homosexual contact. In many countries including the US, Brazil, UK, Australia and New Zealand, this is still the main route of transmission ((UNAIDS), 2012). However, in regions such as Africa, the rate of HIV infection is high and the prevalence of infection is similar in males and females. This is a direct consequence of primarily heterosexual transmission in this area.

Transmission in China, Pakistan, Viet Nam and Indonesia is dominated by injecting drug users. This includes increasing numbers of women, and transmission through unprotected sex with non-regular partners, sex workers or between men (2007). Heterosexual transmission accounted for the majority of infections in Argentina, and increasing numbers of infections in the UK, US and NZ ((UNAIDS), 2012).

The introduction of routine blood screening in 1985 has all but eliminated transmission during blood transfusions (Curran, 1985). Use of HIV infected blood products (containing factor

VIII) led to infection of many people with haemophilia and other blood disorders.

Likewise, while transmission can occur from mother to child during pregnancy, at the time of birth or through breast milk, the majority of neonatal cases can be prevented. Introduction of

Zidovudine in 1994, a reverse transcriptase (RT) inhibitor, significantly decreased neonatal transmission (Connor et al., 1994). In 2003, only 1% of women in severely affected countries had access to drug therapy (2003; Pomerantz & Horn, 2003). By 2005, treatment was available to 15% of women in low or middle income countries (2007). In 2009, 53% of women in low or middle income countries, including 54% of women in sub-Saharan Africa,

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received anti-retrovirals to lower the risk of mother to child transmission ((UNAIDS), 2010).

WHO aim to eliminate infections in children by 2015 with the introduction of new treatment options (WHO, 2012a). Amendments have been made to current protocols which aim to simplify treatment regimens. Previous treatment plans were based on CD4 cell count and involved multiple drug combinations throughout pregnancy and breastfeeding. Instead, a single regimen is being implemented regardless of CD4 cell count. This recommends life long fully suppressive triple antiretroviral drug regimens for the mother, and daily nevirapine

(NVP) or azidothymidine (AZT) from birth to 4-6 weeks of age for the infant (WHO, 2012a; b).

1.2.3 Pathogenesis

HIV-1 infects cells expressing the CD4 receptor; this is largely restricted to CD4+ T lymphocytes and macrophage (Dalgleish et al., 1984). HIV-1 entry is mediated by virus attachment to the CD4 receptor on the host cell surface, in conjunction with additional chemokine receptors which act as co-receptors (Alkhatib et al., 1996a; Alkhatib et al., 1996b;

Dalgleish et al., 1984; Feng et al., 1996; Klatzmann et al., 1984; McDougal et al., 1985).

Initially viral isolates which replicate in peripheral blood T lymphocytes and monocyte- derived macrophage, but not T cell lines, were deemed macrophage (M) tropic. Viral isolates that replicated in T cell lines were deemed T cell (T) tropic (Collman et al., 1989). The discovery of isolates which could replicate in all these cell types were designated dual (D) tropic and indicated limitations to this classification system. Tropism mostly correlates with co-receptor usage (Deng et al., 1996; Doranz et al., 1996; Feng et al., 1996). Thus viral isolates were designated R5 (chemokine (C-C motif) receptor 5, CCR5), X4 (chemokine (C-

X-C motif) receptor 4, CXCR4) or R5X4 (CCR5 and CXCR4) based on co-receptor usage

(Berger et al., 1998). Dual tropic R5X4 viruses may be further classified as dual-R or dual-X, referring to more efficient use of CCR5 over CXCR4, or CXCR4 over CCR5 respectively, in the target cell population (Huang et al., 2007; Irlbeck et al., 2008; Yi et al., 1999).

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R5 strains predominate during early and chronic HIV-1 infection (Keele et al., 2008), and are important for initiating infection and establishing persistence (Miedema et al., 1994; Penn et al., 1999). R5 isolates dominate throughout disease progression in the majority of patients. As patients progress to AIDS, the viral population can diversify to use multiple co-receptors, changing from R5 to R5X4, or exclusively X4 Schuitemaker, 1992 #494}. Although the prevalence of CXCR4 usage appears to be continually changing, pure X4 strains are rare and usually restricted to later stages of disease (Melby et al., 2006; Miedema et al., 1994; Penn et al., 1999).

HIV-1 has also been found in CD4 negative cells including astrocytes, glial cells and dendritic cells (Gorry et al., 2003; Patterson & Knight, 1987). These cell types appear to predominantly trap or harbour virus particles in vivo, but evidence suggests that they may also support limited viral replication (Clarke et al., 2006; Gorry et al., 2003).

During the first 2-6 weeks after infection there is dramatic viral replication (Figure 1.2)

(Schacker et al., 1996). Approximately 10-20% of patients exhibit an acute syndrome characterised by fever, lymphadenopathy and occasional meningitis. This period is asymptomatic in most patients.

The initial burst of viral replication is controlled by a CD8+ MHC class I-restricted anti-HIV-

1 cytotoxic T lymphocyte (CTL) response predominantly against the envelope glycoprotein

(gp120), as well as Gag. This results in a long asymptomatic period. However, HIV-1 persistence is ensured by rapid replication by HIV-1 throughout infection (up to 1010 virions are produced per day). The entrapment of virions within germinal centres of follicular dendritic cell networks and HIV infected cells that harbour provirus without producing viral proteins also ensure persistence (Heath et al., 1995).

During the asymptomatic stage of infection viral load remains low, but constant and rapid

HIV replication destroys nearly a billion T cells each day (Wei et al., 1995). Productive infection of CD4+ cells with HIV is sufficient to induce cell death of both infected and uninfected cells (Finkel et al., 1995; Sylwester et al., 1998). HIV infection induces cell death

8

Figure 1.2 HIV pathogenesis Upon initial HIV infection there is substantial virus replication. Following the initial burst of replication HIV individuals experience an asymptomatic period, viral load drops, but viral replication remains constant. HIV infection causes extensive death of CD4+ T cells resulting in impaired immune responses. AIDS develops when CD4 T cell counts drop below 200 cells/µL and patients become susceptible to opportunistic infections. Rowland-Jones, Nature Reviews Immunology, 2003. directly through increased permeability of cellular membranes and loss of cell integrity

(Leonard et al., 1988). Cell death can be induced by cytotoxic T cell killing and various apoptosis mechanisms. Syncytia, which is the result of fusion between infected cells with uninfected cells to create giant multinucleated cells, also promotes the loss of cell viability

(Lifson et al., 1986). HIV-1 specific antibodies can lead to the activation of the complement cascade which results in lysis of infected cells, and accounts for further cell death.

Within 6-8 weeks following infection, an antibody response is initiated predominantly against the viral envelope glycoprotein gp120 and p24 capsid (CA) protein. Detection of p24 CA antibodies is used as a clinical diagnostic marker eight weeks following suspected infection with HIV-1. The slow production of HIV-specific antibodies, together with rapid mutation allows HIV to evade host cell control mechanisms (Poignard et al., 1999). In addition to destruction of CD4+ T cells, macrophage function is disrupted significantly by HIV-1 infection. This impacts on T cell function and results in impaired immune responses to intracellular pathogens. De-regulation of cytokine networks promotes increased production of

HIV-1 inducing pro-inflammatory cytokines (e.g. TNF-α and IL-1β) and decreased production of immunoregulatory cytokines (e.g. IL-2 and IL-12) (Clerici et al., 1993; Cohen et al., 1997). As disease progresses, the ability to mount HIV-1 specific humoral responses reduce until the destruction of lymphoid tissue and immune dysfunction result in the onset of

AIDS.

The time of the asymptomatic period in untreated patients may vary from months to years, with an average 10 years before the onset of symptomatic AIDS (Borrow et al., 1994). Anti- retroviral therapy improves patients’ clinical prognoses by extending the asymptomatic period. The US CDC defines three stages of disease following laboratory confirmation of HIV infection, based on the level of immunosuppression. Stage 1: CD4+ T-lymphocyte count of ≥

500 cells/µL, stage 2: CD4+ T-lymphocyte count of 200-499 cells/µL and stage 3: which defines AIDS, a CD4+ T-lymphocyte count of < 200 cells/µL and the appearance of one or more of 27 AIDS-defining conditions (Table 1.1) (Schneider et al., 2008). The inability to

9

Table 1.1 AIDS defining conditions Following a laboratory confirmed case of HIV infection and a CD4+ T lymphocyte count less than 200 cells/µL, AIDS is defined by the acquisition of one or more of the following conditions.

Schneider et al, MMWR Recomm Rep. 2008 Appendix A : AIDS defining conditions AIDS defining conditions

Bacterial infections, multiple or recurrent Candidiasis of bronchi, trachea, or lungs Candidiasis of esophagus Cervical cancer, invasive Coccidioidomycosis, disseminated or extrapulmonary Cryptococcosis, extrapulmonary Cryptosporidiosis, chronic intestinal (>1 month's duration) Cytomegalovirus disease (other than liver, spleen, or nodes), onset at age >1 month Cytomegalovirus retinitis (with loss of vision) Encephalopathy, HIV related Herpes simplex: chronic ulcers (>1 month's duration) or bronchitis, pneumonitis, or esophagitis (onset at age >1 month) Histoplasmosis, disseminated or extrapulmonary Isosporiasis, chronic intestinal (>1 month's duration) Kaposi sarcoma Lymphoid interstitial pneumonia or pulmonary lymphoid hyperplasia complex Lymphoma, Burkitt (or equivalent term) Lymphoma, immunoblastic (or equivalent term) Lymphoma, primary, of brain Mycobacterium avium complex or Mycobacterium kansasii, disseminated or extrapulmonary Mycobacterium tuberculosis of any site, pulmonary, disseminated, or extrapulmonary Mycobacterium, other species or unidentified species, disseminated or extrapulmonary Pneumocystis jirovecii pneumonia Pneumonia, recurrent Progressive multifocal leukoencephalopathy Salmonella septicemia, recurrent Toxoplasmosis of brain, onset at age >1 month Wasting syndrome attributed to HIV mount an effective immune response, primarily the result of T cell depletion, allows acquisition of opportunistic infections (fungal, viral, protozoan or bacterial) and cancers. Loss of at least 10% body mass or wasting syndrome and dementia may also occur. A formal diagnosis of AIDS is made when these clinical criteria are met and death usually follows within 2 years.

1.2.4 HIV-1 Prevention and Therapy

Although HIV and AIDS were first reported more than twenty-five years ago, worldwide efforts to generate an effective HIV-1 vaccine have not been successful. Efforts to generate either a therapeutic or prophylactic HIV vaccine have highlighted the importance of determining the type of immune response necessary to overcome HIV infection. Vaccines targeting HIV have been designed to generate strong antibody-mediated immune responses or alternatively cell-mediated responses. However, it appears both types of immune responses are required (Corey et al., 1998). Non-human primate studies suggest high levels of antibodies may be capable of protecting against HIV, but current vaccines have not been able to induce sufficient antibody levels. The level of antibodies required in vivo may be up to

1000-fold higher than levels required for neutralisation in vitro.

Currently, protocols for HIV-1 therapy aim at decreasing virus replication thereby helping maintain or increase T cell numbers in order to delay opportunistic infections (McMichael &

Hanke, 2003).

Treatment of HIV infected individuals relies on anti-retroviral therapy. There are currently close to thirty anti-retroviral compounds (Table 1.2) approved by the US Food and Drug

Administration (FDA). These target different stages of the HIV life cycle including viral entry, reverse transcription, integration and maturation (FDA, 2013).

Anti-retroviral therapy became available for HIV infected individuals in 1987 with the approval of Azidothymidine (AZT), the first of the nucleoside reverse transcriptase inhibitors

(NRTI). These inhibitors are nucleoside analogues lacking the 3’OH group required for addition of subsequent DNA nucleotides, thereby terminating synthesis of nascent viral DNA

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Brand Name Generic Names Approval Date

Multi-class Combination Products Atripla efavirenz, emtricitabine and tenofovir disoproxil fumarate 12-Jul-2006 emtricitabine, rilpivirine, and tenofovir disoproxil Complera 10-Aug-2011 fumarate elvitegravir, cobicistat, emtricitabine, tenofovir disoproxil Stribild 27-Aug-2012 fumarate

Nucleoside Reverse Transcriptase Inhibitors (NRTIs) Combivir lamivudine and zidovudine 27-Sep-1997

Emtriva emtricitabine, FTC 02-Jul-2003

Epivir lamivudine, 3TC 17-Nov-1995

Epzicom abacavir and lamivudine 02-Aug-2004

Hivid zalcitabine, dideoxycytidine, ddC 19-Jun-1992

Retrovir zidovudine, azidothymidine, AZT, ZDV 19-Mar-1987

Trizivir abacavir, zidovudine, and lamivudine 14-Nov-2000

Truvada tenofovir disoproxil fumarate and emtricitabine 02-Aug-2004

Videx EC enteric coated didanosine, ddI EC 31-Oct-2000

Videx didanosine, dideoxyinosine, ddI 09-Oct-1991

Viread tenofovir disoproxil fumarate, TDF 26-Oct-2001

Zerit stavudine, d4T 24-Jun-1994 Ziagen abacavir sulfate, ABC 17-Dec-1998

Nonnucleoside Reverse Transcriptase Inhibitors (NNRTIs) Edurant rilpivirine 20-May-2011

Intelence etravirine 18-Jan-2008

Rescriptor delavirdine, DLV 04-Apr-1997

Sustiva efavirenz, EFV 17-Sep-1998

Viramune nevirapine, NVP 21-Jun-1996

Viramune XR nevirapine, NVP 25-Mar-2011

Protease Inhibitors (PIs) Agenerase amprenavir, APV 15-Apr-1999

Aptivus tipranavir, TPV 22-Jun-2005

Crixivan indinavir, IDV, 13-Mar-1996

Fortovase saquinavir (no longer marketed) 7-Nov-1997

Invirase saquinavir mesylate, SQV 6-Dec-1995

Kaletra lopinavir and ritonavir, LPV/RTV 15-Sep-2000

Lexiva Fosamprenavir Calcium, FOS-APV 20-Oct-2003

Norvir ritonavir, RTV 01-Mar-1996

Prezista darunavir 23-Jun-2006

Reyataz atazanavir sulfate, ATV 20-Jun-2003

Viracept nelfinavir mesylate, NFV 14-Mar-1997

Fusion Inhibitors Fuzeon enfuvirtide, T-20 13-Mar-2003

Entry Inhibitors - CCR5 Co-Receptor Antagonist Selzentry maraviroc 06-Aug-2007

HIV Integrase Strand Transfer Inhibitors Isentress raltegravir 12-Oct-2007

Table 1.2 FDA approved anti-retroviral drugs used in the treatment of HIV infection. Brand names and generic names for each approved drug are indicated, along with the date of US FDA approval. Information in this table is taken from the US FDA website. (http://www.fda.gov/ForConsumers/byAudience/ForPatientAdvocates/HIVandAIDSActivities/ucm 118915.htm) Current as of June 2013.

(De Clercq, 1988; Nakashima et al., 1986; Richman, 1988). Non-nucleoside reverse transcriptase inhibitors (NNRTI), while also targeting reverse transcription, inhibit HIV replication via an alternative mode. NNRTIs bind a hydrophobic pocket in RT preventing enzymatic activity through allosteric changes at the polymerase active site (Merluzzi et al.,

1990).

A number of inhibitors also target maturation of the HIV virion. This can be achieved in two ways; protease inhibitors (PI) target the viral protease preventing essential processing of viral proteins. The resulting virions exhibit an immature phenotype and are not infectious

(Wlodawer et al., 1989). More recently other maturation inhibitors have been shown to induce an immature phenotype, while they do not directly target the viral protease enzyme, they also prevent proper processing. An example is the compound Bevirimat which specifically prevents cleavage at the CA-p2 cleavage site in the immature virion (Li et al., 2003; Zhou et al., 2004).

Other anti-retroviral inhibitors target viral entry. The first entry inhibitor designed, Fuzeon, was released in the US in 2003 (FDA, 2013). This compound blocks fusion between the viral envelope and cellular membranes by targeting conformations of the viral transmembrane protein, thereby preventing entry of the viral core (Fauci, 2003; Pomerantz &

Horn, 2003). Other entry inhibitors act as co-receptor antagonists, and block binding between the viral receptors and host cell chemokine receptors CCR5 and CXCR4 (Murakami et al.,

1997).

An inhibitor targeting the viral integrase protein, Isentress, was released in October 2007

(FDA, 2013). Inhibition of strand transfer mediated by integrase prevents integration of viral genomic DNA into the host cell chromosome precluding further viral replication.

HIV-1 treatment is complicated by the appearance of drug resistant virus mutants which rapidly evolve during the course of therapy. Combination anti-retroviral therapy or highly active anti-retroviral therapy (HAART) was introduced in 1996. HAART involves administration of a combination of three or more drugs belonging to different classes of HIV-

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1 inhibitors. The most common combination includes two NRTIs and a PI or NNRTI.

Combination therapy is considerably more effective than monotherapy and extends the potential for anti-retroviral treatment. However this therapy relies on the identification of novel inhibitors. HAART is typically initiated in asymptomatic patients whose CD4+ T cell counts drop below 350 cells/µL (Agrawal et al., 2006). This regimen is successful in reducing viral load to an undetectable level in both plasma and tissue, and increasing CD4+ T cell counts in peripheral blood. Additionally HAART promotes restoration of immune function, including CD4 and CD8 cellular responses to antigen and normal regulation of cell apoptosis after 12 months (Ensoli & Cafaro, 2002). However, while improving patient survival,

HAART does not eliminate HIV replication. HIV virions persist in cellular reservoirs and further resistant strains often emerge with long term therapy (Agrawal et al., 2006). Other difficulties that HIV patients face include the complexity of drug administration regimens, adverse side effects and complications, and for some patients a complete lack of an appropriate immune response (Pomerantz & Horn, 2003).

Programs such as UNAIDS endeavour to provide anti-retroviral treatment to more people worldwide; however, most HIV-1 infected individuals, particularly in third world nations, do not have access to, or cannot afford treatment. In developed countries the cost of some anti- retroviral treatments have dropped from $US10000 to as little as $US100 per year, still unobtainable by many in resource-poor countries ((UNAIDS), 2013). A WHO/UNAIDS initiative was successful in placing 3 million individuals in low and middle income countries on anti-retroviral treatment regimens by 2007 (WHO/UNAIDS, 2008). As of 2013, 7 million people across Africa have access to anti-retroviral treatment ((UNAIDS), 2013). Anti- retroviral drug therapy is a great success and has enabled HIV to become a treatable disease.

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1.3 HIV-1 virion

1.3.1 HIV-1 virion

Analysis of mature HIV-1 virions by electron microscopy (EM) demonstrates virions approximately 100-140 nm in diameter (Figure 1.3) (Briggs et al., 2006; Briggs et al., 2003;

Fuller et al., 1997; Gelderblom, 1991). Mature HIV-1 virions are defined by a host-derived outer lipid membrane studded with an average of 14 trimeric envelope glycoprotein spikes

(ranging from 4-35 spikes per virion) (Zhu et al., 2006). The p17 matrix (MA) protein lines the interior of the viral envelope. Inside the viral envelope, the viral core structure is defined by a conical shell composed of the p24 capsid (CA) protein and includes the internal viral ribonucleoprotein complex. This definition of the HIV viral core will be used throughout this thesis. The internal complex at the centre of the viral core comprises the dimerised HIV-1 RNA genome bound by nucleocapsid protein and additionally the viral enzymatic proteins reverse transcriptase (RT), protease (PR) and integrase (IN). HIV-1 accessory proteins Vpr, Vif, Nef and a number of cellular proteins are also incorporated into the virion during assembly. Two-dimensional electrophoresis studies have detected 25-250 proteins on both the interior and exterior of HIV virions, suggesting the incorporation of a vast number of cellular proteins (Chertova et al., 2006; Fuchigami et al., 2002; Ott et al.,

2000). The CA region in Gag (residues 221 and 223) is responsible for incorporation of cyclophilin A (CypA) into HIV virions at a ratio of 1:10 (CypA:CA) (Franke & Luban, 1995).

CA also assists incorporation of lysyl-tRNA synthetase (LysRS) into HIV-1 virions. In turn,

LysRS directs the incorporation of human tRNALys3, the primer required for initiation of HIV-

1 reverse transcription (Kovaleski et al., 2007). Additionally of interest, several kinases are also incorporated into the HIV-1 virion (Lck, ERK2, NDR1/2, C-PKA (Cartier et al., 1997;

Cartier et al., 2003; Devroe et al., 2005; Ott et al., 2000)). In contrast, the regulatory proteins

Tat and Rev are not packaged into virions, but are transcribed and translated inside the cell when required for regulation of HIV-1 gene expression.

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Figure 1.3 The HIV virion Schematic representation of the (A) immature and (B) mature HIV virion. The virion is defined by an outer lipid membrane studded with surface envelope , gp120. The p55 Gag precursor is arranged radially in the immature virion and is cleaved to produce mature Gag proteins in the mature virion, generating the cone-shaped virion core which encapsulates the viral ribonucleoprotein complex.

Cold Spring Harbour Laboratory Press, 1997 A

B 1.3.2 HIV-1 genome

Each HIV virion possesses two copies of the HIV-1 genome, a positive-sense single stranded

RNA molecule approximately 9.2 kb in length. Characteristic of retroviruses, the HIV-1 genome encodes three main coding regions: group-specific antigen (gag), polymerase (pol) and envelope (env). gag encodes the viral structural proteins, matrix (MA), capsid (CA), nucleocapsid (NC), p6 and two spacer proteins p2 and p1. The env region encodes the viral envelope glycoprotein gp160, which is cleaved by a cellular protease to produce the surface envelope glycoprotein gp120 (SU) and transmembrane glycoprotein gp41 (TM). pol encodes the viral enzymatic proteins protease (PR), reverse transcriptase (RT) and integrase (IN).

The HIV-1 genome also contains additional overlapping open reading frames encoding proteins that possess accessory functions. The ability to encode these additional proteins classifies HIV-1 as a complex retrovirus. The HIV-1 genome encodes three regulatory proteins: Tat, Rev and Nef, and additionally three accessory proteins: Vif, Vpr and Vpu.

During reverse transcription, the incorporation of regions termed long terminal repeats

(LTRs) results in the generation of a 9.7 kb transcriptionally active proviral DNA genome.

LTRs are generated at either end of the proviral genome by duplication of short direct repeats

(R) alongside non-coding regions (U5 and U3), at the 5’ and 3’ ends of the viral genome respectively (Figure 1.4).

The viral DNA provirus is initially transcribed by the host cell RNA polymerase II to produce mRNA. Spliced transcripts are transported from the nucleus for translation of viral proteins.

Unspliced transcripts are exported from the nucleus and either translated into structural proteins, or incorporated into nascent virions as genomic RNA.

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Figure 1.4 The HIV genome Representation of the HIV proviral genome indicating the 3 major regions: gag, pol and env. gag and env encode the structural proteins. pol encodes the viral enzymatic proteins. Additionally HIV is a complex retrovirus and encodes 6 accessory or regulatory proteins : Vif, Vpr, Vpu, Rev, Nef and Tat.

Human Retroviruses and AIDS, 1998 http://www.hiv.lanl.gov/content/sequence/HIV/MAP/landmark.html

1.4 HIV-1 Replication cycle

1.4.1 Overview of HIV-1 replication cycle

Fusion between the viral and host cell membranes facilitates entry of HIV virions into the host cell cytoplasm (Figure 1.5). The CA shell encapsulating the viral core is restructured to reveal the reverse transcription complex (RTC). This complex contains the RT enzyme responsible for reverse transcribing the single stranded RNA genome into dsDNA. RTCs shed some viral proteins in preparation for integration of newly synthesised DNA into the host cell chromosome. This newly formed complex is termed the preintegration complex (PIC). PICs translocate from the cytoplasm into the nucleus where nascent DNA integrate into the host cell chromosome forming the provirus. mRNA produced from the provirus serve as transcripts for the production of viral proteins or genomes for progeny virus. Viral proteins accumulate at the plasma membrane and assemble into new virions. Newly formed virions leave the cell via budding.

1.4.2 Entry

HIV-1 entry is mediated by binding of the trimeric envelope glycoprotein structures (gp120), which protrude from the viral envelope, and cell surface receptors on the target cell. In 1984,

CD4 was identified as the major host cell surface receptor by studies demonstrating that CD4 antibodies could block viral infection and virus-mediated cell fusion (Dalgleish et al., 1984;

Klatzmann et al., 1984; McDougal et al., 1985). The function of CD4 as the major HIV-1 cell receptor was confirmed by transfection of CD4 negative cells (usually resistant to HIV-1 infection) with CD4, rendering them susceptible to HIV-1 infection (Maddon et al., 1986).

HIV-1 virion binding also involves a co-receptor on the target cell. The identity of the co- receptor is dependent on Env (gp120) and determines the cell tropism of individual viral isolates. The major co-receptors for CD4+ T lymphocytes and primary macrophage are the chemokine receptors CXCR4 and CCR5.

15

Figure 1.5 The HIV-1 life cycle Following receptor mediated entry the viral core enters the host cell cytoplasm, CA protein dissociates from the core releasing the RTC, and enables efficient reverse transcription. dsDNA integrates into the host chromosome in the nucleus to form the provirus. Proviral DNA is transcribed and translated by host proteins. Assembly of new virus particles occurs at the plasma membrane where virions exit the cell via budding. Furtado et al, NEJM, 1999

The heteromeric GTP-binding protein (G protein)-coupled receptor, fusin, later designated

CXCR4, was identified as an HIV-1 co-receptor in T lymphocytes by screening cDNA expression libraries (Feng et al., 1996). Expression of both recombinant CD4 and CXCR4 on non-human cells, but not CD4 or CXCR4 alone, allowed fusion with cells expressing Env, confirming the requirement of the CXCR4 receptor (Feng et al., 1996).

Likewise, suppression of HIV-1 infection in the presence of chemokines RANTES, MIP-1α and MIP-1β, suggested an alternative chemokine receptor was required for viral entry by certain isolates. Cells expressing CD4 with or without the G-protein-coupled receptor C-C chemokine receptor 5 (CCR5) demonstrated the requirement of the CCR5 receptor for infection by these viral isolates (Alkhatib et al., 1996a; Alkhatib et al., 1996b).

Initial interactions between the HIV gp120 glycoprotein and CD4 cell surface receptor induce conformational changes in gp120 exposing the chemokine receptor binding surface. This allows binding of the relevant co-receptor (Safai et al., 1984; Sattentau, 1992; Sattentau &

Moore, 1993), and further conformational changes expose the fusion domain of the transmembrane gp41 protein. gp41 mediates fusion between the lipid membrane of the viral envelope and plasma membrane of host cells, priming the virion for release of the viral core

(Marcon & Sodroski, 1994).

1.4.3 Core disassembly

The HIV-1 CA shell that surrounds the internal ribonucleoprotein complex defining the viral core must disassemble to allow complete reverse transcription and nuclear translocation

(Arhel et al., 2007). Initially it was presumed all CA spontaneously dissociated from the HIV core upon entry into the host cell cytoplasm (Grewe et al., 1990). Further studies have suggested that during release of CA from the viral core, while the typical cone shaped core is lost, a proportion of CA remains associated with the RTC/PIC where it helps to facilitate nuclear entry (Iordanskiy et al., 2006; McDonald et al., 2002). Disassembly of the conical CA shell is highly regulated, albeit by an unknown mechanism (discussed in more detail in section 1.6) (Auewarakul et al., 2005; Forshey et al., 2002).

16

Conditions used to enhance intravirion reverse transcription in vitro, also induce the concurrent release of viral core proteins (CA, NC, RT). The similarity to disassembly of the viral core suggests these viral processes are intrinsically linked (Warrilow et al., 2008; Zhang et al., 2000). While natural endogenous reverse transcription (NERT) reactions can occur in vitro without the use of detergent to permeabilise the virion, these reactions are very limited and incomplete. Complete reverse transcription only occurs in the cytoplasm of infected cells.

The implications of this are two-fold. At first it suggests uncoating of the core is unnecessary for initiation of reverse transcription. However, at the same time is suggests complete reverse transcription is linked to activation of uncoating of the core. Uncoating assays demonstrate that p24 CA is detected in the cytoplasm following entry and released from the core 1-4 hr post-fusion (Hulme et al., 2011; Yamashita et al., 2007). Inhibition of reverse transcription with an RT inhibitor prevents uncoating of the core in this assay (Hulme et al., 2011).

Whether uncoating activates reverse transcription, or the contrary, i.e. reverse transcription induces uncoating, is still a topic of debate. Research is yet to uncover what activates and regulates uncoating of the core.

1.4.4 Reverse transcription

Reverse transcription of the HIV positive sense RNA genome into dsDNA is mediated by the virally encoded reverse transcriptase (RT) enzyme. The active RT heterodimer (p66/p51) possesses RNA-dependent and DNA-dependent DNA polymerase activity (Poch et al., 1989).

RT facilitates synthesis of viral DNA from genomic RNA or newly transcribed DNA templates. Additionally, RT possesses RNase H activity, mediating degradation of the genomic RNA template in the RNA/DNA duplex to allow synthesis of dsDNA (Hansen et al.,

1987).

Reverse transcription takes place in the host cell cytoplasm in a complex defined as the reverse transcription complex (RTC). Immunoprecipitation studies demonstrate that the RTC contains the viral enzymatic proteins RT, IN, PR, small amounts of MA and CA, Vpr and cellular histones, complexed with the viral RNA genome and newly synthesised DNA

17

(Bukrinsky et al., 1993; Fassati & Goff, 2001; Karageorgos et al., 1993; Nermut & Fassati,

2003).

Initiation of HIV DNA synthesis requires a tRNALys3 primer provided by the host cell. The tRNALys3 primer attaches to an 18nt primer binding site located immediately 3’ of the U5 region on the HIV-1 RNA genome (Figure 1.6). The first viral DNA transcript produced is minus strand strong stop DNA, a 181nt DNA intermediate beginning from the tRNALys3 mediated initiation site and terminating at the 5’ end of the viral genome. Following its synthesis, the strong stop DNA intermediate is displaced and then hybridises to the inverted repeat (R) sequence at the 3’ end of the viral genome. Here it serves as a primer for completion of negative strand DNA synthesis. Concomitant with DNA synthesis, RNase H activity mediates degradation of the RNA template. Positive strand synthesis is initiated at two distinct polypurine tracts (PPTs). PPTs serve as RNA primers and are protected from

RNase H degradation. Displacement of the 3’ positive strand intermediate by the growing plus strand, located further upstream, results in a second strand transfer to the 5’ end of the genome. When the remainder of the plus and minus strands are synthesised, they form a complete dsDNA genome containing duplicated U3 and U5 sequences in the LTRs at either end of the molecule (Karageorgos et al., 1995; Li et al., 1993).

A lack of 3’-5’ exonuclease proof-reading activity, in combination with illegitimate template switches, results in high rates of DNA mutation (estimated rate of 3 x 10-5 per replication cycle in vivo). Factors such as these contribute to the vast heterogeneity of viral isolates and generation of drug resistant mutants in HIV-1 infected patients undergoing anti-retroviral treatment.

Although all the proteins which are known to constitute the RTC are present within the viral core, reverse transcription within intact virions is limited. NERT reactions containing salts and physiological polyamines are able to stimulate intravirion reverse transcription (Zhang et al., 2000). However, synthesis of late reverse transcription intermediates (second strand transfer and full-length minus strand) in artificial ERT reactions is greatly enhanced by

18

Figure 1.6 Reverse transcription HIV reverse transcription initiates when a primer provided by the host cell binds the primer binding site (PBS). This generates the first intermediate PBS-U5. Strand displacement and RNaseH mediated degradation of the RNA genomic template are required for plus strand synthesis. A second strand transfer is required for completion of viral DNA synthesis.

RNA and DNA are denoted by thin and bold black lines respectively. (R = repeat region, U5 = 5’ untranslated region, U3 = 3’ untranslated region, PPT = polypurine tract, = tRNALys3 primer)

Adapted from Li et al, Virology, 1993 R U5 PBS PPT PPT U3 R 5’ 3’ 3’ R’ U5’

R U5 PBS PPT PPT U3 R 5’ 3’

R’ U5’

R U5 PBS PPT PPT U3 R PBS 5’ 3’ 3’ U3’ R’ U5’

PPT U3 PPT R PBS 5’ 3’ 3’ R U5 PBS U3’ R’ U5’

U3 R U5 PBS U3 R U5 PBS 5’ 3’ 5’ 3’ 3’ 5’ R’ U5’

U3 R U5 PBS 5’ 3’ 3’ 5’ U3’ R’ U5’

U3 R U5 PBS 5’ 3’ 5’ 3’ 3’ 5’ R’ U5’ PBS’ U3 R U5

U3 R U5 PBS PPT PPT U3 R U5 5’ 3’ 3’ 5’ U3’ R’ U5’ PBS’ U3’ R’ U5’ factors present in cell lysates (Warrilow et al., 2008). Nonetheless, ERT is still less efficient than reverse transcription in infected cells. This suggests the involvement of processes occurring during entry, such as disassembly of CA from the viral core, coupled with activating factors found within the cytoplasmic milieu, are necessary for efficient reverse transcription (Warrilow et al., 2008).

1.4.5 Nuclear translocation and integration

In contrast to many retroviruses, lentiviruses are able to infect non-dividing and terminally differentiated cells utilising the nuclear pore to gain access to genomic DNA within the host cell (Bukrinsky et al., 1992).

RTCs associate with microtubule networks in the cell cytoplasm and use dynein motors to traffic replicating complexes towards the nucleus prior to integration (McDonald et al., 2002).

Following the completion of reverse transcription in the cytoplasm of infected cells, many

RTC proteins are released generating the preintegration complex (PIC). The PIC contains

MA, CA, RT, IN, PR, Vpr, viral RNA and DNA, as well as cellular proteins (Iordanskiy et al., 2006; Miller et al., 1997). Nuclear localisation has been linked with viral determinants including IN, MA, the central DNA flap and also CA (discussed in more detail in section 1.6)

(Fassati, 2006).

HIV PICs are actively imported into the nucleus through the nuclear pore complex (NPC)

(Bukrinsky et al., 1992; Katz et al., 2003). The NPC is made up from ~30 different nucleoporins and provides bi-directional transport between the cytoplasm and the nucleus for molecules that are too large to freely diffuse through the pore (Allen et al., 2000; Gorlich &

Mattaj, 1996; Terry et al., 2007). Numerous nucleoporins have been associated with HIV nuclear import and infectivity (Brass et al., 2008; Ebina et al., 2004; Konig et al., 2008; Lee et al., 2010; Matreyek & Engelman, 2011; Woodward et al., 2009; Zhang et al., 2010).

Notably, several genome-wide siRNA screens have identified nuclear pore proteins, such as

Nup358, Nup153, and TNPO3, which are crucial for HIV-1 infectivity (Brass et al., 2008;

Konig et al., 2008; Yeung et al., 2009; Zhou et al., 2011). There is mounting data implicating

19

CA, not only in the PIC, but at the nuclear pore (discussed in more detail in 1.6.6) (Arhel et al., 2007; Dismuke & Aiken, 2006; Krishnan et al., 2010; Lee et al., 2010; Matreyek &

Engelman, 2011; Yamashita & Emerman, 2004). CA mediates interactions with nuclear pore complex (NPC) proteins directing the import pathway utilised in a cell type dependent fashion. These interactions not only affect efficient nuclear entry, but they can also alter the targeting and preference of integration sites (Lee et al., 2010; Schaller et al., 2011). CA/CypA interactions in the target cell may also affect which nuclear import pathways are used

(Matreyek & Engelman, 2011; Schaller et al., 2011).

Once the PIC reaches the nucleus, IN catalyses integration of newly synthesised viral DNA into the host cell chromosome. HIV IN belongs to the protein superfamily of nucleotidyl transferases, a family of enzymes that break or join nucleic acids via the phosphodiester backbone. In the host cell cytoplasm, IN binds to attachment (‘att’) sites located in the U3 and

U5 regions of the HIV LTR (Katzman & Katz, 1999). IN then mediates 3’ end processing of newly synthesised viral DNA by cleaving 2-3nt before a conserved dinucleotide, exposing a

3’OH group (Leavitt et al., 1996; Masuda et al., 1995). Once the PIC reaches the nucleus, IN, assisted by host proteins, mediates strand transfer of viral DNA onto a 5’ PO4 of the phosphate backbone, into the major groove of the host cell DNA (Leavitt et al., 1996; Masuda et al., 1995). IN requires sequences in chromatin to catalyse DNA strand transfer. Although these target sites are limited, HIV-1 IN favours integration into gene-enriched transcriptionally active regions of chromatin (Holman & Coffin, 2005). Integration site preference is influenced by host cell proteins including (LEDGF)/p75 (Ciuffi et al., 2005;

Koh et al., 2013; Marshall et al., 2007; Shun et al., 2007).

Purified IN is capable of mediating 3’ processing and DNA strand transfer utilising recombinant DNA substrates that mimic linear viral DNA transcripts in vitro (Bushman &

Craigie, 1991; Cherepanov et al., 1999). However, cellular factors are required during infection of target cells to stabilise the PIC, nuclear import and targeted access to chromatin.

Integration defines completion of the early phase of the HIV-1 life cycle.

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1.4.6 Transcription, translation, assembly and budding

Once HIV DNA is integrated into the host chromosome forming a transcriptionally active provirus, full-length mRNAs are transcribed by the host RNA polymerase II enzyme. Viral mRNA transcripts are capped at the 5’ end and a poly (A) tail is added at the 3’ termini.

Initially, these transcripts are multiply spliced into 2kb mRNA species by host-cell RNA splicing enzymes. These viral RNA species encode the regulatory proteins Tat, Rev and Nef.

Tat mediates viral transcription of the HIV provirus by dramatically increasing processivity of cellular RNA polymerase II (Ammosova et al., 2003). Tat binds to the transactivation response (TAR) RNA secondary structure element, a stem loop structure on nascent RNA transcripts, and recruits cellular proteins including cdk9 (cyclin dependent protein kinase 9) and cyclinT. cdk9 phosphorylates the RNA polymerase II transcription complex further stimulating elongation. cyclinT enhances the interaction between Tat and TAR, present at the

5’ termini of viral transcripts, stimulating gene expression (Arya et al., 1985).

Viral mRNA export is biphasic. Early stage mRNA export is Rev-independent. Fully spliced viral mRNAs (2kb; Rev, Tat and Nef), accumulate in the nucleus and then exit using cellular mRNA export pathways. Rev protein then mediates later phase export of unspliced (9kb; encoding Gag and Gag-Pol proteins) and singly spliced (4kb; Env, Vif, Vpr and Vpu) mRNAs.

Cellular mRNAs are retained in the nucleus by their interaction with splicing factors and then either rapidly fully spliced and exported to the cytoplasm, or degraded. To overcome this,

Rev-mediated nuclear export utilises the Rev response element (RRE), a secondary structure located within env. The RRE is present in unspliced or singly spliced viral RNA transcripts, but not fully spliced transcripts which lack env (Malim & Cullen, 1993; Malim et al., 1991).

A nuclear export signal located at the C-terminus of Rev interacts with nuclear shuttling proteins to direct nuclear export of Rev dependent mRNAs from the nuclear pore to the cytosol for protein synthesis (Fouts et al., 1997; Malim & Cullen, 1993; Malim et al., 1991).

RRE directed mRNA transport is energy-dependent. As GTP is hydrolysed to GDP, the

21

complex dissociates and Rev is imported back into the nucleus directed by an N-terminal nuclear localisation signal (NLS) (Emerman & Malim, 1998).

The majority of viral protein synthesis, in particular Gag synthesis, occurs on cytosolic ribosomes. Subsequently, Gag precursor proteins are transported to the cell membrane for viral particle assembly. A frame shift occurs in 5-10% of Pr55Gag translation products resulting in the synthesis of Pr160GagPol, the precursor for pol encoded enzymes. PR is autocatalytically cleaved from the GagPol precursor and is responsible for cleavage of the

Gag and GagPol precursors. Gag generates the viral structural proteins, MA, CA, NC and p6.

GagPol generates the viral enzymatic proteins, PR, RT and IN. Env precursors (gp160) are synthesised in the endoplasmic reticulum and golgi (Wyatt & Sodroski, 1998). Cleavage by a cellular protease produces the non-covalently associated (TM-SU)3 trimeric glycoproteins which are transported to the cell membrane for virus assembly.

HIV assembly is directed by the p55Gag precursor protein. p55Gag is capable and sufficient to produce assembling and budding VLPs (Fuller et al., 1997), however other host cell and viral proteins are required for the production of infectious HIV particles.

Virus assembly includes five basic steps: Gag multimerisation, binding of Gag p55 complexes to genomic viral RNA, formation of Gag/GagPol complexes and formation, then transport, of preassembled complexes to the plasma membrane.

Gag multimerisation is required for plasma membrane targeting. This involves two domains within Gag, the NC region and the CA/p2 domain. NC possesses two zinc binding motifs and a large proportion of basic amino acid residues which mediate high affinity for binding RNA.

NC uses the genomic viral RNA as a scaffold, promoting further Gag-Gag association. Gag multimerisation facilitates binding of Gag to the plasma membrane exposing a myristate moiety. Myristoylation of MA within the Gag precursor together with hydrophobic residues and a polybasic region (Arg15-Lys30) in MA regulate the intracellular localisation of the Gag precursor (Bryant & Ratner, 1990; Zhou et al., 1994). This targets Gag and GagPol polyproteins to lipid rafts (membrane domains enriched with cholesterol, glycosphingolipids

22

and sphinomyelin) in the plasma membrane and late endosomal compartments (Brown,

2002). Further Gag precursors are added in a radial fashion, with MA at the external membrane and p6 at the centre, generating new virions which protrude from the cell membrane. Virus maturation occurs as the virion leaves the cell. The viral PR cleaves the viral Gag and GagPol precursors generating the mature structural proteins (MA, CA, NC, p6) and enzymatic proteins (PR, RT, IN).

HIV virions acquire a host cell-derived lipid envelope as they leave the infected cell in a process called “budding”. Once preassembled complexes form, budding is directed by the L domain at the amino terminus of p6 which assists detachment of nascent virions from the cell surface. It has been suggested p6 interacts with cellular ubiquitination and endosomal sorting machinery to facilitate the release of immature virions from the cell.

1.5 The HIV-1 Core

This thesis investigates in vitro properties of HIV-1 core stability and disassembly. Therefore a proper definition of the viral core is important to the understanding of this thesis. The p24

CA protein monomer forms a shell which encapsulates the internal ribonucleoprotein complex. In this thesis this entire structure is termed the viral core. Uncoating or disassembly of the viral core describes the process of rearrangement and release of CA from around the viral core structure thereby revealing the RTC. In this process, while the familiar conical capsid shell may disassemble, a proportion of CA remains associated with the resulting

RTC/PIC complexes.

1.5.1 CA protein

The HIV capsid (CA) protein is initially translated as part of the p55 Gag polyprotein (Figure

1.7). Cleavage by PR generates the 231 amino acid capsid protein, p24 CA (Gag residues

133-363), which forms the conical shell that defines the boundary of the viral core. The CA protein has a molecular weight of approximately 25kDa (Lanman et al., 2004) and is post-

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Figure 1.7 The p55 Gag precursor Cleavage of p55 Gag precursor (A) generates the three major proteins matrix, MA p17 (1-132), capsid, CA p24 (133-363), and nucleocapsid, NC p7 (377- 432); and p6 (449-500), p2 (364-376) and p1 (433-448) spacer peptides. The Gag precursor (B) assembles in radial pattern to form the immature virus (C), upon maturation and cleavage by the viral protease (PR) the Gag precursor is cleaved generating the individual Gag proteins and formation of the conical CA core (D).

Von Schwedler et al, EMBO, 1998 A

B

C D translationally modified by formylation (Fuchigami et al., 2002) and phosphorylation (section

1.7.4.2) (Cartier et al., 1999; Laurent et al., 1989; Mervis et al., 1988; Veronese et al., 1988).

The CA protein monomer is composed of two independently folded domains, the amino terminal domain (NTD, residues 1-146) and the carboxyl terminal domain (CTD, residues

147-231), which are joined by a flexible interdomain linker sequence (Gamble et al., 1996;

Gamble et al., 1997; Gitti et al., 1996; Worthylake et al., 1999). The correct folding and self assembly of the CA protein is critical to the structure and function of CA within the viral core, and consequently HIV virion integrity and infectivity (von Schwedler et al., 2003).

The three dimensional structure of the HIV-1 CA protein has been solved by nuclear magnetic resonance (NMR) (Gitti et al., 1996) and X-ray crystallography (Gamble et al., 1996;

Momany et al., 1996). This showed that the HIV CA protein is primarily helical in nature

(Figure 1.8). The NTD is shaped like an arrowhead, comprising five coiled-coil α-helices, a proline rich region and two short α-helices (Berthet-Colominas et al., 1999; Gamble et al.,

1996; Gamble et al., 1997; Ganser-Pornillos et al., 2007; Gitti et al., 1996; Momany et al.,

1996). The CTD domain is globular and comprises a 310 helix, followed by an extended strand and four α-helices connected by short loops (Berthet-Colominas et al., 1999; Gamble et al., 1997; Ganser-Pornillos et al., 2007; Worthylake et al., 1999).

All retroviral CA sequences (except spumaviruses) contain a unique 20 amino acid stretch

(residues 153-172) in the CTD designated the major homology region (MHR). This region is conserved across lentiviruses, oncoviruses, and is also found in yeast retrotransposon TY-3.

The MHR is essential for viral replication, regulating particle formation and morphogenesis of the mature cone shaped core (Dorfman et al., 1994; Mammano et al., 1994). Within the

CTD, the MHR forms a compact helix-turn-strand that packs against helix 9. The MHR contributes to an intricate series of hydrogen bonding networks between helices 1 and 2.

Mutation of the residues that forming these bonds disrupts this highly ordered structure and the folding that is essential for the function of the MHR. While structural modelling suggests that the MHR does not make any important intermolecular contacts, the high degree of

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Figure 1.8 Ribbon diagram of the crystal structure of the full-length HIV-1 CA monomer (PDB entry 1e6j) CA amino-terminal domain (NTD) and carboxyl terminal domains (CTD) are indicated. α-helices are numbered 1-7 (NTD) and 8-11 (CTD). Figure drawn using PyMol version 1.0r1. PDB entry 1e6j Monaco-Malbet, Structure, 2000 1

2 9 N-terminus 3

7 8 6 5 11 4 10

CypA loop C-terminus

NTD CTD conservation suggests this region may interact with cellular factors or other molecules (Li et al., 2000). Additionally, it has been speculated that the MHR may be involved in genome organisation or organisation of the RTC (Mammano et al., 1994).

1.5.2 The HIV-1 Core

The HIV-1 capsid (CA) protein shares limited sequence homology with other retroviruses

(e.g. HIV-2, MLV, SIV, FIV, and EIAV), yet despite this, retroviral capsid proteins share similar tertiary structure. HIV-1 and MLV capsid proteins demonstrate high levels of structural homology, exhibiting similar protein folding and lattice formation within the retroviral core (Li et al., 2000). Although retroviruses exhibit a variety of core formations, lentivirus cores are predominantly cone shaped.

Approximately 1000-1500 CA monomers self assemble to form the protein shell that encapsulates the internal viral ribonucleoprotein complex, defining the boundary of the viral core (Briggs et al., 2004; Briggs et al., 2003).

The mature HIV core is a conical or fullerene-like structure that is capped at both ends

(Briggs et al., 2003). The CA shell that frames the viral core is composed of a hexameric lattice network. Initially, six CA monomers bind forming hexameric rings with an exterior diameter of approximately 100 Å or 9.6 nm (Briggs et al., 2003; Li et al., 2000)). The NTD-

NTD dimerisation interface at the centre of the hexamer is formed through binding of helices

1, 2 and 3 generating 18-helix (hexamer) or 15-helix (pentamer) bundles. CTD dimerisation mediated by helix 9 (Gamble et al., 1997; Worthylake et al., 1999; Yoo et al., 1997) links adjacent hexamers (Ganser-Pornillos et al., 2007; Li et al., 2000) generating a hexameric lattice network (Figure 1.9). Intra-hexameric contacts are further stabilised by helix 4 from the NTD of one molecule with the groove formed by helix 11 in the CTD of its neighbouring

CA molecule (Briggs & Krausslich, 2011). Twelve pentameric defects are required within the hexameric lattice to close the conical viral core, five at the narrow end and seven at the wide end of the core (Ganser-Pornillos et al., 2007; Ganser et al., 1999). These pentameric defects

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Figure 1.9 CA forms a hexameric lattice network to form the core Six CA monomers (indicated by different colours) bind generating hexameric rings which form a hexameric lattice. Approximately 1500-2000 CA monomers are required to produce the fullerene-like or cone shaped core.

Ganser-Pornillos and Yeager, 2007 www.scripps.edu/newsandviews/e 20071015/hiv.html A B are thought to be controlled by an electrostatic switch involving Arg18 (Briggs & Krausslich,

2011).

Stoichiometric data suggests that greater than 5000 Gag protein molecules are present per virion (Briggs et al., 2004), implying that not all the CA protein within the virion participates in formation of the core. Supporting this, hydrogen deuterium exchange analysis of CA from immature and mature virus-like particles (VLP) demonstrates two distinct populations of CA

(Lanman et al., 2004). CA displays varying affinities for different lipids found in the viral envelope, prompting suggestions that a proportion of CA may become embedded in the internal side of the viral envelope rather than contributing to core formation (Barrera et al.,

2008).

Early EM studies suggested the narrow end of viral core was linked to the viral envelope, designated the core-envelope linkage (CEL) (Hoglund et al., 1992; Nilsson et al., 1992;

Zhang et al., 2000). However subsequent studies demonstrated a 12 nm gap between the outer

CA shell of the viral core and the viral envelope that is maintained by steric hindrance between the MA and CA proteins (Benjamin et al., 2005; Briggs et al., 2006).

Cores have been successfully isolated from many viruses. However, mature HIV cores are intrinsically unstable and extremely detergent sensitive. Therefore viral core isolation methods that have been successful for viruses such as AMV (Stromberg, 1972) are much too harsh for HIV, causing the majority of input virus to be lost. However, modification of these methods has allowed isolation of viral core structures. This technique combines sucrose gradient ultracentrifugation with the ‘spin-thru’ technique; virions are briefly exposed to detergent to remove the outer viral envelope prior to sedimentation of viral core particles.

Isolation of authentic viral cores derived from HIV virions using this technique demonstrates a vast array of shapes and sizes (Briggs et al., 2003; Welker et al., 2000). The dimensions of

HIV-1 cores analysed by negative staining EM demonstrated an average length of 103 nm, diameter of 52 nm with an internal angle of 21.3º (Welker et al., 2000). CryoEM produces slightly larger measurements, with an average length of 119 nm, 61 nm in width and similar

26

angle 22.3º (Briggs et al., 2003), perhaps due to less dehydration of samples using this method. Vast heterogeneity of viral cores was observed using both methods. The majority

(63%) of HIV-1 cores visualised are cone-shaped (Briggs et al., 2006). However, cylinders and aberrant particles are often observed. This is likely to reflect the low infectivity of HIV-1 preparations. p24 CA is the major protein present in HIV core preparations. In addition to CA, isolation of

HIV-1 viral cores has demonstrated that MA, NC, RT, IN, Vpr, PR, Vif and the cleaved fragment of Nef are found within the viral core (Accola et al., 2000; Forshey & Aiken, 2003;

Welker et al., 2000). Some groups detected Gag intermediates, which could potentially be explained by the greater relative stability exhibited by immature HIV cores versus mature cores, and the large proportion of immature virus often found in viral preparations (Welker et al., 2000).

The viral core must contain all the necessary viral enzymes for the virus to carry out reverse transcription and integration of the viral genome. The majority of the proteins known to form the viral core are present in the same location within the core as within mature virus. Yet although large amounts of MA have been reported in other retroviral core preparations (SIV,

EIAV, MMTV, AMV, FeLV), only low levels of MA are detected in HIV cores (Accola et al., 2000; Welker et al., 2000). The minor amount of MA remaining may assist trafficking of the RTC through the cytoplasm.

Unexpectedly, p6 and CypA are not found within the HIV core (Accola et al., 2000; Kondo &

Gottlinger, 1996). The radial organisation of Gag proteins within the virion dictates that p6 would be in the centre of virion and hence form part of the core. Additionally, the association with p6 mediates the incorporation of Vpr into the virion, which remains associated with the core following entry (Accola et al., 2000; Kondo & Gottlinger, 1996). The corresponding p9 protein of EIAV is also found outside the core (Roberts & Oroszlan, 1989).

Similarly, although CypA is incorporated into HIV-1 virions via its association with CA, virtually no CypA is detected in immature or mature cores (Accola et al., 2000; Welker et al.,

27

2000). Structural studies indicate that the CypA binding loop is situated on the outside of the capsid lattice and therefore CypA may be easily lost from the core during isolation. The absence of these proteins from the viral core could potentially be explained by rearrangement of the proteins which form the core following maturation, or alternatively, a weak association with the viral core proteins. The absence of CypA might be explained by its importance in the target cell rather than the producer cell.

While studies demonstrate the ability to isolate HIV cores (Accola et al., 2000; Forshey et al.,

2002; Welker et al., 2000), it is uncertain if these structures represent viral complexes capable of contributing to productive infection, or represent defective dead-end structures incapable of further replication. Cores isolated from HIV-1 viruses with mutations at charged residues in

CA demonstrate altered properties when compared to wild type (WT) HIV-1 cores (Dismuke

& Aiken, 2006; Forshey et al., 2002). Substitution of charged residues within CA resulted in viral cores displaying altered core stability and core yields. These CA mutant viruses also displayed blocks in replication (Dismuke & Aiken, 2006; Forshey et al., 2002). Regardless of whether isolated cores are replication competent or otherwise, biochemical characterisation of

HIV cores derived from CA mutant viruses demonstrates that proper maintenance of the viral core has major implications on HIV infectivity (see core stability section 1.6.3) (Arhel et al.,

2007; Dismuke & Aiken, 2006; Douglas et al., 2004; Forshey et al., 2002; Scholz et al., 2005;

Tang et al., 2003; Wacharapornin et al., 2007; Warrilow et al., 2008).

1.6 Assembly and Disassembly of the HIV core

1.6.1 Immature virion

Immature HIV virions are morphologically distinct from mature HIV virions (see Figure 1.3).

Immature virus particles are not infectious until proteolytic cleavage (by PR) induces maturation concomitant with structural changes in virion morphology, condensation of the genomic RNA and encapsulation by capsid.

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Core assembly during virus morphogenesis is a carefully regulated process. It is important to consider with respect to creating mutations in HIV-1 CA, that as a region in p55 Gag, it is involved in both assembly and disassembly of the viral core. Therefore mutations in CA may affect either, or both, assembly and disassembly, thereby contributing to altered virus infectivity.

The radial arrangement of the Gag p55 polyprotein in immature particles allows tight association of the MA domain at the N-terminus of the Gag precursor with the inner leaflet of viral membrane, contributing to the stability of the membrane. Inside the viral membrane, the

C terminus of MA is observed as a low density region, situated outside an electron dense ring formed by the CA proteins. Together the CA and NC domains form regular layers that contribute to radial density of the immature particles.

PR mediates cleavage of Gag p55 in a sequential manner: p2-NC, MA-CA and p1-p6, NC-p1, then CA-p2 junctions (Pettit et al., 1998). The rate of cleavage at each junction is mediated by the sequence and structure of the processing site, and accessibility by the viral protease (Pettit et al., 1994).

Proteolysis at the first junction, p2-NC, is essential for dimerisation of the genomic RNA.

This facilitates the incorporation of two identical copies of RNA into the virion, and condensation of the viral ribonucleoprotein complex (Shehu-Xhilaga et al., 2001).

Next PR mediates cleavage of the MA-CA and p1-p6 junctions. Proper cleavage is critical; the presence of just four MA residues on the N-terminus of CA promotes the assembly of spheres (reminiscent of immature virions), instead of cylinders (von Schwedler et al., 1998).

Cleavage of the MA/CA junction creates a new CA-CA interface essential for assembly of the conical core. Biochemical studies demonstrate the importance of two regions that make up the

NTD interfaces of CA for immature virion formation. Helices 1 and 2 form the centre of the hexamer. Helices 4 and 7 are required for linking adjacent hexamers. TEM analysis shows mutations in residues on this new CA-CA interface prevent formation of conical cores and also reduces infectivity (von Schwedler et al., 1998).

29

Final cleavage of Gag at the CA-p2 junction is essential for formation of the mature CA lattice and the characteristic cone-shaped shell (de Marco et al., 2010; Pettit et al., 1994;

Wiegers et al., 1998). Cleavage of p2 transforms the form of assembly of recombinant HIV-1

Gag proteins, from immature-like spherical structures to cylinders displaying CA binding similar to mature particles (Gross et al., 2000; Wilk et al., 2001). This suggests p2 cleavage may act as a switch for virion morphogenesis.

1.6.2 Core disassembly following viral entry

HIV core disassembly is the process of rearrangement and release of the p24 CA shell that surrounds the viral ribonucleoprotein complex. This process reveals the RTC within the host cell cytoplasm following viral entry (Figure 1.10). During disassembly of the core, the familiar conical capsid shell disappears, but some CA remains associated with the RTC/PIC.

The mechanisms that regulate viral core disassembly are unclear. Initially it was proposed that the core undergoes spontaneous dissociation following entry into the target cell, due to the inability to detect significant amounts of CA in replicating complexes extracted shortly after infection (Fassati & Goff, 2001; Miller et al., 1997). Other studies suggested detachment of the core-envelope linkage (CEL) during entry may promote core disassembly (Zhang et al.,

2000). However, the observation of a 12 nm gap between the core and the viral envelope indicate this is unlikely to regulate disassembly of the core (Briggs et al., 2006).

In vitro analysis suggests HIV-1 core disassembly is biphasic. Initially CA was released rapidly from isolated cores following exposure to 37ºC. After 1 hr approximately 55% of CA was released, yet after 2½ hr, only 80% of CA was released (Forshey et al., 2002). Incomplete dissociation of CA from the core is supported by the observation that a proportion of CA protein remains associated with the RTC (McDonald et al., 2002). Supporting this, an uncoating assay revealed 62.6% of viral complexes contained CA in the cytoplasm 1 hr post- fusion. The amount of CA associated with these viral complexes decreased progressively, and by 4 hr post-fusion only 33.3% of viral complexes contained CA (Hulme et al., 2011).

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Figure 1.10 HIV core disassembly Following receptor mediated entry, the HIV core is released into the host cell cytosol. The core undergoes a regulated dissociation of CA monomers. This complex undergoes restructuring to generates the viral reverse transcription (RTC) and preintegration complexes (PIC).

Adapted from Bukrinsky, Molecular Medicine, 2004

The interaction between multimeric, but not monomeric, CA and the retroviral restriction factor TRIM5α indicates that CA does not dissociate from the core immediately upon virus entry, and that CA interacts with host cell factors in the cytoplasm (Forshey et al., 2005;

Sebastian & Luban, 2005; Shi & Aiken, 2006; Stremlau et al., 2006; Yang et al., 2013).

Mutational analysis of the CA protein demonstrates the requirement for core disassembly for infectivity, and that the proper regulation of core disassembly is essential (Forshey et al.,

2002). Reduced ability of cores to disassemble is associated with blocks during replication, while increased rates of core disassembly also result in non-infectious virus (Dismuke &

Aiken, 2006; Forshey et al., 2002). This indicates core disassembly is a highly regulated and critical step in the HIV-1 life cycle.

1.6.3 Core stability and core yield

Disassembly of the HIV-1 core following virus entry has proven a difficult stage of the virus life cycle to investigate. The in vitro stability of the HIV-1 core can be measured to predict defects in viral core disassembly.

Immature virus cores are relatively stable in the presence of detergent, while mature HIV-1 cores are detergent sensitive (Welker et al., 2000). Core stability is measured by isolating virus cores following a brief exposure to detergent (spin-thru method). Measuring the resulting free (dissociated) versus complex-associated CA (cores) gives an indication as to the extent of disassembly. Subjecting cores to various biochemical treatments (e.g. temperature, pH, salt ion concentrations, detergent or a range of cellular factors) can also provide an indication of the relative core stability (Auewarakul et al., 2005; Forshey et al., 2002;

Warrilow et al., 2008). The spin-thru method, first described for isolation of AMV cores

(Stromberg, 1972), utilises Triton-X-100 detergent overlayed on a sucrose gradient or cushion. Purified virus is sedimented at high speeds through the detergent into the gradient.

The amphipathic nature of both the detergent and viral lipid envelope encourages hydrophobic interactions and removal of the viral envelope. Utilising the detergent in the

31

spin-thru layer dramatically increases the recovery of cores compared with similar methods involving incubation of cores in detergent prior to ultracentrifugation (Yu et al., 1993).

In vitro studies suggest HIV-1 cores disassemble rapidly at 37ºC, but remain relatively intact between 4ºC and 25ºC. Core stability is reduced at low pH, but cores are stabilised at high pH.

Increases in ionic strength are also reported to promote disintegration of the core (Forshey et al., 2002).

Mutational analysis of the CA protein has demonstrated that viruses containing cores which display altered cores properties in comparison to WT, including core stability and core yield, are defective during reverse transcription or subsequent nuclear translocation (Dismuke &

Aiken, 2006; Forshey et al., 2002). Forshey et al (2002) showed that substitution of charged residues across CA (helices 1, 2, 4, 7, 8, 10 and 11) altered core stability, suggesting that regulation of charge within the core could play a role in core stability, and potentially in regulation of core disassembly (Forshey et al., 2002). This indicates that bonding throughout the core is critical for proper maintenance of the core structure, and argues that selective maintenance of the core, indicating a delicate balance between stability and instability is essential for viral replication.

The involvement of viral proteins, other than CA, in core stability has not been fully elucidated. The viral proteins Nef and Vif are detected within the core (Forshey & Aiken,

2003; Kotov et al., 1999; Liu et al., 1995), but Nef does not appear to affect the stability of the HIV core (Forshey & Aiken, 2003). In contrast, Vif has been suggested to increase the stability of WT cores in the presence of high pH (pH9), cytosolic cell fractions (cytosol S100) and dNTPs, when compared to cores from Vif-defective virus (Ohagen & Gabuzda, 2000). In addition, the stability of isolated cores is compromised in several virus matrix (MA) protein mutants, which replicate poorly in primary cells and are impaired in single cycle infectivity assays (Davis & Aiken, 2004). Likewise, several IN mutants show decreased core stability.

These IN mutants were also associated with a decrease in intravirion CypA, and it was

32

hypothesised that IN effects disassembly by assisting interactions between CA and CypA

(Briones et al., 2010).

Therefore, while many factors have been shown to alter core disassembly, what regulates core stability is not understood. However, proper stability of the core is essential for virus infectivity.

1.6.4 Cellular factors, restriction factors and core disassembly

The regulation of core disassembly and altered infectivity has been associated with several host cell factors including unidentified cell components, heat shock protein 70 (Hsp70), and peptidyl-prolyl isomerases (PPIases) Pin1 and cyclophilin A (CypA). Additionally, the restriction factor Trim5alpha and its association with CA & CypA provide some interesting insights into uncoating of the core.

Auewarakul et al (2005) showed that a cellular factor which is required for HIV-1 core disassembly is found in activated CD4+ lymphocytes (Auewarakul et al., 2005). Lysates of activated (HLA-DR+) cells, but not quiescent (HLA-DR-) cells, were able to induce dissociation of CA from the core to a similar extent as a p24 CA disruption buffer. This was in contrast to the limited dissociation induced by a hypotonic lysis buffer. Fractionation following gel filtration on a fast phase liquid chromatography (FPLC) system demonstrated the majority of CA protein was released as monomeric CA, with a molecular mass close to

25kDa (Auewarakul et al., 2005). This observation lead to suggestions that either a factor required for core disassembly was present in activated cells, or that a factor that inhibited core disassembly was present in quiescent cells (Auewarakul et al., 2005). Unsorted cells containing a greater number of quiescent cells still maintained activity capable of dissociating

CA from the core, supporting the proposal of a positively acting factor in activated cells

(Auewarakul et al., 2005).

Quiescent cells and cells blocked during cell division do not support HIV infection (Zack et al., 1990). Consistent with these findings, full-length reverse transcription intermediates were isolated from cores incubated in lysates from activated but not quiescent cells (Auewarakul et

33

al., 2005). Furthermore, lysates from cells arrested in G0-G1a could not induce dissociation of CA from the core in uncoating assays. However, cells arrested in G1b could induce uncoating. This suggests one or more factors required for core disassembly are cell cycle dependent (Auewarakul et al., 2005). Progression through the cell cycle to G1b is also required for completion of reverse transcription (Auewarakul et al., 2005; Korin & Zack,

1998).

In efforts to identify a potential cell factor displaying uncoating activity activity, fractionation of H9 cell lysates was performed by gel filtration on an FPLC system. Uncoating assays on the resulting fractions demonstrated major and minor peaks of uncoating activity corresponded to proteins with apparent molecular masses of 60kDa and 160kDa (Auewarakul et al., 2005). However, the cellular factor responsible has not been identified. Several cellular proteins that have been proposed to effect core disassembly are discussed below.

1.6.4.1 Heat Shock Protein 70 (HSP70)

Hsp70 is incorporated into the virion and was proposed to play a role in core disassembly analogous to that for clatherin coating cages (Gurer et al., 2002; Gurer et al., 2005). Inhibition of Hsp70 by ATPγS reduced infectivity and blocked reverse transcription. However, this was attributed to a defect in core morphology rather than core disassembly (Gurer et al., 2005).

1.6.4.2 Peptidyl-prolyl isomerase NIMA-interacting 1 (Pin1)

Misumi et al (2010) proposed that the prolyl isomerase Pin1 is required for uncoating

(Misumi et al., 2010). Pin1 recognises phosphorylated serine/threonine-proline motifs. There is a single target site within HIV-1 CA, Ser16-Pro17. Mutation to S16A/P17A alters infectivity, resulting in decreased, or inhibition of replication in a cell type dependent manner.

Substitution of S16E shows WT infectivity, while S16A or S16T show reduced infectivity.

This indicates that phosphorylation or negative charge is required at this site. Pin1 knockdown, heat inactivation and mutation of Pin1 prevent HIV infection and increase pelletable CA in a cell-based fate-of-capsid assay, further supporting a role for Pin1 in uncoating or early infection (Misumi et al., 2010). 34

However, it is noted that substitution of S16A by Cartier et al (1999) did not affect either CA phosphorylation in mature virus, or replication by this virus (Cartier et al., 1999), therefore it was not examined in the present study.

1.6.4.3 Peptidyl-prolyl isomerase A (CyclophilinA, CypA)

The host cell protein CypA is a peptidyl-prolyl isomerase involved in protein folding (Braaten et al., 1996; Endrich et al., 1999; Gamble et al., 1996; Grattinger et al., 1999). CypA binds the CA protein in both immature and mature virions (Endrich et al., 1999) and is required for virus infectivity. Structural modelling indicates that CypA binds a loop on the exterior of the

CA protein monomer away from intermolecular CA interfaces (G89-P90). Isomerisation has been suggested to play a role in uncoating of the core (Bosco et al., 2002; Bosco & Kern,

2004; Braaten et al., 1996; Gitti et al., 1996). However, in vitro assembly assays demonstrated CypA did not destabilise the mature core (Grattinger et al., 1999; Li et al.,

2000; Wiegers et al., 1999). As such, CypA was deemed unlikely to affect disassembly.

Furthermore, these in vitro assembly assays suggested CypA aided assembly, possibly by acting as a chaperone.

However, it has become apparent that binding of CypA in the target cell may indeed impact on core disassembly. CypA appears to regulate uncoating in a dose dependent and cell type dependent manner (Gamble et al., 1996; Grattinger et al., 1999; Li et al., 2009; Shah et al.,

2013 ; Wiegers et al., 1999). It is hypothesised that CypA binds the incoming capsid preventing uncoating, thereby allowing efficient reverse transcription in the cytoplasm or during transport to the nucleus (Shah et al., 2013). CypA binds progressively to the core, and once it reaches a critical concentration, CA subunits separate and the core begins to collapse

(Fassati, 2012). CypA association with dynein protein complexes along microtubules might help the viral complex traffic towards the nucleus (Galigniana et al., 2004). CA-CypA engages the NPC protein TNPO3, which can stimulate core disassembly in vitro, but is thought to exert its role removing excess CA from the PIC in the nucleus (Shah et al., 2013;

Zhou et al., 2011). CA-CypA interactions appear to influence CA interactions with NPC

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proteins and potentially the choice of which nuclear transport pathway is utilised (Lee et al.,

2010; Matreyek & Engelman, 2011; Schaller et al., 2011; Shah et al., 2013; Zhou et al.,

2011). A role for CypA is further supported by two IN mutant viruses. Cores from these viruses display accelerated uncoating, but decreased levels of intravirion CypA. Similarly, cores isolated from virus depleted of CypA also display unstable cores. It is hypothesised IN acts to maintain CA-CypA interactions that contribute to optimal core stability (Briones &

Chow, 2010; Briones et al., 2010).

1.6.4.4 Tripartite motif-containing 5 (Trim5α)

The association of HIV CA with the cellular TRIM5 proteins provides important insights into

HIV core disassembly and replication.

TRIM5 analogues, TRIM5-Cyp and TRIM5alpha, restrict HIV infection in non-human primate cells (Sayah et al., 2004; Stremlau et al., 2004). TRIM proteins are comprised of three domains: the N terminal RING domain, B-box and coiled-coil domain. These domains are conserved across all TRIM family proteins (Ganser-Pornillos et al., 2011). The B-box and coiled-coil domains promote multimerisation required for restriction activity (Diaz-Griffero et al., 2009; Li & Sodroski, 2008), while the RING domain displays E3 Ubiquitin ligase activity

(Xu et al., 2003).

The owl monkey orthologue TRIM5-Cyp targets CA via a C-terminal Cyp domain (Nisole et al., 2004; Sayah et al., 2004). While the rhesus macaque orthologue TRIM5alpha interacts with CA via its C-terminal PRY-SPRY domain (Stremlau et al., 2004). CA-CypA binding is required for HIV-1 restriction by Old World monkey orthologues of TRIM5 (Berthoux et al.,

2005). Knockdown of CypA or TRIM5, or treatment of rhesus cells with CsA results in increased infectivity, indicating these factors are likely to be involved in the same pathway

(Berthoux et al., 2005).

The capsid lattice is recognised by the active TRIM5 multimer rapidly following release of the core into the cytoplasm (Battivelli et al., 2013; Sebastian & Luban, 2005; Stremlau et al.,

2006). TRIM5 dimerises and forms a hexagonal structure on the viral capsid. Spacing and

36

symmetry analogous to hexagonal CA structures aids pattern recognition (Ganser-Pornillos et al., 2011; Pornillos et al., 2011). Ultimately, retroviral restriction results from premature uncoating of the core. This is associated with degradation of CA, abrogation of reverse transcription and loss of infectivity (Black & Aiken., 2010; Chatterji et al., 2006; Ganser-

Pornillos et al., 2011; Stremlau et al., 2006).

Sucrose gradient sedimentation of viral complexes from TRIM5 restricted cell indicates loss of CA and IN, decreased viral RNA and reverse transcription intermediates. In the absence of

TRIM5 restriction, inhibition of reverse transcription does not affect CA and IN in these complexes. However, inhibition of reverse transcription in combination with TRIM5 restriction results in the loss of viral RNA. Although the association of CA, IN, viral RNA and reverse transcription intermediates are restored in the presence of the proteosome inhibitor MG132, infection is not restored. This suggests TRIM5 restriction results from modification of the core which allows degradation of IN and viral RNA (Kutluay et al.,

2013).

Trim5 restriction of HIV infection is saturated by high levels of input virus or virus-like particles. One interesting insight, is that viruses displaying altered core stability, also exhibit differences in the ability to saturate TRIM5 restriction (Forshey et al., 2005). This suggests that in addition to the specific binding interfaces which mediate this interaction, the overall stability of the core is able to influence interactions with cellular factors in the cytoplasm.

Human TRIM5 however, which potently restricts EIAV, FIV and N-tropic MLV, exhibits weak anti-HIV activity. This species specificity is explained by polymorphisms in the capsid recognition domain (Luban, 2007; Sawyer et al., 2005; Stremlau et al., 2005; Yap et al.,

2005). Mutation of several CA residues renders HIV susceptible to human TRIM5alpha. This effect is compounded by preventing interactions between CA and CypA (Maillard et al.,

2011). It has been hypothesised that CA:CypA prevents interactions with putative restriction factors in human cells that could promote premature uncoating, and restrict viral infection

(Sayah & Luban, 2004; Towers et al., 2003).

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The involvement of CA with multiple cellular factors suggests HIV-1 core disassembly is an active multistep process.

1.6.5 Capsid (CA) and reverse transcription

Many studies imply an important relationship between core disassembly and reverse transcription, however what this link is, and how it is maintained has yet to be fully elucidated.

Reverse transcription is capable of initiating inside intact virions (Lori et al., 1992; Trono,

1992). However, the majority of reverse transcription occurs within the cytoplasm of infected cells. Within 2 hr following infection RTCs are detected at the perinuclear region at the microtubule-organising centre (MTOC) (McDonald et al., 2002). Following this, the RTC translocates into the nucleus to allow integration of the provirus.

The presence of CA in RTCs and PICs was controversial for a long time. While many biochemical analyses of viral complexes were unable to detect CA (Fassati & Goff, 2001;

Karageorgos et al., 1993), many of the methods used relatively high concentrations of detergents in either lysis buffers or during sucrose gradient sedimentation. These factors may have detrimental effects on the integrity and retention of CA within these complexes.

Subsequently, several studies detected CA in RTCs (Iordanskiy et al., 2006; McDonald et al.,

2002; Miller et al., 1997; Schweitzer et al., 2013). McDonald et al (2002) tracked VSV-G pseudotyped live virus using TEM, and demonstrated that approximately 67% of RTCs trafficking along microtubule networks contained CA. RTCs lacking CA were also visualised migrating along microtubule networks (McDonald et al., 2002). Additionally, Iordanskiy et al

(2006) reported CA in RTCs (Iordanskiy et al., 2006). Miller et al (1997) also reported detection of minor amounts of CA in PIC using low salt conditions, but not in PIC isolated using high salt conditions (Miller et al., 1997). This is consistent with reports that high ionic strength promotes disassembly of CA from the viral core in vitro (Forshey et al., 2002). It is now accepted that CA is retained in viral complexes post-entry and is necessary for nuclear

38

import (Lee et al., 2010; Matreyek & Engelman, 2011; Schaller et al., 2011; Shah et al.,

2013; Zhou et al., 2011).

Although intravirion reverse transcription occurs, the levels of DNA synthesis inside the virion are insignificant compared to high levels of DNA synthesis observed following virus entry into the host cell cytoplasm. This led to the suggestion that factors present in the intracellular cytoplasmic milieu are associated with core disassembly and release of the RTC, and that these are required for activation of reverse transcription and synthesis of full length reverse transcription intermediates. The addition of dNTPs and polyamines enhances reverse transcription in NERT reactions (Zhang et al., 2000) and results in the dissociation of viral core proteins (CA, RT and NC), similar to the effect of treating virions with the cytosolic cell fraction (cytosol S100) (Ohagen & Gabuzda, 2000).

While mutagenesis studies also show that substituting residues within the CA protein can block reverse transcription of viral DNA (Cartier et al., 1999; Dismuke & Aiken, 2006;

Fitzon et al., 2000; Forshey et al., 2002; Tang et al., 2001; Tang et al., 2003). The CA protein has been shown to interact (via CA helix 4) with the host cell factor (lysyl-tRNA synthetase

(LysRS)) (Kovaleski et al., 2007) responsible for incorporating the tRNA (tRNALys3) required for initiation of reverse transcription (Jiang et al., 1993). However, many defects associated with CA core stability mutant viruses occur at late stages during reverse transcription

(Forshey et al., 2002), which is known to be affected by cellular factors (Warrilow et al.,

2008). This suggests that the defect may be in part due to the function of CA in the core and interruption of orderly core disassembly.

It is unclear at what stage during replication dissociation of the majority of CA is necessary, considering not all actively replicating complexes migrating towards the nucleus contain CA

(McDonald et al., 2002). However, alterations in core stability properties which result in aberrant loss or retention of CA can result in blocks during reverse transcription (Dismuke &

Aiken, 2006; Forshey et al., 2002). Likewise, blocks during reverse transcription can prevent

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proper core disassembly (Arhel et al., 2007; Hulme et al., 2011). Both these scenarios result in detrimental effects on infectivity.

This relationship between core disassembly and reverse transcription is supported by studies demonstrating that inhibition of RT activity, through mutation of the active site or inhibition by antivirals, prevents properly regulated uncoating and results in the persistence viral core complexes containing CA (Hulme et al., 2011; Yang et al., 2013).

Intact cores from virus mutants that are arrested at the final stage of reverse transcription

(central flap formation) accumulate at the nuclear membrane longer than WT cores (Arhel et al., 2007). This suggested, in contrast to popular dogma, that the majority of reverse transcription can occur inside an intact capsid, and also that intact cores are capable of docking at the nuclear membrane (Arhel et al., 2007). This also indicated that the absence of complete reverse transcription is associated with increased core stability. Although, whether the intact cores at the nuclear membrane represent actively replicating viral complexes or dead end products is unclear. Additionally, this supports a link between reverse transcription and core disassembly, and indicates that proper regulation of CA disassembly is necessary to permit efficient nuclear translocation (Dismuke & Aiken, 2006; Forshey et al., 2002).

1.6.6 Capsid (CA), nuclear import and integration

The size permitted to cross the nuclear envelope through the nuclear pore complex is 40-45 nm (Pante & Kann, 2002). During HBV replication, intact HBV (~35 nm (Crowther et al., 1994)) are permitted to pass through the nuclear pore. In contrast, the average HIV core is 103 nm in length and 52 nm in diameter (Benjamin et al., 2005; Briggs et al., 2003;

Hoglund et al., 1992; Welker et al., 2000). Passive import of intact HIV cores is restricted because of the size limitations of the nuclear pore. Thus, in the absence of spontaneous disassembly of the core, the observation of intact cores accumulating at the nuclear pore is not completely unexpected (Arhel et al., 2007).

The diameter of the interior ‘hole’ (see Figure 1.9) in HIV-1 helical capsid assemblies is similar in size to the holes in HBV cores (2-3 nm (Crowther et al., 1994; Kenney et al., 1995;

40

Li et al., 2000)). These holes are large enough to allow entry of ATP into the intact core (Hui,

2002). However, Warrilow et al (2008) reported the factor required for efficient synthesis of late reverse transcription intermediates is greater than 3.5 kDa, suggesting restructuring of the core is required for efficient elongation (Warrilow et al., 2008).

Several studies indicate a proportion of CA remains associated with WT PICs (Iordanskiy et al., 2006; Miller et al., 1997; Schweitzer et al., 2013.). However, defects in nuclear localisation in some CA mutant viruses (e.g. Q63A/Q67A) have been attributed to the retention of an unusually high proportion of CA in PICs (Dismuke & Aiken, 2006). This implicated a role for CA, via core stability and the necessity for properly regulated core disassembly, in the successful nuclear import of viral complexes. It is now widely accepted that CA is present in complexes that reach the nuclear pore (Lee et al., 2010; Matreyek &

Engelman, 2011; Schaller et al., 2011; Shah et al., 2013; Zhou et al., 2011).

Several viral proteins, including IN and MA, have been reported to impact the active nuclear transport of HIV replication complexes. Additionally, genome-wide siRNA screens have identified several nuclear pore proteins which are required for HIV replication; e.g.

NUP358/RanBP2, NUP153 and TNPO3 (Brass et al., 2008; Konig et al., 2008; Yeung et al.,

2009; Zhou et al., 2011). The dependence of HIV replication on these host cell factors appears to be mediated by p24 CA.

1.6.6.1 NUP358

Depletion of NUP358 results in diminished HIV replication. The production of late reverse transcription intermediates are unaffected, but the number of 2LTR circles and integrated proviruses are reduced (Schaller et al., 2011). Di Nunzio et al (2012) reported that depletion of Nup358/RanBP2 impairs arrival of the PIC at the nuclear envelope (Di Nunzio et al.).

NUP358 binds CA through a Cyp domain at its C-terminus on the cytoplasmic side of the

NPC (Bichel et al., 2013; Di Nunzio et al., 2012; Fassati, 2012). Crystal structures of the CA

NTD in complex with the Cyp domain of NUP358 reveal CA residue P90 binds a hydrophobic pocket it NUP358 (Bichel et al., 2013). Interestingly, NUP358 binding is

41

influenced by the ability of CA to bind CypA, but is unaffected by cyclosporine A (CsA) treatment (Schaller et al., 2011). Although NUP358 and CypA bind the same region, NUP358 binds a larger interface. Like CypA, NUP358 is an active prolyl isomerase, and has reignited the possibility that isomerisation of CA may act as a trigger for uncoating of the viral core.

NUP358 possesses weaker isomerase activity compared to CypA on synthetic substrates (Lin et al., 2013). However, NUP358 is able to isomerise CA substrates more efficiently than

CypA can (Bichel et al., 2013). Viruses with mutations in CA at G89V and P90A (and therefore cannot bind CypA) are not affected by depletion of NUP358, but these viruses are sensitive to the depletion of the nuclear pore protein TNPO3 (Schaller et al., 2011). These CA mutant viruses also displayed altered integration targeting preference, showing a greater frequency of integration events in regions with a higher density of transcription units

(Schaller et al., 2011). This suggests that in the absence of an interaction with primary NPC protein partners, CA can interact with alternative NPC proteins. The interaction of CA with selected NPC proteins may direct transport through different nuclear import pathways with downstream affects on integration, and ultimately infectivity.

1.6.6.2 NUP153

Similarly, NUP153 depletion also reduces HIV infectivity (Schaller et al., 2011). Further characterisation revealed that while production of late reverse transcription products were unaffected, a moderate reduction in the number of 2LTR circles and a large reduction in the number of integrated provirus were observed (Matreyek & Engelman, 2011). Nup153 interacts with chromatin (Capelson et al., 2010). HIV PICs are thought to exploit this capability when targeting integration sites (Konig et al., 2008; Matreyek & Engelman, 2011;

Woodward et al., 2009). However, comprehensive characterisation of these pathways is still required.

1.6.6.3 TNPO3

TNPO3 is a member of the Importin β family of proteins and promotes the transfer of serine/arginine rich splicing proteins through the nuclear pore (Kataoka et al., 1999). WT HIV 42

requires TNPO3 in both dividing and non-dividing cells (Christ et al., 2008). Viruses with mutations in CA that exhibit WT core stability are dependent on TNPO3 for infection (e.g.

Q4A, G116A, T119A, R143A, P207A).

Interestingly, TNPO3 depletion does not affect several CA mutants viruses which exhibit altered core stability (e.g. K203A, E45A, Q63A/Q67A) (Lee et al., 2010; Shah et al., 2013).

The mutation in CA do not localise to a specific region, suggesting that this interaction doesn’t necessarily effect a binding surface, but rather implies a potential link between intrinsic core stability and the requirement for TNPO3.

TNPO3 primarily localises to the cytoplasm (Shah et al., 2013; Valle-Casuso et al., 2012)

Depletion of TNPO3 inhibits HIV infection, but does not affect the accumulation of proviral

DNA in the nucleus. However, depletion of TNPO3 results in the accumulation of CA protein in the nucleus (Shah et al., 2013; Zhou et al., 2011).

Additionally, TNPO3 knockdown redirects localisation of CPSF6 (an SR protein) that binds to CA to the cytoplasm. Cytoplasmic CPSF6 blocks HIV replication by preventing nuclear import and is associated with stabilisation of the viral core (De Iaco et al., 2013).

Furthermore, purified recombinant TNPO3 stimulates uncoating of HIV cores in vitro, while addition of CypA inhibits uncoating and reduces the effect of TNPO3 (Shah et al., 2013).

Enhanced HIV infection due to CsA treatment of TNPO3 depleted cells suggests the requirement for TNPO3 is regulated by CypA-CA interactions (Shah et al., 2013).

Shah et al (2013) proposed that CypA binds the incoming capsid and prevents premature uncoating of the core. This permits efficient reverse transcription in the cytoplasm and during intracytoplasmic transport towards the nucleus. CA-CypA engages with TNPO3, the complex enters the nucleus, where upon CypA dissociates, allowing TNPO3 to induce uncoating prior to integration (Shah et al., 2013).

This ability to use multiple import pathways may help to explain the cell type dependent patterns of infection for many virus mutants, and the different requirements for successful infection between dividing and non-dividing cells.

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1.7 Protein phosphorylation and HIV

1.7.1 Protein Phosphorylation

Protein phosphorylation is the ATP-dependent transfer of a phosphate group onto the hydroxyl group of serine, threonine and tyrosine amino acid residues. Phosphorylation is mediated by ubiquitous enzymes called kinases. Each kinase mediates phosphorylation within a specific kinase consensus sequence; although they are often able to recognise a minor variant consensus sequence (e.g. casein kinase II recognises the sequence S/T-x-x-E/D).

Phosphorylation can alter the reactivity of protein molecules through the addition of negative charge, and it is instrumental in the control of cell signalling pathways and regulation of protein function.

1.7.2 Virion incorporated kinases

The incorporation of kinases has been reported for numerous DNA and RNA, enveloped and non-enveloped viruses; including HBV, HSV, VSV, , sendai, FMDV, and other poxviruses and adenoviruses.

Viruses such as HSV and VSV incorporate kinases into their tegument region, while other viruses, such as HBV, incorporate a host cell kinase into the nucleocapsid allowing these proteins to carry out different roles. Phosphorylation of the HSV tegument proteins by the virus-associated protein kinase (VAPK) has been suggested to initiate dissociation of the tegument.

The HIV virion has been reported to incorporate several host cell protein kinases, including

ERK2 (Cartier et al., 1997; Jacque et al., 1998), PKA (Cartier et al., 2003), DR1/2 (Devroe et al., 2005), an unidentified 53kDa kinase (Cartier et al., 1999), LCK, NME1 (Ott et al., 2000),

PKC, PGK1, CDC42 and STAT1 (Chertova et al., 2006).

Cartier et al (1997) described a series of phosphorylated proteins in purified HIV virions, two of which showed strong auto-phosphorylation. These proteins demonstrated kinase activity,

44

and phosphoamino acid composition analysis demonstrated these kinases phosphorylate protein substrates at serine and tyrosine residues (Cartier et al., 1997). The first virion incorporated kinase was identified as the extracellular signal-regulated kinase (ERK2) mitogen activated kinase (MAPK). While the second kinase (53kDa) was not initially identified (Cartier et al., 1997), a later study suggests this may represent one of, or both the nuclear Dbf2 related (NDR1 and NDR2) kinases (~54kDa) (Devroe et al., 2005).

Additionally, the catalytic subunit of the cAMP dependent protein kinase (C-PKA) is also detected in HIV-1 virions (Cartier et al., 2003).

While many of the kinases identified in HIV virions have been shown to effect HIV replication upon activation or depletion within the target cell, their specific role within the virion is unclear. The functions of several well described kinases are discussed in relation to

HIV-1 and CA below.

1.7.2.1 Mitogen activated protein kinase (MAPK)

Mitogen activated protein kinase (MAPK) (also known as ERK2) is present in all cell types and is instrumental in the regulation of cell proliferation and differentiation (Yang &

Gabuzda, 1999). MAPK is activated by threonine and tyrosine phosphorylation by the MAPK

ERK2 kinase (MEK). Both MAPK and MEK are activated by extracellular stimuli including mitogens, cytokines and growth factors. Inhibition of MAPK signalling in virus producing cells with specific MEK inhibitors reduced RT activity in culture supernatants by 75-90%.

Constitutive activation of MAPK signalling in virus producing cells resulted in replication competence five times greater than WT virions in a single cycle infectivity assay.

Additionally, factors such as serum and PMA that stimulate MAPK have also been shown to stimulate HIV replication, further implicating MAPK, or MAPK regulated signalling in HIV infectivity (Yang & Gabuzda, 1999).

1.7.2.2 Nuclear Dbf2 related (NDR) kinases

Cartier et al (1997) described a 53kDa auto-phosphorylated protein found inside HIV-1 virions consistent in size with both the NDR1 or NDR2 kinases (both approximately 54kDa) 45

(Cartier et al., 1997). NDR1 is readily detected in subtilisin protease digested and sucrose gradient purified HIV-1 particles (Devroe et al., 2005). NDR1 and NDR2 kinases are incorporated into HIV-1 virions, and are substrates of the HIV-1 PR. NDR1 accumulates in the nucleus regardless of cleavage by HIV-1 PR. Truncation of NDR2 by HIV-1 PR alters its subcellular distribution, redirecting localisation from the nucleus to the cytoplasm (Devroe et al., 2005). While the role of these proteins in HIV-1 replication is unclear, cleavage by PR appears to modify their enzymatic activity. Following cleavage by PR both kinases are capable of autophosphorylation, but are unable to phosphorylate heterologous substrates

(Devroe et al., 2005).

1.7.2.3 cAMP-dependent protein kinase (C-PKA)

The 40kDa catalytic subunit of the cAMP-dependent protein kinase is also detected in subtilisin-treated purified virus (Cartier et al., 2003). While two regulatory subunits usually associated with C-PKA in the cell were not detected in HIV-1 particles, PKA retained enzymatic activity within subtilisin-treated virions. Inhibition of C-PKA activity results in significantly lowered RT activity, and reduced replication competence in a single cycle infectivity assay. Additionally, virus production is delayed, but virions exhibit normal morphology. While no direct role for C-PKA was identified, active recombinant C-PKA can phosphorylate recombinant HIV-1 CA. Notably, C-PKA and CA co-precipitate following immunoprecipitation of p24 CA from H9 cells expressing NL4-3 virus (Cartier et al., 2003).

1.7.3 Phosphorylation of HIV proteins

Phosphorylation plays an integral role throughout the HIV-1 life cycle, regulating the function of many viral and cellular proteins, and normal cellular pathways, often to the advantage of viral replication.

Attachment of virus at the cell surface activates numerous intra-cellular signal transduction pathways mediated by the interactions between gp120 and CD4 and CCR5. Cross-linking of the HIV envelope with CD4+ T cells induces phosphorylation of pyk2, paxillin, LCK, CCR5 and FAK in primary human CD4+ T cells (Cicala et al., 1999). 46

An essential regulatory mechanism employed by the host cell, phosphorylation is thought to play a role regulating the function of nearly all the proteins encoded by the HIV-1 genome.

Some examples are described below.

1. Tyrosine phosphorylation of approximately 1% of HIV-1 MA occurs in the virion during assembly. Subsequently MA becomes associated with the viral IN enzyme in the nucleoprotein complex, where it aids targeting of the PIC to the nucleus (Gallay et al., 1995a;

Gallay et al., 1995b).

2. Phosphorylation of the remaining MA proteins at virus entry is thought to be important for facilitating dissociation of the incoming viral core from the intracellular viral membrane

(Bukrinskaya et al., 1996; Kaushik & Ratner, 2004). Phosphorylation could change the balance of charge on the surface of MA proteins required for release from the envelope

(Kaushik & Ratner, 2004).

3. The HIV p6 protein represents the major phosphoprotein in HIV-1 particles and may have a potential role in budding (Muller et al., 2002).

4. HIV-1 RT, like many viral and cellular polymerases (e.g. adenovirus DNA polymerase and

DNA polymerase α (Donaldson & Gerner, 1987; Ramachandra et al., 1993)), is regulated by phosphorylation. HIV-1 RT is a substrate for in vitro phosphorylation by a number of mammalian kinases (AK, MBPK, CPK, CK-II and PKC) (Idriss et al., 1999). Phosphorylation of recombinant RT by CKII stimulates RT RNA-dependent (Harada et al., 1998) and DNA- dependent DNA polymerase and RNase activities in vitro (Harada et al., 1999).

5. The HIV-1 regulatory protein Rev undergoes serine phosphorylation in the nucleus of infected cells (Hauber et al., 1988; Yang & Gabuzda, 1999). Phosphorylation of Rev induces conformational changes resulting in an enhanced RNA binding state. However, truncation of the C-terminal 25 amino acids, which includes sites of phosphorylation, is dispensable for regulation of viral gene expression (Fouts et al., 1997).

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6. Tat stimulates HIV transcription elongation by interacting with the TAR stem loop structure at the 5’ end of all viral transcripts (Arya et al., 1985). Tat also interacts with the

RNA dependent cellular protein kinase (PKR) (McMillan et al., 1995), with several outcomes. Phosphorylation of Tat by PKR results in greatly enhanced binding to TAR RNA

(Endo-Munoz et al., 2005). Additionally Tat binding to PKR also assists evasion of host anti- viral mechanisms. Normally, PKR binds dsRNA (including the TAR stem loop) and induces phosphorylation of the alpha subunit of eIF2, which results in a block in translation. Tat binding to PKR prevents eIF2 phosphorylation, thereby allowing protein synthesis and viral replication (Endo-Munoz et al., 2005).

7. The HIV-1 Nef protein is essential for high levels of HIV replication and disease progression. Phosphorylation enhances Nef-mediated down regulation of CD4 receptors. A proline rich motif in Nef interacts with several host cell kinases. Binding Lck or MAPK inhibits kinase activity. Alternatively Nef-kinase interactions appear to mediate phosphorylation of the .

8. Vpr is a small accessory protein involved in nuclear import, transcriptional transactivation and growth arrest of cells. Analysis of Vpr in both purified virions and infected cells demonstrates approximately 5% of total virion associated Vpr is phosphorylated (Muller et al., 2002; Muller et al., 2000). Vpr is phosphorylated at three sites, serine residues S79, S94 and S96. Substitution of serine with alanine demonstrates that phosphorylation of S79 is required for cell cycle arrest, while mutation of all three serines attenuates HIV-1 replication in macrophage (Zhou & Ratner, 2000).

9. Vpu is phosphorylated by casein kinase II (Schubert et al., 1992) and is detected in infected cell lysates (Muller et al., 2002).

10. Vif phosphorylation is essential for Vif function and HIV-1 replication. The 23kDa Vif protein is phosphorylated at five sites (Yang & Gabuzda, 1998; Yang et al., 1996), including four residues in the C-terminus, which is known to be critical for membrane association of

Vif. Phosphorylation of Vif at T96 and S144 are critical, as substitution of either amino acid

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results in loss of 90% of Vif function and inhibits HIV-1 replication (Yang & Gabuzda, 1998;

Yang et al., 1996).

These examples show phosphorylation plays a varied, but critical role in regulation of HIV-1 protein function.

1.7.4 CA phosphorylation & potential kinases

1.7.4.1 Phosphorylation of viral capsid proteins

The major viral capsid protein is phosphorylated in many viruses, including both mammalian and plant viruses. Capsid phosphorylation occurs in enveloped (HIV, SIV, HBV, DHBV,

HCV, HSV, rubella virus and M-MuLV) and non-enveloped viruses (HPV, polyomavirus, poliovirus, adenovirus, CaMV and PVA).

Capsid phosphorylation has various roles including promoting DNA synthesis (DHBV) (Kock et al., 2003; Yu & Summers, 1994), nuclear import, RNA encapsidation (HBV) (Kann &

Gerlich, 1994; Kann et al., 1999; Rabe et al., 2003), RNA binding (PVA) (Ivanov et al.,

2003; Ivanov et al., 2001) and potentially virus phosphorylation might aid cell binding activity (polyomavirus) (Anders & Consigli, 1983; Li et al., 1995). Additionally CA phosphorylation is involved in destabilisation of the capsid in poliovirus (Ratka et al., 1989;

Scharli & Koch, 1984).

1.7.4.2 Phosphorylation of the HIV-1 CA protein

Phosphorylation of HIV-1 CA has been demonstrated in infected cells, transfected cells and purified virus (Cartier et al., 1999; Laurent et al., 1989; Mervis et al., 1988; Muller et al.,

32 2002; Veronese et al., 1988). p24 CA was immunoprecipitated from Pi-labelled Gag proteins using pooled sera from AIDS patients, or antibodies directed against Gag proteins

(Mervis et al., 1988; Veronese et al., 1988). Whether CA is phosphorylated while it is part of

Gag (Veronese et al., 1988), or after it has been cleaved to produce the mature p24 CA protein (Mervis et al., 1988; Muller et al., 2002) is uncertain, however several studies suggest

49

CA is phosphorylated before entry into the target cell (Cartier et al., 1999; Swingler et al.,

1997; Veronese et al., 1988)

Phosphorylation is present on approximately 5% of CA molecules within the virion (Muller et al., 2000). Analysis of phosphoamino acid content of the CA protein showed greater than

90% phosphoserine and minor amounts phosphothreonine (Mervis et al., 1988). 2D PAGE analysis has demonstrated four CA isoforms present in purified virus and cells infected with the HIV-1 BRU isolate, with isoelectric points (pI) of 6.8, 6.6, 6.5, and 6.3 (Laurent et al.,

1989). All four species were detected in culture medium, however only the first two species

32 were present in the virion pellet. PO4 labelling showed the 6.6 and 6.3 species were phosphorylated. This was confirmed by bacterial alkaline phosphatase (BAP) treatment, which resulted in a decrease in phosphorylated isoforms (pI 6.6 and 6.3) and an increase in the 6.8 and 6.5 species (Laurent et al., 1989). The presence of two species following BAP treatment suggests a further modification to the CA protein.

HIV-1 CA is phosphorylated by a virion-incorporated kinase. Studies have demonstrated CA is phosphorylated in vitro by the cellular kinase Casein Kinase II (CKII) (Harada et al., 1998;

Ohtsuki et al., 1998), and virion incorporated cAMP-dependent protein kinase (C-PKA)

(Cartier et al., 2003). While the function associated with HIV-1 CA phosphorylation has not been identified, inhibition of C-PKA reduces replication competence in a single cycle infectivity assay, reduces RT activity and delays virus replication kinetics (Cartier et al.,

2003).

Substitution with alanine at each of the nine serine (S) residues of HIV-1 CA identified five

CA mutant viruses (S41A, S109A, S146A, S149A and S178A) that showed altered virus

32 replication (Cartier et al., 1999). Analysis of radioactive phosphate ( Pi) incorporation in virus lysates revealed that three CA mutant viruses (S109A, S149A and S178A) displayed

32 reduced Pi incorporation compared with WT virus, indicating phosphorylation at those sites in mature virions (Cartier et al., 1999). Infection of primary cells (PBMC) and T cell lines

(H9, C8166 and SupT1) with these CA mutant viruses demonstrated that two CA mutants

50

(S109A and S149A) were not infectious in any cells tested, while the third (S178A) was not infectious in any T cell line but exhibited delayed replication in PBMCs, suggesting the importance of phosphorylation at these residues for HIV replication (Cartier et al., 1999). CA mutant S41A virus replicated normally in H9 cells but was delayed in PBMC, while S146A showed the opposite pattern (Cartier et al., 1999). Analysis of reverse transcription intermediates demonstrated a block in reverse transcription following second strand transfer by S109A and S149A CA mutant viruses (Cartier et al., 1999). The S178A CA mutant virus synthesised full length viral cDNA, but not 2-LTR forms indicative of nuclear import (Cartier et al., 1999). In summary, ablating potential phospho-acceptor sites in CA results in various defects during reverse transcription and nuclear translocation.

In contrast, exogenous RT activity, as assessed by measuring the incorporation of [3H]-dTTP into a poly(A)-dT template-primer mix, was not significantly reduced in these CA mutant viruses compared with WT virus. Furthermore, substitution of S109, S149 and S178 with aspartic acid to mimic phosphorylation also results in dramatically reduced production of virus and blocks in replication characterised by severely reduced reverse transcription (Brun et al., 2010). This led to the question whether CA phosphorylation plays a modulating role in reverse transcription either directly or indirectly by affecting viral core disassembly.

1.8 Aims and scope of thesis.

The aims of this thesis are to investigate early stages of HIV infection on a series of CA mutant viruses. Specifically, this thesis examines the in vitro stability of the HIV viral core, core disassembly and reverse transcription of these viruses.

Little is known about the general mechanisms involved in regulating dissociation of the CA protein from the viral core during core disassembly. Early studies detected little CA protein within isolated replication complexes, suggesting rapid dissociation of CA from the viral core soon after entry. Further studies suggested some CA may remain associated with actively replicating RTCs and PICs. Replacing three serine residues (S109, S149 and S178) by 51

substitution with alanine has been shown to result in a block early in replication, during or following reverse transcription. Substituting three additional residues in CA (S41A, S146A and T188V) results in altered virus replication. This study aimed to investigate whether the block in replication was mediated by a defect in core disassembly.

This thesis investigates HIV-1 core stability, core disassembly and replication competence.

The following chapters describe:

1. characterisation of replication competency of HIV-1 CA mutant viruses (Chapter 3).

2. analysis of in vitro core stability of CA mutant viruses (Chapter 4).

3. use of the ‘fate-of-capsid’ assay following cell infection to study core disassembly

(Chapter 5).

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CHAPTER 2 - Materials and methods

2.1 Materials

2.1.1 Cell culture

All cell lines were maintained in a 37ºC incubator with 5% CO2. 293T cells were obtained from the ATCC. 293T cells are a clone of the 293 cell line derived from human embryonic kidney. 293 cells have been transformed with adenovirus 5 DNA, and the 293T clone contains the temperature sensitive gene for the SV40 T-antigen. 293T cells are adherent and highly transfectable. 293T cells were passaged in Dulbecco’s Minimal Essential Medium (DMEM) complete medium; DMEM containing 12 μg/mL Penicillin, 16μg/mL Gentamycin, 2 mM L-

Glutamine, 10 mM HEPES and 10% Foetal Bovine Serum. Cells were seeded at 1.6 x 106 in

100 mm dishes, or 4 x 105 in 6 well plates, and transfected on the following day when they had reached approximately 70% confluency.

HuT-78 cells were obtained through the NIH AIDS Research and Reference Reagent

Program. These cells were derived from a Human cutaneous T cell lymphoma from the peripheral blood of a patient with Sezary syndrome. These cells are negative for secreted (slg) and plasma-membrane-bound (mlg) forms of the complement receptor and EBNA. HuT-78 cells possess the IL-2 receptor and secrete IL-2 and migration inhibition factor. HuT-78 cells were passaged in complete RPMI medium, containing 12 μg/mL Penicillin, 16 μg/mL

Gentamycin, 2 mM L-Glutamine, 20 mM HEPES, 7.5 mL of sodium bicarbonate and 10%

Foetal Bovine Serum. HuT-78 cells were resuspended at 1 x 106 cells/mL the day before infection.

HeLa-CD4-LTR-β-gal cells were obtained through the NIH AIDS Research and Reference

Reagent Program. These cells are human cervical carcinoma cells (HeLa) that have been infected with a retroviral vector expressing CD4, and stably transfected with a truncated HIV-

1 LTR-β-gal plasmid. These cells express high levels of CD4, and contain one integrated copy

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of the HIV-1 LTR linked to the β-galactosidase gene. HeLa-CD4-LTR-β-gal reporter cells were passaged in medium A, containing 12 μg/mL Penicillin, 16 μg/mL Gentamycin, 2 mM

L-Glutamine, 10 mM HEPES, 10% Foetal Bovine Serum, 200 μg/mL G418 Sulfate

(Geneticin®) and 100 μg/mL Hygromycin B. For virus infection, HeLa-CD4-LTR-β-gal cells were seeded at 2 x 104 cells per well of a 48 well plate one day before infection in medium A.

Virus inoculum was diluted in medium B; 50 mL DMEM and 100 µL 10 mg/mL DEAE

Dextran.

2.1.2 Plasmid vectors pNL4-3 plasmid vector encodes the NY5/BRU (LAV-1) recombinant clone of HIV-1, NIH accession number M19921 (Appendix 1). pNL4-3 p24 CA region was cloned into pET-32a (Novagen) (using BssHII/ApaI and

SphI/ApaI restriction enzymes respectively) to generate pET-32-CA(BssHII/ApaI) and pET-

32-CA(SphI/ApaI) (Appendix 2). CA mutants in this study have been named S*A, where ‘*’ denotes the changed amino acid position in CA.

2.1.3 Virus stocks

Virus stocks were produced from 293T cells transfected with pNL4-3, or plasmids encoding

CA mutant viruses. H3B virus supernatant was harvested from H3B cells, which are H9 cells persistently infected with the HXB2 strain of HIV-1 (Li et al., 1992).

2.1.4 Bacterial culture

Escherichia coli (E. coli) strain DH5α were made competent using the Calcium Chloride method (Protocol II (Sambrook et al., 1989)). Commercially available NovaBlueTM competent cells (Novagen) were also used. Glycerol stocks of bacterial cultures were prepared by adding 15% sterile glycerol and were stored at -80ºC.

2.1.5 Common solutions

10x DNA loading buffer: 60% glycerol; 100 mM ethylenediaminetetraacetic acid (EDTA)

(pH 8.0); 100 mM Tris (pH 7.5); Bromophenol blue; Xylene cyanol

54

4x SDS protein loading buffer: 0.25 M Tris-HCl, 8% (w/v) SDS, 20% (v/v) glycerol,

Bromophenol blue pH 6.8, β-mercaptoethanol (βME) (50 µL/mL)

Coomassie Blue Stain: 50% methanol, 10% acetic acid, 0.2% (w/v) Coomassie Brilliant Blue

Destain: 50% methanol, 10% acetic acid

5% TCA solution: 5% TCA, 0.2 M Na2HPO4

HIRT solution I: 5 mM Tris pH 7.7, 10 mM EDTA

HIRT solution II: 5 mM Tris pH 7.7, 10 mM EDTA, 1.2% SDS

TBS-T: 25 mM Tris pH 7.6, 137 mM NaCl, 0.1% Tween-20

PBS-T: 1x PBS, 0.05% Tween-20

Blocking Buffer: 5% skim milk powder, TBS-T

TAE Buffer: 40 mM Tris-acetate, 1 mM EDTA

Luria-Bertani Broth (LB): 1% (w/v) bacto-tryptone, 0.5% (w/v) bacto-yeast extract, 0.17 M

NaCl

LB Ampicillin plates: LB broth, 100 μg/mL Ampicillin, 3 g/100mL Bacto-agar

TNE Buffer: 100 mM NaCl, 1 mM EDTA, 10 mM Tris-HCl pH 7.2

Modified RIPA Buffer: 0.07 mM Tris pH 7.4, 0.15 mM NaCl, 1 mM EDTA, 0.25% Na deoxycholate, 1% NP-40, 1 protease inhibitor cocktail tablet (Roche)/10 mL

Hypotonic Buffer (5x): 50 mM Tris, 250 mM KCl, 25 mM MgCl2, 1 protease inhibitor cocktail tablet (Roche)/10 mL

Protein Gel Running Buffer: 0.3% (w/v) Tris, 1.44% (w/v) Glycine, 0.1% SDS

PVDF Transfer Buffer: 1.44% (w/v) Glycine, 0.3% (w/v) Tris, 20% Methanol

SOC medium: 0.5% Yeast Extract, 2% Tryptone, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2,

10 mM MgSO4, 20 mM Glucose

2.1.6 Antibodies

Mouse anti-p24 monoclonal antibody (#6458 or #4121, NIH AIDS Research and Reference

Reagent Program) was used at 1/5000 in TBS-T. Anti-gp120 (obtained from the National

Serology Reference Laboratory) was used at 1/2000 in TBS-T in western blot. Anti-p17

55

antibody was used at 1/5000 in TBS-T in western blot. Goat anti-mouse HRP (Pierce) and goat anti-rabbit HRP (Pierce) were used at 1/100000. Anti-p24, anti-gp120 and goat anti- mouse HRP were all used in ELISA at 1/1000.

2.2 DNA cloning and extraction

2.2.1 Restriction digestion

DNA was digested using 0.5-1 μL of restriction enzyme (New England BioLabs, NEB) with the appropriate enzyme buffer, and 0.1 mg/mL BSA when required, in a 20 µL reaction.

Digests were incubated for 1½ hr at 25-50ºC as directed. Two-enzyme digests were performed concurrently where possible. Restricted DNA fragments were analysed on 1% agarose gels.

2.2.2 Agarose gel electrophoresis

1% agarose gels were prepared using 1 g of DNA grade agarose powder (Progen) melted into

100 mL of 1x TAE Buffer, poured into plastic gel trays and allowed to set. DNA samples were loaded in 10x DNA loading buffer and electrophoresed at 100 V for approximately 1 hr.

The gels were stained with Ethidium Bromide for 10 min and destained in water for 15 min before being visualised using a UV transilluminator and photographed. DNA required for gel extraction and cloning was visualised using a hand-held ULTRA-LUM low wave-length lamp to minimise damage of DNA.

2.2.3 DNA extraction from agarose gels

DNA bands were excised from agarose gels using a sterile scalpel blade. DNA was extracted using a QIAquick Extraction Kit (QIAGEN) following the protocol in the QIAquick

Extraction Kit handbook. DNA was eluted using 50 μL of EB Buffer as described.

56

2.2.4 Ligation

DNA ligation reactions were performed using T4 DNA ligase (New England BioLabs) in 20

μL reactions. 100 ng total DNA was used at 1:1 and 2:1 insert to vector ratios. Ligation reactions were incubated at 16ºC overnight, transformed into competent E. coli (DH5α or

NovaBlue), and grown on LB bacto-agar plates containing 100 μg/mL ampicillin or 50 μg/mL kanamycin.

2.2.5 Transformation

DNA was transformed into NovaBlue chemically competent E. coli (Invitrogen).

Transformations were incubated on ice for 5 min and heat-shocked at 42ºC for 30 sec, incubated on ice for a further 2 min before addition of 250 µL volume of LB Broth or SOC medium. Bacterial cultures were then plated onto LB bacto-agar ampicillin or kanamycin plates, and incubated overnight at 37ºC. Bacterial colonies were picked the following day and grown overnight in LB broth containing the appropriate antibiotics, and plasmid DNA was isolated using QIAGEN Mini-prep or Maxi kits.

2.3 DNA extraction from cells

2.3.1 Plasmid extraction and purification

Plasmid DNA was extracted and purified from bacterial cultures using the QIAprep Spin

Miniprep Kit (QIAGEN) and Plasmid Midi and Maxi Kit (QIAGEN), following the manufacturer’s instructions. The extracted DNA was resuspended in double distilled water at

1-3 μg/μL.

2.3.2 HIRT DNA extraction

Extrachromosomal cellular DNA was separated from chromosomal DNA using the HIRT method (Hirt, 1967). Briefly, infected cells were washed in PBS and the pellets resuspended in 160 μL HIRT cell lysis solution I, followed by the addition of 20 μL Proteinase K, then 200

μL HIRT solution II. Tubes were inverted 10 times slowly to minimise chromosomal DNA 57

shearing, then incubated at 37ºC overnight. 100 μL of 5 M NaCl was added and tubes inverted twice gently to mix, then placed at 4ºC overnight. Samples were centrifuged at 17000 x g for

45 min at 4ºC, and the supernatant (extrachromosomal DNA) separated from the pellet

(chromosomal DNA) fraction. DNA in the HIRT supernatant was purified by adding an equal volume of phenol/chloroform/isoamylalcohol, and then precipitated in 2x volume 100% ethanol in the presence of glycogen. DNA pellets were washed in 70% ethanol, air dried and resuspended in 50 μL of water.

2.3.3 Spectrophotometry and DNA quantitation

Plasmid DNA concentrations were measured using the UV/Visible Spectrophotometer,

Ultrospec 300 (Pharmacia Biotech). Samples were diluted 1/50, and the optical density (OD) measured at 260 nm and 280 nm. An OD260/280 ratio between 1.6 and 1.8 was routinely observed, and DNA concentration determined at OD260 nm. For estimation of DNA amounts in ethidium bromide stained agarose gels, the intensity of DNA bands was compared with

Spp1 DNA standards of known molecular mass.

2.4 Construction of capsid phosphorylation mutants

2.4.1 PCR mutagenesis of the CA region

Serine to alanine mutations at HIV-1 CA phosphorylation sites were introduced by PCR mutagenesis of pNL4-3 (Appendix 1) using 2 strategies: site-directed mutagenesis (2.4.2) or overlap extension PCR mutagenesis (2.4.3) (Figure 2.1).

Mutagenic primers were designed to create six CA mutants, S41A, S109A, S146A, S149A,

S178A and T188V; and two control mutants E128A/R132A and K203A (Table 2.1). Each primer pair (designated forward (F) and reverse (R)) incorporated the appropriate amino acid substitution, and base changes specifying a unique restriction site without altering the amino acid sequence. This enabled identification of mutant clones by restriction digest. Primers were synthesised as 34-45 base oligonucleotides by GeneWorks.

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Figure 2.1 Schematic representation of CA mutagenesis strategies (A) Site Directed PCR Mutagenesis. (i) pNL4-3 CA was sub-cloned into (ii) pET-32a vector (either as BssHII/ApaI or SphI/ApaI fragments, pET32a (BA) or (SA), see Appendix 2). (iii) Mutagenic primers (designated with “x”, see Table 2.1) were used to incorporate the desired amino acid change giving (iv) pET32a-CAx. CAx was then sub-cloned back into pNL4-3 giving (v) pNL4-3- CAx. (B) Over-lap Extension Mutagenesis. (i) 1st round PCR used a primer encoding the desired amino acid change (designated with “x”, see Table 2.1) and a primer outside the CA region (see Table 2.3) to generate fragments encoding the desired amino acid substitution using full length pNL4-3 as the template. (ii) 2nd round PCR was performed on annealed 1st round products and utilised primers outside CA region to generate the entire CA region containing the desired amino acid substitution CAx. CAx was then cloned back into full length pNL4-3 using BssHII or SphI and ApaI to give (iv) pNL4-3- CAx. CA

A BssHII SphI ApaI X X BssHII SphI712 1448 i ApaI 2011 ii iii

pNL4-3 pET32a-CA pET32a-CA pNL4-3 ~6000bp 14879bp14879 bp

Sub-cloning of Site-directed CA region mutagenesis

x

BssHII SphI712 1448 ApaI 2011 x X iv v

x pET32a-CA x pNL4pNL4-3-3-CA 14879 bp Sub-cloning of CA back into infectious clone

B A1 A2 B1 B2 X X i CA Full length pNL4-3

A1 B2

X SphI BssHII ApaI ii X BssHII SphI712 1448 ApaI 2011 X

BssHII SphI ApaI

x X pNL4pNL4-3-3-CA iii iv 14879 bp

Mutagenic Primers

S41A 5' GTAATACCCATGTTTGCAGCGCTATCAGAAGGAGCCA 3' 5' TGGCTCCTTCTGATAGCGCTGCAAACATGGGTATTAC 3' S109A 5' ATAGCAGGAACTACTGCTACCCTTCAGGAACA 3' 5' TGTTCCTGAAGGGTAGCAGTAGTTCCTGCTAT 3' KpnI S109A 5’ AAGTGACATAGCAGGTACCACTGCGACCCTTCAGGAACAA 3' 5' TTGTTCCTGAAGGGTCGCAGTGGTACCTGCTATGTCACTT 3' GCG S109A 5' ATAGCAGGAACTACTGCGACCCTTCAGGAAC 3' 5' GTTCCTGAAGGGTCGCAGTAGTTCCTGCTAT 3' S146A 5' ATAGTAAGAATGTATGCCCCTACATCGATTCTGGACATAAGA 3' 5' TCTTATGTCCAGAATCGATGTAGGGGCATACATTCTTACTAT 3' S149A 5' ATGTATAGCCCTACCGCCATTCTAGACATAAGACAAGGA 3' 5' TCCTTGTCTTATGTCTAGAATGGCGGTAGGGCTATACAT 3' S178A 5' AGAGCCGAGCAAGCTGCACAAGAGGTAAAAA 3' 5' TTTTTACCTCTTGTGCAGCTTGCTCGGCTCT 3' T188V 5' AATTGGATGACAGAAGTCCTACTAGTCCAAAATGCGAAC 3' 5' GTTCGCATTTTGGACTAGTAGGACTTCTGTCATCCAATT 3' E128A/R132A 5' CTATCCCAGTAGGAGCAATCTATAAAGCTTGGATAATCCTGGGA 3' 5' TCCCAGGATTATCCAAGCTTTATAGATTGCTCCTACTGGGATAG 3' K203A 5' TGTAAGACTATTTTAGCAGCATTGGGCCCAGGAGCGACACTA 3' 5' TAGTGTCGCTCCTGGGCCCAATGCTGCTAAAATAGTCTTACA 3'

Table 2.1 Mutagenic primer pairs used for site-directed mutagenic or overlap extension PCR Forward and reverse sense primer pairs used to generate each CA mutant are presented.

BOLD/Underlined indicates nucleotide substitutions.

Highlight indicates the amino acid residue substituted.

2.4.2 Site-directed PCR mutagenesis of CA in pET-32a

The CA region from pNL4-3 was cloned into the pET-32a vector (as it contained the necessary restriction sites) using restriction enzymes BssHII/ApaI or SphI/ApaI to generate pET-32a-CA (BA) or (SA) respectively (Appendix 2). These plasmids were used to generate

S41A (pET-32a-CA (BA)), S109A, S149A and S178A (pET-32a (SA)) by site-directed PCR mutagenesis (Figure 2.1A). This approach used specific mutagenic primers (Table 2.1) to introduce the desired amino acid substitution and a unique restriction site in the DNA sequence for identification of CA mutants. The desired region of CA was subcloned into pNL4-3 to produce full length infectious CA mutants. pET32-CA PCR replicons were purified by gel extraction from 1% agarose gels and amplified by transformation into E. coli

DH5α or NovaBlue cells.

2.4.3 Overlap extension PCR mutagenesis

Additionally, CA mutants were generated by overlap extension PCR mutagenesis and the

PCR products subcloned back into pNL4-3 to generate full length infectious CA mutants

(Figure 2.1B). Specific mutagenic primers and a primer outside the CA region were used to generate two PCR fragments containing the desired mutation. The outside primers were then used to amplify the entire CA region, which was then ligated into pNL4-3. S109AKpnI,

S109AGCG, S146A, E128A/R132A, T188V and K203A were produced by overlap extension

PCR directly from pNL4-3.

Due to difficulties demonstrating viral protein expression for CA mutant virus S109A, this clone was generated using three different approaches each using alternative sets of primers with altered codon usage.

2.4.4 PCR cycling parameters

Each 50 μL reaction PCR was performed using 5 U/μL high fidelity Platinum® Taq DNA

Polymerase (Invitrogen) in 0.2 mL thin wall microcentrifuge tubes (Axygen Scientific). PCR reactions contained DNA polymerase, 10x PCR buffer, primers, MgSO4, DMSO and water.

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PCR amplification was carried out in a GeneAmp® PCR System 9700 thermocycler (Perkin

Elmer).

Following PCR amplification, the reactions were digested with Dpn1. Dpn1 specifically digests methylated GATC sequences present in bacterially derived plasmid template, but not newly synthesised PCR plasmid replicons.

2.4.5 Subcloning mutant CA sequences to generate full length infectious pNL4-3

To minimise the amount of PCR amplified DNA sub-cloned back into the pNL4-3 construct,

CA S41A was sub-cloned from pET-32a (BA) by BssHII/SphI digestion and ligated into pNL4-3 (BssHII/SphI). S109A, S149A and S178A were sub-cloned from pET-32a (SA) by

SphI/ApaI digestion before ligation into pNL4-3 (SphI/ApaI) (Figure 2.2).

Similarly, S109AKpnI, S109AGCG, S146A, E128A/R132A and T188V CA clones produced by overlap extension PCR, were digested with SphI/ApaI and ligated back into pNL4-3

(SphI/ApaI). K203A PCR amplified DNA was digested with SphI/SbfI before ligation into pNL4-3 (SphI/SbfI) (Figure 2.2).

Following ligation of each CA mutant DNA region back into pNL4-3, sequence analysis of the entire CA sequence and the flanking ligation junctions for each CA mutant clone was again performed to ensure no further mutations were introduced (Appendix 3). 10 CA mutants were successfully generated as full length infectious clones: S41A, S109A,

S109AKpnI, S109AGCG, S146A, S149A, S178A, T188V, E128A/R132A and K203A.

2.4.6 Screening CA mutant clones by restriction digest analysis

Following PCR mutagenesis by either strategy above, the pNL4-3 CA mutant plasmid was transformed into competent Escherichia coli, transformants were selected and the desired CA mutants were identified by their unique restriction enzyme cleavage pattern compared to WT pNL4-3 (Table 2.2).

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a Restriction Unique Restriction Site CA Mutation Enzyme Introduced Site Removed site S41A AfeI 1311 - S109A SpeI - 1508 S109AKpnI KpnI 1506 1508 S109AGCG SpeI - 1508 S146A ClaI 1631 - S149A XbaI 1636 - S178A HindIII - 1713 T188V SpeI 1753 - E128A/R132A HindIII 1577 - K203A ApaI 1804 -

Table 2.2 Positions of unique restriction sites in pNL4-3 CA mutants Restriction sites were introduced or removed from CA mutant nucleotide sequences without altering the coding amino acid sequence. This enabled identification of mutated clones through the unique restriction enzyme patterns produced compared to WT pNL4-3 virus. a nucleotide numbering in accordance with HIV pNL4-3 GenBank accession number M19921.

Figure 2.2 Schematic representation of the sub-cloning strategy for the mutagenic CA region HIV DNA including the CA region (indicated by the red box) containing amino acid substitutions was amplified in pET32a(BA), pET32a(SA) or by overlap extension PCR (indicated by extended dotted line). DNA containing the desired substitutions were sub-cloned back into pNL4-3 using either BssHII/ApaI, SphI/ApaI or SphI/SbfI restriction digest as indicated, to create infectious clones. BssHII SphI ApaI SbfI (712) (1448) (2011) (2844)

WT NL4-3 CAPSID S41A * S109A * S109AKpn * S109AGCG * S146A * S149A * S178A * T188V * E128A/R132A ** K203A * 2.5 Confirmation of mutations by sequence analysis

Potential CA mutant clones identified in 2.4.6 were further analysed by sequencing the entire region amplified during the PCR mutagenesis protocol. It was essential that CA mutants did not contain other spurious mutations in the CA region that could have been introduced during

PCR mutagenesis. Sequences of each CA mutant aligned with the theoretical WT pNL4-3 derived sequence are presented in Appendix 3.

2.5.1 Sequencing primers

Sequencing primers were designed using PRIMER software (Primer Designer Version 2.0), based on criteria to avoid dimers and hairpin formation and 3 identical base runs (e.g. GGG), while maintaining greater than 50% GC content and Tm (melting temperature) value greater than 65ºC. 12 forward primers (single strand sequence specific) overlapping the entire HIV-1

CA sequence in pNL4-3 or pET-32a-CA at intervals of 500 bp were synthesised

(GeneWorks) (Table 2.3).

2.5.2 Sequencing PCR

Mutated pNL4-3-CA clones propagated from E. coli DH5α transformed with pNL4-3-CA

PCR replicons were sequenced using the dye terminator method. Briefly, 500 ng of DNA was added to a 10 μL reaction containing 4 μL of Big Dye III Sequencing Mix (provided in the

ABI Prism Dye Primer Cycle Sequencing Ready Reaction Kit), and 2 μL of sequencing primer (1 pmol/μL). PCR reactions were performed in the GeneAmp® PCR System 9700 thermocycler at 96ºC for 1 min, 25 cycles of 96ºC for 10 sec, 50ºC for 5 sec and 60ºC for 4 min, and then held at 4ºC. PCR products for sequencing were precipitated using isopropanol, and sequenced by the Sequencing Centre of the Division of Molecular Pathology, IMVS, SA,

Australia. Sequencing results were analysed using BioEdit Sequence Alignment Editor software version 6.0.7 using the pNL4-3 CA sequence (NIH accession number M19921).

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NL4-3 Primers

Eco682 5' CACACAGAATTCTCTCTCGACGCAGGA 3' 843 5' AATTCGGTTAAGGCCAGG 3' 1369 5' CATCAAGCAGCCATGCAA 3' 1803 5' ACCAGGAGCGACACTAGA 3' 1880 5' CTGAAGCAATGAGCCAAG 3' 2365 5' TGCCAGGAAGATGGAAAC 3' r816 5' GCTTAATACCGACGCTCT 3' r2081 5' GCCTGTCTCTCAGTACAA 3' Bam2107 5' CACACAGGCTCCAAGGCCAGATCTTCC 3' r3760 5' TCTGTCCACCATGCTTCC 3'

pET32a Primers pET-870 5' TATATCGCCGACATCACC 3' pET-r1982 5' ACTACCGAGATGTCCGCA 3'

Table 2.3 DNA primer sequences Primers used for sequencing of CA clones in pET32a or pNL4-3 plasmid or used in conjunction with mutagenic primers (listed in Table 2.1) for overlap extension PCR.

2.6 Cell transfections and virus isolation

2.6.1 Transfection

293T cells were seeded at 1.6 x 106 cells in 100 mm dishes. Cells were transfected the following day when they had reached 70% confluency with 10 μg plasmid DNA and

SuperFect® Transfection Reagent (QIAGEN) as specified in the manufacturer’s handbook.

Briefly, cells were incubated in 3 mL antibiotic-free and serum-free DMEM medium containing plasmid DNA and SuperFect® reagent for 2½-3 hr at 37ºC in a CO2 incubator.

Cells were washed with PBS and cultured in 10 mL complete DMEM media (2.1.1) for a further 24-48 hr prior to harvesting cells and virus.

2.6.2 Recovery of virus for infections

Virus-containing supernatants from transfections were treated with 50 µg/mL DNase1

(Roche) in the presence of 10 mM MgCl2 for 30 min at room temperature and filtered through a 0.2 µm filter. Supernatants were aliquotted and stored at -80ºC for infection experiments.

285 µL supernatant (plus 15 µL 10% Triton-X-100) was stored at 4ºC for later measurement of virus protein p24 CA levels by p24 ELISA.

2.6.3 Concentration of virus preparations

Virus-containing supernatant was filtered through a 0.2 µm filter. Supernatant was then centrifuged through an Amicon® Ultra Centrifugal Filter Device 300K molecular weight cut- off (MWCO) (Millipore) or Vivaspin20 100K MWCO (Sartorius) for 30 min at 3000 rpm per

15-20 mL. A maximum volume of 50 mL was concentrated through a single filter device to a final volume of 500 µL.

2.6.4 Pelleting virus preparations

Virus was further purified by pelleting concentrated virus through 25% sucrose in TNE in a

Beckman TLA-100 rotor at 40000 rpm for 90 min at 4ºC.

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Virus pellets were resuspended in 40 μL protein loading buffer (2.1.5) and stored at -20ºC for analysis by western blot, or resuspended in 200 μL 1x TNE for immediate sucrose gradient ultracentrifugation.

2.6.5 Harvesting transfected cell lysates

Following removal of virus supernatant, the transfected cell monolayers were washed in 1x

PBS. Cells were lysed in 1 mL modified RIPA buffer (2.1.5) for 1 hr at 4ºC before centrifugation at 12000 x g for 30 min at 4ºC to pellet cell debris. Clarified cell lysates were stored in aliquots in protein loading buffer at -20ºC for western blot analysis.

2.7 Cell infections

2.7.1 Virus CA p24 measurement

Virus capsid (CA) p24 protein concentration was measured using a commercially available

HIV-1 p24 ELISA kit from NEN™ Life Sciences Products (PerkinElmer™). Virus- containing supernatants from transfected 293T cells were diluted to 1/25, 1/250, 1/2 500 and

1/25000 (in 1x PBS containing 0.5% Triton-X-100 to inactivate virus). p24 protein content was measured to determine the amount of virus present in each culture supernatant. This was used normalise the amount of virus inoculum used for infection experiments.

2.7.2 HuT-78 cell infection

In triplicate, 2 x 106 HuT-78 cells were resuspended in 200 μL virus inoculum containing 50 ng p24 in each virus-containing supernatant (2.6.2). Cells were incubated with virus at 4ºC for

30 min, mixing every 10 min, followed by centrifugation at 3300 rpm for 1 hr at 37ºC. This protocol, known as centrifugal enhancement, has been estimated to enhance the effective MOI

10-fold (Pietroboni et al., 1989). Virus inoculum was removed; cells were gently resuspended in complete RPMI media and incubated at 37ºC for 20 min. Finally, cells were washed twice by centrifugation at 3000 rpm for 1 min to remove traces of virus inoculum. Infected cells were then pooled and resuspended at 2 x 106 cells/mL. 3 x 105 infected cells (150 μL) were 63

seeded per well (in a 48 well plate) in triplicate for 8 hr and 24 hr time points. Cells were seeded into microcentrifuge tubes (in duplicate only) for 0 hr time points. At each time point, cells were harvested into microcentrifuge tubes, washed in ice cold PBS, and DNA extracted by the HIRT method (2.3.2).

2.7.3 Single cycle infectivity assay

HeLa-CD4-LTR-β-gal cells were seeded 2 x 104 cells per well (in a 48-well plate) in 300 µL of medium A the day before infection. Cell monolayers were approximately 60% confluent on the day of infection. Virus inoculum from transfected cell supernatant (2.6.2) (containing 20 ng of p24 diluted in 150 μL medium B), was adsorbed on to HeLa-CD4-LTR-β-gal cell monolayers in triplicate at 37ºC for 2 hr. Virus inoculum was removed and replaced with 200

μL medium A, and cells were incubated for a further 38 hr. At 40 hr post infection, cells were washed three times with PBS to remove virus inoculum, and fixed in 1% formaldehyde in

PBS for 10 min. Fixing solution was removed, cells were washed three times with PBS, followed by the addition of X-gal stain solution (made by mixing Solution A: 1x PBS

K4Fe(CN)6.3H20 0.02g, K3Fe(CN)6, 1 M MgCl2, with Solution B: 50 mg/mL X-gal stock solution in DMSO), and incubated for a further 1½ hr. Finally, the stain solution was removed, cells were washed three times in PBS, and the number of blue cells per cell monolayer was counted.

2.7.4 Fate-of-capsid infection

Virus-containing supernatants from HIV plasmid transfected cells (2.5 mL, 200 ng/mL) were mixed with 5 x 106 HuT-78 cells and incubated at 23ºC for 3 hr (temperature arrested state,

TAS), centrifugally enhanced (3300 rpm at 37ºC for 1 hr), then moved to 37ºC. Samples were taken at the time of virus and cell mixing (0 hr), after TAS, and after centrifugal enhancement.

Cells were washed twice in room temperature PBS to remove virus inoculum, resuspended in fresh RPMI and incubated at 37ºC. Further samples were harvested after incubation for 2 hr, 4 hr or 6 hr at 37ºC (including 1 hr during centrifugal enhancement).

64

Samples were centrifuged to pellet cells and washed twice in ice cold PBS. Cells were resuspended in serum-free RPMI and treated with 0.5% trypsin for 3 min at room temperature to remove virus that may have adhered to the outside of cells. Trypsin was inactivated by adding 50% volume of complete RPMI. Cells were washed twice more in ice cold PBS. Cells were lysed in modified RIPA buffer and analysed by western blot for total CA. Alternatively, cells were lysed with disposable plastic pestles in hypotonic buffer for analysis on sucrose gradients.

2.8 Real-Time PCR

Extra-chromosomal DNA extracted by the HIRT procedure from infected HuT-78 cells was analysed by real-time PCR in the Rotor Gene, using primers for Mitochondrial DNA and

HIV-1 reverse transcription intermediate products (Table 2.4). 2 reverse transcription intermediates were detected by real-time PCR: minus strand strong stop DNA (SS1 and SS2b) and a late stage double stranded DNA product (GAG5 and GAG6). Real-time PCR reaction mixtures contained 5 μL Syber Green PCR Mix (includes buffers and enzyme), 1 μL each of

(10 pmol/μL) forward and reverse primers, 1 μL of water and 2 μL of extrachromosomal

HIRT DNA. Results were analysed using Rotor-Gene Real-Time Analysis Software 6.0.14.

2.9 Protein analysis

2.9.1 Protein quantitation assay

Protein was quantitated using a Bio-Rad DC Protein Assay (Bio-Rad Laboratories).

Routinely, lysates were diluted and assayed against known quantitative bovine serum albumin

(BSA) protein standards to allow determination of protein levels in lysates.

65

Primers for real-time PCR Coordinates

a SS1 5' CTAACTAGGGAACCCACTGC 3' nt 498-517 SS2b 5' ACTAGAGATCCCTCAGACCC 3' nt 606-587a GAG5 5' GGTACATCAGGCCATATCAC 3' nt 1215-1234 GAG6 5' GTTCCTGCTATGTCACTTCC 3' nt 1505-1486 MIT1 5' GACCTTAGGTCAAGGTGTAG 3' n/a MIT2 5' GGTTGTCTGGTAGTAAGGTG 3' n/a

Table 2.4 DNA primer sequences Primers used for detecting HIV DNA intermediates in real-time PCR reactions. aPositions in the HIV-1 pNL4-3 DNA sequence are indicated (GenBank accession number M19921).

2.9.2 Protein precipitation of sucrose gradient fractions

One tenth the volume of 100% TCA was added to each gradient fraction, and incubated at 4ºC for >1 hr. Samples were centrifuged at 13000 rpm at 4ºC for 30 min. Supernatants were carefully removed to avoid residual sucrose, and pellets washed three times in acetone plus

0.07% βME. Protein pellets were air dried and resuspended in 15 uL of 4x SDS LB.

2.9.3 Preparation of cell lysates from fate-of-capsid infections

Total protein from infected cell lysates was quantitated using BioRad Dc Protein assay

(2.9.1). Equal amounts of protein were loaded onto large format 12.5% acrylamide gels and analysed by western blot for p24 CA.

Infected cell lysates to be analysed on sucrose gradients were lysed and stored at -20ºC overnight. The following day, cell lysates were loaded onto 15-40% sucrose gradients and ultracentrifuged at 35000 rpm for 40 min at 4ºC. Fractions were taken from the top of the gradient, protein was precipitated (2.9.2) and fractions analysed by western blot or p24

ELISA (NEN™ Life Science Products, PerkinElmer™).

2.10 Western Blot Analysis

Protein samples (50-100 μg total protein) in 4x protein loading buffer were boiled for 5 min and loaded onto 12.5% acrylamide gels (12.5% resolving, 5% stacking), alongside

BenchMark™ Prestained Protein markers (Invitrogen). Proteins were electrophoresed at 100

V for 1-1½ hr. Protein was transferred to Hybond™-P PVDF Transfer Membrane (Amersham

Biosciences) using a TRANS-BLOT® SD, Semi-Dry Transfer Cell (BIO-RAD) (0.8 mA/cm2 for 1½ hr) or BioRad Mini-Protean III wet transfer cell (BIO-RAD) (100 V for 1 hr).

Membranes were soaked in protein blocking buffer for 1 hr before incubation with primary antibody (1/5000 monoclonal anti-p24 antibody) in TBS-T, at 4ºC overnight. Membranes were washed four times in TBS-T for 15 min with shaking, then incubated in the appropriate

HRP conjugated secondary antibody (1/100000 goat anti-mouse-HRP (Pierce) for p24 protein

66

detection) for 1 hr at room temperature with shaking. Membranes were washed four times in

TBS-T for 15 min with shaking, before addition of SuperSignal® West Dura Extended

Duration Substrate (PIERCE) or ECL (Amersham) for 3 min, and finally exposed to X-

OMAT K film (Kodak) or Fuji film (Fuji) for 2-20 min before development.

2.11 Sucrose Gradient Ultracentrifugation

2.11.1 Sucrose solutions

Sucrose solutions were prepared in 1X TNE buffer and stored at 4ºC or 25ºC (room temperature). Sucrose percentage was checked by measuring the refractive index of each solution with a refractometer. Solutions were adjusted by adding further 1X TNE buffer or sucrose where appropriate. One protease inhibitor tablet (Pierce) was dissolved per 10 mL sucrose prior to pouring gradients.

2.11.2 Preparation of gradients

Sucrose gradients were prepared in 12.5 mL ultra-clear ultracentrifuge tubes (Beckman). Each gradient contained a 500 µL 60% sucrose cushion and a 10 mL linear 15-40% gradient.

Gradients were poured using a gradient forming chamber. Equal volumes of low and high percentage sucrose were poured into the chambers and mixed with a magnetic flea. 500 µL sucrose containing detergent was laid on top of the gradient. 15% sucrose was added to balance the weight of tubes before ultracentrifugation. Ultracentrifugation was performed in a

Beckman SW-42Ti centrifuge.

2.11.3 Collection of fractions

Following ultracentrifugation of virus through sucrose gradients, 1 mL fractions were collected from the top of the gradient using a 1 mL Gilson pipette. Fraction 1 represents the top of the gradient, i.e. lowest percentage sucrose. Fraction 12 represents the bottom of the gradient, i.e. highest percentage sucrose.

67

2.11.4 Measuring refractive index

Refractive index measurements were recorded for each fraction using a refractometer. Values were converted to percentage sucrose. This enabled slight variations in sucrose percentage between gradients to be taken into account when examining migration of virus components in the gradient.

68

CHAPTER 3 - Characterisation of capsid mutant viruses

3.1 Introduction

The assembly of a stable HIV core requires the generation of hexameric CA protein “rings” which link together generating the hexameric CA lattice network. This relies primarily on the formation of two protein dimerisation interfaces between CA monomers, the NTD-NTD dimerisation interface (to generate hexameric rings) and the CTD-CTD dimerisation interface

(to link adjacent hexamers). Helices 1 and 2 contribute to the NTD interface at the centre of

CA hexamers, stabilising the structure. Mutations which disrupt the NTD dimerisation interface (e.g. A22D, E28A/E29A, M39D, A42D, and D51A) prevent formation of a conical core, and reduce virus production and infectivity (von Schwedler et al., 1998). Likewise, alanine substitution at key CTD dimerisation residues (e.g. W184 and M185 in helix 9) prevents CTD dimerisation and results in the production of non-infectious virus (Gamble et al., 1997).

The helices within CA monomers are packed together by extensive hydrophobic interactions

(Gitti et al., 1996). These interactions maintain the proper stability of the immature core structure, but must also allow for rearrangement of the CA monomers following proteolytic processing during maturation. It is therefore possible, that residues may take part in interactions in the immature core that are no longer present in the mature core.

Forshey et al (2002) introduced mutations at charged HIV-1 CA residues which resulted in altered core stability (e.g. R18A/N21A, P38A, E45A, Q63A/67A, L136A, R143A, K170A and Q219A). Notably, two HIV-1 CA mutant viruses, E128A/R132A and K203A, exhibited hyperstable and hypostable cores respectively in an in vitro core stability assay (Dismuke &

Aiken, 2006; Forshey et al., 2002; Hulme et al., 2011). This study highlighted the contribution of charged residues in the formation of a sufficiently stable HIV core. In addition to altered stability, these viruses display defects in virus replication at reverse transcription or nuclear translocation. 69

The importance of charge within the viral core led to consideration whether modification of the major HIV-1 core protein, p24 CA, by phosphorylation might play a role in regulating the stability of the core and potentially affect release of CA from the core during core disassembly.

In vitro phosphorylation demonstrates that the HIV-1 CA protein present in mature virions is phosphorylated on three serine residues (S109, S149 and S178) (Cartier et al., 1999). In over

100 HIV-1 strains in the HIV Sequence Database these three serine residues are either highly conserved (S109:110/110, S149:106/110 sequences) or conservatively substituted by another phospho-acceptor amino acid (threonine) (S178:33/110 sequences, 76 sequences contain a threonine at position 178) (2008a). Experimental substitution of serine with alanine identified five serine residues that are required for optimal viral replication: S41, S109, S146, S149 and

S178. S109A, S149A and S178A CA mutant viruses produced non-infectious virions that had normal RT activity and normal virion morphology, but were blocked at an undefined early stage of HIV replication prior to integration (Cartier et al., 1999). This suggests an important role for these CA residues in the HIV life cycle, perhaps via protein phosphorylation at these sites. The location of S109, S149 and in particular S178 in the CA protein monomer suggests they could provide important contributions to HIV-1 core structure and stability (Figure 3.1).

S178 forms part of HIV-1 CA α-helix 9, which forms the main CTD dimer interface between adjacent CA monomers (del Alamo et al., 2003; Gamble et al., 1997; Lidon-Moya et al.,

2005). This residue also makes intersubunit contacts with CTD residues on adjacent hexameric ring structures, forming a hydrogen bond with E180 (del Alamo & Mateu, 2005; del Alamo et al., 2003; Worthylake et al., 1999). Therefore S178 may affect HIV-1 CA dimer formation, and regulation of charge by phosphorylation may be important for stabilising the core structure. S109 is located between helices 5 and 6 in the NTD, and S149 is located immediately prior to the C10 helix in the CTD (Gamble et al., 1996; Kelly et al., 2006). The functions of these regions are unknown. They are not part of the major homology region

(MHR), or sites known to interact with other viral or host cell proteins or nucleic acids, but

70

Figure 3.1 Ribbon diagram of the crystal structure of the HIV-1 CA monomer (PDB entry 1e6j) CA amino-terminal domain (NTD) and carboxyl terminal domains (CTD) are indicated. α-helices are numbered 1-7 (NTD) and 8-11 (CTD). Serine residues substituted to alanine are indicated by spheres. Hydroxyl groups removed during mutagenesis are represented by red/gray spheres. (Carbon = green, Hydrogen = gray, Nitrogen = blue, Oxygen = red)

Figure drawn using PyMol version 1.0r1. PDB entry 1e6j Monaco-Malbet, Structure, 2000 S178

1

2 S41 N-terminus S109 3 9

7 8 S146 5 S149 11 6 4 10

CypA loop C-terminus

NTD CTD nevertheless S109 and S149 residues are still essential for HIV-1 infectivity (Cartier et al.,

1999). CA mutant viruses S41A and S146A were reported to show delayed replication kinetics in several cell types (Cartier et al., 1999). These residues are situated in helix 2 and immediately following helix 7 in the NTD and make intramolecular and intermolecular contacts in formation of the mature and immature (helix 7) virion (Wright et al., 2007). This suggests the location these residues may also be important for stability of the viral core. While the substitution of S41 and S146 with alanine did not reduce CA protein phosphorylation in mature virions (Cartier et al., 1999), this does not rule out the potential for phosphorylation at these sites once virus enters the cell.

T188, like S178, is situated on the CTD dimerisation interface. Substitution at T188 to alanine increases dimerisation affinity at the CTD dimerisation interface (del Alamo et al., 2003), and thus it was of interest to consider if the enhanced stability of the CA dimer afforded by this substitution led to enhanced stability of the virus core.

Additionally, two CA mutant viruses were generated to be used as controls in Chapter 4. The

CA mutant viruses E128A/R132A and K203A are reported to exhibit hyperstable and hypostable cores respectively (Dismuke & Aiken, 2006; Forshey et al., 2002; Hulme et al.,

2011).

Therefore CA mutant viruses were created at S109, S149 and S178 to potential phosphorylation sites, and additionally S41A, S146A, T188V, E128A/R132A and K203A

(Figure 3.2) to investigate the effect on core stability and the early post-entry steps in the

HIV-1 life cycle.

Chapter 3 describes the characterisation of these CA mutant viruses. This chapter demonstrates that viral protein processing in these virions is normal, with the exception of

S109A, and analyses the reverse transcription and replication competence of these viruses following infection of target cells.

3.2 Results

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Figure 3.2 Representation of amino acid substitutions within the CA sequence The residue number and amino acid substitutions are indicated on the left. Their positions are indicated within the CA sequence. Conservative changes were made: serine, glutamic acid, arginine and lysine were substituted for alanine. Threonine was substituted with valine. 1 231

WT NL4-3 P------S------S----E-R---S-S------S---T------K----L

S41A ------A------

S109A ------A------

S146A ------A------

S149A ------A------

S178A ------A------

T188V ------V------

E128A/R132A ------A-A------

K203A ------A------3.2.1 Analysis of viral protein profiles of CA mutant viruses

3.2.1.1 Viral proteins in virus producer cells

To analyse the expression of viral proteins by the CA mutant viruses, 293T cells were transfected with DNA encoding WT NL4-3, or each of the NL4-3 CA mutants. 48 hr post transfection, cells were lysed and total protein analysed for Gag proteins by SDS PAGE and western blot. Gag proteins were detected using two p24 CA specific antibodies, #6458 and

#4121 (NIH AIDS Research and Reference Reagent Program). These antibodies also detect the p55 Gag precursor and proteolytic cleavage intermediates. A major band of 24kDa was detected in cells expressing WT NL4-3, and all NL4-3 CA mutant viruses generated, with the exception of S109A. No protein was detected in mock transfected cells (Figure 3.3). Two minor bands at 55kDa and 37/41kDa were also detected, consistent with the p55 Gag precursor protein and p37/p41 proteolytic cleavage intermediates.

Using #6458 anti-CA antibody only a minor amount of 24kDa CA was detected in cells expressing the NL4-3 CA mutant S109A (Figure 3.3A). However CA proteins could be detected at 24kDa, 37/41kDa and 55kDa using an alternative anti-CA antibody (#4121). The level of p24 CA protein produced was lower than WT NL4-3 expressing cells (Figure 3.3B).

To improve S109A CA protein expression an alternative S109A CA mutant clone was generated with different codon usage (S109AKpnI). Transfection of cells with this infectious clone still yielded lower levels of S109A CA protein than the equivalent number of WT NL4-

3 or other CA mutant transfected cells (Figure 3.3C). For WT NL4-3 CA protein, anti-CA antibody #4121 did not detect p24 CA protein as efficiently as anti-CA antibody #6458. In contrast, detection of S109A was improved with anti-CA antibody #4121 compared with anti-

CA antibody #6458 (Figure 3.3C). Altered ability of the anti-CA antibody #6458 to recognise protein from this CA mutant virus suggests the amino acid substitution may interfere with substrate recognition by this antibody. However, as the epitopes these antibodies bind have not been mapped it is difficult to speculate about this possibility further.

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Figure 3.3 HIV Gag protein levels in transfected cell lysates 293T cells were transfected with HIV-1 NL4-3 WT or mutant plasmid as indicated. Cells were lysed at 48 hr post-transfection and normalised for total protein. 50 µg total protein was analysed by western blot with two monoclonal antibodies specific for p24 CA (A) #6458 and (B) #4121. (C) 50 µg total protein expressing S109A and S109AKpn CA mutants were compared with WT NL4-3. Lanes 1-4 were probed with anti-CA #6458, lanes 5-8 were probed with anti-CA #4121. C B A

NL4-3 WT NL4-3 anti - S109A CA #6458 S41A

S109AKpnI S109A anti anti No DNA S146A - - S149A CA #4121 CA #6458

S178A NL4-3 WT anti - S109A CA #4121 T188V

S109AKpnI E128A/R132A K203A No DNA No DNA p24 CA p55 Gag p55 Gag p24 CA p55 Gag p24 CA Creation of a third S109A mutant (S109AGCG) again with alternative codon usage still did not improve viral CA protein production in transfected cells (data not shown).

3.2.1.2 Viral proteins in isolated virions

Next, to analyse the incorporation of CA mutant proteins into virions, 293T cells were transfected with the respective infectious NL4-3 CA clones, and the culture medium was collected 48 hr post transfection, filtered and pelleted through 25% sucrose. Pelleted virus was then analysed by SDS PAGE and western blot using antibody specific for HIV-1 p24 CA

(Figure 3.4). Consistent with observations in cell lysates, two minor CA specific bands corresponding in size with p55 Gag and p37/p41 proteolytic cleavage intermediates, and the major p24 CA protein were detected in virus particles produced from cells transfected with

WT NL4-3 and each of the CA mutants, with the exception of S109A (Figure 3.4A). No viral

CA protein was detected in cell culture medium harvested from mock transfected cells.

Concentration (using Amicon centrifugal filtration devices) of a twice the amount of S109A virus prior to sucrose purification yielded CA specific bands of 55kDa and 24kDa, consistent with p55 Gag and p24 CA proteins. Again, S109A CA proteins were detected at much lower levels than WT NL4-3 virus (Figure 3.4B), consistent with a reduction in viral protein expression or impaired detection of protein expression due to antibody binding as previously observed in transfected cells (Figure 3.3).

Virions from all CA mutant viruses contained less unprocessed forms of Gag relative to the mature p24 CA protein compared with transfected cell lysates, reflecting processing of Gag during virus maturation.

3.2.1.3 Quantitation of p24 CA in culture medium

The level of p24 CA present in the cell culture supernatant was also quantitated by HIV-1 p24

ELISA (NEN™ Life Science Products, PerkinElmer™). All CA mutant constructs produced p24 CA at levels comparable to WT NL4-3, except S109A CA mutant virus (approximately

30% of WT virus); although again this may be artificially low due to inefficient antibody detection of the S109A CA (Figure 3.5). 73

Figure 3.4 HIV Gag protein levels in purified virus 293T cells were transfected with HIV-1 NL4-3 WT or CA mutant plasmid as indicated. Cell culture supernatant was collected at 48 hr post-transfection and quantitated by ELISA. Virus (100 ng p24) was concentrated and then purified through 25% sucrose and analysed by western blot with antibody specific for p24 CA (#6458) (A). WT NL4-3 and S109A virus containing supernatant (200 ng p24) was concentrated and then purified through 25% sucrose and analysed by western blot with antibody specific for p24 CA (#6458) (B). 100 ng virus purified from H3B cells was run as a positive control. B A

NL4-3 S41A

NL4-3 WT S109A anti S146A anti

Kpn - CA #6458 S109A #1 - S149A CA #6458

S109AKpn #2 S178A

T188V +ve control E128A/R132A p55 Gag

p24 CA K203A No DNA p24 CA p55 Gag Figure 3.5 Quantitation of p24 CA in transfection supernatants Virus-containing supernatants were collected 48 hr post-transfection and p24 CA quantitated by ELISA. Results presented are taken from 2 separate transfections, the values are presented as the average ± standard deviation. These are typical of trends in p24 levels between WT NL4-3 and CA mutant viruses used in all experiments. 600

500

400

300

p24 CA (ng/mL) 200 p24 CA p24CA (ng/mL) 100

0 NL4-3 S41A S109A S146A S149A S178A T188V Mock

CA mutant virus

CA mutant virus 3.2.2 Some CA mutant viruses have altered replication competence and reverse transcription ability

3.2.2.1 Replication competence of CA mutant virus in a single cycle infectivity assay

To investigate the replication competence of the CA mutant viruses in a single cycle infectivity assay, HeLa-CD4-LTR-β-gal reporter cells were infected with equivalent amounts of WT NL4-3 or CA mutant viruses (20 ng p24 by p24 ELISA). Infection was quantitated at

48 hr post infection (p.i.) by enumeration of blue cells following fixation and staining for β- galactosidase, or β-galactosidase colourimetric assay (Promega). β-galactosidase production demonstrated a comparable level of infection by WT NL4-3, S41A and S146A viruses, approximately 30-40% of WT NL4-3 infection by S178A and T188V viruses, while S109A and S149A showed near background levels of β-galactosidase activity (Figure 3.6). These results suggest that S41A and S146A mutant viruses replicate with comparable ability to WT

NL4-3 virus, while replication competence was reduced for S178A and T188V CA mutant viruses, and severely reduced for S109A and S149A CA mutant viruses in comparison to WT

NL4-3 virus. A reduction in β-galactosidase production in this single cycle infectivity assay is indicative of a block at, or prior to Tat activation of LTR transcription. Next, it was investigated if this block was mediated during reverse transcription.

3.2.2.2 Efficiency of reverse transcription by CA mutant viruses

To quantitate the ability of WT NL4-3 and CA mutant viruses to perform reverse transcription in infected cells, HuT-78 cells were infected (with 50 ng p24 by p24 ELISA) and at 0, 8 and

24 hr p.i. the cells were lysed and unintegrated DNA was isolated by phenol/chloroform extraction. Viral DNA in infected cell lysates was analysed by real-time PCR using primers specific for HIV-1 reverse transcription intermediates representing early (R-U5) and late

(PBS-Gag) stages of HIV-1 reverse transcription (Table 2.4). Mitochondrial DNA levels were also quantitated to normalise for cytoplasmic DNA input between samples. DNA values are expressed as a ratio of viral DNA: mitochondrial DNA.

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Figure 3.6 Replication competence of CA mutant viruses in a single cycle infectivity assay 2 x 104 HeLa-CD4-LTR-β-gal cells were infected with 20 ng p24 of virus or an equivalent amount of untransfected control cell supernatant. 48 hr following infection cells were lysed and β-galactosidase measured using the Beta-Red assay. Infections were performed in triplicate, the values are presented as the average ± standard deviation. Results are representative of two independent infections. 7 6 5 4 3

Beta-gal (mg/mL) 2

beta-gal (mg/mL) 1 0 NL4-3 S41A S109A S146A S149A S178A T188V Mock -3 41A 09A 46A 49A S 1 1 CA mutant1 virus T188 NL4 S S S S178A o DNA N To prevent false results due to viral plasmid DNA from the initial transfection, all virus stocks were filtered and DNase treated (2.6.2) to remove contaminating plasmid prior to infection.

Active replication was demonstrated by an increase in viral DNA copy numbers between 0-8 hr and a greater increase between 8-24 hr (Figure 3.7). At 0 hr p.i. some unintegrated HIV

DNA was observed (Figure 3.8), possibly due to intravirion DNA within virus inoculum that had absorbed to the outside of the cell.

To compare reverse transcription between CA mutant viruses, results are expressed relative to the amount of WT NL4-3 virus DNA intermediates at each time point. S149A showed levels of both early and late DNA products that were comparable to WT NL4-3 virus at all time points analysed (Figure 3.8). S109A showed similar amounts of early reverse transcription products at 0 hr p.i., possibly due to similar intravirion HIV DNA, but reduced levels of early products at 8-24 hr p.i. All other CA mutants (S41A, S146A, S178A and T188V) displayed a reduction in reverse transcription intermediates for both early and late products over the time course studied. These data demonstrate that reverse transcription is not affected by alanine substitution at position S149A. However, S109A CA mutant virus initiates reverse transcription but is blocked late during the process, while S41A, S146A, S178A and T188V

CA mutant viruses show a more severe defect prior to or during reverse transcription.

3.3 Discussion

In this chapter CA mutants S41A, S109A, S146A, S149A, S178A and T188V were analysed for viral CA protein expression, replication competence and reverse transcription. All CA mutant viruses, with the exception of S109A, demonstrated WT levels of viral protein expression and normal processing and packaging of p24 CA into virions. This indicates that the amino acid substitutions made in the CA region did not adversely affect virus production and processing.

S109A CA mutant virus showed low levels of viral CA protein production following transfection in both cell lysates and culture supernatant as determined by western blot and

ELISA. Generation of two further S109A CA mutant viruses that incorporated alternative 75

Figure 3.7 Increase in reverse transcription intermediates over 24 hr following infection of HuT-78 cells 2 x 106 HuT-78 cells were infected with 50 ng p24 of virus or an equivalent amount of mock transfected control cell supernatant. Early reverse transcription intermediates were analysed 0, 8 and 24 hr following infection. DNA copy numbers indicate the increase in viral DNA produced after infection. 250000

200000

150000

100000 DNA copy number DNA copy 50000

0 0hr 8hr 24hr Time post-infection

NL4-3 NoMock DNA Figure 3.8 Quantitation of reverse transcription ability of CA mutants 2 x 106 HuT-78 cells were infected with 50 ng of WT NL4-3 and CA mutant virus. Early (A) and late (B) reverse transcription intermediates were analysed 0, 8 and 24 hr following infection. Data has been taken from two separate experiments each comprising three infections. Values are presented as average ± standard deviation. Values represent viral DNA copies as a percentage of WT viral DNA copies at each time point. A

A

B

B codon usage for alanine resulted in similarly low levels of CA protein production. Detection of S109A CA protein by western blot with two different antibodies directed against HIV-1

CA or Gag proteins, or by p24 ELISA showed lower CA protein production than WT NL4-3 and all other CA mutant viruses. The reduction in S109A CA protein levels detected by western blot and p24 ELISA could be attributed to reduced antibody recognition. In support of this improved detection of S109A was achieved with anti-CA antibody #4121. However the level of S109A CA detected using anti-CA antibody #4121 was still reduced compared to

WT NL4-3 and other CA mutants, suggesting alanine substitution at this site may also affect virus production. Potential explanations for reduced CA production by S109A are discussed further in section 6.2.

Single cycle infectivity experiments using virus inoculum normalised by p24 CA levels demonstrated that three CA mutant viruses, S109A, S149A and S178A, were not replication competent, in line with previous literature (Cartier et al., 1999). Additionally, replication competence was reduced for T188V CA mutant virus. S41A and S146A showed comparable replication competence to WT NL4-3 virus in this assay, consistent with previously published results (Cartier et al., 1999).

The single cycle infectivity assay used in this study requires that virus complete cell entry, reverse transcription, integration, transcription and translation of Tat to activate the HIV LTR promoter and express β-galactosidase. Hence, the defect(s) observed in these three viruses must occur somewhere along this path. The reverse transcription intermediates produced by these viruses were analysed to examine if the defect in replication occurred during reverse transcription. CA mutant viruses demonstrating severe reductions in replication competence

(S109A, S149A, S178A and T188V) were all capable of reverse transcribing viral RNA and producing early and late HIV DNA intermediates.

For S109A virus, normal reverse transcription products were observed at early time points, but HIV DNA was reduced at later time points and S109A was not replication competent in a single cycle infectivity assay. This suggests that the S109A mutant is capable of initiating

76

reverse transcription at normal levels but during later time points reverse transcription, and hence replication, are blocked.

Reverse transcription intermediates from S178A and T188V CA mutant viruses were reduced at all time points analysed, suggesting a defect at or before reverse transcription, which could contribute to reduced replication competence in the single cycle infectivity assay.

For S41A and S146A CA mutant viruses, reverse transcription intermediates were severely reduced over the first 24hr. However these viruses demonstrated comparable replication competence to WT NL4-3 virus at 48 hr p.i. in the single cycle infectivity assay. This is consistent with previous reports that these CA mutant viruses exhibit delayed replication kinetics (Cartier et al., 1999) and suggests that while the number of reverse transcribing complexes or total yield of HIV DNA may be reduced, reverse transcription can produce enough complete DNA products capable of integration, transcription and translation, to achieve a similar signal to WT NL4-3 virus in the single cycle infectivity assay.

Alternatively, investigation of these mutants in a previous study suggests difference in infectivity may depend on cell type.

In contrast, the CA mutant virus S149A did not display a defect during reverse transcription, but was not replication competent in single cycle infectivity assays. This suggests that S149A displays a defect after reverse transcription but prior to Tat activation of LTR transcription.

Hence, the S149A CA mutant may represent a defect in the later stages of early HIV replication such as nuclear translocation of the PIC or integration. Studies have shown that blocks to nuclear translocation or integration have been associated with CA mutant viruses with altered core stability (Arhel et al., 2007; Dismuke & Aiken, 2006). CA assists nuclear translocation of the PIC through binding nuclear pore proteins (Bichel et al., 2013; Schaller et al., 2011; Shah et al., 2013). Mutation at S149A may interfere with restructuring of CA within the core and hence subsequent replication.

In summary, some CA mutations can block reverse transcription and may have a minor

(S41A, S146A) or major (S109A, S178A, T188V) effect on infectivity, or may act at a post-

77

reverse transcription stage of infection (S149A). How these CA mutations affect the early events in HIV infections is not yet obvious, but suggests that CA protein function, potentially via core disassembly is intrinsically linked to reverse transcription, nuclear translocation and ultimately infectivity. The effect of these CA mutations on core stability and core disassembly was assessed next (Chapter 4).

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CHAPTER 4 - In vitro analysis of core stability as a measure of core disassembly

4.1 Introduction

In this thesis the term ‘core’ is used to describe the structure that is defined by the conical p24

CA protein shell and includes the internal ribonucleoprotein complex enclosed within this shell. HIV-1 cores have been isolated and viewed by EM, and while most cores were cone- shaped or cylindrical, aberrantly shaped cores were often observed (Accola et al., 2000;

Briggs et al., 2003; Welker et al., 2000). EM demonstrates extreme variation in both the shape and size of cores, with widths ranging from 34.2-69.1 nm, length 80.9-151.3 nm and base angle ranging from 11.2-44.0º (Accola et al., 2000; Briggs et al., 2003; Welker et al.,

2000). Some virions contain multiple cores or empty cores without encapsidated HIV RNA.

Most critically, in the absence of proteolytic cleavage of Gag during maturation to produce a mature conical core, virions are not infectious.

The HIV-1 core structure exhibits a selectively maintained stability. It must be sufficiently stable to permit both orderly assembly and disassembly of the core at the appropriate stages of the HIV replication cycle. Amino acid substitution within p24 CA, the major core protein, can alter the stability of both individual CA protein monomers and cores derived from CA mutant viruses (Dismuke & Aiken, 2006; Forshey et al., 2002; Tang et al., 2001). Site-directed mutagenesis throughout the HIV-1 CA protein, including residues situated on the amino terminal domain (NTD) and carboxyl terminal domain (CTD) dimerisation interfaces, leads to changes in the kinetic energy and dimerisation affinity of the HIV-1 CA monomer. This indicates these substitutions are able to change properties, including the stability, of the p24

CA protein monomer (del Alamo & Mateu, 2005; del Alamo et al., 2003; Ganser-Pornillos et al., 2007). Additionally, substitution of CA residues can result in changes to stability of the resulting viral core. It is not unexpected that CA substitutions that alter in vitro core stability can also have detrimental effects on virus infectivity. Substitution of charged residues K203

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(in the CTD) and E128/R132 (in the NTD) with alanine within the CA protein, result in altered in vitro core stability (Dismuke & Aiken, 2006; Forshey et al., 2002; Hulme et al.,

2011). These defects were determined by measuring the amount of core-associated pelletable

CA following detergent exposure of virions and isopycnic ultracentrifugation (Forshey et al.,

2002). CA mutant virus K203A showed reduced amounts of detectable core-associated CA relative to WT virus. Cores from this virus were classified as hypostable. In contrast, core yields from CA mutant E128A/R132A were double those of WT virus and cores from this mutant virus are described as hyperstable (Forshey et al., 2002). The stability of cores from

CA mutant viruses were further analysed by quantitation pelletable core-associated CA after incubation under different pH, ionic strength or temperature conditions. This also indicated that cores from these CA mutant viruses behaved differently from WT virus cores. A greater yield of cores did not necessarily correspond with slower dissociation of cores, nor did low core yields always correspond with increased rates of core dissociation (Forshey et al., 2002).

However, in terms of function, replication of CA mutant viruses that displayed altered core properties compared to WT virus were all blocked at reverse transcription or nuclear translocation (Dismuke & Aiken, 2006; Forshey et al., 2002; Tang et al., 2003). This link between altered in vitro core stability and impaired replication in newly infected cells demonstrates that proper maintenance of the core structure and regulated core disassembly are essential for HIV infectivity.

Some of the CA mutant viruses (S109A, S149A, S178A and T188V) examined in Chapter 3 demonstrate a post entry block in HIV replication, either during reverse transcription (S109A,

S178A, T188V) or post reverse transcription (S149A). During early HIV infection, the main function of the p24 CA protein is to form the HIV core. In this study, it was hypothesised that the blocks to early infection shown by CA mutant viruses may relate to CA function in the

HIV core such as core stability or regulation of core disassembly. Following rearrangement of the iconic fullerene shaped core en route to the nucleus, CA in the RTC/PIC is implicated in the interactions with nuclear pore proteins that are critical for nuclear import and subsequent

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integration (Bichel et al., 2013; Schaller et al., 2011; Shah et al., 2013). Thus in turn, impaired core disassembly could impact reverse transcription and nuclear translocation.

Each of the CA serine-alanine substitution mutant viruses, S109A, S149A and S178A, examined in Chapter 3 removes a site of CA phosphorylation (Cartier et al., 1999). The addition or subtraction of a negatively charged phosphate group has the potential to change the overall charge of the CA protein monomer and alter protein-protein interactions that may be important in maintaining the optimal stability of the HIV-1 virus core. In silico modelling suggests phosphorylation of S149 for example, creates sufficient repulsion to induce disassembly of the conical core (Giroud et al., 2011).

In this chapter, in vitro core stability was examined for CA mutant viruses S109A, S149A or

S178A, which prevent phosphorylation, and also S41A, S146A and T188V. This required development of an assay to analyse in vitro core stability of these CA mutant viruses. In order to distinguish between WT, hyperstable and hypostable mutant virus cores; previously described ‘spin-thru’ ultracentrifugation methods were modified and optimised using the CA mutant viruses K203A (hypostable) and E128A/R132A (hyperstable) as controls (Accola et al., 2000; Forshey & Aiken, 2003; Forshey et al., 2002; Kotov et al., 1999; Ohagen &

Gabuzda, 2000; Scholz et al., 2005; Stromberg, 1972; Stromberg et al., 1974; Welker et al.,

2000). The detergent conditions defined were used to analyse the in vitro core stability of CA mutant viruses characterised in Chapter 3.

4.2 Results

4.2.1 Methods to identify core structures.

Several factors need to be considered for the separation of viral components by ultracentrifugation. These include (i) the preparation of virus samples to be analysed, (ii) the design of centrifugation and gradient used (i.e. isopycnic versus rate zonal

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ultracentrifugation), (iii) the sedimentation parameters (force, duration and temperature during centrifugation) and (iv) the detergent and salt conditions used in the spin-thru layer.

4.2.1.1 Preparation and purification of virus samples.

NL4-3 virus was produced by transfection of 293T cells (2.6.1) and harvesting supernatant 48 hr post transfection. Virus-containing supernatant was filtered through 0.2 µm filters and concentrated using Amicon (Millipore) or Sartorius centrifugal filter devices (2.6.3)

(concentrated virus). Virus preparations were further purified by pelleting concentrated virus through a 25% sucrose cushion (2.6.4). Pelleted virus was washed and resuspended in TNE buffer (concentrated purified virus). 3.5µg p24 CA as determined by ELISA (approximately

5-10 mL virus containing supernatant), was sedimented by rate zonal ultracentrifugation on sucrose gradients. The appropriate preparation of virus was essential to observe detergent disruption of virus or cores when using the spin-thru centrifugation method. Using 1.5%

Triton-X-100 in a spin-thru procedure, virions from concentrated purified WT virus were effectively disrupted. This resulted in complexes containing CA presumed to be cores and free CA. Virus was not disrupted in this amount of detergent if virus was concentrated but not purified (Figure 4.1). Analysis of concentrated virus samples that were not purified resulted in similar CA distribution in the gradient. This profile was observed regardless of the presence or absence of up to 3% Triton-X-100 detergent using the spin-thru layer method, or by incubation of virus separately in detergent prior to sedimentation (data not shown). Thus the additional sucrose cushion purification step was necessary to achieve tight sedimentation profiles of CA protein across the sucrose gradient and effective detergent treatment of virions.

It is presumed that pelleting virus through sucrose removes contaminating proteins (e.g. serum) from the cell culture media used during production of virus stocks. As a result, detergent activity is not saturated by these contaminating proteins and can interact more effectively with virus. Therefore, virus was concentrated, pelleted through sucrose, washed and resuspended in TNE buffer immediately before ultracentrifugation in all subsequent experiments.

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Figure 4.1 In vitro core stability assays investigating the effect sucrose purification of virus samples on WT NL4-3 virus sedimentation Concentrated, or concentrated and sucrose purified virus was spun-thru 0% or 1.5% Triton-X-100 onto 15-40% linear sucrose gradients overlaid onto a 60% sucrose cushion. Ultracentrifugation was performed at 35000 rpm for 60 min in a Beckman SW-42Ti centrifuge (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24 CA. Fraction 1 represents the top of the gradient, fraction 11 represents the bottom of the gradient. The percentage sucrose of each fraction was determined by measuring the refractive index (B). A Concentrated virus Concentrated & pelleted virus 1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 1.5 % Triton-X-100

50

B 45

40

35

30

25

% sucrose 20

15

10

5

0 1 2 3 4 5 6 7 8 9 10 11 Fraction

Conc 0% Triton-X-100 x Pellet 0% Triton-X-100 Conc Conc 1.5% Pellet Pellet 1.5% Conc 1.5% Triton-X-100 x Pellet 1.5% Triton-X-100 4.2.1.2 Centrifugation Method

4.2.1.2.1 Isopycnic Centrifugation

Initial attempts were made to isolate HIV-1 cores using the isopycnic ultracentrifugation methods that were published by Forshey et al (2002) (1% Triton-X-100 spin-thru layer, 30-

65% linear sucrose gradient, 100000 x g, 16 hr, 4ºC) (Forshey et al., 2002). Concentrated purified virus was analysed by spin-thru isopycnic ultracentrifugation in the presence or absence of detergent, followed by detection of CA protein in gradient fractions by western blot and ELISA. In the absence of detergent, CA protein sedimented to fraction 4 within the gradient (1.12-1.16g/mL sucrose), consistent with the density of intact HIV-1 virions (Figure

4.2). In the presence of 0.05% Triton-X-100 detergent (Figure 4.2), or 1% Triton-X-100 (data not shown) as used by Forshey et al (2002), no CA protein was detected in fractions at the density consistent with HIV-1 cores (1.24-1.28g/mL, fractions 7-9). Thus alternative gradient systems were investigated.

4.2.1.2.2 Rate Zonal Centrifugation

Next, the separation of HIV-1 cores was investigated utilising the detergent spin-thru method coupled with rate zonal ultracentrifugation through 15-40% sucrose gradients on top of a 60% sucrose cushion (Figure 4.3). Triton-X-100 detergent (0-3%) was diluted in sucrose and laid above pre-formed linear 15-40% gradients. Sucrose solutions were made in TNE (2.1.6) to keep pH levels and salt concentrations optimal for core recovery (Forshey et al., 2002;

Welker et al., 2000). Virus was prepared as in section 4.2.1.1. Fractions were collected from the top of the gradient (fractions 1-12 represent low to high percentage sucrose). The distribution of CA protein in gradient fractions was analysed by SDS PAGE and western blot.

4.2.1.3 Rate zonal centrifugation parameters

4.2.1.3.1 Force and duration of centrifugation

To determine optimal centrifugation conditions to separate cores from free CA protein, concentrated and purified virus was sedimented in the absence of detergent on 15-40%

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Figure 4.2 Isolation of virus and cores by isopycnic ultracentrifugation Purified virus was spun-thru 0% or 0.5% Triton-X-100 onto 10 mL 30-65% linear sucrose gradients at 24100 rpm for 18 hr. Fractions (1 mL) were collected from the top of the gradient and analysed for p24 CA by ELISA. The amount of CA in each fraction is indicated on the primary y-axis and density of each fraction is indicated on the secondary y-axis. 16000 1.3

14000 1.28 1.26 12000 1.24 10000 1.22 8000 1.2

6000 1.18 Density

p24 CAp24(pg/mL) 1.16 4000 1.14 2000 1.12 0 1.1 1 2 3 4 5 6 7 8 9 10 11 Fraction WT0.5% 0% TritonTriton-X-100-X-100 WT0% 0.5% Triton-X-100 Triton-X-100 Density Density Figure 4.3 Schematic representation of rate-zonal sucrose gradients used to assess in vitro core stability Gradients were prepared in 12.5 mL ultracentrifuge tubes and contained a 0.5 mL 60% sucrose cushion at the bottom of the tube. A 10 mL 15-40% linear gradient was topped with a 0.5 mL detergent spin-thru layer made up of Triton- X-100 in 15% sucrose. Concentrated and pelleted virus was loaded on to the top of the gradient immediately before ultracentrifugation. Centrifugation was performed at 35000 rpm for 40 min in a Beckman SW-42Ti centrifuge. Pelleted virus (0.2mL) Top Triton- X-100 (0.5mL) (fraction 1)

15-40% sucrose gradient (10mL)

Bottom 60% sucrose (0.5mL) (fraction 12) sucrose gradients at 35000 rpm for 40 min, 50 min or 60 min at 4ºC in a Beckman SW-42Ti centrifuge. The distribution of CA protein across the gradient was analysed by SDS PAGE and western blot. Sedimentation for 40 min resulted in optimal distribution of CA across the gradient, allowing differentiation between dissociated free CA protein at the top of the gradient and CA-containing structures in the middle of the gradient (data not shown).

Accumulation of CA protein at the bottom of the gradient was observed when gradients were centrifuged for longer than 40 min. All further gradients were sedimented for 40 min.

4.2.1.3.2 Temperature

Due to the intrinsic instability of mature HIV-1 cores, all sucrose solutions and gradients were prepared and stored at 4ºC, with the aim of minimising protein degradation during preparation and subsequent ultracentrifugation of virus. However, Triton-X-100 is a non-ionic detergent and works more efficiently at higher temperatures. To assess the affect of temperature on the efficiency of detergent treatment, purified virus was sedimented through Triton-X-100 in the spin-thru layer (concentrations ranging from 0-1%) at either 4ºC or 25ºC (room temperature).

The increase in temperature resulted in tighter banding of particulate CA (data not shown).

Therefore gradients were prepared and centrifuged at 25ºC. An exception was made for experiments investigating hypostable (K203A) cores which were performed at 4ºC to reduce potential premature core dissociation.

4.2.1.4 Detergent Conditions

4.2.1.4.1 Detergent-free sucrose “barrier” layer

To minimise virus exposure to detergent, some groups utilised a layer of sucrose free of detergent as a barrier between the virus and the underlying spin-thru layer containing detergent (Kewalramani & Emerman, 1996; Kotov et al., 1999; Warrilow et al., 2008). A detergent-free low concentration sucrose barrier layer (5-10%) was added above the spin thru detergent layer containing Triton-X-100 in 15% sucrose at the top of the gradient. However, even when low concentration sucrose was used in the barrier layer (e.g. 10% as used by

84

Kotov et al (1999)) this sucrose was visually observed to deposit under the detergent layer.

Thus regulated exposure of virus to detergent could not be achieved in this manner and the use of the extra barrier layer between the detergent and virus was not pursued further.

To minimise the extended exposure of virus to detergent, virus was added to gradients immediately before ultracentrifugation.

4.2.1.4.2 Detergent concentration in the spin-thru layer

A study by Forshey et al (2002) utilised the spin-thru technique in combination with isopycnic ultracentrifugation to remove the viral envelope, yet still allow identification of core structures based on density (Forshey et al., 2002). Published methods used Triton-X-100 concentrations of up to 1% (Forshey et al., 2002; Kewalramani & Emerman, 1996; Kotov et al., 1999; Yu et al., 1993). However, in this study sedimentation of virus through 1.5%

Triton-X-100 resulted in almost complete loss of core structures, with the majority of CA detected as free protein sedimenting at the top of the gradient (Figure 4.1). This loss of cores indicates 1.5% Triton-X-100 detergent treatment was too harsh. In subsequent experiments concentrations between 0.01-1% Triton-X-100 were utilised in the spin-thru layer (as indicated in figures).

4.3 Analysis of detergent treated virus by rate zonal ultracentrifugation

4.3.1 Increasing detergent concentration results in release of free CA protein from core structures

To determine conditions that could separate free CA protein and virus cores, WT NL4-3 virus was sedimented through a range of detergent concentrations and the distribution of CA protein throughout the gradient was analysed by SDS PAGE and western blot of gradient fractions.

In the absence of detergent, CA protein sedimented to gradient fractions 7-9 (Figure 4.4) representing the sedimentation profile of intact virus. Increasing detergent concentrations

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Figure 4.4 In vitro core stability assay demonstrating the effect of increasing concentrations of Triton-X-100 detergent on WT NL4-3 virus WT NL4-3 virus was concentrated and pelleted through 25% sucrose before ultracentrifugation through 0%, 0.1%, or 1.0% Triton-X-100 onto 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24 CA. The percentage sucrose of each fraction was determined by measuring the refractive index (B). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. WT NL4-3 A 1 2 3 4 5 6 7 8 9 10 11 12 0% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 0.1% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 1.0% Triton-X-100

50 B 45 40 35 30 25

Sucrose 20 % % 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

WT 0% Triton-X-100 WT0% 0.1%0.1% Triton-X-1001.0% x WT 1.0% Triton-X-100 between 0-1% Triton-X-100 resulted in an increased proportion of CA protein in the fractions at the top of the gradient, indicative of increased dissociation of CA from larger, faster sedimenting complexes.

At 0.1% and 1.0% Triton-X-100, the majority of CA sedimented within the top half of the gradient as free protein, in fractions 2-7 and 1-6 respectively. However, the peak of CA protein within these fractions was distinctly different; shifting from fractions 3-4 for 0.1%

Triton-X-100, to fractions 1-2 when exposed to 1% Triton-X-100 (Figure 4.4). This demonstrated that the increase in detergent caused a shift in the peak of CA protein.

The results from different detergent concentrations are summarised in Figure 4.5. Due to the nature of these experiments, some variability was observed between gradients runs. The

Triton-X-100 treatments chosen were those deemed to be the most reproducible. An incremental increase of 0.01% Triton-X-100 at a low concentration (i.e. from 0-0.03% Triton-

X-100) made a considerable impact on the profile of low and high molecular mass complexes containing CA. Whereas between 0.1-1.0% Triton-X-100, the impact on CA profiles was difficult to detect even with incremental increases of 0.1% Triton-X-100. These experiments indicate that treatment using concentrations of detergent greater than 0.1% Triton-X-100 is sufficient to disrupt WT NL4-3 cores.

Next, I aimed to verify if the faster sedimenting complexes containing CA represented virions or cores based on the presence of the HIV envelope protein.

4.3.2 Development of conditions to remove the viral envelope.

Addition of detergent in the spin thru layer at the top of the gradient aimed to remove the outer viral envelope (gp120), while leaving the internal core (the p24 CA shell and its contents) of the virus intact. Virions and cores can be identified by their density when separated on isopycnic gradients. However, the use of rate zonal centrifugation required another characteristic to distinguish between virus and cores. To differentiate between intact virions and cores on the gradient, gradient fractions were also analysed by SDS PAGE and western blot or ELISA using antibody specific for the viral envelope glycoprotein gp120, and

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Figure 4.5 CA distribution profiles following in vitro core stability assays WT NL4-3 virus was concentrated and pelleted through 25% sucrose before rate zonal ultracentrifugation spun-thru 0-1.0% Triton-X-100 onto 15-40% sucrose gradients at 35000 rpm for 40 min as indicated. Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for CA (#6458). The fractions containing the majority of CA at each concentration of Triton-X- 100 from multiple experiments (n = 1-5 for each concentration) are indicated on the graph. 1 0.2 0.1 0.08 0.05 0.04

0.03 % Triton-X-100 0.02 0.01 0 1 2 3 4 5 6 7 8 9 10 11 12

Fraction compared with p24 CA protein sedimentation. Only minor levels of gp120 could be detected in the absence of detergent using either western blot or ELISA. This was not unexpected since

HIV-1 virions possess very few (4-35, averaging 14) envelope spikes per virion (Zhu et al.,

2006), compared with approximately 2000 CA monomers per virion (Briggs et al., 2004). In the presence of detergent (0.03% Triton-X-100) gp120 was faintly detected only in the top fraction, while in contrast p24 CA was detected in the middle fractions (Figure 4.6). At concentrations of Triton-X-100 above 0.03% p24 CA begins to shift to the top fractions, suggesting that detergent treatment under these conditions promotes release of the viral envelope.

4.3.3 0.1% Triton-X-100 distinguishes between WT and hyperstable virus

Next, detergent treatment and sedimentation conditions were required that would allow differentiation of WT NL4-3, hyperstable and hypostable CA mutant virus cores. Previous studies have shown CA mutant viruses E128A/R132A and K203A produce hyperstable and hypostable cores respectively (Dismuke & Aiken, 2006; Forshey et al., 2002; Hulme et al.,

2011). WT NL4-3 virus and hyperstable CA E128A/R132A mutant virus stocks were prepared as described previously (4.2.1.1) and analysed on rate zonal sucrose gradients using the spin-thru method with varying concentrations of detergent. Gradient fractions were analysed for p24 CA by SDS PAGE and western blot.

We found it helpful to examine the gradient profiles in thirds. Some variability was observed in CA profiles between runs, and there was often a small shift in the fractions that CA was detected in. However, overall the CA profiles produced at each condition were similar.

Fractions 1-4 represent the top of the gradient. CA present in those fractions is likely to present free CA released from higher molecular mass complexes. Fractions 5-8 represent the middle of the gradient. This is where cores and intact virus are expected. While fractions 9-12 represent the bottom of the gradient. CA detected at the bottom of the gradient is mostly non- specific high molecular mass CA. By examining the CA present in each third of the gradient,

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Figure 4.6 In vitro core stability assay demonstrating the effect of detergent on WT virus envelope and core stability WT NL4-3 virus was concentrated and pelleted through 25% sucrose before ultracentrifugation through 0.03% Triton-X-100 onto 15-40% sucrose gradients at 35000 rpm for 40 min. Fractions (1 mL) were collected from the top of the gradient and analysed for gp120 and p24 CA by western blot (A) or ELISA (B). The percentage sucrose of each fraction was determined by measuring the refractive index (C). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. A WT NL4-3 1 2 3 4 5 6 7 8 9 10 11

gp120 0.03% Triton-X-100 CA p24

B

C 50 45 40 35 30 25 20 15 % Sucrose% 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction WT 0.03% Triton-X-100 it is easier to identify differences in the CA profiles between WT virus and CA mutant viruses.

In the absence of detergent, WT NL4-3 and E128A/R132A produced similar sedimentation profiles. CA protein from both viruses was detected in the middle of the gradient (fractions 6-

9) representing intact virions (Figure 4.7). Spin-thru sedimentation through 0.5% and 1%

Triton-X-100 resulted in comparable profiles for WT NL4-3 and hyperstable E128A/R132A virus; the peak of CA present at the top of the gradient (fractions 1-3) indicated disruption of cores from both viruses under these conditions (data not shown).

However, when detergent in the spin-thru layer was reduced to 0.1% Triton-X-100 differences were observed between the CA profiles for WT NL4-3 and E128A/R132A hyperstable viruses. Following sedimentation of WT NL4-3 virus the majority of CA protein showed a noticeable shift to the top fractions (fractions 1-4), consistent with the release of CA from cores to free protein (Figure 4.7). In contrast, the majority of CA protein from the

E128A/R132A hyperstable CA mutant again sedimented to the middle of the gradient

(fractions 5-9) and only a minor amount of CA protein sedimented in the top four fractions.

Therefore a spin-thru rate zonal sucrose sedimenting system with 0.1% Triton-X-100 was able to differentiate CA protein profiles from WT NL4-3 and E128A/R132A viruses.

4.3.4 WT and hypostable virus cores could not be distinguished using rate zonal ultracentrifugation

Similarly, conditions were required to distinguish WT NL4-3 virus and the hypostable CA mutant virus K203A. Virus was produced from WT NL4-3 and hypostable CA mutant K203A and sedimented through a 0.1% Triton-X-100 layer as described in 4.3.3. For both viruses, the majority of CA protein was detected by western blot in the top gradient fractions, indicating these conditions disrupt cores from both WT NL4-3 and K203A virus (data not shown). To identify conditions that could dissociate the hypostable virus core, but would leave WT NL4-

3 virus cores intact, the concentration of detergent in the spin-thru layer was reduced until CA protein sedimentation profiles were similar in both the presence and absence of detergent.

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Figure 4.7 In vitro core stability assay to distinguish WT NL4-3 and E128A/R132A hyperstable mutant virus Virus was concentrated and pelleted through 25% sucrose before rate zonal ultracentrifugation ‘spun-thru’ 0% or 0.1% Triton-X-100 onto 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24 CA. The percentage sucrose of each fraction was determined by measuring the refractive index using a refractometer and is indicated on the graph (B). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. HYPERSTABLE A WT NL4-3 E128A/R132A 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0.1% Triton-X-100

50 B 45 40 35 30 25

20 % Sucrose % 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

x WT 0% WT 0%WT Triton 0.1%-X-100E128A/R132A E128A/R132A 0% E128A/R132A 0% Triton-X-100 0.1% WT 0.1% Triton-X-100 x E128A/R132A 0.1% Triton-X-100 Spin-thru sedimentation of WT NL4-3 virus through 0.01% or 0.02% Triton-X-100 had little effect on CA protein sedimentation in comparison to no detergent at all (Figure 4.8), with CA detected in fractions 5-8 and fractions 5-7 respectively. Similarly, hypostable mutant K203A spun-thru 0.01-0.02% Triton-X-100 also produced CA sedimentation profiles comparable to those observed in the absence of detergent. Thus 0.01-0.02% Triton-X-100 was not sufficient to disrupt either WT NL4-3 virus or hypostable K203A mutant cores. When WT NL4-3 and

K203A virus were spun-thru 0.03% Triton-X-100, both viruses showed a marked shift in the distribution of CA protein, with the majority of CA protein sedimenting at the top of the gradient representing free protein. This indicated disruption of both WT NL4-3 and K203A cores in 0.03% Triton-X-100.

Although the proportion of CA detected in top fractions (1-3) versus middle fractions (6-7) were slightly different between WT NL4-3 and K203A, the differences were subtle and not definitive. Detergent conditions could not be determined that disrupted K203A cores, but not

WT NL4-3 cores. Even when virus was treated as gently as possible, WT NL4-3 and K203A showed similar CA distribution. This suggests K203A cores display WT core stability in this in vitro assay.

4.4 Analysis of core stability of CA mutants

4.4.1 CA mutants S41A, S109A, S146A, S149A, S178A and T188V are not hyperstable

Next, the in vitro core stability of the CA mutants generated in Chapter 3 that contained substitutions at phosphorylation sites (S109A, S149A, S178A) or additional sites (S41A,

S146A, T188V) were examined. Rate zonal gradient analysis was performed using the conditions determined in section 4.3.3. Virus was prepared from the CA mutant viruses and analysed in the absence of Triton-X-100 or following exposure to 0.1% Triton-X-100 in the spin-thru layer.

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Figure 4.8 In vitro core stability assay to distinguish WT NL4-3 and K203A hypostable mutant virus Virus was concentrated and pelleted through 25% sucrose before ultracentrifugation through 0.01%, 0.02% or 0.03% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. A WT NL4-3 HYPOSTABLE K203A 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0.01% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0.02% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0.03% Triton-X-100

B 50 45 40 35 30 25

20 % Sucrose 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

WTWT 0.01%0.01% Triton-X-100WT x0.02% K203A 0.01% WTTriton 0.03%-X-100 WTK203A 0.02% 0.01% Triton-X-100K203A 0.02% K203A 0.02% K203ATriton- X0.03%-100 x WT 0.03% Triton-X-100 K203A 0.03% Triton-X-100 In the absence of detergent, the majority of CA protein from each CA mutant virus (S41A,

S146A, S149A, S178A, T188V) was present in the middle of the gradient (Figure 4.9-4.11), consistent with WT NL4-3 virus profiles without detergent (Figure 4.7). A minor proportion of CA was observed in the very top fraction for some CA mutant viruses (e.g. S178A), although this was also occasionally observed for WT NL4-3 virus. When exposed to 0.1%

Triton-X-100, the majority of CA protein from these viruses appeared in the top gradient fractions (1-3), the same as for WT NL4-3 virus. This indicates these CA mutants show similar detergent susceptibility compared to WT NL4-3 and were not hyperstable. The absence of p24 CA present in the middle of the gradient for S109A, S149A and S178A viruses may suggest an increased sensitivity to this detergent treatment.

Due to earlier difficulties detecting S109A p24 CA protein by western blot (section 3.2.1),

S109A sedimentation profiles were analysed by p24 ELISA. In the absence of detergent, a broad peak of CA protein was detected in fractions 6-8 (Figure 4.12). When exposed to 0.1%

Triton-X-100 the peak of CA protein shifted to fractions 1-4 at the top of the gradient. Thus, similar to the other CA mutant viruses analysed, S109A CA mutant cores are not hyperstable.

4.4.2 CA mutants exhibit WT (S41A, S146A, T188V) or hypostable (S109A, S149A, S178A) cores

Although the K203A mutant that had been generated as a hypostable control could not be distinguished from WT NL4-3 virus (4.3.4), the CA mutants (S41A, S109A, S146A, S149A,

S178A and T188V) were also analysed under lower detergent conditions. Their susceptibility to these lower detergent concentrations was then compared to WT NL4-3 virus to identify viruses potentially displaying hypostable cores. As previously described, virus was generated, purified and the CA mutant viruses were analysed by rate zonal ultracentrifugation on sucrose gradients with a layer of 0.01%, 0.02% or 0.03% Triton-X-100 at the top of the gradient.

Following sedimentation of S41A, S146A, S149A, S178A and T188V CA mutants or WT

NL4-3 virus through 0.01% Triton-X-100, analysis of gradient fractions by SDS PAGE and western blot demonstrated that the majority of CA protein banded in the middle of the

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Figure 4.9 In vitro core stability assay to investigate if S149A and S178A CA mutant viruses contain hyperstable cores CA mutant viruses were concentrated and pelleted through 25% sucrose before ultracentrifugation through 0% or 0.1% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. A S149A S178A

1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0.1% Triton-X-100

50 B 45 40 35 30 25

20 % Sucrose % 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

x S149A S149A 0% 0% TritonS149A-X-100 0.1% S178AS178A 0%0% TritonS178AS149A-X-100 0.1% S149A 0.1% Triton-X-100 x S178A 0.1% Triton-X-100 Figure 4.10 In vitro core stability assay to investigate if S41A and S146A CA mutant viruses contain hyperstable cores CA mutant viruses were concentrated and pelleted through 25% sucrose before ultracentrifugation through 0% or 0.1% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. A S41A S146A 1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 1 2 3 4 5 6 7 8 9 10 11 12 0.1% Triton-X-100

B 50 45 40 35 30 25 20 15 % Sucrose% 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction S41A 0% Triton-X-100 S146A 0% Triton-X-100 S41A 0.1% Triton-X-100 S146A 0.1% Triton-X-100 Figure 4.11 In vitro core stability assay to investigate if investigate T188V CA mutant virus contains hyperstable cores CA mutant viruses were concentrated and pelleted through 25% sucrose before ultracentrifugation through 0% or 0.1% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). Note if the volume of the final fraction (e.g. fraction 12) was less than 1 mL a refractive index measurement was not taken. A T188V 1 2 3 4 5 6 7 8 9 10 11 12 0% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 0.1% Triton-X-100

B 50 45 40 35 30 25 20 15 % Sucrose% 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction T188V 0% Triton-X-100 T188V 0.1% Triton-X-100 Figure 4.12 In vitro core stability assay to investigate if S109A CA mutant virus contains hyperstable cores S109A CA mutant virus was concentrated and pelleted through 25% sucrose before ultracentrifugation through 0% or 0.1% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by CA p24 ELISA. The percentage sucrose of each fraction was determined by measuring the refractive index (B). A 8000 7000 6000 5000 4000

3000 p24 pg/mL p24 2000 1000 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

S109A 0%0% Triton Triton-X-100-X-100 S109A0.1% Triton-X-100 0.1% Triton-X-100

B 50 45 40 35 30 25

20 % Sucrose % 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

S109A 0% TritonS109A-X -0%100 S109A S109A 0.1% 0.1% Triton-X-100 gradient (Figure 4.13-4.15). Sedimentation through 0.01-0.02% Triton-X-100 resulted in the release of a proportion of CA protein that sedimented towards the top of the gradient. In most viruses, CA was also still present in the middle of the gradient. Finally, sedimentation through

0.03% Triton-X-100 resulted in the detection of the majority of CA in the top fractions. While the proportion was small, some CA remained in the middle of the gradient these CA mutant viruses (S41A, S146A, T188V) indicating that these viruses display similar detergent sensitivity to WT NL4-3 virus in this assay.

Following sedimentation through 0.03% Triton-X-100 the majority of CA from S149A and

S178A was present in the top four fractions. CA was mostly absent from the middle of the gradient for these viruses. Additionally, elevated amounts of CA were present in the top fractions at each detergent concentration from 0.01-0.03% Triton-X-100. This contrasts sedimentation of WT NL4-3 virus under these conditions. This suggests S149A and S178A display a decrease in in vitro core stability compared with WT NL4-3 virus.

Once again due to limited detection of S109A CA by western blot analysis, this virus was analysed by p24 ELISA (Figure 4.16). S109A demonstrated similar results to S149A and

S178A at each of the detergent concentrations tested. When S109A virus was sedimented through 0.01% or 0.02% Triton-X-100 a large proportion of CA was detected as free protein at the top of the gradient and a peak of CA was observed in fractions 6-8 in the middle of the gradient. When S109A was sedimented through 0.03% Triton-X-100, CA was detected at the top of the gradient in fractions 1-3. The peak of CA in the middle of the gradient was not observed at this concentration of detergent. These results suggest that S109A also shows some hypostability in comparison to WT NL4-3 virus under these detergent conditions.

Although the lack of a hypostable core control made it difficult to determine hypostability relative to WT NL4-3 virus with confidence, the analysis of in vitro core stability showed CA mutants S41A, S146A and T188V demonstrated comparable stability to WT NL4-3 virus, while S109A, S149A and S178A CA mutant viruses were more susceptible to low levels of

Triton-X-100 detergent than WT NL4-3 virus.

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Figure 4.13 In vitro core stability assay to investigate if S149A and S178A CA mutant viruses contain hypostable cores CA mutant viruses were concentrated and pelleted through 25% sucrose before ultracentrifugation through 0.01%, 0.02% or 0.03% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). A S149A S178A 1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0.01% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0.02% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0.03% Triton-X-100

B 50 45 40 35 30 25 20 % Sucrose% 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction

S149AS149A 0.01% 0.01% Triton-X-100S149A x 0.02% S178A 0.01%S149A Triton 0.03%-X-100 S149AS178A 0.02% 0.01% Triton-X-100S178A 0.02% S178A 0.02%S178A Triton 0.03%-X-100 x S149A 0.03% Triton-X-100 S178A 0.03% Triton-X-100 Figure 4.14 In vitro core stability assay to investigate if S41A and S146A CA mutant viruses contain hypostable cores CA mutant viruses were concentrated and pelleted through 25% sucrose before ultracentrifugation through 0.01%, 0.02% or 0.03% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). A S41A S146A 1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0.01% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0.02% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 1 2 3 4 5 6 7 8 9 10 11 0.03% Triton-X-100

50 B 45 40 35 30 25 20 15 % Sucrose% 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction S41A 0.01% Triton-X-100 S146A 0.01% Triton-X-100 S41A 0.02% Triton-X-100 S146A 0.02% Triton-X-100 S41A 0.03% Triton-X-100 S146A 0.03% Triton-X-100 Figure 4.15 In vitro core stability assay to investigate if T188V CA mutant virus contains hypostable cores CA mutant viruses were concentrated and pelleted through 25% sucrose before ultracentrifugation through 0.01%, 0.02% or 0.03% Triton-X-100 into 15- 40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by western blot specific for p24. The percentage sucrose of each fraction was determined by measuring the refractive index (B). T188V A 1 2 3 4 5 6 7 8 9 10 11 12 0.01% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 0.02% Triton-X-100

1 2 3 4 5 6 7 8 9 10 11 12 0.03% Triton-X-100

B 50 45 40 35 30 25 20 15 % Sucrose% 10 5 0 1 2 3 4 5 6 7 8 9 10 11 Fraction T188V 0.01% Triton-X-100 T188V 0.02% Triton-X-100 T188V 0.03% Triton-X-100 Figure 4.16 In vitro core stability assay to investigate if S109A CA mutant virus contains hypostable cores S109A CA mutant virus was concentrated and pelleted through 25% sucrose before ultracentrifugation through 0.01%, 0.02% or 0.03% Triton-X-100 into 15-40% sucrose gradients at 35000 rpm for 40 min (A). Fractions (1 mL) were collected from the top of the gradient and analysed by ELISA specific for p24 CA. The percentage sucrose of each fraction was determined by measuring the refractive index (B). A 8000 7000 6000 5000 4000

3000 p24 pg/mL p24 2000 1000 0 1 2 3 4 5 6 7 8 9 10 11 12 Fraction

S109A 0.01% Triton-X-100 S109A 0.02% Triton-X-100 S109A 0.01% x S109AS109A 0.03% 0.02% Triton-X-100S109A 0.03%

50 B 45 40 35 30 25

20 % Sucrose % 15 10 5 0 1 2 3 4 5 6 7 8 9 10 11 12 Fraction

S109AS109A 0.01% 0.01% Triton-X-S109A100 0.02% S109A 0.02%S109A Triton 0.03%-X-100 x S109A 0.03% Triton-X-100 4.5 Discussion

This chapter aimed to establish an in vitro assay to investigate the stability of the HIV-1 core.

There are inherent difficulties analysing core stability in infected cells, primarily due to the low number of infecting HIV-1 particles and the labile nature of intracellular virus. Hence previous studies have examined the stability of cores derived from virions in vitro to reflect core disassembly within the infected cell (Auewarakul et al., 2005; Forshey et al., 2002;

Wacharapornin et al., 2007).

In the present study, core stability of WT NL4-3 and CA mutant viruses (S41A, S109A,

S146A, S149A, S178A and T188V) were compared during rate zonal ultracentrifugation combined with the spin-thru technique. Optimal sample preparation and centrifugation conditions were determined for analysis of core stability. The concentration of detergent in the spin-thru layer was reduced considerably (≤ 0.1% Triton-X-100) from concentrations described in methods in early literature (Forshey et al., 2002; Kewalramani & Emerman,

1996; Kotov et al., 1999; Yu et al., 1993). Results in this thesis demonstrate that the virus core was completely disrupted in concentrations of detergent greater than 0.03% Triton-X-

100. This is consistent with a publication that reported 0.03% Triton-X-100 had marked affects on virus structure, resulting in the disruption of the intravirion RTC, altered sedimentation of CA and RT on isopycnic gradients and reduction of ERT activity (Warrilow et al., 2007).

Our results demonstrate increasing dissociation of p24 CA from virus cores in the presence of increasing concentrations of detergent, and conditions were successfully established that could distinguish between WT NL4-3 and hyperstable cores. Using these conditions, it was determined that CA mutant viruses S41A, S109A, S146A, S149A, S178A and T188V did not exhibit hyperstable cores.

The CA mutant K203A was generated to be used as hypostable control in this study. A previous study which examined this virus using a combination of the spin-thru method (1%

Triton-X-100) and isopycnic ultracentrifugation, reported detection of only minor amounts of

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CA protein by p24 CA ELISA. This indicated that very few core structures sedimented through the gradient, indicative of extreme instability and hypostable cores (Forshey et al.,

2002). However, in my study, K203A was detected by ELISA and western blot. In fact, it produced nearly identical core stability profiles as WT NL4-3 virus between 0.01-1.0%

Triton-X-100. This suggests that in our hands the K203A CA mutant is not unstable as previously reported (Forshey et al., 2002). However, mature HIV cores are intrinsically unstable, and it may not be possible to find detergent conditions using the current assay that allow differences in core stability to be observed between the CA mutant K203A and WT

NL4-3 virus.

Analysis of CA sedimentation profiles from CA mutants S41A, S146A and T188V demonstrated cores from these viruses behave similarly to WT NL4-3 virus. While CA mutants S109A, S149A and S178A showed some hypostability relative to WT NL4-3 virus.

During the course of this study Wacharapornin et al (2007) reported reduced core stability for three viruses with mutations in CA (S109A, S149A and S178A) identical to those used in the present study. The core stability assay used in the study by Wacharapornin et al (2007) concentrated virions through 10% sucrose containing 0.1% Igepal into 50% sucrose, then quantitated the amount of pelleted CA representing core structures compared with total CA input by ELISA. Their results demonstrated only a very minor (5%) difference in core stability between the CA mutant viruses (4-6%) and WT NL4-3 virus (11%) (Wacharapornin et al., 2007). Results in this thesis indicate CA mutants S109A, S149A and S178A were not hyperstable, and that they showed greater susceptibility to detergent than WT NL4-3 virus

(Figures 4.9, 4.12, 4.13 and 4.16). While the results from the present study are not quantitative, they support the decreased stability observed by Wacharapornin et al (2007).

Additionally Wacharapornin et al (2007) analysed the “uncoating activity” of these CA mutant viruses by incubating isolated cores in the presence of H9 cell lysates (suggested to contain an “uncoating factor”) and measuring the resulting pelletable CA. They demonstrated that these viruses displayed reduced core stability following detergent treatment (i.e. less

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pelletable CA-containing complexes were recovered compared with WT). Paradoxically they displayed reduced susceptibility to the H9 uncoating activity (i.e. they did not release as much

CA from cores as WT virus when treated with H9 cell lysates) (Wacharapornin et al., 2007).

This lack of correlation of core susceptibility to detergent with core dissociation kinetics is supported by the study by Forshey et al (2002), where several CA mutant viruses (E45A,

E128A/R132A) that produced core yields higher than WT virus in the presence of detergent, displayed slower disassembly kinetics during in vitro dissociation assays at 37ºC. Conversely,

Forshey et al (2002) also observed that lower core yields in the presence of detergent

(Q63A/Q67A) but more rapid disassembly kinetics in vitro at 37ºC. In contrast, the same study also reported that CA mutant R143A produced lower core yields than WT virus in the presence of detergent, but slower disassembly at 37ºC compared to WT virus (Forshey et al.,

2002). This suggests the link between core stability in response to detergent treatment, heat or cellular uncoating factors is not a simple linear correlation.

The literature indicates that proper maintenance and regulation of core structures, including both assembly and disassembly of the virus core, is required for efficient reverse transcription and nuclear translocation (Arhel et al., 2007; Dismuke & Aiken, 2006; Forshey et al., 2002).

The change in core stability conferred by these mutations in CA (S109A, S149A and S178A) may potentially explain the block during replication demonstrated in Chapter 3. It is uncertain how this change in core stability would be reflected specifically in core disassembly.

Release or rearrangement of CA protein from the core to generate the RTC is an essential step in HIV replication, yet little is known about the mechanism or regulation of this process.

While most types of virus require some form of core disassembly for genome replication they utilise different mechanisms to achieve this. In HIV infection, the majority of CA was reported to dissociate from the core during an early stage of viral replication in the cytoplasm, the viral genome is reverse transcribed into dsDNA within the RTC, and is subsequently transported into the nucleus within the PIC (Farnet & Haseltine, 1991; Karageorgos et al.,

1993; Miller et al., 1997). It is now apparent that while the conical CA core may be lost,

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rearrangement permits retention of some CA in RTCs and PICs. Studies using fluorescent imaging have shown that some CA protein is still associated with transcriptionally active complexes in the cytoplasm of infected cells (Iordanskiy et al., 2006; McDonald et al., 2002).

However, mutagenesis studies have also demonstrated that CA mutant viruses which retained elevated levels of CA in RTCs or PICs are associated with defective viral replication including blocks in reverse transcription and nuclear translocation (Arhel et al., 2007;

Dismuke & Aiken, 2006; Forshey et al., 2002; Tang et al., 2003). Furthermore, the accumulation of cores from WT virus has been observed at the nuclear membrane 24-48 hr p.i.. While cores from viruses arrested at the final stage of reverse transcription (DNA flap formation) persist for up to 72 hr p.i. (Arhel et al., 2007). This implies that the core disassembly process is not spontaneous post membrane fusion and additionally, while it is still not understood, there is a link between core disassembly and reverse transcription.

Furthermore, defects in disassembly of CA from the core may impact interactions with nuclear pore proteins and subsequent nuclear translocation of the PIC. This highlights the delicate nature of maintenance and dissociation of the core for HIV integrity.

Various mechanisms are utilised by other viruses to disassemble their viral capsids and safely deliver their viral genomes to the nucleus for replication. For HBV, cores are transported directly into the nucleus where replication occurs (Rabe et al., 2003). For other DNA viruses, such as adenovirus and HSV, intact capsids are transported directly towards the nucleus, where they bind to the cytoplasmic face of nuclear pore complexes to release viral DNA into the nucleoplasm (Chardonnet & Dales, 1972; Greber et al., 1997; Morgan et al., 1968;

Morgan et al., 1969; Ojala et al., 2000). The release of viral RNA directly into the cytoplasm is sufficient for many RNA viruses to initiate infection.

Common mechanisms that regulate the release of capsid protein from the core vary from pH acidification in endosomes to proteosomal degradation of viral proteins surrounding the genome. For example, Influenza enters the cell via endosomes and an influx of protons through membrane channels are thought to promote disassembly of the nucleocapsid,

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allowing release of the segmented genome from the and into the cytosol (Zhirnov

& Grigoriev, 1994). Alternatively, following receptor mediated uptake and release from endosomes, adenovirus undergoes a stepwise release of fibres, then stabilising core proteins.

Finally, a viral protease degrades an additional core protein allowing DNA import into the nucleus (Greber et al., 1997). Poliovirus core disassembly is initiated by Zn2+ ion destabilisation of the capsid protein which subsequently increases the permeability of the core. Poliovirus capsid proteins are also phosphorylated by a virion associated kinase which induces further destabilisation of the core to allow release of the poliovirus RNA (Ratka et al.,

1989). Partial unfolding of the poliovirus capsid alone is not sufficient to lead to complete capsid dissociation, again indicating that core disassembly is often a stepwise and regulated process (Scharli & Koch, 1984).

Reverse transcription of retroviral RNA genomes into DNA inside the cytoplasm occurs in a replication complex generated by restructuring of the internal viral core and is usually depleted of most of the CA protein, or alternatively in a replication complex with an intact core that somehow disassembles to allow nuclear entry. Our understanding of the retroviral core, its role in reverse transcription and how it disassembles is not clearly defined, but a chain of events, potentially including phosphorylation, could be involved in HIV-1 core disassembly. We are only beginning to understand this process.

In conclusion, in this chapter an in vitro assay was established to assess HIV core stability.

Conditions were established to differentiate between WT NL4-3 virus and the hyperstable CA mutant virus E128A/R132A. CA mutant virus K203A exhibits comparable core stability to

WT virus in our assay. CA mutant viruses S41A, S146A and T188V showed similar detergent susceptibility when compared to WT NL4-3 virus in the core stability assay. CA mutant viruses S109A, S149A and S178A, with defined defects in infectivity (Chapter 3), showed decreased in vitro core stability relative to WT NL4-3 virus. This suggests that the defect during the reverse transcription (S109A, S178A) or post reverse transcription (S149A) steps in these viruses was not due to increased core stability or the formation of hyperstable cores,

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but may be due to reduced core stability. Altered regulation of core disassembly early during infection can have dramatic effects on HIV-1 replication.

Whether core stability measured in vitro accurately reflects core disassembly within the host cell is still unknown. To address this, Chapter 5 investigates core disassembly inside an infected cell using a fate-of-capsid assay to measure levels of CA associated in low and high molecular mass complexes following viral infection.

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CHAPTER 5 - Persistence of CA following infection as a measure of core disassembly in the infected cell

5.1 Introduction

Following fusion mediated entry into the host cell cytoplasm the HIV-1 core was presumed to rapidly disassemble. Early biochemical analyses under detergent conditions were unable to detect CA protein in replicating complexes, including reverse transcription complexes (RTCs)

(Fassati & Goff, 2001) and preintegration complexes (PICs) (Karageorgos et al., 1993;

McDonald et al., 2002; Miller et al., 1997), suggesting dissociation of CA from cores soon after entry into the cell.

The inability of early EM studies to visualise intact core structures in newly infected cells seemed to confirm that the viral core structure disintegrates soon after cell entry (Grewe et al.,

1990). However EM has identified intact core structures inside the cell, suggesting the core may remain intact upon entry (Arhel et al., 2007; Jun et al.). Furthermore, these core structures accumulate in a mutant virus defective for completion of reverse transcription. This suggests that reverse transcription can progress within an intact core structure, but that core disassembly may require completion of reverse transcription. While it also indicates that PICs cannot enter the nucleus without disassembly or restructuring of the virus core. Reverse transcription intermediates up to second strand transfer are efficiently synthesised in vitro, suggesting reverse transcription requires additional factors provided by the cellular milieu for completion (Hooker & Harrich, 2003; Warrilow et al., 2008). Therefore, the CA shell that defines the perimeter of the core would need to be shed, restructured, or at least become permeable to cellular factors, to allow reverse transcription to proceed. A proportion of CA persists longer in the host cell cytoplasm as part of the RTC (Iordanskiy et al., 2006;

McDonald et al., 2002). CA is detected within the cell cytosol by confocal fluorescent microscopy within 30 min of infection (Marechal et al., 1998). Within 2 hr of infection of cells with GFP-Vpr labelled virus, viral complexes were identified containing GFP-Vpr and

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labelled dUTP, identifying the complexes as active RTCs. These high molecular mass complexes, which also contained CA, were detected by immuno-staining around the perinuclear region (McDonald et al., 2002). In that study 67% of RTCs trafficking through the host cell cytoplasm via microtubules were reported to contain levels of CA protein similar to that found in intact virions (McDonald et al., 2002). Additionally, intact cores from WT virus have been visualised at the nuclear pore (Arhel et al., 2007). The presence of substantial amounts of CA in these complexes does not necessarily indicate that the conical core structure present in the mature virion is maintained. Restructuring of the core to generate an RTC permeable to factors required for efficient reverse transcription, may allow CA to remain inside the cytoplasm as part of an actively replicating complex much longer than initially thought. Furthermore, reports that CA interacts with nuclear pore proteins, including Nup358 and TNPO3, indicate that CA may enter the nucleus as a constituent of the PIC (Schaller et al., 2011; Shah et al., 2013). Altered uncoating of the viral core may impact these interactions; hence the frequent identification of reverse transcription and nuclear import defects in virus mutants displaying altered core stability (Cartier et al., 1999; Dismuke &

Aiken, 2006; Forshey et al., 2002).

There is limited knowledge about what regulates core disassembly largely due to the fact that analysing the HIV core and its disassembly remains difficult. The previous chapter and other studies in the literature have tried to analyse core stability by treating viral cores under in vitro conditions (including pH, temperature, ionic strength, detergent and cell components) and using this as a means to mimic core disassembly inside the cell (Auewarakul et al., 2005;

Forshey et al., 2002; Ohagen & Gabuzda, 2000; Wacharapornin et al., 2007). However, in vitro core stability assays do not take into account cell factors, or cellular responses to HIV infection, such as post-translational modifications (i.e. phosphorylation of viral proteins), or activation of cells which may be involved in core disassembly.

To address this, alternative methods in the literature were used to analyse core disassembly by measuring the presence and persistence of CA proteins inside infected cells, referred to as the

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‘fate-of-capsid’ assay (Chatterji et al., 2006; Stremlau et al., 2006). This assay does not directly measure core disassembly, but reflects naturally occurring CA degradation which is thought to follow on from dissociation of CA from the protective core structure (Chatterji et al., 2006; Stremlau et al., 2006). A fate-of-capsid assay has been used in human HeLa cells infected with HIV-1 in the presence of TRIM5α from old world monkeys to demonstrate that

TRIM5α mediates viral restriction by accelerating dissociation of viral cores and subsequent degradation of the HIV-1 CA protein (Chatterji et al., 2006; Stremlau et al., 2006). In the absence of TRIM5α, a reduction in the amount of particulate CA protein was still observed two to six hours following infection, representing the normal kinetics of core disassembly

(Chatterji et al., 2006). I aimed to adapt these methods to investigate HIV-1 core disassembly and relate this to the in vitro core stability of CA mutant viruses examined in Chapter 4

(Chatterji et al., 2006; Marechal et al., 1998; Stremlau et al., 2006). To reflect infection within the cell as closely as possible, the pattern of CA protein release from the viral core was investigated following HIV env-mediated infection rather than infection by pseudotyped viruses.

HuT-78 cells were infected with WT NL4-3 and CA mutant viruses (E128A/R132A and

K203A) with altered core stability and the change in CA protein levels associated with whole infected cell lysates were analysed by western blot. Additionally, infected cell lysates were subjected to rate zonal ultracentrifugation (as described in Chapter 4) to separate core structures and were to investigate possible restructuring and dissociation of CA from the core during early infection.

Results indicate that following infection there is a decrease in total CA protein levels and subtle changes to the sedimentation properties of CA-containing complexes in infected cell lysates. However, the results were inconsistent between experiments and the in vitro core stability of WT NL4-3 and CA mutant controls observed in Chapter 4 could not be correlated with persistence of CA protein or core structures observed inside infected cells.

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5.2 Results

5.2.1 Optimisation of cell-free infection for fate-of-capsid assay

The fate-of-capsid assay follows the kinetic decay of total intracellular HIV-1 CA protein following infection of target cells (Chatterji et al., 2006; Stremlau et al., 2006). This gives an indication of how long CA protein is detected inside the infected cell, and if it remains associated with replication complexes. HIV virions enter target cells via several routes. Entry mediated by specific cell surface receptors results in productive infection, while virus particles internalised by endocytic vesicles are not thought to result in non-productive infection (Chatterji et al., 2006; Marechal et al., 1998). The proportion of virus that enters the cell cytoplasm utilising cell surface receptors is smaller, likely reflecting the low number of infecting HIV particles. Fractionation of infected cell lysates to differentiate cytosolic and vesicular components can be used to separate virus that has entered via receptor mediated entry versus endocytosis. However, a reduction in the CA protein levels in whole cell lysates is also observed (Chatterji et al., 2006; Marechal et al., 1998). Many factors, including the low number of infecting HIV particles, make actively tracking virus that go on to productive infection a very difficult aspect of studying HIV core disassembly. While we hope to gain valuable knowledge about the fate of the total virions that enter the cell, it is acknowledged there is the potential for contaminating endocytic vesicles. Fate-of-capsid assays describe a time-course experiment measuring CA protein levels present in whole cell lysates from infected cells. This allows the level of CA associated in high and low molecular weight complexes to be followed as a means to determine how mutations in CA affect the ability of

CA to associate with post-entry complexes, such as RTCs and PICs.

5.2.2 Virus Stocks

Cell-free virus stocks for WT NL4-3 and CA mutant (E128A/R132A and K203A) viruses were generated by transfecting pNL4-3 plasmid into 293T cells and collecting virus- containing supernatants 48 hr post transfection (2.6.1). To determine the optimal amount of

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viral inoculum required, 5 x 106 HuT-78 cells were infected with clarified and concentrated

H3B cell culture supernatant (used as a high titre positive control [concentration not determined]) or 0.875 µg (in 2.5 mL), 1.75 µg (in 5 mL) or 3.75 µg (in 10 mL) p24 (measured by HIV-1 p24 ELISA (NEN™ Life Science Products, PerkinElmer™)) WT NL4-3 virus inoculum (Figure 5.1). Cells were resuspended in virus inoculum and incubated at 4ºC for 2 hr to allow virus to bind to cell receptors. Cells were centrifuged at 37ºC for 1 hr at 3300 rpm

(centrifugal enhancement, see 5.2.3.1), then washed to remove virus and resuspended in fresh

RPMI media and moved to 37ºC. At 2 hr p.i. cells were washed and lysed in RIPA buffer.

Cell lysates were clarified, samples were normalised for total protein, and the CA protein content analysed by SDS PAGE and western blot. Infection with clarified and concentrated

H3B cell culture supernatant resulted in detection of high levels of p24 CA 2 hr p.i. Infection of cells with the 0.875 µg p24 (2.5 mL) NL4-3 virus resulted in the greatest level of CA detection in infected cell lysates compared with infection of cells with higher amounts (1.75

µg or 3.75 µg p24) of NL4-3 virus (Figure 5.1). The dilution of virus and cell populations likely results in decreased proximity between cells and virus potentially reducing infection efficiency. Thus infections were performed using 0.875 µg in a total volume of 2.5 mL for fate-of-capsid infections. However, the level of NL4-3 CA protein detected was low. Next, methods were investigated to increase the detection of CA protein in infected cell lysates.

5.2.3 Enhancing the fate-of-capsid assay

Two steps were employed to enhance the efficiency of infection: (i) centrifugal enhancement, to improve the efficiency of cell-free infection and (ii) temperature arrested state (TAS) of fusion, to increase the synchronicity of infection.

5.2.3.1 Centrifugal Enhancement to improve cell-free infection

Centrifugal enhancement describes the process of centrifuging virus and target cells at 37ºC, at a low speed (3300 rpm) (O'Doherty et al., 2000; Pietroboni et al., 1989). The centrifugal force applied is not sufficient to sediment virus onto cells. However, this process has been suggested to alter cell morphology making them more susceptible to virus entry (O'Doherty et 102

Figure 5.1 Amount of virus inoculum required to detect intracellular CA following infection 5x106 HuT-78 cells were infected with clarified H3B virus supernatant [positive control, concentration not determined] or 0.875 µg, 1.75 µg or 3.75 µg p24 of WT NL4-3 virus. Virus was absorbed to cells by incubation at 4ºC for 2 hr, then centrifugally enhanced at 3300 rpm at 37ºC for 1 hr. Virus was removed, cells resuspended in fresh RPMI and incubated for a further 1 hr at 37ºC. Infected cells were washed, lysed in RIPA buffer and 200 µg total protein analysed by western blot specific for CA (#6458). H3B NL4-3

CA

+ve 0.875 1.75 3.75 Virus inoculum (µg) al., 2000). While the details of virus entry utilising this process are not clear, the increase in efficiency of infection afforded by centrifugal enhancement has been estimated to be equivalent to a 10-fold increase in multiplicity of infection. The use of centrifugal enhancement of infection has been previously optimised for HIV (O'Doherty et al., 2000;

Pietroboni et al., 1989).

To adsorb virus, cells were incubated at 4ºC for 1 hr. To assess the effect of centrifugal enhancement of infection, cells were either incubated at 37ºC without centrifugation, or centrifugally enhanced (1 hr at 3300 rpm). Next, cells were washed twice in room temperature

PBS and resuspended in fresh RPMI media for incubation at 37ºC for the remainder of the infection. Cells were washed and lysed at 0 hr, 2 hr, 4 hr and 6 hr p.i. and analysed for HIV-1

CA protein by SDS PAGE and western blot. In the absence of centrifugal enhancement, CA protein was barely detectable 4-6 hr p.i. (Figure 5.2A). In contrast, centrifugal enhancement of infection resulted in detection of high levels of CA protein in cell lysates 2-6 hr p.i.

(Figure 5.2B).

Therefore centrifugal enhancement of infection was used for subsequent fate-of-capsid assays.

5.2.3.2 Temperature Arrested State (TAS)

Temperature Arrested State (TAS) describes incubation of virus and cells at 23-25ºC for 3 hr prior to infection (Markosyan et al., 2003). This process of temperature arrest allows virus to bind cell receptors but doesn’t allow entry of virus into the cell. This results in accumulation of virus at the cell membrane and increased synchronisation of viral infection. Upon increasing the temperature to 37ºC, infection proceeds with an increased efficiency due to synchronisation of viral infection.

5.2.4 Protein Analysis of infected cell lysates

5.2.4.1 Cell lysis for SDS PAGE and western blot

To detect changes in total CA protein levels, infected cell lysates were analysed by SDS

PAGE and western blot. Detection of CA protein by western blot using antibody specific for

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Figure 5.2 Centrifugal enhancement is required to detect CA following infection 5x106 HuT-78 cells were infected with clarified H3B virus supernatant. Virus was absorbed to cells by incubation at 4ºC for 1 hr (“0 hr”), incubated at 37ºC (A), or centrifugally enhanced at 3300rpm at 37ºC for 1 hr (B), cells were washed and incubated at 37ºC for the remainder of the infection. Infected cells were washed, lysed in RIPA buffer and 200 µg total protein and analysed by western blot specific for CA (#6458). H3B

A CA 0 2 4 6 Time post-infection (hr)

B CA 0 2 4 6 Time post-infection (hr) p24 CA (#6458) was compared following cell lysis in modified RIPA buffer (2.1.5) and dilution of cell lysates directly in 4x SDS loading buffer, or following cell lysis in RIPA buffer followed by total protein precipitation (TCA/acetone precipitation). Following infection (as described in 5.2.2) HIV-1 CA protein was readily detected in clarified cell lysates diluted directly in 4x SDS loading buffer, while TCA/acetone precipitation resulted in significant loss of detectable CA protein (data not shown). Therefore cells were lysed in RIPA buffer, diluted with 4x SDS loading buffer and analysed for p24 CA by SDS PAGE and western blot.

5.3 Analysis of CA persistence in infected cells

5.3.1 CA is detected within infected target cells up to 6 hr post-infection.

The persistence of CA within infected cells was analysed to determine differences in the release of CA protein from cores following infection. HuT-78 cells were infected with WT

NL4-3 virus. Whole cell lysates were collected following cell mixing (‘0hr’), after TAS incubation (3 hr at 23ºC) and after incubation at 37ºC for 2 hr, 4 hr, and 6 hr p.i.. Cells were washed, trypsinised and lysed in RIPA buffer and CA protein was analysed by SDS PAGE and western blot using antibody specific for p24 CA.

At the time of cell and virus mixing (0hr), no CA protein was detected by western blot

(Figure 5.3). Minor amounts of CA were detected when cells and virus were incubated at

23ºC for 3 hr (TAS). When virus and cells were incubated at TAS then shifted to 37ºC to allow virus cell fusion and entry, intracellular CA protein was detected in most cell lysates.

However, as the results presented from three different infections in Figure 5.3 suggest, CA profiles between experiments were inconsistent. Within 2-6 hr following infection high levels of CA protein were detected within the cell. In 2 out of 3 infections, intracellular levels of CA declined by 6 hr p.i. (Figure 5.2A and B).

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Figure 5.3 Persistence of intracellular CA in infected cells 5x106 HuT-78 cells were infected with 1 µg WT NL4-3 virus. Virus was absorbed to cells by incubation at 4ºC for 1 hr (0 hr), incubated at 23ºC for 3 hr (TAS), then centrifugal enhancement was performed at 3300 rpm at 37ºC for 1 hr. Virus was removed, cells were washed twice in PBS, resuspended in fresh RPMI and incubated at 37ºC as indicated. Infected cells were washed and trypsinised to remove external virus, lysed in RIPA buffer and 200 µg total protein and analysed by western blot specific for CA (#6458). A-C represent triplicate infections. NL4-3

A CA 0 T 2 4 6 Time post-infection (hr)

B CA 0 T 2 4 6 Time post-infection (hr)

C CA 0 T 2 4 6 Time post-infection (hr) 5.3.2 CA complexes change subtly 2-6 hr post infection

To investigate if CA protein remains associated with, or is released from cores during early infection, the sedimentation of high molecular mass CA-containing complexes was analysed by rate zonal sucrose gradient sedimentation.

HuT-78 cells were infected with WT NL4-3 virus as previously described (section 5.3.1).

Cells were collected at 0 hr, after TAS incubation, and 2 hr, 4 hr and 6 hr p.i. and trypsinised to remove external virus. Infected cells were lysed in a hypotonic lysis buffer (2.1.6) using a disposable plastic pestle in a microcentrifuge tube. The hypotonic lysis buffer did not contain detergent to reduce the potential effects of detergent on any high molecular mass complexes containing CA. Cell lysates were loaded onto 15-40% sucrose gradients and ultracentrifuged at 35000 rpm for 40 min at 4ºC (as described in Chapter 4). Fractions (1 mL) were collected from the top of the gradient and analysed for p24 CA protein. Western blotting specific for p24 CA (#6458) only weakly detected CA protein in fractions 1 and 6 (data not shown).

Therefore fractions were also analysed by p24 ELISA. CA was detected in fractions 6-9 in lysates collected 2 hr, 4 hr and 6 hr p.i. (Figure 5.4). At 0 hr and following TAS low levels of

CA were detected in the top fraction of the gradient representing free CA protein, with negligible CA detected in the rest of the gradient. Total intracellular CA levels peaked at 2 hr p.i.. CA was primarily present as a broadly sedimenting peak in the middle of the gradient

(fractions 6-10), representing high molecular mass complexes containing CA. Non-specific high molecular mass CA was detected at the bottom of the gradient. At 4-6 hr p.i., there was a small decrease in the total amount of CA protein detected in the gradient, consistent with analysis of total cell lysates by SDS PAGE and western blot (Figure 5.3). At 4 hr and 6 hr p.i.

CA protein primarily sedimented in fractions 6-8. The peak of high molecular mass complexes containing CA was concentrated in fraction 7. The proportion of non-specific high molecular mass CA detected in the bottom fraction increased 4-6 hr p.i..

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Figure 5.4 Analysis of CA structures in infected cells 5x106 HuT-78 cells were infected with 1µg WT NL4-3 virus. Virus was absorbed to cells by incubation at 4ºC for 1 hr (0 hr), incubated at 23ºC for 3 hr (TAS), followed by centrifugal enhancement at 3300 rpm at 37ºC for 1 hr. Virus was removed, cells were washed twice in PBS, resuspended in fresh RPMI and incubated at 37ºC as indicated. Infected cells were washed and trypsinised to remove external virus, then lysed under detergent free conditions and p24 CA analysed by ELISA. Faster sedimenting CA containing structures were separated from free CA protein by rate zonal ultracentrifugation on 15- 40% sucrose gradients at 35000 rpm for 40 min. Fractions (1 mL) were collected from the top of the gradient and analysed by p24 CA ELISA. Total CA in each gradient is represented in (A), sedimentation of CA throughout gradients is represented in (B). A 16000 14000

12000

10000 0hr TAS 8000 2hr 4hr

p24 CAp24(pg/mL) 6000 6hr

4000

2000

0 0hr TAS 2hr 4hr 6hr

Post TAS

5000

B 4500

4000

3500 0hr 3000 TAS 2500 2hr 4hr 2000

p24 CA (pg/mL) CA p24 6hr 1500

1000

500

0 1 2 3 4 5 6 7 8 9 10 11 12 Fraction 5.3.3 CA persistence inside infected cells does not correlate with in vitro core stability

Next, we assessed if defects in in vitro core stability correlated with altered release of CA from the core following infection. To examine this, hyperstable and hypostable CA mutant viruses E128A/R132A and K203A were compared with WT NL4-3 virus in the fate-of-capsid assay.

In Chapter 4, we assessed the in vitro core stability of E128A/R132A and K203A.

E128A/R132A cores displayed greater resistance to detergent treatment compared with cores from WT NL4-3 virus. No difference was observed between K203A and WT NL4-3 virus in our in vitro core stability assay. However, other studies report CA mutant K203A exhibits decreased core stability with respect to WT virus (Dismuke & Aiken, 2006; Forshey et al.,

2002; Hulme et al., 2011).

HuT-78 cells were infected with WT NL4-3 and CA mutant viruses E128A/R132A and

K203A (as described in section 5.3.1). Total cell lysates were analysed by SDS PAGE and western blot for p24 CA. No intracellular CA was detected at 0 hr p.i or following TAS treatment (Figure 5.5). Intracellular CA protein was detected 2-6 hr p.i. for WT NL4-3 and

CA mutant viruses K203A and E128A/R132A. In comparison with WT virus, lower levels of

CA protein were detected for K203A. For K203A, maximum CA protein levels were observed at 2 hr p.i., and decreased levels of CA were observed at 4 hr p.i.. Whereas for WT

NL4-3 virus, we observed two different patterns of intracellular CA. CA protein was detected at similar intensities at 2 hr and 4 hr p.i., but was not detected 6 hr p.i.. Alternatively, CA protein was detected at a similar intensity at 2 hr, 4 hr and 6 hr p.i.. CA protein profiles for

K203A during this fate-of-capsid assay suggest slightly decreased stability of this CA mutant virus.

For CA mutant E128A/R132A, high levels of CA protein were detected early after infection

(2-4 hr). CA protein was not detectable by 6 hr p.i. This CA protein profile was similar to some CA protein profiles observed for WT NL4-3 virus (Figure 5.3A & 5.3B). However,

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Figure 5.5 Persistence of intracellular CA in cells infected with CA mutants displaying defects in core stability 5x106 HuT-78 cells were infected with 1µg (A) WT NL4-3, (B) hyperstable CA mutant E128A/R132A or (C) hypostable CA mutant K203A viruses. Virus was absorbed to cells by incubation at 4ºC for 1 hr (0 hr), incubated at 23ºC for 3 hr (TAS), then centrifugal enhancement was performed at 3300 rpm at 37ºC for 1 hr. Virus was removed, cells were washed twice in PBS, resuspended in fresh RPMI and incubated at 37ºC as indicated. Infected cells were washed and trypsinised to remove external virus, lysed in RIPA buffer and 200 µg total protein and analysed by western blot specific for CA (#6458). NL4-3 A CA 0 T 2 4 6 Time post-infection (hr)

E128A/R132A B CA 0 T 2 4 6 Time post-infection (hr)

K203A

C CA 0 T 2 4 6 Time post-infection (hr) WT NL4-3 is also observed to show consistent levels of CA 2-6 hr p.i. (Figures 5.3C and

5.5A), suggesting that no difference could be shown between these viruses.

CA protein profiles from these fate-of-capsid assays were inconsistent from experiment to experiment. The CA mutants showed different patterns of CA persistence in subsequent fate- of-capsid assays making it impossible to draw firm conclusions. For this reason, CA substitution mutants (S41A, S109A, S146A, S149A, S178A) were not assessed using this fate-of-capsid assay.

5.4 Discussion

Analysis of in vitro detergent susceptibility of virus cores (Chapter 4), while indicating potential defects in the viral core, may not truly reflect intracellular core stability. Thus the

‘fate-of-capsid’ assay, which has been used to demonstrate accelerated release of CA protein from the core mediated by host-cell restriction factors (Chatterji et al., 2006; Stremlau et al.,

2006), was adapted to investigate CA release from the core of WT NL4-3 virus and the previously characterised CA mutants exhibiting altered core properties (E128A/R132A and

K203A).

The low number of infectious HIV particles in any virus preparation makes studying early

HIV entry events difficult. The use of HIV pseudotyped with VSV-G or MLV Env to maximise viral entry in other studies utilising the fate-of-capsid assay (Chatterji et al., 2006;

Iordanskiy et al., 2006; Marechal et al., 1998; McDonald et al., 2002)) overcomes the problem of poor virus entry into the cell. However, this alters the native entry process used by

HIV which is crucial when investigating subsequent CA release from the core during core disassembly. Therefore, in the present study NL4-3 virus with HIV Env proteins was used to reflect true changes occurring following infection. This required several optimisation steps to increase the efficiency of infection, including temperature arrested state (TAS) and centrifugal enhancement of infection.

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Cells and virus were incubated at 4˚C to allow virus attachment and aid synchronous infection. Use of a temperature arrested state (TAS) has been reported to result in greater efficiency and synchronicity of infection. The TAS incubation step allows co-receptor binding, but virus entry is prevented until the temperature is raised above 23ºC (Markosyan et al., 2003). Fusion then occurs quickly, resulting in greater synchronicity of infection and consequently greater detection of high molecular mass complexes containing CA after infection at 37ºC.

Centrifugal enhancement of infection has been demonstrated to be as effective as a 10-fold increase in the multiplicity of infection. However the mechanism underlying this process is still unclear (Pietroboni et al., 1989). While this step was not included in similar protocols in other studies (Chatterji et al., 2006; Marechal et al., 1998; Stremlau et al., 2006), centrifugal enhancement was necessary to visualise intracellular CA protein in infected cell lysates in the present study (Figure 5.2).

Together TAS and centrifugal enhancement of infection aimed to prepare virus for entry into the cell, after which incubation at 37ºC with centrifugal enhancement should allow rapid entry of virus into the target cell. Maximising the synchronicity of entry enables both increased detection of CA protein and ensures the majority of CA detected at each time point is representative of virus at the same stage of infection.

WT NL4-3, E128A/R132A and K203A CA mutant virus stocks were produced by transfection of plasmid DNA into 293T cells. HuT-78 cells were infected with each virus and whole cell lysates were collected from infected cells. Total CA protein was analysed by western blot, ELISA or by rate zonal ultracentrifugation. Minimal CA protein was detected at early time points following virus inoculation but prior to virus-cell fusion (0 hr, TAS).

Infected cells were treated with trypsin to remove any virus that had adsorbed to the outside of the cell. Therefore the presence of minor levels of CA protein is likely to be due to endocytosis of viral particles.

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Results presented in this chapter suggest that the fate-of-capsid assay is variable and care should be exercised when making quantitative conclusions. CA profiles were inconsistent between experiments; however, in most experiments large amounts of WT NL4-3 CA protein was observed 2-4 hr p.i. in total infected cell lysates, followed by a decrease in CA protein 6 hr p.i. (Figure 5.3). Although in some experiments CA protein was detected at considerable levels 6 hr p.i.. Chatterji et al (2006) also measured CA protein in infected cells from 0.5-6 hr p.i., and showed that CA protein levels peaked 1-2 hr p.i. and were still detectable 6 hr p.i., consistent with results presented in this chapter (Chatterji et al., 2006). In the Chatterji et al study, CA (from VSV-G pseudotyped virus) was detected in the cytoplasm as soon as 30 min following infection (Chatterji et al., 2006). CA protein has been observed at perinuclear regions within 2 hr (McDonald et al., 2002). The decrease in intracellular CA protein levels observed 6 hr p.i. in most experiments is consistent with release of CA protein from high molecular mass complexes containing CA and subsequent CA protein degradation.

To specifically investigate this possibility, CA-containing complexes from infected cell lysates were analysed by rate zonal ultracentrifugation to detect changes in sedimentation that may reflect alterations in the structure of the core.

Cell lysates were collected from cells infected with WT NL4-3 virus at 0 hr (1 hr incubation at 4ºC), after TAS (3 hr incubation at 23ºC), and 2 hr, 4 hr or 6 hr p.i at 37ºC. To conserve the structure and integrity of CA-containing complexes, cells were lysed in a hypotonic lysis buffer in the absence of detergent using a disposable plastic pestle. Infected cell lysates were subjected to rate zonal ultracentrifugation, and fractions were collected and tested for the presence of CA protein by p24 CA ELISA. CA-containing complexes were present in the middle gradient fractions (7-9) from 2 hr to 6 hr p.i. (Figure 5.4), similar to sedimentation profiles obtained in the in vitro core stability assay (Chapter 4). At 2 hr p.i. CA protein was detected in fractions 6-10, and also as non-specific high molecular weight CA in the bottom gradient fractions potentially representing aggregates. At 4 hr and 6 hr p.i., the CA protein peak was sharper at a slightly slower sedimenting position (fraction 7) and increasing CA

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protein was present at the bottom of the gradient. Analysis of CA-containing complexes produced following infection by rate zonal ultracentrifugation demonstrated a relatively uniform range of CA-containing complexes, suggesting although the total levels of CA protein in the cell varied considerably, the size and shape of the structures present were quite similar. The complexes recovered from cells appeared more uniform than those detected using our in vitro core stability assay directly from transfected virus stocks. These complexes may represent only a subset of structures which are competent for infection, while it is likely the broader range of complexes containing CA detected in vitro will not all be competent for replication in host cells. Whether the complexes at the bottom of the gradient were representative of true high molecular mass complexes containing CA inside the cell or represented non-specific high molecular mass CA resulting for technical reasons following lysis was uncertain, but the latter was expected.

To investigate the relationship between altered in vitro core stability measured in Chapter 4 and core disassembly within the cell, the core stability control CA mutants were also analysed in the fate-of-capsid assay. When whole cell lysates from cells infected with WT NL4-3 virus or the hyperstable CA mutant (E128A/R132A) were compared, the results were similar

(Figure 5.5). As expected no CA protein was detected at 0 hr p.i., since virus has not entered the cell. Minor amounts of CA protein were detected following incubation at TAS. This could be due to trypsin treatment not completely removing bound virus from the cell surface or endocytosis of virus. At 2 hr and 4 hr p.i. a strong CA signal was detected, followed by a marked decline in CA protein levels observed 6 hr p.i.. Forshey et al (2002) reported that cores from the E128A/R132A CA mutant disassembled more slowly than WT NL4-3 following incubation at 37ºC, and in Chapter 4 this mutant showed some hyperstability to detergent compared with WT NL4-3 virus (Figure 4.7). Assuming that CA protein is resistant to degradation when it is part of the core structure, a consistent strong CA signal in cell lysates was expected for the E128A/R132A CA mutant throughout the time course. This

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suggests that in vitro core stability does not necessarily reflect the kinetics and susceptibility of core disassembly inside the infected cell.

The fate-of-capsid assay was also performed for the hypostable CA mutant K203A. CA protein was not detected in cell lysates taken at 0 hr or TAS time points. While CA was readily detected in cell lysates collected at 2 hr, 4 hr and 6 hr p.i. (Figure 5.5). Previous studies have reported that cores from the CA mutant K203A are intrinsically unstable, and that virtually no CA protein could be detected following detergent treatment and isopycnic gradient ultracentrifugation conditions that successfully isolate cores from WT NL4-3 virus

(Forshey et al., 2002). The analysis of in vitro core stability for the CA mutant K203A in

Chapter 4 (Figure 4.8) however, suggests that this CA mutant possesses greater stability than previously reported and cannot be distinguished from WT NL4-3. The results of the fate-of- capsid assay in this chapter suggested that K203A CA mutant cores do not disassemble more rapidly than WT NL4-3 cores. While levels of CA associated with high molecular mass complexes reduced more quickly that WT virus, the differences were minimal. Again this suggested that caution be taken when analysing these results.

Similar levels and persistence of intracellular CA protein in cells infected with WT NL4-3 or

CA mutants E128A/R132A and K203A suggests that while these CA mutants may exhibit altered core properties in vitro, core disassembly within the cell may be modified by association or interactions with other cellular factors or by localisation to specific compartments within the cell. Therefore more information might be gained by tracking the migration of viral complexes containing CA through the cell, as these complexes may show defects early following entry or alternatively later, such as at the nuclear pore, similar to virus blocked at the final stage of reverse transcription (Arhel et al., 2007). An uncoating assay that labels viral complexes following infection with VSV-G pseudotyped virus suggests the majority of p24 CA associated with the core is lost 1-4 hr post-fusion (Campbell et al., 2007;

Hulme et al., 2011; Yamashita et al., 2007). In this assay 62.6% of intracellular fused particles contained CA 1 hr p.i.. This reduced to 42.4% after 2 hr, and approximately 30% of

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particles contained CA after 4 hr (Hulme et al., 2010; Yamashita et al., 2007). Analysis of CA mutant viruses E128A/R132A (hyperstable), K203A (hypostable) and R143A (hypostable) demonstrated observable differences reflecting their altered stability 1-2 hr post-infection.

However, any differences were no longer apparent by 4 hr p.i.. This suggests the timing of analysis of viral complexes to assess core disassembly following infection is crucial, and since the earliest time point analysed in our assay was 2 hr p.i. it is possible we were outside of this window. However, this timing refers to infection by VSV-G pseudotyped virus. It also must be noted that since VSV-G pseudotyping can rescue defects in some virus (e.g. S149A), this may alter the effect of CA during core disassembly and replication (Brun et al., 2008).

In summary, a decrease was observed in the amount of intracellular CA protein present following infection. Additionally, subtle changes were observed in the sedimentation of CA- containing complexes from WT NL4-3 virus. In vitro core stability properties did not correlate with CA persistence profiles. The results of this assay varied greatly, CA profiles were inconsistent between experiments, suggesting that analysis of the persistence of CA protein using the current fate-of-capsid assay is not a reliable indicator of altered HIV core disassembly.

It is presumed that prior to nuclear translocation, CA protein must be released from the core as free protein, or the core must somehow rearrange into a complex capable of nuclear translocation (Dismuke & Aiken, 2006; Forshey et al., 2002). Reports suggest CA protein is present in RTCs and PICs (Iordanskiy et al., 2006; McDonald et al., 2002) implying remodelling of these complexes occurs after virus entry, potentially reflected by the shift in sedimentation of the majority of CA-containing structures. An immediate reduction in intracellular CA protein levels in WT NL4-3 virus infection was not observed, reinforcing the concept that CA is not released from the core immediately upon entry into the host cells.

Inconsistent CA protein profiles between experiments, suggested even tight control of all factors (virus stocks, cell stocks, temperature differences etc) could not overcome the variability involved in the timing of core entry into the cell (when detecting via CA protein)

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and thus more advanced methodology is needed to study core disassembly following HIV infection. The fate-of-capsid assay was not a reliable indicator of defects in HIV core disassembly, as in addition to the variability observed, in vitro core stability properties did not correlate with profiles of CA release from core structures. Due to the variability observed with WT NL4-3, E128A/R132A and K203A CA mutant viruses, the CA mutant viruses

(S41A, S109A, S146A, S149A, S178A and T188V) were not analysed using this method, therefore the fate of intracellular cores within these viruses early after infection is still uncertain.

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CHAPTER 6 - General Discussion

6.1 The role of the viral core in the early stages of HIV replication

The mechanisms which control disassembly of the HIV core remain to be unravelled. What promotes the rearrangement of CA within the core? How does core disassembly effect reverse transcription and viral replication?

Following fusion-mediated entry into the host cell, the virus core is released into the cytoplasm. The interaction between multimeric CA and the restriction factor TRIM5α in old world monkey cells implies the HIV viral core enters the host cell cytoplasm at least partially intact and that there it interacts with host cell factors (Shi & Aiken, 2006). Properly regulated core disassembly is required for efficient and complete reverse transcription. Very limited reverse transcription occurs inside the intact virion (Zhang et al., 1994). While NERT assays stimulate an increase in intravirion DNA synthesis, levels of early reverse transcription intermediates are significantly lower than those within infected cells (Hooker & Harrich,

2003). Later stages of viral DNA synthesis appear to rely on factors present in the intracellular cytoplasmic milieu. NERT assays have been reported to stimulate reverse transcription and also induce dissociation of viral core proteins (CA, NC, RT), suggesting these processes are linked (Zhang et al., 2000). Additionally, release of core proteins following incubation of virions (Ohagen & Gabuzda, 2000), or cores (Auewarakul et al.,

2005) in cell lysates indicates that host cell factors impact HIV core disassembly.

Substituting charged residues in CA, particularly on the CTD dimerisation interface, alters core stability, an in vitro measure of core disassembly (Forshey et al., 2002). This indicates charge also plays an important role stabilising the viral core. This led to the hypothesis that modification of the CA protein, and regulation of charge by phosphorylation, might be involved in the regulation of core disassembly. To investigate this, we examined viruses with mutations in CA at serine (S41A, S109A, S146A, S149A and S178A) and threonine (T188V) residues that display defects during replication. Reports indicate that the HIV-1 CA protein is 114

post-translationally modified by phosphorylation at three serine residues (S109, S149 and

S178) in the virion by an undefined kinase (Cartier et al., 1999). Substitution of serine at three sites (S109A, S149A S178A), and threonine (T188V) blocks replication at an early step in the viral life cycle ((Figure 3.6) and (Cartier et al., 1999)). Substitution at two additional serines

(S41A and S146A) alters, but does not terminate viral replication ((Figure 3.6) and (Cartier et al., 1999)).

6.2 Replication of HIV CA mutant viruses

This study aimed to examine the effect of a series of CA mutant viruses on in vitro core stability and the HIV core in early infection. To examine these properties, viruses with mutations in CA at three phosphorylation sites (S109A, S149A, S178A) and other sites of interest displaying altered replication (S41A, S146A, T188V) were generated by PCR mutagenesis. Conservative mutations were made to minimise disruption to the CA protein structure, and sequencing was undertaken to ensure no other mutations were introduced.

Analysis of Gag processing in transfected cell lysates and virions demonstrated normal viral protein expression, and processing and packaging of p24 CA into virions in all CA mutants generated, with the exception of mutant S109A.

In the current study, CA mutant S109A consistently showed lower levels of protein expression in comparison to WT NL4-3 virus. One possibility is that the mutation at this site affects recognition by our p24 CA antibody (#6458). Increased detection of CA was achieved with an alternative antibody (#4121) for this mutant. However, neither of the epitopes recognised by these antibodies have been mapped, so it is not possible to speculate on this further. Results published by Cartier et al (1999) and subsequently Brun et al (2008) showed relatively normal viral protein expression from CA mutant S109A (Brun et al., 2008; Cartier et al., 1999). To ensure lower expression was not a result of cloning, PCR mutagenesis was repeated with additional sets of primers encoding alternative codon usage. However, similar results were obtained. This also suggested that this reduction was specific to the S109A 115

mutant and not simply a result of lower transfection efficiency. Furthermore, S109A CA protein was detected with three different CA antibodies by either western blot or ELISA. This confirmed that viral proteins from the S109A CA mutant were detected at lower levels than

WT NL4-3 virus in the producer cell, but that S109A CA did not appear to be packaged normally into virions. Of note, Brun et al (2008) demonstrated limited proteolytic processing relative to WT virus (Brun et al., 2008). S109A virus exhibited greater levels of p55 Gag precursor and cleavage intermediates than processed p24 CA (Brun et al., 2008). Brun et al

(2008) reported that S109A mutant virions did not contain cone shaped cores indicating an apparent assembly defect in this mutant, potentially consistent with the results in my study

(Brun et al., 2008). In contrast, Cartier et al (1999) reported that S109A produced morphologically normal virions, although EM data was not shown (Cartier et al., 1999).

Although, EM analysis cannot tell us about the rate at which these viruses assemble, and if it is comparable to WT virus. Together these results suggest this residue is indeed critical to virus infectivity. However, the mutation appears to exert an effect prior to disassembly of the core. While we confirmed reduced core stability in this mutant, we believe this to be a downstream effect of a potential assembly defect. A more rigorous analysis of assembly of the S109A CA mutant needs to be carried out for further insights as to how this residue is involved in assembly. Marked impairment of assembly by virus substituted with aspartic acid at this site indeed suggests S109 is critical for formation of the CA hexamer and core (Brun et al., 2010).

We also performed more detailed analysis of the replication competence of these CA mutant viruses, including the detection of both early (R-U5) and late (PBS-Gag) reverse transcription intermediates. A defect was found prior to translation of Tat in some CA mutants (S109A,

S149A, S178A and T188V), although the replication capacity for each of these viruses varied subtly (Figures 3.6, 3.8). CA mutants S41A and S146A showed replication competence comparable to WT NL4-3 virus in a single cycle infectivity assay (Figure 3.6), but lower levels of early reverse transcription intermediates early after infection (Figure 3.8).

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While S109A and S149A CA mutant viruses were able to reverse transcribe early and late viral DNA intermediates, single cycle infectivity assays showed replication was blocked prior to Tat activation of the LTR promoter (Figure 3.8, 3.6). Additionally, virion production was reduced for the S109A mutant (as described above) (Figure 3.3 & 3.4). In this study early and late intermediates were detected from both CA mutants S109A and S149A (Figure 3.8).

This was in consistent with reports by Cartier et al (1999) that reverse transcription intermediates up until second strand transfer were detected (Cartier et al., 1999), but in contrast to Brun et al (2008) who reported these mutants showed severe reductions in the ability to synthesise both early and late reverse transcription intermediates (Brun et al., 2008).

In my study, CA mutant S149A produced equal to greater amounts of reverse transcription intermediates than WT NL4-3 virus. Interestingly, Brun et al (2008) reported that, using an

ERT assay, viral DNA synthesis by the S149A mutant was increased 2.5 fold in comparison to WT NL4-3 virus (Brun et al., 2008). This indicated RT activity is not significantly adversely affected alanine substitution at these sites, but that following infection of the cell a defect relating to the function of the CA protein manifests during reverse transcription.

Findings by Warrilow et al (2008) suggest a cellular factor is required for synthesis of late reverse transcription intermediates (Warrilow et al., 2008), and restructuring of CA that forms the core is likely to be required to permit access of these factors to the replication complex.

Arhel et al (2007) reported the accumulation of intact cores at the nuclear pore derived from virus which cannot complete the final step of reverse transcription (central flap formation).

While we cannot be certain that these cores represent active replication competent structures, if reverse transcription can proceed almost to completion in an intact core structure, it does not rule out the possibility that these viruses might still display a defect in core disassembly that prevents subsequent nuclear translocation of the PIC (Arhel et al., 2007). CA interacts with several nuclear pore proteins during entry to the nucleus. No replication profiling has been carried out in combination with knockdown of nuclear pore proteins such as TNPO3, with the CA mutant viruses in this study to determine if there viruses display altered

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susceptibility to restriction in these cells. Thus without adequate disassembly of the core, it may not be in a position to undergo efficient nuclear translocation.

CA mutants S178A and T188V demonstrated reduced replication competence in a single cycle infectivity assay (Figure 3.6) and a comparable reduction in early and late reverse transcription intermediates (Figure 3.8). Residues S178 and T188 are both situated on the CA

CTD dimerisation interface and hence similar replication profiles were not unexpected. T188 participates in hydrophobic contacts with three residues (T148, I150, L151 (Worthylake et al.,

1999)), suggesting it is important during CA CTD dimerisation to link adjacent hexamers during the formation of the viral core. Analytical gel filtration chromatography demonstrated that replacing T188 with alanine reduced the dimerisation affinity of the CA CTD (del Alamo et al., 2003). Therefore, it may have been expected to reduce the stability of the core.

Alternatively replacing S178 with alanine increases dimerisation affinity of the CA monomer

(del Alamo et al., 2003), and might be expected to increase the stability of the core. S178 forms a hydrogen bond with residue E180 on adjacent CA protein hexamers (Worthylake et al., 1999). Substitution at position E180 with alanine results in a compensatory mutation at position 178 to aspartic acid (negatively charged), further indicating the importance of negative charge at position 178 (del Alamo & Mateu, 2005). Negative charge (e.g. phosphorylation) at positions 178 and 180 may generate repulsive forces which might be required to promote disassembly of the core. CA CTD dimerisation is relatively stable, allowing the CA protein to form dimers but not hexamers in solution. This suggests that since

CA NTD dimerisation is easily disrupted, altering the charge on the CTD dimerisation interface (potentially by phosphorylation of S178) might be necessary to promote core disassembly. Hence substitution at these sites may result in altered core disassembly, and subsequently impact efficient reverse transcription by these mutant viruses.

CA mutants S41A and S146A synthesised reduced levels of early and late reverse transcription intermediates between 0-24 hr following infection. Yet replication competence was comparable to WT virus in a single cycle infectivity assay (Figure 3.6), consistent with

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reports these CA mutants undergo delayed replication (Cartier et al., 1999). This suggests that delayed or reduced levels of viral DNA synthesis early during infection can be overcome by these viruses. Further characterisation of this might be achieved by investigated infection at multiple and extended time points. Alternatively, this may be the result of differences in infectivity in different cell types.

The results in this study demonstrate that viruses containing mutations in CA at sites shown to be phosphorylated in the virion (S109A, S149A and S178A) exhibit reduced replication competence and a block in replication prior to integration (Figure 3.6). These results confirm a prior study, although there were some minor differences to the reported replication profiles

(Cartier et al., 1999).

The HIV CA protein is the major constituent of the viral core. Exactly how mutation of the

CA protein contributes to defects during reverse transcription and nuclear translocation remains uncertain. However, it appears likely that core disassembly, reverse transcription and nuclear translocation are tightly linked and mutually required.

6.3 HIV p24 CA and core stability p24 CA is the major structural protein present within the viral core in mature virions prior to infection. However, early biochemical analysis studies were unable to detect CA protein in late replication complexes within the cell (Karageorgos et al., 1993). While this led to suggestions all CA protein was lost from the core soon after entry, uncoating assays demonstrate CA is associated within viral complexes up to 4 hr following fusion (Hulme et al., 2010; Yamashita et al., 2007). Subsequently, CA protein has been detected in RTCs/PICs

(Dismuke & Aiken, 2006; McDonald et al., 2002) and intact cores have been visualised at the nuclear pore inside infected cells (Arhel et al., 2007).

The isolation of cores from numerous viruses, such as AMV, Semliki Forest virus and HBV, have been achieved utilising sucrose gradient sedimentation in conjunction with disrupting agents, including both ionic and non-ionic detergents. The extreme instability and sensitivity 119

to detergent displayed by HIV-1 cores presents considerable disadvantages compared to studying many other viruses. This is further complicated by the increased stability of immature cores which can also be present in core preparations. Thus, while abundant preparations of intact viral cores can be generated from many viruses, studying HIV cores has proven to be considerably more difficult.

One method of analysing HIV-1 cores utilises a modified spin-thru method first described for

AMV (Stromberg, 1972). This involves sedimentation of purified virus stocks through a layer of detergent into a sucrose gradient. Sedimenting virus through a layer of detergent rather than incubation directly in detergent prior to sedimentation aims to remove the viral envelope, while minimising the exposure of the virus to detergent to prevent premature dissociation of the core. Variations of this method have been successful in isolating HIV-1 cores to analyse biochemical properties (Auewarakul et al., 2005; Forshey et al., 2002; Wacharapornin et al.,

2007), to demonstrate constituents of the viral core and to investigate core structures via EM

(Accola et al., 2000; Welker et al., 2000).

In the current study, a modified version of the spin-thru technique involving rate zonal ultracentrifugation was designed to investigate the effect of mutations in CA on HIV-1 core stability and to study a potential role for these sites in disassembly of the viral core after entry into the cell.

A spin-thru system was optimised to differentiate dissociated (free) CA versus complex associated CA, representing cores or intact virions. While free CA and complex-associated

CA sedimented differently within the gradient (Figure 4.1), neither sedimentation profiles nor the presence of the viral envelope were completely successful in distinguishing viral cores from intact virions. However, dissociation of gp120 suggests these detergents treatments are capable of removing the viral envelope (Figure 4.6).

CA mutants E128A/R132A and K203A were generated as controls for the analysis of in vitro core stability (Chapter 4), as a previous study reported these viruses possess hyperstable and hypostable virus cores respectively (Forshey et al., 2002). K203 (like S178 and T188) is

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situated on the CTD dimerisation interface and is involved in hydrophilic interactions with other residues (forming a salt bridge with D152 (Worthylake et al., 1999)). However, unlike

S178A and T188A, analytical gel filtration chromatography showed replacing K203 with alanine does not affect CA CTD dimerisation (del Alamo et al., 2003). The E128A/R132A

CA mutant includes two substitutions within helix 7 which is suggested to be involved in

NTD interactions in the immature virion (Kelly et al., 2006). Analysis of the rate of assembly of purified CA protein indicates that the double mutant assembles at the same rate as WT virus. Single mutant E128A assembles at twice the rate of WT virus, while single mutant

R132A assembles at half the rate of WT virus. These mutations did not affect secondary structure, stability or dimerisation of the CA monomer, but the rate of assembly changed with altered salt concentrations suggesting a role for electrostatic effects (Douglas et al., 2004).

Both these CA mutant viruses displayed severely reduced replication competence and reduced early and late reverse transcription intermediates in comparison to WT NL4-3 virus (data not shown). This effect on viral replication further implies the importance of charged residues, and especially those residues which form the NTD and CTD dimerisation interfaces.

In my study, the spin-thru assay generated CA protein profiles for WT NL4-3 virus which were distinct from those for the hyperstable E128A/R132A virus (Figure 4.7). However, CA mutant virus K203A, which were previously characterised as highly unstable following spin- thru ultracentrifugation (Forshey et al., 2002; Wacharapornin et al., 2007), behaved similarly to WT NL4-3 virus under all conditions tested in the present study (0-1% Triton-X-100)

(Figure 4.8). As CA mutant K203A did not appear to be hypostable, it could not be used as a control to distinguish virus cores that showed reduced in vitro core stability compared to WT

NL4-3 virus.

Analysis of in vitro core stability indicated mutations in CA at S41A, S146A and T188V did not alter effect the core stability of these viruses (Figure 4.10-11 and 4.14-15).

S41A showed replication competence similar to WT, although the production of reverse transcription intermediates was reduced. While reports suggest this virus may display varied

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replication capacity in different cell types, this does not appear to be the result of altered core stability. S146A has been examined previously as part of a study investigating the residues in the flanking region between the CA NTD and CTD domains. In agreement with our study,

S146A showed WT core stability and additionally TEM analysis demonstrated normal virion morphology (Jiang et al., 2011). It has been suggested that regulation of charge at the CA

CTD interface might be able to induce disassembly of the core. Although T188A substitution has been shown to result in increased affinity at the CTD dimerisation interface (del Alamo et al., 2003), T188V substitution did not appear to affect the stability of the core in this study.

While analysis of in vitro core stability for CA mutant viruses S109A, S149A and S178A, indicated these viruses were not hyperstable compared to WT virus. These viruses showed greater susceptibility to higher concentrations of detergent than WT virus (Figure 4.9 and

4.12). Additionally, these three CA mutant viruses also showed an increase in the amount of

CA protein released at low concentrations of Triton-X-100 in comparison to WT NL4-3 virus

(Figure 4.13 and 4.16). This implies these viruses exhibit decreased in vitro core stability.

This finding was confirmed in reports by Wacharapornin et al (2007) and Jiang et al (2011)

(Jiang et al., 2011; Wacharapornin et al., 2007). CA mutants S109A, S149A and S178A showed decreased core yield (recovery of less cores suggests lower core stability), but is not consistent with a report of decreased uncoating activity (less CA protein released during exposure to detergent) when compared with WT virus (Wacharapornin et al., 2007) (see summary Table 6.1).

Brun et al (2008) analysed the CA protein released following incubation at 37ºC after detergent stripping the viral envelope (Brun et al., 2008). Somewhat inconsistent with results from Wacharapornin et al (2007), Brun et al (2008) reported WT virus and S149A showed similar uncoating activity; but consistent with data in this thesis, S178A showed increased release of CA (Brun et al., 2008; Wacharapornin et al., 2007). The CA mutant S109A was not analysed in the Brun et al (2008) study. Furthermore, data presented in this thesis is consistent with the report that none of these viruses (S109A, S149A and S178A) could saturate TRIM5α

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Core Core Uncoating Uncoating CA Mutant Stability a Yield b Activity b Activity c WT ++ ++ ++ ++ S109A + + + n/a S149A + + + ++ S178A + + + +++

Table 6.1 Summary of core stability and CA dissociation for CA mutant viruses +++ = increase compared to WT, ++ = WT, + = reduction compared to WT, – = severe reduction compared to WT.

Definitions: a Results from this thesis (Chapter 4) Core Stability: as determined by detergent susceptibility profiles of CA mutant viruses. CA release following exposure to detergent. b Wacharapornin et al (2007) Virology Core Yield: concentrated virions were spun through 10% sucrose containing 0.1% Igepal into a 50% cushion. Pelletable CA is quantitated by ELISA to give core yield. Uncoating Activity: release of CA following incubation of purified cores with a cell lysate. c Brun et al (2008) Retrovirology Uncoating Activity: release of CA following incubation of purified cores in buffer.

restriction activity following challenge with GFP-virus, indicative of altered CA structures and lowered stability of the core in comparison to WT NL4-3 virus (Brun et al., 2008; Jiang et al., 2011). Analysis of core stability profiles for each of the viruses using TRIM5alpha restriction assays would allow confirmation of core stability characterisation for the viruses in our study.

The contrasting results across these three studies are indicative of the difficulties analysing this stage of the HIV life cycle. However, overall this thesis showed that introducing mutations in CA at S109A, S149A and S178A does indeed alter the in vitro core stability of these viruses, possibly by preventing phosphorylation of HIV-1 CA.

6.4 HIV core disassembly during infection

Analysis of in vitro core stability indicates CA mutations leading to viral core defects were also associated with blocks in viral replication. Viruses that display altered core stability properties in vitro are often impaired during replication, e.g. during reverse transcription or nuclear translocation (Brun et al., 2008; Dismuke & Aiken, 2006; Forshey et al., 2002;

Wacharapornin et al., 2007). However, core stability in vitro may not necessarily reflect the natural process of core disassembly inside the infected cell. The viral core enters the cell at least partially intact and may remain present for a period of time inside the cytoplasm before regulated disassembly (Arhel et al., 2007; McDonald et al., 2002). Some CA protein remains in RTCs (Iordanskiy et al., 2006; McDonald et al., 2002) and PICs to assist with nuclear localisation (Dismuke & Aiken, 2006). In Chapter 5, the ‘fate-of-capsid’ assay was adapted to investigate complex-associated CA and any defects during disassembly of the viral core. This assay was originally used to investigate the method of restriction employed by the retroviral restriction factor TRIM5α. TRIM5α restricts HIV infection by accelerating HIV-1 core disassembly (Chatterji et al., 2006; Stremlau et al., 2006). Analysis of intracellular CA demonstrated decreasing levels of CA protein following infection. This decrease was exaggerated in the presence of TRIM5α (Chatterji et al., 2006). It was reasoned that by using 123

this approach, viruses with altered core disassembly properties might also be identified following infection by tracing the fate of CA protein within the infected cell. Complex- associated CA was identified following sucrose gradient sedimentation, SDS PAGE and western blot, while CA released from the core is assumed to be rapidly degraded by cellular mechanisms.

Thus cores which disassemble at different rates compared to WT NL4-3 virus could be identified by their CA profile in this assay. Intracellular CA protein was detected in cells 2-6 hr following infection and CA levels generally decreased within this time period. However, results were inconsistent between experiments and firm conclusions could not be drawn from the data obtained. While other studies in the literature demonstrated a consistent decrease in intracellular CA present following infection, experiments presented in this thesis were not able to detect a substantial or reproducible decrease within 6 hr of infection by WT NL4-3 virus (Figure 5.3). The sedimentation of core-associated CA did not change substantially within 6 hr p.i. (Figure 5.4), somewhat unexpectedly since restructuring to generate the RTC and loss of the majority of CA would be expected to occur within the early hours following infection.

Analysis of intracellular CA following infection with the K203A and E128A/R132A CA mutant viruses, which have been previously reported to exhibit hypostable and hyperstable cores respectively, demonstrated only slight difference in CA profiles compared to WT NL4-

3 virus (Figure 5.5). While the amount of intracellular CA detected decreased 2-6 hr p.i. for each of these viruses, CA profiles did not correspond with characterisation of in vitro core stability (Figure 4.7 and 4.8).

Similar studies in the literature use VSV-G pseudotyped viruses to increase viral entry.

However, VSV-G mediates virus entry by endocytosis while HIV utilises fusion-mediated entry, and therefore VSV-G would not accurately reflect HIV Env-mediated entry and any associated defects in core disassembly of these viruses. Additionally, uncoating defects can be rescued by VSV-G pseudotyping (e.g. S149A) (Brun et al., 2008).

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Rather than using pseudotyped virus, other methods were considered to enhance HIV infection. Centrifugal enhancement of infection and temperature-arrested state of fusion

(TAS) conditions were used to increase the efficiency of infection (Figure 5.2). While these increased the detection of intracellular CA protein following infection, the levels of CA protein detected varied significantly between infections (Figure 5.3). The variation might be explained by a lack of synchronous infection or alternatively a proportion of virus that do not undergo productive infection, which may also be detected in this assay unless they are rapidly targeted for degradation.

The persistence of cores from viruses that are blocked before the final stage of reverse transcription (central flap formation) (Arhel et al., 2007), suggests that viral complexes which are not directed to the correct compartment, or take part in the proper interactions with host cell factors within the cell cytoplasm, may remain intact for sometime before being targeted for degradation. Such complexes would also contribute to detection of CA in this assay.

Alternatively, core disassembly inside the cell could be followed using super resolution or live cell microscopy techniques to track the persistence and dynamics of CA inside the cell post-fusion (Koch et al., 2009; Pereira et al., 2012).

6.5 Concluding remarks

HIV-1 CA residues S109, S149, S178 and T188 play critical roles in HIV-1 infection and are required during the early steps in viral replication prior to integration. While S41A and

S146A show mild defects in reverse transcription that do not block replication.

The results from this thesis (Chapter 3) indicate substitution of these CA residues affect reverse transcription. Substitution at two sites (S109A and S178A) reduced the synthesis of reverse transcription intermediates. Substitution at S149 appeared to enhance reverse transcription. Literature suggests CA is present in RTCs, and mutation of the HIV-1 CA protein can block viral replication at reverse transcription and nuclear translocation (Dismuke

& Aiken, 2006; Forshey et al., 2002). A role for CA has not been definitively linked with 125

initiation, elongation or completion of reverse transcription. However, CA within the viral complexes is associated with nuclear pore proteins that facilitate nuclear import and integration. Most likely CA exerts these effects through regulation of core disassembly prior to or concomitantly with these steps. Premature uncoating can result damage to viral complexes through loss of viral core proteins and viral transcripts (Kutluay et al., 2013).

This study (Chapter 4) and studies by others (Brun et al., 2008; Jiang et al., 2011;

Wacharapornin et al., 2007) indicate alanine substitution at serine residues S109, S149 and

S178 alters in vitro core stability, but results are somewhat conflicting. Viruses displaying both increased and reduced kinetics of release of CA from isolated cores also show blocks in replication (reverse transcription and nuclear translocation) (Dismuke & Aiken, 2006;

Forshey et al., 2002). How CA affects reverse transcription is poorly understood; but it is probable that any disruption to core disassembly is likely to block replication by altering CA interactions with cellular factors or disrupting the reorganisation of replication complexes in the infected cell.

This study (Chapter 4) suggests increased release of CA from cores in these CA mutant viruses. Recent reports that VSV-G pseudotyping rescues infectivity of CA mutants S149A and S178A suggests that substitution at these sites disrupts the early stages of replication, in particular core disassembly (Brun et al., 2008).

Regulation of core disassembly is tightly controlled. Reverse transcription relies upon access to dNTPs and cellular factor(s) which may require dissociation of CA or at the very least some restructuring of the core. An important insight from core stability studies is that altering charge impacts on stability of the HIV-1 core, and thus regulation of charge by phosphorylation could indeed play a role in regulation of disassembly of the HIV core.

As it stands, more knowledge is needed about the regulation of HIV-1 core disassembly. In vitro core stability and core disassembly studies will help further our knowledge regarding

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this stage of the viral life cycle and development of antiretroviral inhibitors to restrict HIV replication.

6.6 Future work

Further characterisation of the methods used in this thesis would help to understand the process of core disassembly.

This thesis investigated six HIV-1 CA mutant viruses, including three sites that are phosphorylated in the virion and that display defects in the virus life cycle (S41A, S109A,

S146A, S149A, S178A and T188V). It is just as likely that the defects mediated by alanine substitution result from alterations to bonding within the core, as from phosphorylation. Any substitutions introduced into CA might also affect assembly of these viruses. It must not be excluded that these mutations influence core stability, and this is just as likely to reflect defects during formation of the core. Various assays exist to investigate assembly of CA monomers, and also the transformation of CA into higher order structures, such as cylinders and spheres. However, substitution with negatively charged amino acids such as aspartic acid might to mimic phosphorylation, or combinations of these mutations might also provide further insights into a role for these residues and the potential effect of phosophorylation.

We assumed that the concentration of HIV virions in our preparations were not great enough to allow detection and isolation of cores during isopycnic centrifugation experiments. Greater quantities of input virus and further concentration of virus preparations should enable generation of a detectable amount of cores. Without relying on the density of the resulting complexes, other characteristics are essential to identify the resulting CA-containing complexes as cores. Greater quantities of input virus would also aid detection of gp120 in core preparations, vital to ensure proper characterisation of viral cores versus virions.

The isolation of cores without the use of detergent is suggested to be a superior method in comparison to the spin thru method, and should avoid the artificial loss of cores proteins.

Generation of sufficient core preparations using these methods might enable analysis of CA 127

phosphorylation in isolated cores. Several CA isoforms are detected in whole virus.

Examining the phosphorylation status of CA in isolated cores and following infection using techniques such as 2D PAGE or mass spectrometry, from both WT virus and CA mutants in this study, could shed light on modifications to CA after entry that might be associated with core disassembly. This would help to clarify if the effect is potentially mediated through phosphorylation rather than disruption of core formation due to amino acid substitution.

Rate zonal ultracentrifugation in combination with spin-thru sedimentation successfully demonstrated increasing release of CA in response to increasing concentrations of detergent.

Analysis of the ability of each virus to saturate TRIM5α restriction would provide confirmation of in vitro core stability profiles. This has already been performed for some of the CA mutant viruses in this study. Additionally, infection in combination with depletion of

CypA or nuclear pore proteins e.g. TNPO3, that are associated with CA and maintenance of the core might yield further insights as to the role played by these reisudes during early infection.

What triggers disassembly of the core and the step-by-step mechanisms involved are still unknown. However, current literature is beginning to outline factors which may prompt disassembly of the viral core: dNTPs and other cellular factors can induce release of some viral core proteins. Viral complexes containing CA associate with actin filaments en route to the nucleus. CA interactions with some cellular factors appear to be linked to the intrinsic stability of the core, further implying the significance of this viral characteristic. As investigation of the viral core progresses, attention has again been drawn to isomerisation of

CA as a potential trigger of uncoating of the core, and CA is associated with several promising candidates which include CypA, NUP358 & PIN1.

CypA appears to be instrumental in maintenance of the core. CA interactions mediated through CypA appear to protect the viral core from premature disassembly in response to some cellular factors, e.g. TRIM5 and CPF56. CypA association with dynein protein

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complexes might tether viral complexes to the cytoskeleton and allow trafficking through the cytoplasm toward the nucleus. Where interactions with nuclear pore proteins such as NP358 and TNPO3 facilitate nuclear import and reorganisation of the PIC and influence integration of proviral DNA, but the story is not yet complete.

Slowly but surely, a model of the basic mechanisms involved in disassembly of the core is being developed. This will prove a useful tool for furthering our understanding of this step in the HIV life cycle, and additionally potential ways to exploit this process to control viral replication.

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Appendix 1. Plasmid map of HIV-1 pNL4-3, NY5/BRU (LAV-1) recombinant clone. Accession number M19921 HIV-1 pNL4-3 sequence is represented by the white box. The CA region (nt 1186-1878) is represented by the red box. Relevant restriction enzyme sites used for cloning and for identifying CA mutant clones (see Table 2.2) are indicated. HindIII 532 BssHIIBssHII 712 712 SphI SphI1448 1448 SpeI 1508 CAPSID nt 1186-1878 HindIII 1713 ApaI 2011 2011 SbfI 2844

pNL4-3 14879 bp KpnI 3831 KpnI 4159

HindIII 9607

KpnI 9010 HindIII 6027

HindIII 8132 KpnI 6348 Appendix 2. CA constructs generated by PCR mutagenesis (A) pNL4-3 sequence was cloned into pET-32a vectors using the restriction enzymes indicated. (B) pET-32-CA(BA) contains BssHII/ApaI HIV-1 pNL4-3 CA insert (nt 712- 2011, 1300 bp insert), this includes entire CA region (amino acids 1-231). (C) pET-32-CA(SA) contains SphI/ApaI HIV-1 CA pNL4-3 insert (nt 1448- 2011, 564 bp insert), contains only amino acids 89-231 of CA. A BssHII 712 ApaI 2011

CAPSID

SphI 1448

BssHII3965 BssHII (pNL4-3 nt 712) B

SphI4701 (pNL4SphI -3 nt 1448) CAPSID

(pNL4-3 nt 1186-1878) 6991 bp 6991

ApaI 5264 ApaI pET32a(BA) (BA)pET32a ApaI (pNL4-3 nt 2011) 6991bp

SphI (pNL4-3 nt 1448) C SphI 997 CAPSIDCAPSID (pNL4-3 nt 1448-1878)

ApaI 1560 (pNL4-3 nt 2011)

pET32a (SA) pET32a5727bp5727 (SA) bp Appendix 3. Sequence analysis of pNL4-3 CA mutant clones CA mutant clones generated by PCR mutagenesis were sequenced to ensure only the desired nucleotide changes. Sequences are aligned using pNL4-3 as the reference sequence. Relevant restriction sites are indicated with an arrow, unique restriction sites with an arrow and asterisk. Sequencing primers are indicated above the sequence. BssHII 710 720 730 740 750 760 770 780 790 800 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 GCTTGCTGAA GCGCGCACGG CAAGAGGCGA GGGGCGGCGA CTGGTGAGTA CGCCAAAAAT TTTGACTAGC GGAGGCTAGA AGGAGAGAGA TGGGTGCGAG S41A ...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------s816 s843 810 820 830 840 850 860 870 880 890 900 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 AGCGTCGGTA TTAAGCGGGG GAGAATTAGA TAAATGGGAA AAAATTCGGT TAAGGCCAGG GGGAAAGAAA CAATATAAAC TAAAACATAT AGTATGGGCA S41A ...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------

910 920 930 940 950 960 970 980 990 1000 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 AGCAGGGAGC TAGAACGATT CGCAGTTAAT CCTGGCCTTT TAGAGACATC AGAAGGCTGT AGACAAATAC TGGGACAGCT ACAACCATCC CTTCAGACAG S41A ...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------1010 1020 1030 1040 1050 1060 1070 1080 1090 1100 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 GATCAGAAGA ACTTAGATCA TTATATAATA CAATAGCAGT CCTCTATTGT GTGCATCAAA GGATAGATGT AAAAGACACC AAGGAAGCCT TAGATAAGAT S41A ...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------

1110 1120 1130 1140 1150 1160 1170 1180 1190 1200 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 AGAGGAAGAG CAAAACAAAA GTAAGAAAAA GGCACAGCAA GCAGCAGCTG ACACAGGAAA CAACAGCCAG GTCAGCCAAA ATTACCCTAT AGTGCAGAAC S41A ...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------

1210 1220 1230 1240 1250 1260 1270 1280 1290 1300 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 CTCCAGGGGC AAATGGTACA TCAGGCCATA TCACCTAGAA CTTTAAATGC ATGGGTAAAA GTAGTAGAAG AGAAGGCTTT CAGCCCAGAA GTAATACCCA S41A ...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------AfeI* s1369 1310 1320 1330 1340 1350 1360 1370 1380 1390 1400 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 TGTTTTCAGC ATTATCAGAA GGAGCCACCC CACAAGATTT AAATACCATG CTAAACACAG TGGGGGGACA TCAAGCAGCC ATGCAAATGT TAAAAGAGAC S41A .....G.... GC...... S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ------SphI* 1410 1420 1430 1440 1450 1460 1470 1480 1490 1500 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 CATCAATGAG GAAGCTGCAG AATGGGATAG ATTGCATCCA GTGCATGCAG GGCCTATTGC ACCAGGCCAG ATGAGAGAAC CAAGGGGAAG TGACATAGCA S41A ...... S109A ------...... S146A ------...... S149A ------...... S178A ------...... T188V ------...... E128A/R132A ------...... K203A ------...... SpeI(-)* 1510 1520 1530 1540 1550 1560 1570 1580 1590 1600 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 GGAACTACTA GTACCCTTCA GGAACAAATA GGATGGATGA CACATAATCC ACCTATCCCA GTAGGAGAAA TCTATAAAAG ATGGATAATC CTGGGATTAA S41A ...... ------S109A ...... G C...... S146A ...... S149A ...... S178A ...... T188V ...... E128A/R132A ...... C...... GC T...... K203A ...... ClaI* XbaI* 1610 1620 1630 1640 1650 1660 1670 1680 1690 1700 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 ATAAAATAGT AAGAATGTAT AGCCCTACCA GCATTCTGGA CATAAGACAA GGACCAAAGG AACCCTTTAG AGACTATGTA GACCGATTCT ATAAAACTCT S41A ------S109A ...... S146A ...... GC...... AT CG...... S149A ...... G C...... A...... S178A ...... T188V ...... E128A/R132A ...... K203A ...... * HindIII(-) SpeI* 1710 1720 1730 1740 1750 1760 1770 1780 1790 1800 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 AAGAGCCGAG CAAGCTTCAC AAGAGGTAAA AAATTGGATG ACAGAAACCT TGTTGGTCCA AAATGCGAAC CCAGATTGTA AGACTATTTT AAAAGCATTG S41A ------S109A ...... S146A ...... S149A ...... S178A ...... G...... T188V ...... GT.C .AC.A...... E128A/R132A ...... K203A ...... GC...... s1880 1810 1820 1830 1840 1850 1860 1870 1880 1890 1900 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 GGACCAGGAG CGACACTAGA AGAAATGATG ACAGCATGTC AGGGAGTGGG GGGACCCGGC CATAAAGCAA GAGTTTTGGC TGAAGCAATG AGCCAAGTAA S41A ------S109A ...... S146A ...... S149A ...... S178A ...... T188V ...... E128A/R132A ...... K203A ..C...... 1910 1920 1930 1940 1950 1960 1970 1980 1990 2000 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 CAAATCCAGC TACCATAATG ATACAGAAAG GCAATTTTAG GAACCAAAGA AAGACTGTTA AGTGTTTCAA TTGTGGCAAA GAAGGGCACA TAGCCAAAAA S41A ------S109A ...... S146A ...... S149A ...... S178A ...... T188V ...... E128A/R132A ...... K203A ...... ApaI s2081 2010 2020 2030 2040 2050 2060 2070 2080 2090 2100 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 TTGCAGGGCC CCTAGGAAAA AGGGCTGTTG GAAATGTGGA AAGGAAGGAC ACCAAATGAA AGATTGTACT GAGAGACAGG CTAATTTTTT AGGGAAGATC S41A ------S109A ...... S146A ...... ------S149A ...... S178A ...... T188V ...... ------E128A/R132A ...... K203A ......

2110 2120 2130 2140 2150 2160 2170 2180 2190 2200 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 TGGCCTTCCC ACAAGGGAAG GCCAGGGAAT TTTCTTCAGA GCAGACCAGA GCCAACAGCC CCACCAGAAG AGAGCTTCAG GTTTGGGGAA GAGACAACAA S41A ------S109A ...... ------S146A ------S149A ...... ------S178A ...... ------T188V ------E128A/R132A ...... ------K203A ...... 2210 2220 2230 2240 2250 2260 2270 2280 2290 2300 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 CTCCCTCTCA GAAGCAGGAG CCGATAGACA AGGAACTGTA TCCTTTAGCT -TCCCTCAGA TCACTCTTTG GCAGCGACCC CTCGTCACAA TAAAGATAGG S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ...... -...... s2365 2310 2320 2330 2340 2350 2360 2370 2380 2390 2400 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 GGGGCAATTA AAGGAAGCTC TATTAGATAC AGGAGCAGAT GATACAGTAT TAGAAGAAAT GAATTTGCCA GGAAGATGGA AACCAAAAAT GATAGGGGGA S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ......

2410 2420 2430 2440 2450 2460 2470 2480 2490 2500 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ... .|....| ....|....| NL4-3 ATTGGAGGTT TTATCAAAGT AGGACAGTAT GATCAGATAC TCATAGAAAT CTGCGGACAT AAAGCTATAG GTACAGTATT AGTAGGACCT ACACCTGTCA S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ...... A...... 2510 2520 2530 2540 2550 2560 2570 2580 2590 2600 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 ACATAATTGG AAGAAATCTG TTGACTCAGA TTGGCTGCAC TTTAAATTTT CCCATTAGTC CTATTGAGAC TGTACCAGTA AAATTAAAGC CAGGAATGGA S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ......

2610 2620 2630 2640 2650 2660 2670 2680 2690 2700 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 TGGCCCAAAA GTTAAACAAT GGCCATTGAC AGAAGAAAAA ATAAAAGCAT TAGTAGAAAT TTGTACAGAA ATGGAAAAGG AAGGAAAAAT TTCAAAAATT S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ......

2710 2720 2730 2740 2750 2760 2770 2780 2790 2800 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 GGGCCTGAAA ATCCATACAA TACTCCAGTA TTTGCCATAA AGAAAAAAGA CAGTACTAAA TGGAGAAAAT TAGTAGATTT CAGAGAACTT AATAAGAGAA S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ...... SbfI 2810 2820 2830 2840 2850 2860 2870 2880 2890 2900 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 CTCAAGATTT CTGGGAAGTT CAATTAGGAA TACCACATCC TGCAGGGTTA AAACAGAAAA AATCAGTAAC AGTACTGGAT GTGGGCGATG CATATTTTTC S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ......

2910 2920 2930 2940 2950 2960 2970 2980 2990 3000 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 AGTTCCCTTA GATAAAGACT TCAGGAAGTA TACTGCATTT ACCATACCTA GTATAAACAA TGAGACACCA GGGATTAGAT ATCAGTACAA TGTGCTTCCA S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ......

3010 3020 3030 3040 3050 3060 3070 3080 3090 3100 ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| ....|....| NL4-3 CAGGGATGGA AAGGATCACC AGCAATATTC CAGTGTAGCA TGACAAAAAT CTTAGAGCCT TTTAGAAAAC AAAATCCAGA CATAGTCATC TATCAATACA S41A ------S109A ------S146A ------S149A ------S178A ------T188V ------E128A/R132A ------K203A ...... BIBLIOGRAPHY

(1981). Pneumocystis pneumonia--Los Angeles. MMWR Morb Mortal Wkly Rep 30, 250-252.

(2003). AIDS epidemic update: December 2003, p. ~30: Joint United Nations Programme on

HIV/AIDS & World Health Organisation.

(2007). AIDS epidemic update December 2007: Joint United Nations Programme on

HIV/AIDS (UNAIDS) and World Health Organization (WHO) 2007.

(2008a). HIV Sequence Compendium 2008

Edited by F. B. Leitner T, Hahn B, Marx P, McCutchan F, Mellors J, Wolinsky S, and Korber

B,: Theoretical Biology and Biophysics Group, Los Alamos National Laboratory, NM, LA-

UR 06-0680.

(2008b). The Official Web Site of the Nobel Foundation: Nobel Foundation.

(2012a). HIV, viral hepatitis and sexually transmissible infections in Australia Annual

Surveillance

Report 2012, p. 148. Edited by A. McDonald. Sydney: The Kirby Institute.

(2012b). October 2012 Australian HIV Surveillance Report: The Kirby Institute.

(UNAIDS), J. U. N. P. o. H. A. (2010). Global report: UNAIDS report on the global AIDS epidemic 2010.

(UNAIDS), J. U. N. P. o. H. A. (2012). Global report: UNAIDS report on the global AIDS epidemic 2012, p. 212.

(UNAIDS), J. U. N. P. o. H. A. (2013). UNAIDS Special report How Africa turned AIDS around. 56.

Accola, M. A., Ohagen, A. & Gottlinger, H. G. (2000). Isolation of human immunodeficiency virus type 1 cores: retention of Vpr in the absence of p6(gag). Journal of virology 74, 6198-

6202.

Agrawal, L., Lu, X., Jin, Q. & Alkhatib, G. (2006). Anti-HIV therapy: Current and future directions. Curr Pharm Des 12, 2031-2055.

140

Alkhatib, G., Broder, C. C. & Berger, E. A. (1996a). Cell type-specific fusion cofactors determine human immunodeficiency virus type 1 tropism for T-cell lines versus primary macrophages. Journal of virology 70, 5487-5494.

Alkhatib, G., Combadiere, C., Broder, C. C., Feng, Y., Kennedy, P. E., Murphy, P. M. &

Berger, E. A. (1996b). CC CKR5: a RANTES, MIP-1alpha, MIP-1beta receptor as a fusion cofactor for macrophage-tropic HIV-1. Science (New York, NY 272, 1955-1958.

Allen, T. D., Cronshaw, J. M., Bagley, S., Kiseleva, E. & Goldberg, M. W. (2000). The nuclear pore complex: mediator of translocation between nucleus and cytoplasm. Journal of cell science 113 ( Pt 10), 1651-1659.

Ammosova, T., Jerebtsova, M., Beullens, M., Voloshin, Y., Ray, P. E., Kumar, A., Bollen, M.

& Nekhai, S. (2003). Nuclear protein phosphatase-1 regulates HIV-1 transcription. The

Journal of biological chemistry 278, 32189-32194.

Anders, D. G. & Consigli, R. A. (1983). Comparison of nonphosphorylated and phosphorylated species of polyomavirus major capsid protein VP1 and identification of the major phosphorylation region. Journal of virology 48, 206-217.

Arhel, N. J., Souquere-Besse, S., Munier, S., Souque, P., Guadagnini, S., Rutherford, S.,

Prevost, M. C., Allen, T. D. & Charneau, P. (2007). HIV-1 DNA Flap formation promotes uncoating of the pre-integration complex at the nuclear pore. The EMBO journal 26, 3025-

3037.

Arya, S. K., Guo, C., Josephs, S. F. & Wong-Staal, F. (1985). Trans-activator gene of human

T-lymphotropic virus type III (HTLV-III). Science (New York, NY 229, 69-73.

Auewarakul, P., Wacharapornin, P., Srichatrapimuk, S., Chutipongtanate, S. & Puthavathana,

P. (2005). Uncoating of HIV-1 requires cellular activation. Virology 337, 93-101.

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

Dauguet, C., Axler-Blin, C., Vezinet-Brun, F., Rouzioux, C., Rozenbaum, W. & Montagnier,

L. (1983). Isolation of a T-lymphotropic retrovirus from a patient at risk for acquired immune deficiency syndrome (AIDS). Science (New York, NY 220, 868-871.

141

Barrera, F. N., del Alamo, M., Mateu, M. G. & Neira, J. L. (2008). Envelope lipids regulate the in vitro assembly of the HIV-1 capsid. Biophys J 94, L8-10.

Battivelli, E., Lecossier, D., Clavel, F. & Hance, A. J. (2013). Delaying reverse transcription does not increase sensitivity of HIV-1 to human TRIM5alpha. PloS one 8, e52434.

Benjamin, J., Ganser-Pornillos, B. K., Tivol, W. F., Sundquist, W. I. & Jensen, G. J. (2005).

Three-dimensional structure of HIV-1 virus-like particles by electron cryotomography.

Journal of molecular biology 346, 577-588.

Berger, E. A., Doms, R. W., Fenyo, E. M., Korber, B. T., Littman, D. R., Moore, J. P.,

Sattentau, Q. J., Schuitemaker, H., Sodroski, J. & Weiss, R. A. (1998). A new classification for HIV-1. Nature 391, 240.

Berthet-Colominas, C., Monaco, S., Novelli, A., Sibai, G., Mallet, F. & Cusack, S. (1999).

Head-to-tail dimers and interdomain flexibility revealed by the crystal structure of HIV-1 capsid protein (p24) complexed with a monoclonal antibody Fab. The EMBO journal 18,

1124-1136.

Berthoux, L., Sebastian, S., Sokolskaja, E. & Luban, J. (2005). Cyclophilin A is required for

TRIM5{alpha}-mediated resistance to HIV-1 in Old World monkey cells. Proceedings of the

National Academy of Sciences of the United States of America 102, 14849-14853.

Bichel, K., Price, A. J., Schaller, T., Towers, G. J., Freund, S. M. & James, L. C. (2013).

HIV-1 capsid undergoes coupled binding and isomerization by the nuclear pore protein

NUP358. Retrovirology 10, 81.

Black, L. R. & Aiken, C. (2010). TRIM5alpha disrupts the structure of assembled HIV-1 capsid complexes in vitro. Journal of virology 84, 6564-6569.

Borrow, P., Lewicki, H., Hahn, B. H., Shaw, G. M. & Oldstone, M. B. (1994). Virus-specific

CD8+ cytotoxic T-lymphocyte activity associated with control of viremia in primary human immunodeficiency virus type 1 infection. Journal of virology 68, 6103-6110.

Bosco, D. A., Eisenmesser, E. Z., Pochapsky, S., Sundquist, W. I. & Kern, D. (2002).

Catalysis of cis/trans isomerization in native HIV-1 capsid by human cyclophilin A.

142

Proceedings of the National Academy of Sciences of the United States of America 99, 5247-

5252.

Bosco, D. A. & Kern, D. (2004). Catalysis and binding of cyclophilin A with different HIV-1 capsid constructs. Biochemistry 43, 6110-6119.

Braaten, D., Franke, E. K. & Luban, J. (1996). Cyclophilin A is required for an early step in the life cycle of human immunodeficiency virus type 1 before the initiation of reverse transcription. Journal of virology 70, 3551-3560.

Brass, A. L., Dykxhoorn, D. M., Benita, Y., Yan, N., Engelman, A., Xavier, R. J., Lieberman,

J. & Elledge, S. J. (2008). Identification of host proteins required for HIV infection through a functional genomic screen. Science (New York, NY 319, 921-926.

Briggs, J. A., Grunewald, K., Glass, B., Forster, F., Krausslich, H. G. & Fuller, S. D. (2006).

The mechanism of HIV-1 core assembly: insights from three-dimensional reconstructions of authentic virions. Structure 14, 15-20.

Briggs, J. A. & Krausslich, H. G. (2011). The molecular architecture of HIV. Journal of molecular biology 410, 491-500.

Briggs, J. A., Simon, M. N., Gross, I., Krausslich, H. G., Fuller, S. D., Vogt, V. M. &

Johnson, M. C. (2004). The stoichiometry of Gag protein in HIV-1. Nat Struct Mol Biol 11,

672-675.

Briggs, J. A., Wilk, T., Welker, R., Krausslich, H. G. & Fuller, S. D. (2003). Structural organization of authentic, mature HIV-1 virions and cores. The EMBO journal 22, 1707-1715.

Briones, M. S. & Chow, S. A. (2010). A new functional role of HIV-1 integrase during uncoating of the viral core. Immunologic research 48, 14-26.

Briones, M. S., Dobard, C. W. & Chow, S. A. (2010). Role of human immunodeficiency virus type 1 integrase in uncoating of the viral core. Journal of virology 84, 5181-5190.

Broder, S. & Gallo, R. C. (1984). A pathogenic retrovirus (HTLV-III) linked to AIDS. N Engl

J Med 311, 1292-1297.

143

Brown, D. (2002). Structure and function of membrane rafts. Int J Med Microbiol 291, 433-

437.

Brown, F. (1986). Human immunodeficiency virus. Science (New York, NY 232, 1486.

Brun, S., Chaloin, L., Gay, B., Bernard, E., Devaux, C., Lionne, C., Chazal, N. & Briant, L.

(2010). Electrostatic repulsion between HIV-1 capsid proteins modulates hexamer plasticity and in vitro assembly. Proteins 78, 2144-2156.

Brun, S., Solignat, M., Gay, B., Bernard, E., Chaloin, L., Fenard, D., Devaux, C., Chazal, N.

& Briant, L. (2008). VSV-G pseudotyping rescues HIV-1 CA mutations that impair core assembly or stability. Retrovirology 5, 57.

Bryant, M. & Ratner, L. (1990). Myristoylation-dependent replication and assembly of human immunodeficiency virus 1. Proceedings of the National Academy of Sciences of the United

States of America 87, 523-527.

Bukrinskaya, A. G., Ghorpade, A., Heinzinger, N. K., Smithgall, T. E., Lewis, R. E. &

Stevenson, M. (1996). Phosphorylation-dependent human immunodeficiency virus type 1 infection and nuclear targeting of viral DNA. Proceedings of the National Academy of

Sciences of the United States of America 93, 367-371.

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

Haggerty, S. & Stevenson, M. (1992). Active nuclear import of human immunodeficiency virus type 1 preintegration complexes. Proceedings of the National Academy of Sciences of the United States of America 89, 6580-6584.

Bukrinsky, M. I., Sharova, N., McDonald, T. L., Pushkarskaya, T., Tarpley, W. G. &

Stevenson, M. (1993). Association of integrase, matrix, and reverse transcriptase antigens of human immunodeficiency virus type 1 with viral nucleic acids following acute infection.

Proceedings of the National Academy of Sciences of the United States of America 90, 6125-

6129.

144

Bushman, F. D. & Craigie, R. (1991). Activities of human immunodeficiency virus (HIV) integration protein in vitro: specific cleavage and integration of HIV DNA. Proceedings of the

National Academy of Sciences of the United States of America 88, 1339-1343.

Campbell, E. M., Perez, O., Melar, M. & Hope, T. J. (2007). Labeling HIV-1 virions with two fluorescent proteins allows identification of virions that have productively entered the target cell. Virology 360, 286-293.

Capelson, M., Liang, Y., Schulte, R., Mair, W., Wagner, U. & Hetzer, M. W. (2010).

Chromatin-bound nuclear pore components regulate gene expression in higher eukaryotes.

Cell 140, 372-383.

Cartier, C., Deckert, M., Grangeasse, C., Trauger, R., Jensen, F., Bernard, A., Cozzone, A.,

Desgranges, C. & Boyer, V. (1997). Association of ERK2 mitogen-activated protein kinase with human immunodeficiency virus particles. Journal of virology 71, 4832-4837.

Cartier, C., Hemonnot, B., Gay, B., Bardy, M., Sanchiz, C., Devaux, C. & Briant, L. (2003).

Active cAMP-dependent protein kinase incorporated within highly purified HIV-1 particles is required for viral infectivity and interacts with viral capsid protein. The Journal of biological chemistry 278, 35211-35219.

Cartier, C., Sivard, P., Tranchat, C., Decimo, D., Desgranges, C. & Boyer, V. (1999).

Identification of three major phosphorylation sites within HIV-1 capsid. Role of phosphorylation during the early steps of infection. The Journal of biological chemistry 274,

19434-19440.

Chardonnet, Y. & Dales, S. (1972). Early events in the interaction of adenoviruses with HeLa cells. 3. Relationship between an ATPase activity in nuclear envelopes and transfer of core material: a hypothesis. Virology 48, 342-359.

Chatterji, U., Bobardt, M. D., Gaskill, P., Sheeter, D., Fox, H. & Gallay, P. A. (2006).

Trim5alpha accelerates degradation of cytosolic capsid associated with productive HIV-1 entry. The Journal of biological chemistry 281, 37025-37033.

145

Cherepanov, P., Surratt, D., Toelen, J., Pluymers, W., Griffith, J., De Clercq, E. & Debyser,

Z. (1999). Activity of recombinant HIV-1 integrase on mini-HIV DNA. Nucleic Acids Res 27,

2202-2210.

Chermann, J. C., Barre-Sinoussi, F., Dauguet, C., Brun-Vezinet, F., Rouzioux, C.,

Rozenbaum, W. & Montagnier, L. (1983). Isolation of a new retrovirus in a patient at risk for acquired immunodeficiency syndrome. Antibiot Chemother 32, 48-53.

Chertova, E., Chertov, O., Coren, L. V., Roser, J. D., Trubey, C. M., Bess, J. W., Jr., Sowder,

R. C., 2nd, Barsov, E., Hood, B. L., Fisher, R. J., Nagashima, K., Conrads, T. P., Veenstra, T.

D., Lifson, J. D. & Ott, D. E. (2006). Proteomic and biochemical analysis of purified human immunodeficiency virus type 1 produced from infected monocyte-derived macrophages.

Journal of virology 80, 9039-9052.

Christ, F., Thys, W., De Rijck, J., Gijsbers, R., Albanese, A., Arosio, D., Emiliani, S., Rain, J.

C., Benarous, R., Cereseto, A. & Debyser, Z. (2008). Transportin-SR2 imports HIV into the nucleus. Curr Biol 18, 1192-1202.

Cicala, C., Arthos, J., Ruiz, M., Vaccarezza, M., Rubbert, A., Riva, A., Wildt, K., Cohen, O.

& Fauci, A. S. (1999). Induction of phosphorylation and intracellular association of CC chemokine receptor 5 and focal adhesion kinase in primary human CD4+ T cells by macrophage-tropic HIV envelope. J Immunol 163, 420-426.

Ciuffi, A., Llano, M., Poeschla, E., Hoffmann, C., Leipzig, J., Shinn, P., Ecker, J. R. &

Bushman, F. (2005). A role for LEDGF/p75 in targeting HIV DNA integration. Nat Med 11,

1287-1289.

Clarke, J. N., Lake, J. A., Burrell, C. J., Wesselingh, S. L., Gorry, P. R. & Li, P. (2006).

Novel pathway of human immunodeficiency virus type 1 uptake and release in astrocytes.

Virology 348, 141-155.

Clavel, F., Guetard, D., Brun-Vezinet, F., Chamaret, S., Rey, M. A., Santos-Ferreira, M. O.,

Laurent, A. G., Dauguet, C., Katlama, C., Rouzioux, C. & et al. (1986). Isolation of a new

146

human retrovirus from West African patients with AIDS. Science (New York, NY 233, 343-

346.

Clerici, M., Lucey, D. R., Berzofsky, J. A., Pinto, L. A., Wynn, T. A., Blatt, S. P., Dolan, M.

J., Hendrix, C. W., Wolf, S. F. & Shearer, G. M. (1993). Restoration of HIV-specific cell- mediated immune responses by interleukin-12 in vitro. Science (New York, NY 262, 1721-

1724.

Coffin, J., Haase, A., Levy, J. A., Montagnier, L., Oroszlan, S., Teich, N., Temin, H.,

Toyoshima, K., Varmus, H., Vogt, P. & et al. (1986). Human immunodeficiency viruses.

Science (New York, NY 232, 697.

Cohen, O. J., Kinter, A. & Fauci, A. S. (1997). Host factors in the pathogenesis of HIV disease. Immunol Rev 159, 31-48.

Collman, R., Hassan, N. F., Walker, R., Godfrey, B., Cutilli, J., Hastings, J. C., Friedman, H.,

Douglas, S. D. & Nathanson, N. (1989). Infection of monocyte-derived macrophages with human immunodeficiency virus type 1 (HIV-1). Monocyte-tropic and lymphocyte-tropic strains of HIV-1 show distinctive patterns of replication in a panel of cell types. The Journal of experimental medicine 170, 1149-1163.

Connor, E. M., Sperling, R. S., Gelber, R., Kiselev, P., Scott, G., O'Sullivan, M. J., VanDyke,

R., Bey, M., Shearer, W. & Jacobson, R. L. (1994). Reduction of maternal-infant transmission of human immunodeficiency virus type 1 with zidovudine treatment. Pediatric AIDS Clinical

Trials Group Protocol 076 Study Group. N Engl J Med 331, 1173-1180.

Corey, L., McElrath, M. J., Weinhold, K., Matthews, T., Stablein, D., Graham, B., Keefer,

M., Schwartz, D. & Gorse, G. (1998). Cytotoxic T cell and neutralizing antibody responses to human immunodeficiency virus type 1 envelope with a combination vaccine regimen. AIDS

Vaccine Evaluation Group. The Journal of infectious diseases 177, 301-309.

Crowther, R. A., Kiselev, N. A., Bottcher, B., Berriman, J. A., Borisova, G. P., Ose, V. &

Pumpens, P. (1994). Three-dimensional structure of hepatitis B virus core particles determined by electron cryomicroscopy. Cell 77, 943-950.

147

Curran, J. W. (1985). The epidemiology and prevention of the acquired immunodeficiency syndrome. Ann Intern Med 103, 657-662.

Dalgleish, A. G., Beverley, P. C., Clapham, P. R., Crawford, D. H., Greaves, M. F. & Weiss,

R. A. (1984). The CD4 (T4) antigen is an essential component of the receptor for the AIDS retrovirus. Nature 312, 763-767.

Davis, M. R. & Aiken, C. (2004). Virus replication: Uncoating, RT and Integration. In 11th

Conference on Retroviruses and Opportunistic Infections. San Francisco, CA.

De Clercq, E. (1988). Perspectives for the chemotherapy of AIDS. Chemioterapia 7, 357-364.

De Iaco, A., Santoni, F., Vannier, A., Guipponi, M., Antonarakis, S. & Luban, J. (2013).

TNPO3 protects HIV-1 replication from CPSF6-mediated capsid stabilization in the host cell cytoplasm. Retrovirology 10, 20. de Marco, A., Muller, B., Glass, B., Riches, J. D., Krausslich, H. G. & Briggs, J. A. (2010).

Structural analysis of HIV-1 maturation using cryo-electron tomography. PLoS pathogens 6, e1001215. del Alamo, M. & Mateu, M. G. (2005). Electrostatic repulsion, compensatory mutations, and long-range non-additive effects at the dimerization interface of the HIV capsid protein.

Journal of molecular biology 345, 893-906. del Alamo, M., Neira, J. L. & Mateu, M. G. (2003). Thermodynamic dissection of a low affinity protein-protein interface involved in human immunodeficiency virus assembly. The

Journal of biological chemistry 278, 27923-27929.

Deng, H., Liu, R., Ellmeier, W., Choe, S., Unutmaz, D., Burkhart, M., Di Marzio, P.,

Marmon, S., Sutton, R. E., Hill, C. M., Davis, C. B., Peiper, S. C., Schall, T. J., Littman, D. R.

& Landau, N. R. (1996). Identification of a major co-receptor for primary isolates of HIV-1.

Nature 381, 661-666.

Devroe, E., Silver, P. A. & Engelman, A. (2005). HIV-1 incorporates and proteolytically processes human NDR1 and NDR2 serine-threonine kinases. Virology 331, 181-189.

148

Di Nunzio, F., Danckaert, A., Fricke, T., Perez, P., Fernandez, J., Perret, E., Roux, P., Shorte,

S., Charneau, P., Diaz-Griffero, F. & Arhel, N. J. (2012). Human nucleoporins promote HIV-

1 docking at the nuclear pore, nuclear import and integration. PloS one 7, e46037.

Diaz-Griffero, F., Qin, X. R., Hayashi, F., Kigawa, T., Finzi, A., Sarnak, Z., Lienlaf, M.,

Yokoyama, S. & Sodroski, J. (2009). A B-box 2 surface patch important for TRIM5alpha self-association, capsid binding avidity, and retrovirus restriction. Journal of virology 83,

10737-10751.

Dismuke, D. J. & Aiken, C. (2006). Evidence for a functional link between uncoating of the human immunodeficiency virus type 1 core and nuclear import of the viral preintegration complex. Journal of virology 80, 3712-3720.

Donaldson, R. W. & Gerner, E. W. (1987). Phosphorylation of a high molecular weight DNA polymerase alpha. Proceedings of the National Academy of Sciences of the United States of

America 84, 759-763.

Doranz, B. J., Rucker, J., Yi, Y., Smyth, R. J., Samson, M., Peiper, S. C., Parmentier, M.,

Collman, R. G. & Doms, R. W. (1996). A dual-tropic primary HIV-1 isolate that uses fusin and the beta-chemokine receptors CKR-5, CKR-3, and CKR-2b as fusion cofactors. Cell 85,

1149-1158.

Dorfman, T., Bukovsky, A., Ohagen, A., Hoglund, S. & Gottlinger, H. G. (1994). Functional domains of the capsid protein of human immunodeficiency virus type 1. Journal of virology

68, 8180-8187.

Douglas, C. C., Thomas, D., Lanman, J. & Prevelige, P. E., Jr. (2004). Investigation of N- terminal domain charged residues on the assembly and stability of HIV-1 CA. Biochemistry

43, 10435-10441.

Ebina, H., Aoki, J., Hatta, S., Yoshida, T. & Koyanagi, Y. (2004). Role of Nup98 in nuclear entry of human immunodeficiency virus type 1 cDNA. Microbes and infection / Institut

Pasteur 6, 715-724.

149

Emerman, M. & Malim, M. H. (1998). HIV-1 regulatory/accessory genes: keys to unraveling viral and host cell biology. Science (New York, NY 280, 1880-1884.

Endo-Munoz, L., Warby, T., Harrich, D. & McMillan, N. A. (2005). Phosphorylation of HIV

Tat by PKR increases interaction with TAR RNA and enhances transcription. Virol J 2, 17.

Endrich, M. M., Gehrig, P. & Gehring, H. (1999). Maturation-induced conformational changes of HIV-1 capsid protein and identification of two high affinity sites for cyclophilins in the C-terminal domain. The Journal of biological chemistry 274, 5326-5332.

Ensoli, B. & Cafaro, A. (2002). HIV-1 Tat vaccines. Virus research 82, 91-101.

Essex, M., McLane, M. F., Lee, T. H., Falk, L., Howe, C. W., Mullins, J. I., Cabradilla, C. &

Francis, D. P. (1983). Antibodies to cell membrane antigens associated with human T-cell leukemia virus in patients with AIDS. Science (New York, NY 220, 859-862.

Farnet, C. M. & Haseltine, W. A. (1991). Determination of viral proteins present in the human immunodeficiency virus type 1 preintegration complex. Journal of virology 65, 1910-1915.

Fassati, A. (2006). HIV infection of non-dividing cells: a divisive problem. Retrovirology 3,

74.

Fassati, A. (2012). Multiple roles of the capsid protein in the early steps of HIV-1 infection.

Virus research 170, 15-24.

Fassati, A. & Goff, S. P. (2001). Characterization of intracellular reverse transcription complexes of human immunodeficiency virus type 1. Journal of virology 75, 3626-3635.

Fauci, A. S. (2003). HIV and AIDS: 20 years of science. Nature Medicine 9, 839-843.

FDA (2013). Antiretroviral drugs used in the treatment of HIV infection: US Food and Drug

Administration.

Feng, Y., Broder, C. C., Kennedy, P. E. & Berger, E. A. (1996). HIV-1 entry cofactor: functional cDNA cloning of a seven-transmembrane, G protein-coupled receptor. Science

(New York, NY 272, 872-877.

Finkel, T. H., Tudor-Williams, G., Banda, N. K., Cotton, M. F., Curiel, T., Monks, C., Baba,

T. W., Ruprecht, R. M. & Kupfer, A. (1995). Apoptosis occurs predominantly in bystander

150

cells and not in productively infected cells of HIV- and SIV-infected lymph nodes. Nat Med

1, 129-134.

Fitzon, T., Leschonsky, B., Bieler, K., Paulus, C., Schroder, J., Wolf, H. & Wagner, R.

(2000). Proline residues in the HIV-1 NH2-terminal capsid domain: structure determinants for proper core assembly and subsequent steps of early replication. Virology 268, 294-307.

Forshey, B. M. & Aiken, C. (2003). Disassembly of human immunodeficiency virus type 1 cores in vitro reveals association of Nef with the subviral ribonucleoprotein complex. Journal of virology 77, 4409-4414.

Forshey, B. M., Shi, J. & Aiken, C. (2005). Structural requirements for recognition of the human immunodeficiency virus type 1 core during host restriction in owl monkey cells.

Journal of virology 79, 869-875.

Forshey, B. M., von Schwedler, U., Sundquist, W. I. & Aiken, C. (2002). Formation of a human immunodeficiency virus type 1 core of optimal stability is crucial for viral replication.

Journal of virology 76, 5667-5677.

Fouts, D. E., True, H. L., Cengel, K. A. & Celander, D. W. (1997). Site-specific phosphorylation of the human immunodeficiency virus type-1 Rev protein accelerates formation of an efficient RNA-binding conformation. Biochemistry 36, 13256-13262.

Franke, E. K. & Luban, J. (1995). Cyclophilin and gag in HIV-1 replication and pathogenesis.

Adv Exp Med Biol 374, 217-228.

Fuchigami, T., Misumi, S., Takamune, N., Takahashi, I., Takama, M. & Shoji, S. (2002).

Acid-labile formylation of amino terminal proline of human immunodeficiency virus type 1 p24(gag) was found by proteomics using two-dimensional gel electrophoresis and matrix- assisted laser desorption/ionization-time-of-flight mass spectrometry. Biochem Biophys Res

Commun 293, 1107-1113.

Fuller, S. D., Wilk, T., Gowen, B. E., Krausslich, H. G. & Vogt, V. M. (1997). Cryo-electron microscopy reveals ordered domains in the immature HIV-1 particle. Curr Biol 7, 729-738.

151

Galigniana, M. D., Morishima, Y., Gallay, P. A. & Pratt, W. B. (2004). Cyclophilin-A is bound through its peptidylprolyl isomerase domain to the cytoplasmic dynein motor protein complex. The Journal of biological chemistry 279, 55754-55759.

Gallay, P., Swingler, S., Aiken, C. & Trono, D. (1995a). HIV-1 infection of nondividing cells:

C-terminal tyrosine phosphorylation of the viral matrix protein is a key regulator. Cell 80,

379-388.

Gallay, P., Swingler, S., Song, J., Bushman, F. & Trono, D. (1995b). HIV nuclear import is governed by the phosphotyrosine-mediated binding of matrix to the core domain of integrase.

Cell 83, 569-576.

Gallo, R. C., Robert-Guroff, M., Wong-Staal, F., Reitz, M. S., Jr., Arya, S. K. & Streicher, H.

Z. (1987). HTLV-III/LAV and the origin and pathogenesis of AIDS. Int Arch Allergy Appl

Immunol 82, 471-475.

Gallo, R. C., Salahuddin, S. Z., Popovic, M., Shearer, G. M., Kaplan, M., Haynes, B. F.,

Palker, T. J., Redfield, R., Oleske, J. & Safai, B. (1984). Frequent detection and isolation of cytopathic retroviruses (HTLV-III) from patients with AIDS and at risk for AIDS. Science

(New York, NY 224, 500-503.

Gallo, R. C., Sarin, P. S., Gelmann, E. P., Robert-Guroff, M., Richardson, E., Kalyanaraman,

V. S., Mann, D., Sidhu, G. D., Stahl, R. E., Zolla-Pazner, S., Leibowitch, J. & Popovic, M.

(1983). Isolation of human T-cell leukemia virus in acquired immune deficiency syndrome

(AIDS). Science (New York, NY 220, 865-867.

Gallo, R. C. & Wong-Staal, F. (1985). A human T-lymphotropic retrovirus (HTLV-III) as the cause of the acquired immunodeficiency syndrome. Ann Intern Med 103, 679-689.

Gamble, T. R., Vajdos, F. F., Yoo, S., Worthylake, D. K., Houseweart, M., Sundquist, W. I.

& Hill, C. P. (1996). Crystal structure of human cyclophilin A bound to the amino-terminal domain of HIV-1 capsid. Cell 87, 1285-1294.

152

Gamble, T. R., Yoo, S., Vajdos, F. F., von Schwedler, U. K., Worthylake, D. K., Wang, H.,

McCutcheon, J. P., Sundquist, W. I. & Hill, C. P. (1997). Structure of the carboxyl-terminal dimerization domain of the HIV-1 capsid protein. Science (New York, NY 278, 849-853.

Ganser-Pornillos, B. K., Chandrasekaran, V., Pornillos, O., Sodroski, J. G., Sundquist, W. I.

& Yeager, M. (2011). Hexagonal assembly of a restricting TRIM5alpha protein. Proceedings of the National Academy of Sciences of the United States of America 108, 534-539.

Ganser-Pornillos, B. K., Cheng, A. & Yeager, M. (2007). Structure of full-length HIV-1 CA: a model for the mature capsid lattice. Cell 131, 70-79.

Ganser, B. K., Li, S., Klishko, V. Y., Finch, J. T. & Sundquist, W. I. (1999). Assembly and analysis of conical models for the HIV-1 core. Science (New York, NY 283, 80-83.

Gao, F., Bailes, E., Robertson, D. L., Chen, Y., Rodenburg, C. M., Michael, S. F., Cummins,

L. B., Arthur, L. O., Peeters, M., Shaw, G. M., Sharp, P. M. & Hahn, B. H. (1999). Origin of

HIV-1 in the chimpanzee Pan troglodytes troglodytes. Nature 397, 436-441.

Gelderblom, H. R. (1991). Assembly and morphology of HIV: potential effect of structure on viral function. AIDS (London, England) 5, 617-637.

Gelmann, E. P., Popovic, M., Blayney, D., Masur, H., Sidhu, G., Stahl, R. E. & Gallo, R. C.

(1983). Proviral DNA of a retrovirus, human T-cell leukemia virus, in two patients with

AIDS. Science (New York, NY 220, 862-865.

Giroud, C., Chazal, N. & Briant, L. (2011). Cellular kinases incorporated into HIV-1 particles: passive or active passengers? Retrovirology 8, 71.

Gitti, R. K., Lee, B. M., Walker, J., Summers, M. F., Yoo, S. & Sundquist, W. I. (1996).

Structure of the amino-terminal core domain of the HIV-1 capsid protein. Science (New York,

NY 273, 231-235.

Gorlich, D. & Mattaj, I. W. (1996). Nucleocytoplasmic transport. Science (New York, NY 271,

1513-1518.

Gorry, P. R., Ong, C., Thorpe, J., Bannwarth, S., Thompson, K. A., Gatignol, A., Vesselingh,

S. L. & Purcell, D. F. (2003). Astrocyte infection by HIV-1: mechanisms of restricted virus

153

replication, and role in the pathogenesis of HIV-1-associated dementia. Curr HIV Res 1, 463-

473.

Gottlieb, M. S., Schroff, R., Schanker, H. M., Weisman, J. D., Fan, P. T., Wolf, R. A. &

Saxon, A. (1981). Pneumocystis carinii pneumonia and mucosal candidiasis in previously healthy homosexual men: evidence of a new acquired cellular immunodeficiency. N Engl J

Med 305, 1425-1431.

Grattinger, M., Hohenberg, H., Thomas, D., Wilk, T., Muller, B. & Krausslich, H. G. (1999).

In vitro assembly properties of wild-type and cyclophilin-binding defective human immunodeficiency virus capsid proteins in the presence and absence of cyclophilin A.

Virology 257, 247-260.

Greber, U. F., Suomalainen, M., Stidwill, R. P., Boucke, K., Ebersold, M. W. & Helenius, A.

(1997). The role of the nuclear pore complex in adenovirus DNA entry. The EMBO journal

16, 5998-6007.

Grewe, C., Beck, A. & Gelderblom, H. R. (1990). HIV: early virus-cell interactions. J Acquir

Immune Defic Syndr 3, 965-974.

Gross, I., Hohenberg, H., Wilk, T., Wiegers, K., Grattinger, M., Muller, B., Fuller, S. &

Krausslich, H. G. (2000). A conformational switch controlling HIV-1 morphogenesis. The

EMBO journal 19, 103-113.

Gurer, C., Cimarelli, A. & Luban, J. (2002). Specific incorporation of heat shock protein 70 family members into primate lentiviral virions. Journal of virology 76, 4666-4670.

Gurer, C., Hoglund, A., Hoglund, S. & Luban, J. (2005). ATPgammaS disrupts human immunodeficiency virus type 1 virion core integrity. Journal of virology 79, 5557-5567.

Hansen, J., Schulze, T. & Moelling, K. (1987). RNase H activity associated with bacterially expressed reverse transcriptase of human T-cell lymphotropic virus III/lymphadenopathy- associated virus. The Journal of biological chemistry 262, 12393-12396.

154

Harada, S., Haneda, E., Maekawa, T., Morikawa, Y., Funayama, S., Nagata, N. & Ohtsuki, K.

(1999). Casein kinase II (CK-II)-mediated stimulation of HIV-1 reverse transcriptase activity and characterization of selective inhibitors in vitro. Biol Pharm Bull 22, 1122-1126.

Harada, S., Maekawa, T., Haneda, E., Morikawa, Y., Nagata, N. & Ohtsuki, K. (1998).

Biochemical characterization of recombinant HIV-1 reverse transcriptase (rRT) as a glycyrrhizin-binding protein and the CK-II-mediated stimulation of rRT activity potently inhibited by glycyrrhetinic acid derivative. Biol Pharm Bull 21, 1282-1285.

Hauber, J., Bouvier, M., Malim, M. H. & Cullen, B. R. (1988). Phosphorylation of the gene product of human immunodeficiency virus type 1. Journal of virology 62, 4801-4804.

Heath, S. L., Tew, J. G., Szakal, A. K. & Burton, G. F. (1995). Follicular dendritic cells and human immunodeficiency virus infectivity. Nature 377, 740-744.

Hirt, B. (1967). Selective extraction of polyoma DNA from infected mouse cell cultures.

Journal of molecular biology 26, 365-369.

Hoglund, S., Ofverstedt, L. G., Nilsson, A., Lundquist, P., Gelderblom, H., Ozel, M. &

Skoglund, U. (1992). Spatial visualization of the maturing HIV-1 core and its linkage to the envelope. AIDS Res Hum Retroviruses 8, 1-7.

Holman, A. G. & Coffin, J. M. (2005). Symmetrical base preferences surrounding HIV-1, avian sarcoma/leukosis virus, and integration sites. Proceedings of the

National Academy of Sciences of the United States of America 102, 6103-6107.

Hooker, C. W. & Harrich, D. (2003). The first strand transfer reaction of HIV-1 reverse transcription is more efficient in infected cells than in cell-free natural endogenous reverse transcription reactions. J Clin Virol 26, 229-238.

Huang, W., Eshleman, S. H., Toma, J., Fransen, S., Stawiski, E., Paxinos, E. E., Whitcomb, J.

M., Young, A. M., Donnell, D., Mmiro, F., Musoke, P., Guay, L. A., Jackson, J. B., Parkin,

N. T. & Petropoulos, C. J. (2007). Coreceptor tropism in human immunodeficiency virus type

1 subtype D: high prevalence of CXCR4 tropism and heterogeneous composition of viral populations. Journal of virology 81, 7885-7893.

155

Hui, E. K. (2002). Virion-associated protein kinases. Cell Mol Life Sci 59, 920-931.

Hulme, A. E., Perez, O. & Hope, T. J. (2011). Complementary assays reveal a relationship between HIV-1 uncoating and reverse transcription. Proceedings of the National Academy of

Sciences of the United States of America 108, 9975-9980.

Idriss, H., Kawa, S., Damuni, Z., Thompson, E. B. & Wilson, S. H. (1999). HIV-1 reverse transcriptase is phosphorylated in vitro and in a cellular system. Int J Biochem Cell Biol 31,

1443-1452.

Iordanskiy, S., Berro, R., Altieri, M., Kashanchi, F. & Bukrinsky, M. (2006). Intracytoplasmic maturation of the human immunodeficiency virus type 1 reverse transcription complexes determines their capacity to integrate into chromatin. Retrovirology 3, 4.

Irlbeck, D. M., Amrine-Madsen, H., Kitrinos, K. M., Labranche, C. C. & Demarest, J. F.

(2008). Chemokine (C-C motif) receptor 5-using envelopes predominate in dual/mixed-tropic

HIV from the plasma of drug-naive individuals. AIDS (London, England) 22, 1425-1431.

Ivanov, K. I., Puustinen, P., Gabrenaite, R., Vihinen, H., Ronnstrand, L., Valmu, L.,

Kalkkinen, N. & Makinen, K. (2003). Phosphorylation of the potyvirus capsid protein by protein kinase CK2 and its relevance for virus infection. Plant Cell 15, 2124-2139.

Ivanov, K. I., Puustinen, P., Merits, A., Saarma, M. & Makinen, K. (2001). Phosphorylation down-regulates the RNA binding function of the coat protein of potato virus A. The Journal of biological chemistry 276, 13530-13540.

Jacque, J. M., Mann, A., Enslen, H., Sharova, N., Brichacek, B., Davis, R. J. & Stevenson, M.

(1998). Modulation of HIV-1 infectivity by MAPK, a virion-associated kinase. The EMBO journal 17, 2607-2618.

Jaffe, H. W., Bregman, D. J. & Selik, R. M. (1983). Acquired immune deficiency syndrome in the United States: the first 1,000 cases. The Journal of infectious diseases 148, 339-345.

Jiang, J., Ablan, S. D., Derebail, S., Hercik, K., Soheilian, F., Thomas, J. A., Tang, S.,

Hewlett, I., Nagashima, K., Gorelick, R. J., Freed, E. O. & Levin, J. G. (2011). The

156

interdomain linker region of HIV-1 capsid protein is a critical determinant of proper core assembly and stability. Virology 421, 253-265.

Jiang, M., Mak, J., Ladha, A., Cohen, E., Klein, M., Rovinski, B. & Kleiman, L. (1993).

Identification of tRNAs incorporated into wild-type and mutant human immunodeficiency virus type 1. Journal of virology 67, 3246-3253.

Jun, S., Ke, D., Debiec, K., Zhao, G., Meng, X., Ambrose, Z., Gibson, G. A., Watkins, S. C.

& Zhang, P. Direct visualization of HIV-1 with correlative live-cell microscopy and cryo- electron tomography. Structure 19, 1573-1581.

Kann, M. & Gerlich, W. H. (1994). Effect of core protein phosphorylation by protein kinase

C on encapsidation of RNA within core particles of hepatitis B virus. Journal of virology 68,

7993-8000.

Kann, M., Sodeik, B., Vlachou, A., Gerlich, W. H. & Helenius, A. (1999). Phosphorylation- dependent binding of hepatitis B virus core particles to the nuclear pore complex. The Journal of cell biology 145, 45-55.

Karageorgos, L., Li, P. & Burrell, C. (1993). Characterization of HIV replication complexes early after cell-to-cell infection. AIDS Res Hum Retroviruses 9, 817-823.

Karageorgos, L., Li, P. & Burrell, C. J. (1995). Stepwise analysis of reverse transcription in a cell-to-cell human immunodeficiency virus infection model: kinetics and implications. J Gen

Virol 76 ( Pt 7), 1675-1686.

Kataoka, N., Bachorik, J. L. & Dreyfuss, G. (1999). Transportin-SR, a nuclear import receptor for SR proteins. The Journal of cell biology 145, 1145-1152.

Katz, R. A., Greger, J. G., Boimel, P. & Skalka, A. M. (2003). Human immunodeficiency virus type 1 DNA nuclear import and integration are mitosis independent in cycling cells.

Journal of virology 77, 13412-13417.

Katzman, M. & Katz, R. A. (1999). Substrate recognition by retroviral . Adv Virus

Res 52, 371-395.

157

Kaushik, R. & Ratner, L. (2004). Role of human immunodeficiency virus type 1 matrix phosphorylation in an early postentry step of virus replication. Journal of virology 78, 2319-

2326.

Keele, B. F., Giorgi, E. E., Salazar-Gonzalez, J. F., Decker, J. M., Pham, K. T., Salazar, M.

G., Sun, C., Grayson, T., Wang, S., Li, H., Wei, X., Jiang, C., Kirchherr, J. L., Gao, F.,

Anderson, J. A., Ping, L. H., Swanstrom, R., Tomaras, G. D., Blattner, W. A., Goepfert, P. A.,

Kilby, J. M., Saag, M. S., Delwart, E. L., Busch, M. P., Cohen, M. S., Montefiori, D. C.,

Haynes, B. F., Gaschen, B., Athreya, G. S., Lee, H. Y., Wood, N., Seoighe, C., Perelson, A.

S., Bhattacharya, T., Korber, B. T., Hahn, B. H. & Shaw, G. M. (2008). Identification and characterization of transmitted and early founder virus envelopes in primary HIV-1 infection.

Proceedings of the National Academy of Sciences of the United States of America 105, 7552-

7557.

Kelly, B. N., Howard, B. R., Wang, H., Robinson, H., Sundquist, W. I. & Hill, C. P. (2006).

Implications for viral capsid assembly from crystal structures of HIV-1 Gag(1-278) and

CA(N)(133-278). Biochemistry 45, 11257-11266.

Kenney, J. M., von Bonsdorff, C. H., Nassal, M. & Fuller, S. D. (1995). Evolutionary conservation in the hepatitis B virus core structure: comparison of human and duck cores.

Structure 3, 1009-1019.

Kewalramani, V. N. & Emerman, M. (1996). association with mature core structures of

HIV-2. Virology 218, 159-168.

Klatzmann, D., Champagne, E., Chamaret, S., Gruest, J., Guetard, D., Hercend, T.,

Gluckman, J. C. & Montagnier, L. (1984). T-lymphocyte T4 molecule behaves as the receptor for human retrovirus LAV. Nature 312, 767-768.

Koch, P., Lampe, M., Godinez, W. J., Muller, B., Rohr, K., Krausslich, H. G. & Lehmann, M.

J. (2009). Visualizing fusion of pseudotyped HIV-1 particles in real time by live cell microscopy. Retrovirology 6, 84.

158

Kock, J., Kann, M., Putz, G., Blum, H. E. & Von Weizsacker, F. (2003). Central role of a serine phosphorylation site within duck hepatitis B virus core protein for capsid trafficking and genome release. The Journal of biological chemistry 278, 28123-28129.

Koh, Y., Wu, X., Ferris, A. L., Matreyek, K. A., Smith, S. J., Lee, K., KewalRamani, V. N.,

Hughes, S. H. & Engelman, A. (2013). Differential effects of human immunodeficiency virus type 1 capsid and cellular factors nucleoporin 153 and LEDGF/p75 on the efficiency and specificity of viral DNA integration. Journal of virology 87, 648-658.

Kondo, E. & Gottlinger, H. G. (1996). A conserved LXXLF sequence is the major determinant in p6gag required for the incorporation of human immunodeficiency virus type 1

Vpr. Journal of virology 70, 159-164.

Konig, R., Zhou, Y., Elleder, D., Diamond, T. L., Bonamy, G. M., Irelan, J. T., Chiang, C. Y.,

Tu, B. P., De Jesus, P. D., Lilley, C. E., Seidel, S., Opaluch, A. M., Caldwell, J. S., Weitzman,

M. D., Kuhen, K. L., Bandyopadhyay, S., Ideker, T., Orth, A. P., Miraglia, L. J., Bushman, F.

D., Young, J. A. & Chanda, S. K. (2008). Global analysis of host-pathogen interactions that regulate early-stage HIV-1 replication. Cell 135, 49-60.

Korin, Y. D. & Zack, J. A. (1998). Progression to the G1b phase of the cell cycle is required for completion of human immunodeficiency virus type 1 reverse transcription in T cells.

Journal of virology 72, 3161-3168.

Kotov, A., Zhou, J., Flicker, P. & Aiken, C. (1999). Association of Nef with the human immunodeficiency virus type 1 core. Journal of virology 73, 8824-8830.

Kovaleski, B. J., Kennedy, R., Khorchid, A., Kleiman, L., Matsuo, H. & Musier-Forsyth, K.

(2007). Critical role of helix 4 of HIV-1 capsid C-terminal domain in interactions with human lysyl-tRNA synthetase. The Journal of biological chemistry 282, 32274-32279.

Krishnan, L., Matreyek, K. A., Oztop, I., Lee, K., Tipper, C. H., Li, X., Dar, M. J.,

Kewalramani, V. N. & Engelman, A. (2010). The requirement for cellular transportin 3

(TNPO3 or TRN-SR2) during infection maps to human immunodeficiency virus type 1 capsid and not integrase. Journal of virology 84, 397-406.

159

Kutluay, S. B., Perez-Caballero, D. & Bieniasz, P. D. (2013). Fates of retroviral core components during unrestricted and TRIM5-restricted infection. PLoS pathogens 9, e1003214.

Lanman, J., Lam, T. T., Emmett, M. R., Marshall, A. G., Sakalian, M. & Prevelige, P. E., Jr.

(2004). Key interactions in HIV-1 maturation identified by hydrogen-deuterium exchange.

Nat Struct Mol Biol 11, 676-677.

Laurent, A. G., Krust, B., Rey, M. A., Montagnier, L. & Hovanessian, A. G. (1989). Cell surface expression of several species of human immunodeficiency virus type 1 major core protein. Journal of virology 63, 4074-4078.

Leavitt, A. D., Robles, G., Alesandro, N. & Varmus, H. E. (1996). Human immunodeficiency virus type 1 integrase mutants retain in vitro integrase activity yet fail to integrate viral DNA efficiently during infection. Journal of virology 70, 721-728.

Lee, K., Ambrose, Z., Martin, T. D., Oztop, I., Mulky, A., Julias, J. G., Vandegraaff, N.,

Baumann, J. G., Wang, R., Yuen, W., Takemura, T., Shelton, K., Taniuchi, I., Li, Y.,

Sodroski, J., Littman, D. R., Coffin, J. M., Hughes, S. H., Unutmaz, D., Engelman, A. &

KewalRamani, V. N. (2010). Flexible use of nuclear import pathways by HIV-1. Cell host & microbe 7, 221-233.

Leonard, R., Zagury, D., Desportes, I., Bernard, J., Zagury, J. F. & Gallo, R. C. (1988).

Cytopathic effect of human immunodeficiency virus in T4 cells is linked to the last stage of virus infection. Proceedings of the National Academy of Sciences of the United States of

America 85, 3570-3574.

Levy, J. A., Hoffman, A. D., Kramer, S. M., Landis, J. A., Shimabukuro, J. M. & Oshiro, L.

S. (1984). Isolation of lymphocytopathic retroviruses from San Francisco patients with AIDS.

In Science (New York, NY, pp. 840-842.

Li, F., Goila-Gaur, R., Salzwedel, K., Kilgore, N. R., Reddick, M., Matallana, C., Castillo, A.,

Zoumplis, D., Martin, D. E., Orenstein, J. M., Allaway, G. P., Freed, E. O. & Wild, C. T.

(2003). PA-457: a potent HIV inhibitor that disrupts core condensation by targeting a late step

160

in Gag processing. Proceedings of the National Academy of Sciences of the United States of

America 100, 13555-13560.

Li, M., Lyon, M. K. & Garcea, R. L. (1995). In vitro phosphorylation of the polyomavirus major capsid protein VP1 on serine 66 by casein kinase II. The Journal of biological chemistry 270, 26006-26011.

Li, P., Kuiper, L. J., Stephenson, A. J. & Burrell, C. J. (1992). De novo reverse transcription is a crucial event in cell-to-cell transmission of human immunodeficiency virus. J Gen Virol

73 ( Pt 4), 955-959.

Li, P., Stephenson, A. J., Kuiper, L. J. & Burrell, C. J. (1993). Double-stranded strong-stop

DNA and the second template switch in human immunodeficiency virus (HIV) DNA synthesis. Virology 194, 82-88.

Li, S., Hill, C. P., Sundquist, W. I. & Finch, J. T. (2000). Image reconstructions of helical assemblies of the HIV-1 CA protein. Nature 407, 409-413.

Li, X. & Sodroski, J. (2008). The TRIM5alpha B-box 2 domain promotes cooperative binding to the retroviral capsid by mediating higher-order self-association. Journal of virology 82,

11495-11502.

Li, Y., Kar, A. K. & Sodroski, J. (2009). Target cell type-dependent modulation of human immunodeficiency virus type 1 capsid disassembly by cyclophilin A. Journal of virology 83,

10951-10962.

Lidon-Moya, M. C., Barrera, F. N., Bueno, M., Perez-Jimenez, R., Sancho, J., Mateu, M. G.

& Neira, J. L. (2005). An extensive thermodynamic characterization of the dimerization domain of the HIV-1 capsid protein. Protein Sci 14, 2387-2404.

Lifson, J. D., Reyes, G. R., McGrath, M. S., Stein, B. S. & Engleman, E. G. (1986). AIDS retrovirus induced cytopathology: giant cell formation and involvement of CD4 antigen.

Science (New York, NY 232, 1123-1127.

161

Lin, D. H., Zimmermann, S., Stuwe, T., Stuwe, E. & Hoelz, A. (2013). Structural and functional analysis of the C-terminal domain of Nup358/RanBP2. Journal of molecular biology 425, 1318-1329.

Liu, H., Wu, X., Newman, M., Shaw, G. M., Hahn, B. H. & Kappes, J. C. (1995). The Vif protein of human and simian immunodeficiency viruses is packaged into virions and associates with viral core structures. Journal of virology 69, 7630-7638.

Lori, F., di Marzo Veronese, F., de Vico, A. L., Lusso, P., Reitz, M. S., Jr. & Gallo, R. C.

(1992). Viral DNA carried by human immunodeficiency virus type 1 virions. Journal of virology 66, 5067-5074.

Luban, J. (2007). Cyclophilin A, TRIM5, and resistance to human immunodeficiency virus type 1 infection. Journal of virology 81, 1054-1061.

Maddon, P. J., Dalgleish, A. G., McDougal, J. S., Clapham, P. R., Weiss, R. A. & Axel, R.

(1986). The T4 gene encodes the AIDS virus receptor and is expressed in the immune system and the brain. Cell 47, 333-348.

Maillard, P. V., Zoete, V., Michielin, O. & Trono, D. (2011). Homology-based identification of capsid determinants that protect HIV1 from human TRIM5alpha restriction. The Journal of biological chemistry 286, 8128-8140.

Malim, M. H. & Cullen, B. R. (1993). Rev and the fate of pre-mRNA in the nucleus: implications for the regulation of RNA processing in eukaryotes. Mol Cell Biol 13, 6180-

6189.

Malim, M. H., McCarn, D. F., Tiley, L. S. & Cullen, B. R. (1991). Mutational definition of the human immunodeficiency virus type 1 Rev activation domain. Journal of virology 65,

4248-4254.

Mammano, F., Ohagen, A., Hoglund, S. & Gottlinger, H. G. (1994). Role of the major homology region of human immunodeficiency virus type 1 in virion morphogenesis. Journal of virology 68, 4927-4936.

162

Marcon, L. & Sodroski, J. (1994). gp120-independent fusion mediated by the human immunodeficiency virus type 1 gp41 envelope glycoprotein: a reassessment. Journal of virology 68, 1977-1982.

Marechal, V., Clavel, F., Heard, J. M. & Schwartz, O. (1998). Cytosolic Gag p24 as an index of productive entry of human immunodeficiency virus type 1. Journal of virology 72, 2208-

2212.

Markham, P. D., Sarngadharan, M. G., Salahuddin, S. Z., Popovic, M. & Gallo, R. C. (1984).

Correlation between exposure to human T-cell leukemia-lymphoma virus-III and the development of AIDS. Ann N Y Acad Sci 437, 106-109.

Markosyan, R. M., Cohen, F. S. & Melikyan, G. B. (2003). HIV-1 envelope proteins complete their folding into six-helix bundles immediately after fusion pore formation. Mol

Biol Cell 14, 926-938.

Marshall, H. M., Ronen, K., Berry, C., Llano, M., Sutherland, H., Saenz, D., Bickmore, W.,

Poeschla, E. & Bushman, F. D. (2007). Role of PSIP1/LEDGF/p75 in lentiviral infectivity and integration targeting. PloS one 2, e1340.

Masuda, T., Planelles, V., Krogstad, P. & Chen, I. S. (1995). Genetic analysis of human immunodeficiency virus type 1 integrase and the U3 att site: unusual phenotype of mutants in the zinc finger-like domain. Journal of virology 69, 6687-6696.

Matreyek, K. A. & Engelman, A. (2011). The requirement for nucleoporin NUP153 during human immunodeficiency virus type 1 infection is determined by the viral capsid. Journal of virology 85, 7818-7827.

McDonald, D., Vodicka, M. A., Lucero, G., Svitkina, T. M., Borisy, G. G., Emerman, M. &

Hope, T. J. (2002). Visualization of the intracellular behavior of HIV in living cells. The

Journal of cell biology 159, 441-452.

McDougal, J. S., Mawle, A., Cort, S. P., Nicholson, J. K., Cross, G. D., Scheppler-Campbell,

J. A., Hicks, D. & Sligh, J. (1985). Cellular tropism of the human retrovirus HTLV-III/LAV.

I. Role of T cell activation and expression of the T4 antigen. J Immunol 135, 3151-3162.

163

McMichael, A. J. & Hanke, T. (2003). HIV vaccines 1983-2003. Nature Medicine 9, 874-880.

McMillan, N. A., Chun, R. F., Siderovski, D. P., Galabru, J., Toone, W. M., Samuel, C. E.,

Mak, T. W., Hovanessian, A. G., Jeang, K. T. & Williams, B. R. (1995). HIV-1 Tat directly interacts with the interferon-induced, double-stranded RNA-dependent kinase, PKR. Virology

213, 413-424.

Melby, T., Despirito, M., Demasi, R., Heilek-Snyder, G., Greenberg, M. L. & Graham, N.

(2006). HIV-1 coreceptor use in triple-class treatment-experienced patients: baseline prevalence, correlates, and relationship to enfuvirtide response. The Journal of infectious diseases 194, 238-246.

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

K., Eckner, K., Hattox, S. & Adams, J. (1990). Inhibition of HIV-1 replication by a nonnucleoside reverse transcriptase inhibitor. Science (New York, NY 250, 1411-1413.

Mervis, R. J., Ahmad, N., Lillehoj, E. P., Raum, M. G., Salazar, F. H., Chan, H. W. &

Venkatesan, S. (1988). The gag gene products of human immunodeficiency virus type 1: alignment within the gag open reading frame, identification of posttranslational modifications, and evidence for alternative gag precursors. Journal of virology 62, 3993-4002.

Miedema, F., Meyaard, L., Koot, M., Klein, M. R., Roos, M. T., Groenink, M., Fouchier, R.

A., Van't Wout, A. B., Tersmette, M., Schellekens, P. T. & et al. (1994). Changing virus-host interactions in the course of HIV-1 infection. Immunol Rev 140, 35-72.

Miller, M. D., Farnet, C. M. & Bushman, F. D. (1997). Human immunodeficiency virus type

1 preintegration complexes: studies of organization and composition. Journal of virology 71,

5382-5390.

Misumi, S., Inoue, M., Dochi, T., Kishimoto, N., Hasegawa, N., Takamune, N. & Shoji, S.

(2010). Uncoating of human immunodeficiency virus type 1 requires prolyl isomerase Pin1.

The Journal of biological chemistry 285, 25185-25195.

164

Momany, C., Kovari, L. C., Prongay, A. J., Keller, W., Gitti, R. K., Lee, B. M., Gorbalenya,

A. E., Tong, L., McClure, J., Ehrlich, L. S., Summers, M. F., Carter, C. & Rossmann, M. G.

(1996). Crystal structure of dimeric HIV-1 capsid protein. Nat Struct Biol 3, 763-770.

Morgan, C., Rose, H. M. & Mednis, B. (1968). Electron microscopy of herpes simplex virus.

I. Entry. Journal of virology 2, 507-516.

Morgan, C., Rosenkranz, H. S. & Mednis, B. (1969). Structure and development of viruses as observed in the electron microscope. V. Entry and uncoating of adenovirus. Journal of virology 4, 777-796.

Muller, B., Patschinsky, T. & Krausslich, H. G. (2002). The late-domain-containing protein p6 is the predominant phosphoprotein of human immunodeficiency virus type 1 particles.

Journal of virology 76, 1015-1024.

Muller, B., Tessmer, U., Schubert, U. & Krausslich, H. G. (2000). Human immunodeficiency virus type 1 Vpr protein is incorporated into the virion in significantly smaller amounts than gag and is phosphorylated in infected cells. Journal of virology 74, 9727-9731.

Murakami, T., Nakajima, T., Koyanagi, Y., Tachibana, K., Fujii, N., Tamamura, H., Yoshida,

N., Waki, M., Matsumoto, A., Yoshie, O., Kishimoto, T., Yamamoto, N. & Nagasawa, T.

(1997). A small molecule CXCR4 inhibitor that blocks T cell line-tropic HIV-1 infection. The

Journal of experimental medicine 186, 1389-1393.

Nakashima, H., Matsui, T., Harada, S., Kobayashi, N., Matsuda, A., Ueda, T. & Yamamoto,

N. (1986). Inhibition of replication and cytopathic effect of human T cell lymphotropic virus type III/lymphadenopathy-associated virus by 3'-azido-3'-deoxythymidine in vitro.

Antimicrob Agents Chemother 30, 933-937.

Nermut, M. V. & Fassati, A. (2003). Structural analyses of purified human immunodeficiency virus type 1 intracellular reverse transcription complexes. Journal of virology 77, 8196-8206.

Nilsson, A., Lunqvist, P., Love, A., Torfason, E., Gelderblom, H. R. & Hoglund, S. (1992).

Spatial visualization of progressive states of maturing lentivirus. Vet Microbiol 33, 333-340.

165

Nisole, S., Lynch, C., Stoye, J. P. & Yap, M. W. (2004). A Trim5-cyclophilin A found in owl monkey kidney cells can restrict HIV-1. Proceedings of the National

Academy of Sciences of the United States of America 101, 13324-13328.

O'Doherty, U., Swiggard, W. J. & Malim, M. H. (2000). Human immunodeficiency virus type

1 spinoculation enhances infection through virus binding. Journal of virology 74, 10074-

10080.

Ohagen, A. & Gabuzda, D. (2000). Role of Vif in stability of the human immunodeficiency virus type 1 core. Journal of virology 74, 11055-11066.

Ohtsuki, K., Maekawa, T., Harada, S., Karino, A., Morikawa, Y. & Ito, M. (1998).

Biochemical characterization of HIV-1 Rev as a potent activator of casein kinase II in vitro.

FEBS Lett 428, 235-240.

Ojala, P. M., Sodeik, B., Ebersold, M. W., Kutay, U. & Helenius, A. (2000). Herpes simplex virus type 1 entry into host cells: reconstitution of capsid binding and uncoating at the nuclear pore complex in vitro. Mol Cell Biol 20, 4922-4931.

Oleske, J., Minnefor, A., Cooper, R., Jr., Thomas, K., dela Cruz, A., Ahdieh, H., Guerrero, I.,

Joshi, V. V. & Desposito, F. (1983). Immune deficiency syndrome in children. Jama 249,

2345-2349.

Ott, D. E., Coren, L. V., Johnson, D. G., Kane, B. P., Sowder, R. C., 2nd, Kim, Y. D., Fisher,

R. J., Zhou, X. Z., Lu, K. P. & Henderson, L. E. (2000). Actin-binding cellular proteins inside human immunodeficiency virus type 1. Virology 266, 42-51.

Pante, N. & Kann, M. (2002). Nuclear pore complex is able to transport macromolecules with diameters of about 39 nm. Mol Biol Cell 13, 425-434.

Patterson, S. & Knight, S. C. (1987). Susceptibility of human peripheral blood dendritic cells to infection by human immunodeficiency virus. J Gen Virol 68 ( Pt 4), 1177-1181.

Penn, M. L., Grivel, J. C., Schramm, B., Goldsmith, M. A. & Margolis, L. (1999). CXCR4 utilization is sufficient to trigger CD4+ T cell depletion in HIV-1-infected human lymphoid

166

tissue. Proceedings of the National Academy of Sciences of the United States of America 96,

663-668.

Pereira, C. F., Rossy, J., Owen, D. M., Mak, J. & Gaus, K. (2012). HIV taken by STORM: super-resolution fluorescence microscopy of a viral infection. Virology journal 9, 84.

Pettit, S. C., Moody, M. D., Wehbie, R. S., Kaplan, A. H., Nantermet, P. V., Klein, C. A. &

Swanstrom, R. (1994). The p2 domain of human immunodeficiency virus type 1 Gag regulates sequential proteolytic processing and is required to produce fully infectious virions.

Journal of virology 68, 8017-8027.

Pettit, S. C., Sheng, N., Tritch, R., Erickson-Viitanen, S. & Swanstrom, R. (1998). The regulation of sequential processing of HIV-1 Gag by the viral protease. Adv Exp Med Biol

436, 15-25.

Pietroboni, G. R., Harnett, G. B. & Bucens, M. R. (1989). Centrifugal enhancement of human immunodeficiency virus (HIV) and human herpesvirus type 6 (HHV-6) infection in vitro. J

Virol Methods 24, 85-90.

Poch, O., Sauvaget, I., Delarue, M. & Tordo, N. (1989). Identification of four conserved motifs among the RNA-dependent polymerase encoding elements. The EMBO journal 8,

3867-3874.

Poignard, P., Sabbe, R., Picchio, G. R., Wang, M., Gulizia, R. J., Katinger, H., Parren, P. W.,

Mosier, D. E. & Burton, D. R. (1999). Neutralizing antibodies have limited effects on the control of established HIV-1 infection in vivo. Immunity 10, 431-438.

Pomerantz, R. J. & Horn, D. L. (2003). Twenty years of therapy for HIV-1 infection. Nat Med

9, 867-873.

Popovic, M., Sarngadharan, M. G., Read, E. & Gallo, R. C. (1984). Detection, isolation, and continuous production of cytopathic retroviruses (HTLV-III) from patients with AIDS and pre-AIDS. Science (New York, NY 224, 497-500.

Pornillos, O., Ganser-Pornillos, B. K. & Yeager, M. (2011). Atomic-level modelling of the

HIV capsid. Nature 469, 424-427.

167

Rabe, B., Vlachou, A., Pante, N., Helenius, A. & Kann, M. (2003). Nuclear import of hepatitis B virus capsids and release of the viral genome. Proceedings of the National

Academy of Sciences of the United States of America 100, 9849-9854.

Ramachandra, M., Nakano, R., Mohan, P. M., Rawitch, A. B. & Padmanabhan, R. (1993).

Adenovirus DNA polymerase is a phosphoprotein. The Journal of biological chemistry 268,

442-448.

Ratka, M., Lackmann, M., Ueckermann, C., Karlins, U. & Koch, G. (1989). Poliovirus- associated protein kinase: destabilization of the virus capsid and stimulation of the phosphorylation reaction by Zn2+. Journal of virology 63, 3954-3960.

Ratner, L., Fisher, A., Jagodzinski, L. L., Mitsuya, H., Liou, R. S., Gallo, R. C. & Wong-

Staal, F. (1987). Complete nucleotide sequences of functional clones of the AIDS virus. AIDS

Res Hum Retroviruses 3, 57-69.

Richman, D. D. (1988). The treatment of HIV infection. Azidothymidine (AZT) and other new antiviral drugs. Infect Dis Clin North Am 2, 397-407.

Robert-Guroff, M., Blayney, D. W., Safai, B., Lange, M., Gelmann, E. P., Gutterman, J. W.,

Mansell, P. W., Goedert, J. L., Groopman, J. E., Steigbigel, N. H. & et al. (1984). HTLV-I- specific antibody in AIDS patients and others at risk. Lancet 2, 128-131.

Roberts, M. M. & Oroszlan, S. (1989). The preparation and biochemical characterization of intact capsids of equine infectious anemia virus. Biochem Biophys Res Commun 160, 486-

494.

Safai, B., Sarngadharan, M. G., Groopman, J. E., Arnett, K., Popovic, M., Sliski, A.,

Schupbach, J. & Gallo, R. C. (1984). Seroepidemiological studies of human T-lymphotropic retrovirus type III in acquired immunodeficiency syndrome. Lancet 1, 1438-1440.

Sambrook, J., Fritsch, E. & Maniatis, T. (1989). Molecular Cloning : A Laboratory Manual:

Cold Spring Harbour Laboratory Press.

168

Sarngadharan, M. G., Popovic, M., Bruch, L., Schupbach, J. & Gallo, R. C. (1984).

Antibodies reactive with human T-lymphotropic retroviruses (HTLV-III) in the serum of patients with AIDS. Science (New York, NY 224, 506-508.

Sattentau, Q. J. (1992). CD4 activation of HIV fusion. Int J Cell Cloning 10, 323-332.

Sattentau, Q. J. & Moore, J. P. (1993). The role of CD4 in HIV binding and entry. Philos

Trans R Soc Lond B Biol Sci 342, 59-66.

Sawyer, S. L., Wu, L. I., Emerman, M. & Malik, H. S. (2005). Positive selection of primate

TRIM5alpha identifies a critical species-specific retroviral restriction domain. Proceedings of the National Academy of Sciences of the United States of America 102, 2832-2837.

Sayah, D. M. & Luban, J. (2004). Selection for loss of Ref1 activity in human cells releases human immunodeficiency virus type 1 from cyclophilin A dependence during infection.

Journal of virology 78, 12066-12070.

Sayah, D. M., Sokolskaja, E., Berthoux, L. & Luban, J. (2004). Cyclophilin A retrotransposition into TRIM5 explains owl monkey resistance to HIV-1. Nature 430, 569-

573.

Schacker, T., Collier, A. C., Hughes, J., Shea, T. & Corey, L. (1996). Clinical and epidemiologic features of primary HIV infection. Ann Intern Med 125, 257-264.

Schaller, T., Ocwieja, K. E., Rasaiyaah, J., Price, A. J., Brady, T. L., Roth, S. L., Hue, S.,

Fletcher, A. J., Lee, K., KewalRamani, V. N., Noursadeghi, M., Jenner, R. G., James, L. C.,

Bushman, F. D. & Towers, G. J. (2011). HIV-1 capsid-cyclophilin interactions determine nuclear import pathway, integration targeting and replication efficiency. PLoS pathogens 7, e1002439.

Scharli, C. E. & Koch, G. (1984). Protein kinase activity in purified poliovirus particles and empty viral capsid preparations. J Gen Virol 65 ( Pt 1), 129-139.

Schneider, E., Whitmore, S., Glynn, K. M., Dominguez, K., Mitsch, A. & McKenna, M. T.

(2008). Revised surveillance case definitions for HIV infection among adults, adolescents,

169

and children aged <18 months and for HIV infection and AIDS among children aged 18 months to <13 years--United States, 2008. MMWR Recomm Rep 57, 1-12.

Scholz, I., Arvidson, B., Huseby, D. & Barklis, E. (2005). Virus particle core defects caused by mutations in the human immunodeficiency virus capsid N-terminal domain. Journal of virology 79, 1470-1479.

Schubert, U., Schneider, T., Henklein, P., Hoffmann, K., Berthold, E., Hauser, H., Pauli, G. &

Porstmann, T. (1992). Human-immunodeficiency-virus-type-1-encoded is phosphorylated by casein kinase II. Eur J Biochem 204, 875-883.

Schweitzer, C. J., Jagadish, T., Haverland, N., Ciborowski, P. & Belshan, M. (2013).

Proteomic analysis of early HIV-1 nucleoprotein complexes. Journal of proteome research

12, 559-572.

Sebastian, S. & Luban, J. (2005). TRIM5alpha selectively binds a restriction-sensitive retroviral capsid. Retrovirology 2, 40.

Shah, V. B., Shi, J., Hout, D. R., Oztop, I., Krishnan, L., Ahn, J., Shotwell, M. S., Engelman,

A. & Aiken, C. (2013). The host proteins transportin SR2/TNPO3 and cyclophilin A exert opposing effects on HIV-1 uncoating. Journal of virology 87, 422-432.

Shehu-Xhilaga, M., Kraeusslich, H. G., Pettit, S., Swanstrom, R., Lee, J. Y., Marshall, J. A.,

Crowe, S. M. & Mak, J. (2001). Proteolytic processing of the p2/nucleocapsid cleavage site is critical for human immunodeficiency virus type 1 RNA dimer maturation. Journal of virology

75, 9156-9164.

Shi, J. & Aiken, C. (2006). Saturation of TRIM5 alpha-mediated restriction of HIV-1 infection depends on the stability of the incoming viral capsid. Virology 350, 493-500.

Shun, M. C., Raghavendra, N. K., Vandegraaff, N., Daigle, J. E., Hughes, S., Kellam, P.,

Cherepanov, P. & Engelman, A. (2007). LEDGF/p75 functions downstream from preintegration complex formation to effect gene-specific HIV-1 integration. Genes & development 21, 1767-1778.

170

Siegal, F. P., Lopez, C., Hammer, G. S., Brown, A. E., Kornfeld, S. J., Gold, J., Hassett, J.,

Hirschman, S. Z., Cunningham-Rundles, C., Adelsberg, B. R. & et al. (1981). Severe acquired immunodeficiency in male homosexuals, manifested by chronic perianal ulcerative herpes simplex lesions. N Engl J Med 305, 1439-1444.

Stremlau, M., Owens, C. M., Perron, M. J., Kiessling, M., Autissier, P. & Sodroski, J. (2004).

The cytoplasmic body component TRIM5alpha restricts HIV-1 infection in Old World monkeys. Nature 427, 848-853.

Stremlau, M., Perron, M., Lee, M., Li, Y., Song, B., Javanbakht, H., Diaz-Griffero, F.,

Anderson, D. J., Sundquist, W. I. & Sodroski, J. (2006). Specific recognition and accelerated uncoating of retroviral capsids by the TRIM5alpha restriction factor. Proceedings of the

National Academy of Sciences of the United States of America 103, 5514-5519.

Stremlau, M., Perron, M., Welikala, S. & Sodroski, J. (2005). Species-specific variation in the

B30.2(SPRY) domain of TRIM5alpha determines the potency of human immunodeficiency virus restriction. Journal of virology 79, 3139-3145.

Stromberg, K. (1972). Surface-active agents for isolation of the core component of avian myeloblastosis virus. Journal of virology 9, 684-697.

Stromberg, K., Hurley, N. E., Davis, N. L., Rueckert, R. R. & Fleissner, E. (1974). Structural studies of avian myeloblastosis virus: comparison of polypeptides in virion and core component by dodecyl sulfate-polyacrylamide gel electrophoresis. Journal of virology 13,

513-528.

Swingler, S., Gallay, P., Camaur, D., Song, J., Abo, A. & Trono, D. (1997). The Nef protein of human immunodeficiency virus type 1 enhances serine phosphorylation of the viral matrix.

Journal of virology 71, 4372-4377.

Sylwester, A. W., Grivel, J. C., Fitzgerald, W., Rossio, J. L., Lifson, J. D. & Margolis, L. B.

(1998). CD4(+) T-lymphocyte depletion in human lymphoid tissue ex vivo is not induced by noninfectious human immunodeficiency virus type 1 virions. Journal of virology 72, 9345-

9347.

171

Tang, S., Murakami, T., Agresta, B. E., Campbell, S., Freed, E. O. & Levin, J. G. (2001).

Human immunodeficiency virus type 1 N-terminal capsid mutants that exhibit aberrant core morphology and are blocked in initiation of reverse transcription in infected cells. Journal of virology 75, 9357-9366.

Tang, S., Murakami, T., Cheng, N., Steven, A. C., Freed, E. O. & Levin, J. G. (2003). Human immunodeficiency virus type 1 N-terminal capsid mutants containing cores with abnormally high levels of capsid protein and virtually no reverse transcriptase. Journal of virology 77,

12592-12602.

Terry, L. J., Shows, E. B. & Wente, S. R. (2007). Crossing the nuclear envelope: hierarchical regulation of nucleocytoplasmic transport. Science (New York, NY 318, 1412-1416.

Towers, G. J., Hatziioannou, T., Cowan, S., Goff, S. P., Luban, J. & Bieniasz, P. D. (2003).

Cyclophilin A modulates the sensitivity of HIV-1 to host restriction factors. Nat Med 9, 1138-

1143.

Trainin, Z., Wernicke, D., Ungar-Waron, H. & Essex, M. (1983). Suppression of the humoral antibody response in natural retrovirus infections. Science (New York, NY 220, 858-859.

Trono, D. (1992). Partial reverse transcripts in virions from human immunodeficiency and murine leukemia viruses. Journal of virology 66, 4893-4900.

Valle-Casuso, J. C., Di Nunzio, F., Yang, Y., Reszka, N., Lienlaf, M., Arhel, N., Perez, P.,

Brass, A. L. & Diaz-Griffero, F. (2012). TNPO3 is required for HIV-1 replication after nuclear import but prior to integration and binds the HIV-1 core. Journal of virology 86,

5931-5936.

Veronese, F. D., Copeland, T. D., Oroszlan, S., Gallo, R. C. & Sarngadharan, M. G. (1988).

Biochemical and immunological analysis of human immunodeficiency virus gag gene products p17 and p24. Journal of virology 62, 795-801. von Schwedler, U. K., Stemmler, T. L., Klishko, V. Y., Li, S., Albertine, K. H., Davis, D. R.

& Sundquist, W. I. (1998). Proteolytic refolding of the HIV-1 capsid protein amino-terminus facilitates viral core assembly. The EMBO journal 17, 1555-1568.

172

von Schwedler, U. K., Stray, K. M., Garrus, J. E. & Sundquist, W. I. (2003). Functional surfaces of the human immunodeficiency virus type 1 capsid protein. Journal of virology 77,

5439-5450.

Wacharapornin, P., Lauhakirti, D. & Auewarakul, P. (2007). The effect of capsid mutations on HIV-1 uncoating. Virology 358, 48-54.

Wain-Hobson, S., Sonigo, P., Danos, O., Cole, S. & Alizon, M. (1985). Nucleotide sequence of the AIDS virus, LAV. Cell 40, 9-17.

Warrilow, D., Meredith, L., Davis, A., Burrell, C., Li, P. & Harrich, D. (2008). Cell factors stimulate human immunodeficiency virus type 1 reverse transcription in vitro. Journal of virology 82, 1425-1437.

Warrilow, D., Stenzel, D. & Harrich, D. (2007). Isolated HIV-1 core is active for reverse transcription. Retrovirology 4, 77.

Wei, X., Ghosh, S. K., Taylor, M. E., Johnson, V. A., Emini, E. A., Deutsch, P., Lifson, J. D.,

Bonhoeffer, S., Nowak, M. A. & Hahn, B. H. (1995). Viral dynamics in human immunodeficiency virus type 1 infection. Nature 373, 117-122.

Welker, R., Hohenberg, H., Tessmer, U., Huckhagel, C. & Krausslich, H. G. (2000).

Biochemical and structural analysis of isolated mature cores of human immunodeficiency virus type 1. Journal of virology 74, 1168-1177.

WHO (2012a). A short guide on methods: measuring the impact of national PMTCT programmes: towards the elimination of new HIV infections among children by 2015 and keeping their mothers alive. : World Health

Organization.

WHO (2012b). Use of antiretroviral drugs for treating pregnant women and preventing HIV infection in infants

Programmatic update

World Health Organization.

WHO/UNAIDS (2008). Towards universal access

173

Scaling up priority HIV/AIDS interventions in the health sector

Progress report June 2008.

Wiegers, K., Rutter, G., Kottler, H., Tessmer, U., Hohenberg, H. & Krausslich, H. G. (1998).

Sequential steps in human immunodeficiency virus particle maturation revealed by alterations of individual Gag polyprotein cleavage sites. Journal of virology 72, 2846-2854.

Wiegers, K., Rutter, G., Schubert, U., Grattinger, M. & Krausslich, H. G. (1999). Cyclophilin

A incorporation is not required for human immunodeficiency virus type 1 particle maturation and does not destabilize the mature capsid. Virology 257, 261-274.

Wilk, T., Gross, I., Gowen, B. E., Rutten, T., de Haas, F., Welker, R., Krausslich, H. G.,

Boulanger, P. & Fuller, S. D. (2001). Organization of immature human immunodeficiency virus type 1. Journal of virology 75, 759-771.

Wlodawer, A., Miller, M., Jaskolski, M., Sathyanarayana, B. K., Baldwin, E., Weber, I. T.,

Selk, L. M., Clawson, L., Schneider, J. & Kent, S. B. (1989). Conserved folding in retroviral proteases: crystal structure of a synthetic HIV-1 protease. Science (New York, NY 245, 616-

621.

Wong-Staal, F., Shaw, G. M., Hahn, B. H., Salahuddin, S. Z., Popovic, M., Markham, P.,

Redfield, R. & Gallo, R. C. (1985). Genomic diversity of human T-lymphotropic virus type

III (HTLV-III). Science (New York, NY 229, 759-762.

Woodward, C. L., Prakobwanakit, S., Mosessian, S. & Chow, S. A. (2009). Integrase interacts with nucleoporin NUP153 to mediate the nuclear import of human immunodeficiency virus type 1. Journal of virology 83, 6522-6533.

Worthylake, D. K., Wang, H., Yoo, S., Sundquist, W. I. & Hill, C. P. (1999). Structures of the

HIV-1 capsid protein dimerization domain at 2.6 A resolution. Acta Crystallogr D Biol

Crystallogr 55, 85-92.

Wright, E. R., Schooler, J. B., Ding, H. J., Kieffer, C., Fillmore, C., Sundquist, W. I. &

Jensen, G. J. (2007). Electron cryotomography of immature HIV-1 virions reveals the structure of the CA and SP1 Gag shells. The EMBO journal 26, 2218-2226.

174

Wyatt, R. & Sodroski, J. (1998). The HIV-1 envelope glycoproteins: fusogens, antigens, and immunogens. Science (New York, NY 280, 1884-1888.

Xu, L., Yang, L., Moitra, P. K., Hashimoto, K., Rallabhandi, P., Kaul, S., Meroni, G., Jensen,

J. P., Weissman, A. M. & D'Arpa, P. (2003). BTBD1 and BTBD2 colocalize to cytoplasmic bodies with the RBCC/tripartite motif protein, TRIM5delta. Experimental cell research 288,

84-93.

Yamashita, M. & Emerman, M. (2004). Capsid is a dominant determinant of retrovirus infectivity in nondividing cells. Journal of virology 78, 5670-5678.

Yamashita, M., Perez, O., Hope, T. J. & Emerman, M. (2007). Evidence for direct involvement of the capsid protein in HIV infection of nondividing cells. PLoS pathogens 3,

1502-1510.

Yang, X. & Gabuzda, D. (1998). Mitogen-activated protein kinase phosphorylates and regulates the HIV-1 Vif protein. The Journal of biological chemistry 273, 29879-29887.

Yang, X. & Gabuzda, D. (1999). Regulation of human immunodeficiency virus type 1 infectivity by the ERK mitogen-activated protein kinase signaling pathway. Journal of virology 73, 3460-3466.

Yang, X., Goncalves, J. & Gabuzda, D. (1996). Phosphorylation of Vif and its role in HIV-1 replication. The Journal of biological chemistry 271, 10121-10129.

Yang, Y., Fricke, T. & Diaz-Griffero, F. (2013). Inhibition of reverse transcriptase activity increases stability of the HIV-1 core. Journal of virology 87, 683-687.

Yap, M. W., Nisole, S. & Stoye, J. P. (2005). A single amino acid change in the SPRY domain of human Trim5alpha leads to HIV-1 restriction. Curr Biol 15, 73-78.

Yeung, M. L., Houzet, L., Yedavalli, V. S. & Jeang, K. T. (2009). A genome-wide short hairpin RNA screening of jurkat T-cells for human proteins contributing to productive HIV-1 replication. The Journal of biological chemistry 284, 19463-19473.

Yi, Y., Isaacs, S. N., Williams, D. A., Frank, I., Schols, D., De Clercq, E., Kolson, D. L. &

Collman, R. G. (1999). Role of CXCR4 in cell-cell fusion and infection of monocyte-derived

175

macrophages by primary human immunodeficiency virus type 1 (HIV-1) strains: two distinct mechanisms of HIV-1 dual tropism. Journal of virology 73, 7117-7125.

Yoo, S., Myszka, D. G., Yeh, C., McMurray, M., Hill, C. P. & Sundquist, W. I. (1997).

Molecular recognition in the HIV-1 capsid/cyclophilin A complex. Journal of molecular biology 269, 780-795.

Yu, M. & Summers, J. (1994). Multiple functions of capsid protein phosphorylation in duck hepatitis B virus replication. Journal of virology 68, 4341-4348.

Yu, X., Matsuda, Z., Yu, Q. C., Lee, T. H. & Essex, M. (1993). Vpx of simian immunodeficiency virus is localized primarily outside the virus core in mature virions.

Journal of virology 67, 4386-4390.

Zack, J. A., Arrigo, S. J., Weitsman, S. R., Go, A. S., Haislip, A. & Chen, I. S. (1990). HIV-1 entry into quiescent primary lymphocytes: molecular analysis reveals a labile, latent viral structure. Cell 61, 213-222.

Zhang, H., Bagasra, O., Niikura, M., Poiesz, B. J. & Pomerantz, R. J. (1994). Intravirion reverse transcripts in the peripheral blood plasma on human immunodeficiency virus type 1- infected individuals. Journal of virology 68, 7591-7597.

Zhang, H., Dornadula, G., Orenstein, J. & Pomerantz, R. J. (2000). Morphologic changes in human immunodeficiency virus type 1 virions secondary to intravirion reverse transcription: evidence indicating that reverse transcription may not take place within the intact viral core. J

Hum Virol 3, 165-172.

Zhang, R., Mehla, R. & Chauhan, A. (2010). Perturbation of host nuclear membrane component RanBP2 impairs the nuclear import of human immunodeficiency virus -1 preintegration complex (DNA). PloS one 5, e15620.

Zhirnov, O. P. & Grigoriev, V. B. (1994). Disassembly of influenza C viruses, distinct from that of influenza A and B viruses requires neutral-alkaline pH. Virology 200, 284-291.

Zhou, J., Yuan, X., Dismuke, D., Forshey, B. M., Lundquist, C., Lee, K. H., Aiken, C. &

Chen, C. H. (2004). Small-molecule inhibition of human immunodeficiency virus type 1

176

replication by specific targeting of the final step of virion maturation. Journal of virology 78,

922-929.

Zhou, L., Sokolskaja, E., Jolly, C., James, W., Cowley, S. A. & Fassati, A. (2011).

Transportin 3 promotes a nuclear maturation step required for efficient HIV-1 integration.

PLoS pathogens 7, e1002194.

Zhou, W., Parent, L. J., Wills, J. W. & Resh, M. D. (1994). Identification of a membrane- binding domain within the amino-terminal region of human immunodeficiency virus type 1

Gag protein which interacts with acidic phospholipids. Journal of virology 68, 2556-2569.

Zhou, Y. & Ratner, L. (2000). Phosphorylation of human immunodeficiency virus type 1 Vpr regulates cell cycle arrest. Journal of virology 74, 6520-6527.

Zhu, P., Liu, J., Bess, J., Jr., Chertova, E., Lifson, J. D., Grise, H., Ofek, G. A., Taylor, K. A.

& Roux, K. H. (2006). Distribution and three-dimensional structure of AIDS virus envelope spikes. Nature 441, 847-852.

177