towards the development of synergistic inhibitors that...
TRANSCRIPT
Towards the Development of Synergistic Inhibitors
that Exploit the Replication Strategy of HIV-1
Leonard Keith Pattenden, B. App. Sci. (Hons)
Submitted to
Queensland University of Technology
In Fulfilment of the Requirements for the Degree of
Doctor of Philosophy
May 2003
Abstract HIV-1 has evolved with a great deal of functional complexity contained within a
very small genome by encoding small, but critical viral proteins within larger viral
genes and dividing the replication cycle into early and late phases to differentially
produce all proteins leading to efficient replication and virion release. Early
replication is restricted by the host spliceosome that processes HIV-1 vRNA
transcripts so only the small intragenomic proteins are produced, one of which is
Rev (Regulator of Virion Expression). Rev in turn governs the transition from early
to late replication by interacting with a highly structured region of vRNA termed the
Rev Response Element (RRE). The binding of Rev to the RRE is believed to cause a
change in the vRNA tertiary structure and inhibition of splicing of the vRNA. Once,
a Rev:RRE complex is formed, a nuclear export signal within Rev facilitates the
export of partially spliced and unspliced vRNA to the cytoplasm. During late
replication the partially spliced and unspliced vRNA is translated to polyproteins
and is packaged into a budding virion where the viral aspartyl protease (HIV-1 PR)
autocatalytically excises itself from the larger polyprotein and processes the
remaining polyproteins to release all viral structural and functional proteins to form
a mature and infectious virion.
Since the vRNA salvaged by Rev is translated to the polyproteins containing
HIV-1 PR, the inhibition of Rev function will reduce the amount of HIV-1 PR
available and thereby reduce the amount of HIV-1 PR therapeutics required to elicit a
clinical effect. Therfore a combination approach to HIV-1 treatment using suitably
developed therapeutics that inhibit Rev and HIV-1 PR function represents an attractive
synergistic approach to treating HIV-1 infection in vivo. The work of this thesis was
divided into two parts, the first part was concerned with HIV-1 PR structural biology
and addressing problems encountered with inhibitor design.
A bicyclic peptide (based on inhibitors of analogous structure) was co-
crystallised with active HIV-1 PR to develop an enzyme-product (E-P) complex and
with a catalytically inactive mutant HIV-1 PR to provide an analogy to the enzyme-
substrate (E-S) complex. Both structures of the E-P and E-S complexes were solved to
1.6Å resolution and were compared to a hydroxyethylamine isostere enzyme-inhibitor
complex (E-I), highlighting the similarity of binding mode for all ligands. The inhibitor
in the E-I complex was translated towards the S1 – S3 pockets of the substrate binding
cleft relative to the substrate in the E-S complex due to the increased length of the
II
hydroxylethylamine isostere compared to the peptide backbone, although the inhibitor
“puckered” the isostere linkage and maintains a binding mode similar to the substrate
with very little overall differences in the position of the ligands and surrounding
protein. The similarity of the E-S, E-I and E-P complexes was attributed to the
macrocyclic ligands ordering the surrounding protein environment, especially the
protein β-strand “flap” structures that form a roof over the ligands in the active site but
were not found to close more tightly in any of the trapped catalytic states. The new
structures allowed refinement of details of the mechanism of peptide hydrolysis. The
mechanism relies on the optimal nucleophilic attack of a water molecule on the scissile
amide bond with concerted acid-base catalysis of the active site aspartyl residues
intitiated by D125. The alignment and intrinsic position of the N-terminus of the
bicyclic substrate was interpreted as being critical to facilitate efficient electron
transfer with the bicyclic substrate.
An N-terminal cyclic inhibitor, similar to the N-terminal portion of the bicyclic
substrate, was used to address a major problem in HIV-1 PR drug design termed
“cooperativity,” where the sequential optimisation of an inhibitor (or substrate) to
individual pockets of the substrate binding cleft, can negatively impact on adjacent and
downfield subsites and thereby alter the binding mode of the “optimised” inhibitor.
The technique referred to here as “templating” uses the N-terminal cycle to lock the
binding mode into a known conformation, probing the S1’ and S2’ pockets. The
structure activity relationship suggested that by viewing the S1’ – S3’ pockets as a
single trough, bulky aromatic groups attached to an N-alkyl sulfonamide could be
directed along the line of the trough without adverse interactions with the tops of the
S1’ and S3’ pockets, providing very potent inhibitors. It was also found that specificity
and potency of an inhibitor can be maintained with smaller functionalities that carry
their bulk low and close to the inhibitor backbone in the S2’ pocket, making the P2
functionalities more substrate-like.
The second part of the thesis was concerned with establishing suitable surface
plasmon resonance assays for testing potential inhibitors of Rev function. Recombinant
Rev and its minimal RNA aptamer target (stem loop II of the RRE termed RBE3),
were expressed, purified, and used to develop BIAcore-based assays and test potential
inhibitors of their interaction. The system was applied to screening of aminoglycoside
antibiotics and other small molecules in a competitive assay, and also to quantitative
assay of Neomycin and moderate sized analytes: Rev and three peptidic analogues of
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the high-affinity binding site of Rev - the native peptide, succinylated form of the
peptide and a form incorporating a novel helix-inducing cap. The peptide and protein
assay was undertaken to test the proposition that helix induction of the high-affinity
binding site of Rev can increase affinity for the biologically important RNA target and
thereby form the basis of a new class of inhibitors.
The screen of small molecule antagonists found that Neomycin was the best
inhibitor of the Rev:RBE3 interaction and that efficacy of other aminoglycosides was
due to the neamine-base structure presenting charge to bind to the RNA and blocking
interaction with Rev. The quantitative assay was optimised to reduce non-specific
interactions of Rev protein to allow reliable studies of the analytes with RBE3 by the
sytematic testing of buffers and modifiers. It was found that mutliple analytes bound to
the RBE3 aptamer and a comparison of the KD values found that the native and capped
peptides had similar affinity for RBE3 RNA (native slightly greater at 21 ± 7nM cf
capped 41 ± 10nM) that was greater than the Rev protein (101 ± 19nM), however the
succinylated peptide exhibited stronger binding with a KD ≤8nM and Neomycin had
the lowest affinity (KD 13 ± 3µM). The similarity of the native and capped peptides
may be due to the high concentration of salt in the assay buffers and was necessary for
the stability of the Rev protein, but is sufficient to influence secondary structure of the
peptides. Therefore, it could not be stated that the helix-inducing cap increased the
affinity of the native peptide for the biologically important therapeutic target.
The work conducted in this thesis firmly establishes foundations for the
continued development of inhibitors against both Rev and HIV-1 PR that play key
roles in the HIV-1 replication strategy. It is envisaged this work could lead to a novel
synergistic therapeutic approach to treating HIV-1 infection.
IV
Table of Contents Page No.
Abstract…………………………………….……………..…………………………...II Table of Contents……...………………….……………..…………………………....V List of Figures…...……………………………………..…………………………..VIII List of Tables……………………....…………………………………….………......XI Keywords...……………………………………………...…………...………...…....XII Abbreviations…...…………………………………………………...……………..XIII Acknowledgements…...………………………………..……………………………XV Statement of Original Authorship………….……………….………..……..….....XVI Publications and Presentations…………………………………………………..XVII Chapter 1: Human Immunodeficiency Virus: Background. 1
1.1 Overview of the Human Immunodeficiency Virus. 2 1.2 Viral Characteristics. 3 1.3 HIV-1 Virus Particles. 3 1.4 Replication of HIV. 4 1.5 HIV Accessory Proteins. 8 1.6 Therapeutic Problems. 9 1.7 Hypothesis and Aims of Thesis. 11
Chapter 2: Crystal Structures of HIV-1 Protease Complexed with Substrate
and Products. 15
2.1 Introduction 16 2.1.1 HIV-1 PR Structure 16 2.1.2 HIV-1 PR Function 19 2.1.3 HIV-1 PR Mechanism of Catalysis 21 2.1.4 Protease Activation Assessed by Cryo-crystallography 24 2.1.5 Preliminary Crystallography 26 2.1.6 Aims 28
2.2 Results 30 2.2.1 Crystal Structure of Enzyme-Substrate Complex 30 2.2.2 Crystal Structure of Enzyme-Product Complex 36
2.3 Discussion 42 2.3.1 Insight from Enzyme Kinetics 42 2.3.2 Comparison of Substrate, Product and Analogue 45 2.3.3 Mechanism of Catalysis of HIV-1 PR 52
2.4 Experimental 58 2.4.1 Origin of Protein and Substrate 58 2.4.2 Crystallisation Methods 60 2.4.3 Structure Determination 60 2.4.4 Analysis and Alignment 61
Chapter 3: HIV-1 Protease Inhibitors Based on a Macrocyclic Template. 63
3.1 Introduction 64 3.1.1 The Problems of Current HIV-1 PR Inhibitors. 64 3.1.2 Cooperativity and Drug Design 64 3.1.3 Hypothesis, Aims and Significance 67
V
3.2 Results 67 3.2.1 Design Of A Template For A Focussed Inhibitor Library 67 3.2.2 Inhibitor Design – P1’ and P2’ Variation 71 3.2.3 Synthesis of a Focussed “Templated” Combinatorial Library 73
3.3 Discussion 75 3.3.1 Structure-Activity Relationships at P1' 75 3.3.2 Structure-Activity Relationships at P2' 76 3.3.3 Miscellaneous Compounds 76 3.3.4 Templating and Cooperativity 78 3.3.5 Bioavailability, Resistance and Cooperativity
79 3.3.6 Conclusions 82
3.4 Experimental 84 3.4.1 Purification of HIV-1 PR 84 3.4.2 Inhibition of HIV-1 Protease 85 3.4.3 Computer Assisted Inhibitor Design 88 3.4.4 Inhibitor Synthesis 88 3.4.5 Inhibitor Purification and Characterisation 89
Chapter 4: Expression, Purification and Labelling of Recombinant HIV-1 Rev
and RBE3 RNA. 94
4.1 Introduction 95 4.1.1 HIV-1 Rev – Background 95 4.1.2 The Functional Domains and Function of Rev 97 4.1.3 Aims and Hypothesis 100 4.1.4 The Problems of Labelling and Purifying RNA 100
4.2 Results and Discussion 102 4.2.1 Cloning, Expression and Purification of HIV-1 Rev 102 4.2.2 Refolding Experiments of HIV-1 Rev 107 4.2.3 In Vitro Transcription, Labelling and Purification of RBE3 RNA 108
4.3 Experimental 114 4.3.1 Constructs and Transformation into JM109 Cells 114 4.3.2 Restriction Digests 116 4.3.3 Preparation of Electrocompetent E.coli for Protein Expression 117 4.3.4 LB Antibiotic Media 117 4.3.5 SDS-PAGE and Urea-PAGE Protocols 117 4.3.6 Transfer Protocol for Western Blotting and Biotin Detection 118 4.3.7 Calculation of Protein Concentrations 120 4.3.8 RP-HPLC and Mass Spectrometry 120
Chapter 5: Application of Surface Plasmon Resonance Assays to
Rev:RRE Interactions. 121
5.1 Introduction. 122 5.1.1 Assaying RNA 122 5.1.2 Rev:RRE Inhibition 124
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5.1.2.1 Biological Strategies 124 5.1.2.2 Chemical Strategies 125
5.1.3 Helix Induction of a Decoy Inhibitor 129 5.1.4 Aims 130
5.2 Results and Discussion 131 5.2.1 High Throughput Screening of Aminoglycoside
Antibiotics 131 5.2.2 Assay of Helix Mimetics 139 5.2.3 Does Helix Induction Increase Affinity
for the RBE3 Aptamer? 174 Chapter 6: General Discussion and Conclusions 177
References 186
VII
List of Figures Chapter 1. Fig. 1.1: Structure of HIV-1 Genome and Important RNAs Produced 4 Fig. 1.2: The Major Steps of HIV-1 Replication 7 Chapter 2. Fig. 2.1: Crystal Structures of Aspartic Proteases 17 Fig. 2.2: Structural Features of HIV-1 PR 18 Fig. 2.3: the Processing Profile of HIV-1 PR 20 Fig. 2.4: The Nomenclature of Protease Active Sites 21 Fig. 2.5: The Steps of HIV-1 PR Catalysis 22 Fig. 2.6: The Three Proposed Mechanisms of Catalysis by HIV-1 PR (a) – (c) 23 Fig. 2.7: The Substrate used in Crystallography Experiments 27 Fig. 2.8: Structure of Novel Substrate(1), Products(2,3) and Inhibitor(4)
used in this Study 30 Fig. 2.9: Ramachandran Plot of the E-S Complex for 1 bound to
HIV-1 PR (D25N) 32 Fig. 2.10: Superimposition of the E-S complex with Pepstatin Inhibitor Complex 32 Fig. 2.11: Connolly Surface of the Active Site of the E-S Complex 33 Fig. 2.12: Hydrogen Bonding Pattern of the E-S Complex 34 Fig. 2.13: Electron Density Map of the Substrate within HIV-1 PR 34 Fig. 2.14: The Active Site of the E-S Complex for 1 Bound to HIV-1 PR (D25N) 35 Fig. 2.15: H-Bonding Across the Active Site and Dimer Interface for 1
Bound to HIV-1 PR (D25N) 35 Fig. 2.16: Ramachandran Plot of the E-P Complex for (2+3) Bound to HIV-1 PR 38 Fig. 2.17: H-Bonding Between HIV-1 PR and Product Ligands 39 Fig. 2.18: Electron Density Map of Products 2 and 3 within HIV-1 PR 39 Fig. 2.19: The Two Conformations of the C-Product Ligand 40 Fig. 2.20: The Carboxylate triad of the E-P Complex 41 Fig. 2.21: Bicyclic Substrate, Product and Linear Substrate “Inhibitors” 42 Fig. 2.22: Inhibitor Analogue of the Substrate 45 Fig. 2.23: Superimposition of E-S and E-P Complex 46 Fig. 2.24: Superimposition of E-S and E-I Analogue Complexes 47 Fig. 2.25: Superimposition of the E-S, E-P and E-I Analogue Complexes 48 Fig. 2.26: Possible Active Site Charged States 53 Fig. 2.27: Ionisation State of the Active Site 54 Fig. 2.28: Model of the Active Site at the “Catalytic Event” 55 Fig. 2.29: Alternate Mode of Nucleophilic Attack 55 Fig. 2.30: Modified Mechanism of Cleavage of HIV-1 PR 57 Chapter 3. Fig. 3.1: The Mimicry of Peptides by Macrocycles 66 Fig. 3.2: Potent Peptidomimetic Inhibitor 6 of HIV-1 PR 67 Fig. 3.3: Templating Drug Design 68 Fig. 3.4: The Concept of a Templated Combinatorial Library 70 Fig. 3.5: The S1’ and S2’ Pocket Modelled from the E-I complex of 6 71 Fig. 3.6: Templated S1’ Probes of HIV-1 Protease 72 Fig. 3.7: Templated S2’ Probes of HIV-1 Protease 72 Fig. 3.8: General Synthetic Methods 74 Fig. 3.9: Inhibitors Designed and Synthesised by Dr Robert Reid 74
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Fig. 3.10: Possible Optimisation of the N-alkyl Ureas 79 Fig. 3.11: Schematic of Interactions in HIV-1 PR and Re-shaping of the
Substrate-binding Cleft 80 Fig. 3.12: Mass Spectrometry of Purified HIV-1 PR 85 Fig. 3.13: Hanes Plot of [S]/v vs [S] 86 Fig. 3.14: Dixon Plot Compound 10 87 Fig. 3.15: Henderson Plot of Compound 10 87 Chapter 4. Fig. 4.1: Structure of Rev34-50 Interacting with Stem Loop II of the RRE 96 Fig. 4.2: The Sequence of HIV-1 Rev 98 Fig. 4.3: The In Vitro Cycling of Rev 98 Fig. 4.4: Western Blot Comparison of Different Constructs in Different Cells 105 Fig. 4.5: SDS-PAGE and Western Transfer of the Ni-NTA Purification 105 Fig. 4.6: Image Quantitation Analysis 105 Fig. 4.7: HPLC Profile and Mass Spectrometry of Crude Rev 106 Fig. 4.8: The In Vitro Transcription Reaction 109 Fig. 4.9: The Endlabelling of RNA using T4 RNA Ligase 110 Fig. 4.10: Extracted and Precipitated Labelled RNA 111 Fig. 4.11: Extraction of RNA from Urea-PAGE Gels 114 Fig. 4.12: Restriction Digest of Constructs Obtained 115 Chapter 5. Fig. 5.1: Representative Structures of Potent RNA Export Inhibitors 126 Fig. 5.2: Base Aminoglycoside Structure 127 Fig. 5.3: Representative 4,5-disubstituted 2-DOS Compounds with Improved
Rev:RRE Inhibition over Neomycin B 127 Fig. 5.4: Neomycin-acridine Conjugate 128 Fig. 5.5: Other Inhibitors of Rev:RRE Function 128 Fig. 5.6: Peptides Used in the Study 130 Fig. 5.7: Immobilisation of RBE3 RNA onto SA chips 132 Fig. 5.8: Small Molecule Antagonists of the Rev:RRE Interaction 133 Fig. 5.9: Corrected Sensorgrams of Competitive Inhibition
of Antagonists vs Rev 134 Fig. 5.10: SPR Analysis of Small Molecule Inhibition of the
Rev:RRE interaction 135 Fig. 5.11: Assay Data Expressed as Percentage Inhibition Compared to Neomycin 136 Fig. 5.12: A stylisation of the Surface Plasmon Resonance Assay
Showing Possible Non-specific Interactions 140 Fig. 5.13: A Stylisation of how Additives were Screened for
Reducing Non-specific Binding 143 Fig. 5.14: Replicated Assay of Neomycin Interaction with
B1 Neutravidin:RBE3 Surfaces 146 Fig. 5.15: Uncorrected Non-linear Regression Analysis of
Req vs Concentration for Neomycin:RBE3 Binding (Two Site Model) 148 Fig. 5.16: Non-linear Regression of Neomycin
Req vs Concentration log10 (Below Req = 205) 149
IX
Fig. 5.17: Corrected Non-linear Regression Analysis (One Site Binding Model) of Neomycin Interacting with RBE3 RNA (Corrected Data) 151
Fig. 5.18: Analysis of Flash Folded Rev Protein on Streptavidin-B1-RBE3 Chips 155
Fig. 5.19: Plots of Plasmon Response vs Time for Rev and Peptide interactions with RBE3 RNA 156
Fig. 5.20: Non-linear Regression Analysis of Req vs Concentration for Peptide 2:RBE3 Binding (Two Site Model). 157
Fig. 5.21: Examination of the Low Affinity Site for Peptide 2 (Corrected Data) 158 Fig. 5.22: Non-linear Regression Analysis of Req vs Concentration
for Peptide 3:RBE3 Binding (Two Site Model, All Data) 160 Fig. 5.23: Nonlinear Regression Analysis of Req vs Concentration (log10)
for the Succinylated Peptide 3:RBE3 Binding (All Data, Uncorrected) 161 Fig. 5.24: Nonlinear Regression Analysis of Req vs Concentration
for the Succinylated Peptide 3:RBE3 Binding (Selected Data) 162 Fig. 5.25: Non-linear Regression Analysis of Req vs Concentration
for Peptide 4:RBE3 Binding (Two Site Model) 164 Fig. 5.26: Comparison of High Affinity Sites for Peptide 4 165 Fig. 5.27: Comparison of Low Affinity Sites for Peptide 4 166 Fig. 5.28: Uncorrected and Corrected Non-linear Regression Analysis
of Rev:RBE3 Interaction 168 Fig. 5.29: Rev Interacting with RBE3 RNA [Green] and RNAneg RNA [Pink] 169 Chapter 6. Fig. 6.1: The Possible Future Work of N-terminal Cyclic Inhibitors 181 Fig. 6.2: How a Cyclic Peptide can Serve as a Helix Mimetic 183
X
List of Tables Chapter 2. Table 2.1: Native Cleavage Sites for HIV-1 PR 21 Table 2.2: Diffraction & Refinement Statistics Comparing 290K and 160K Data 37 Table 2.3: Comparison of HIV-1 PR Structures and Ligands 49 Table 2.4: Comparison of HIV-1PR H-bond Contacts to the
Ligands (P3, S3-P3', S3'). 49 Chapter 3. Table 3.1: Inhibitor Potencies vs HIV-1 PR 75 Table 3.2: HIV-1 PR Inhibitors and their Known Mutations 81 Chapter 5. Table 5.1: Effects of Buffer Additives on Non-specific Interactions 144 Table 5.2: Concentration and Req Values for Neomycin
Interactions with Custom RBE3-Neutravin-B1 Surfaces 147 Table 5.3: Nonlinear Regression Analysis of Req vs Concentration for
Neomycin:RBE3 Binding 149 Table 5.4: Nonlinear Regression Analysis of Req vs Concentration for the Native
Peptide 2:RBE3 Binding 158 Table 5.5: Nonlinear Regression Analysis of Req vs Concentration
for the Succinylated Peptide 3:RBE3 Binding (All Data) 161 Table 5.6: Nonlinear Regression Analysis of Req vs Concentration
for the Capped Peptide 4:RBE3 Binding (All Data) 164 Table 5.7: Non-linear Regression Analysis of SPR Data 171
XI
Keywords Aspartyl Protease, HIV-1 Protease, Substrate Complex, Product Complex, X-ray
Crystallography, Cyclic Peptide, β-strand Mimetic, Catalysis, Cooperativity, HIV-1
Rev, RRE, RBE3, RNA, BIAcore, Surface Plasmon Resonance, in vitro transcription,
Aminoglycoside, Helix-inducing Cap, Neomycin, Streptavidin, Neutravidin, Helix
Mimetic.
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Abbreviations º Degrees µ micro ºC Temperature in Degrees Celsius αα(s) Amino Acid(s) Å Angstrom A Absorbance Ab Antibody BSA Bovine Serum Albumin CAPS 3-(Cyclohexylamino)-1-propanesulfonic acid DNA Deoxyribonucleic acid DMSO Dimethylsulfoxide DTT Dithiothreitol [E] Enzyme Concentration EDTA Ethylenediaminetetraacetic acid ESI-MS Electrospray Ionisation Mass Spectrometry EtOH Ethanol g grams Gu Guanidinium HBS Hepes Buffered Saline Hepes N-2-Hydroxyethylpiperazine-N'-2-ethanesulfonic acid h hour(s) HRMS High-Resolution Mass Spectrometry [I] Inhibitor Concentration IC50 Inhibition Concentration (50% inhibition) IPTG isopropyl-β-D-thiogalactopyranoside K Degrees Kelvin KA Association Constant at equilibrium KD Dissociation Constant at equilibrium Ki Inhibition Constant Km Michaelis-Menten Constant L litre m milli M Molar mA milliamperes MBHA 4-Methylbenzhydrylamine MeCN Acetonitrile MeOH Methanol MES 2-(N-Morpholino)-ethanesulfonic acid min minute(s) mRNA Messenger Ribonucleic acid MS Mass Spectrometry MW Molecular Weight n nano Ni-NTA nickel-nitrilotriacetic acid NMR Nuclear Magnetic Resonance pdb Protein Data Bank Poly-A Poly-Adenosine Poly-T Poly-Thymidine Req Response at Equilibrium Rmax Response Maxima at Equilibrium RMS Root Mean Square RMSD Root Mean Square Deviation RNA Ribonucleic acid rNTP Ribonucleotide 5’-triphosphate rpm Revolutions per minute RP-HPLC Reverse Phase High Performance Liquid Chromatography rRNA Ribosomal Ribonucleic acid SDS Sodium Dodecyl Sulfate
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[S] Substrate Concentration SA Streptavidin Sp3 (S-p three) hybrid orbitals snRNA Small Nuclear Ribonucleic acid Sy.x Systematic error in x-axis TBE Tris Buffered Ethylenediaminetetraacetic acid TBS Tris Buffered Saline TEMED N,N,N',N'-Tetramethylethylenediamine TFA Trifluoroacetic acid THF tetrahydrofuran Tris Tris(hydroxymethyl)aminomethane TWEEN polyoxyethylene(20)-sorbitanemonolaureate U Units Us Uridines UTP Uridine 5’-Triphosphate v velocity V Volts VdW Van der Waals vi initial velocity + inhibitor vo initial velocity - inhibitor w watts x g times gravity
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Acknowledgements
First and foremost I thank my wife Caroline, who took this journey and all that it
was with me. Her patience and love has sustained me through the more trying times.
For my wife, “verumtamen neque vir sine muliere neque mulier sine viro in
Domino.”
This project has been a collaborative effort between the University of Queensland's
Institute for Molecular Bioscience (IMB) Chemistry Group and Queensland
University of Technology's Centre for Molecular Biotechnology (CMB) within the
School of Life Science. My principal supervisors for this project were Dr Terry
Walsh (QUT) who oversaw the cloning, expression, purification, labelling and
BIAcore assaying comprising the work of chapters 4 and 5, and Prof David Fairlie
(Principal Investigator/Chemistry Group, IMB, UQ) who has overseen drug design,
organic chemistry, mass spectrometry, NMR, enzymology and HPLC comprising
chapters 2 and 3 and aspects of 4.
From the IMB my thanks go to Dr Jenny Martin for teaching me crystallography
and Dr Shu-Hong Hu for mounting the crystals and collecting the X-ray data, Dr
Michael Kelso for his friendship and collaboration that was mutually beneficial, Dr
Robert Reid for the invaluable lessons in organic chemistry and Dr Doug Bergman
for patiently leading me step-by-step through the facets of enzymology and Alun
Jones for all the mass spectrometry help.
From the CMB my thanks go to Dr Marcus Hastie for taking me under his wing and
providing the support I needed in molecular biology and protein chemistry.
Additional thanks are due to Prof Ross Smith (Dept of Biochemistry UQ) for the
provision of Biotinylated control RNA and Dr Jørgen Kjems for providing all the
constructs that facilitated a great deal of the thesis.
To these people I say thankyou for the sacrifice of your time, the provision of
your aid, the friendship you have shared and the knowledge I have gained.
XV
Statement of Original Authorship
The work contained in this thesis has not been previously submitted for a degree or
diploma at any higher education institution. To the best of my knowledge and
beliefs, the thesis contains no material previously published or written by another
person except where due reference is made.
Signed:………………………………………….
Date:…………………………………………….
XVI
Publications and Presentations During the course of the thesis, several presentations were made to professional
bodies including two oral presentations to the Queensland Protein Group (receiving
a prize in the Student Symposium 1999), a poster presentation at the Australian
Society for Biochemistry and Molecular Biology (ComBio99), a poster was
presented in my behalf at an international conference of proteolytic enzymes 2000
(Florida USA) and I was a speaker and poster presenter at the 3rd Australian
BIAsymposium (2000).
There are several publications that arose during the time work was conducted in this
thesis that is not described within this body of work.
Tyndall, J.D., Reid, R.C., Tyssen, D.P., Jardine, D.K., Todd, B., Passmore, M.,
March, D.R., Pattenden, L.K., Bergman, D.A., Alewood, D., Hu, S.H., Alewood,
P.F., Birch, C.J., Martin, J.L. and Fairlie, D.P. (2000) “Synthesis, stability, antiviral
activity, and protease-bound structures of substrate-mimicking constrained
macrocyclic inhibitors of HIV-1 protease.” J Med Chem. 43: 3495 – 3504.
Where I grew the crystal structures described and
Glenn, M.P., Pattenden, L.K., Reid, R.C., Tyssen, D.P., Tyndall, J.D., Birch, C.J.
and Fairlie, D.P. (2002) “Beta-strand mimicking macrocyclic amino acids: templates
for protease inhibitors with antiviral activity.” J Med Chem. 45: 371 – 381.
Where I assayed the compounds described therein in a fluorometric assay.
There are two papers in preparation from this thesis (from Chapter 2 and 5) and one
that has been published comprising the work of Chapter 3:
Reid, R.C., Pattenden, L.K., Tyndall, J.D., Martin, J.L., Walsh, T and Fairlie, D.P.
(2004) “Countering cooperative effects in protease inhibitors using constrained beta-
strand-mimicking templates in focused combinatorial libraries.” J Med Chem. 47:
1641 – 1651.
XVII
1
Chapter 1: Human Immunodeficiency Virus: Background.
1.1 Overview of the Human Immunodeficiency Virus.
1.2 Viral Characteristics.
1.3 HIV-1 Virus Particles.
1.4 Replication of HIV.
1.5 HIV Accessory Proteins.
1.6 Therapeutic Problems.
1.7 Hypothesis and Aims of Thesis.
2
1.1 Overview of the Human Immunodeficiency Virus.
Human Immunodeficiency Virus type 1 (HIV-1, Virus Code. 61.0.6.5.001) is
lethal to man as the aetiological agent for Acquired Immunodeficiency Syndrome
(AIDS) (Coffin et al., 1995). The most recent statistical estimates are that more than
42 million people worldwide are infected with HIV (UNAIDS/WHO, 2002). AIDS is
characterised by a dysfunctional immune system that enables opportunistic infections
to thrive, leading to death (Ho and Kaydan, 1987). The time period from infection to
loss of immune function - essentially the period of “latency” - is approximately ten
years (Eron, 1997), although this depends upon the status of the immune system of
HIV infected people.
A person infected with HIV shows initial flu-like symptoms such as rashes,
fever and some neurological complaints that disappear after a few weeks as the virus
spreads systemically (Mitsuyasu, 1997). The virus makes complex quasi-steady state
interactions with the various host cells it infects (Eron, 1997). Much of the variability
in viral behaviour can be partly ascribed to host factors such as: the type of cell
infected, the presence or absence of other infectious diseases, the influence of
modulating factors (such as NF-κB, TNF-α, chemokines and CD8 activating factor),
as well as the overall health of the immune system under attack (Mitsuyasu, 1997).
During this period the blood often has little detectable virus present, while the
lymphatic system may have ten fold higher titres, suggesting that HIV-1 is not latent
at all but rather targets the lymphatic system preferentially. It is not understood why
the virus "hides" in the lymphatic system during this stage (Mitsuyasu, 1997; Pantaleo
et al., 1993) though it is conceivable that, like Herpes sp that hides in nerve ganglia,
HIV may use dendritic cells to establish a lymphatic reservoir to develop successful
quasi-species to assault the peripheral blood stream (Turville et al., 2002).
HIV enters the body via intravenous, parenteral or sexual routes and replicates
extensively in bone marrow, nerve, spleen, testicular, brain and intestinal cells as well
as many classes of immune cells including macrophages and T4 lymphocytes (Eron,
1997; Mitsuyasu, 1997). The means by which HIV causes immune-deficiency is hotly
debated (Hellerstein et al., 1999). Most of the current evidence supports the notion
that HIV depletes the number of functional T-cells, although other work suggests that
memory T-cell signalling is actually being manipulated by the virus (Hellerstein et
al., 1999). The host immune system certainly mounts a heroic response to the virus,
3
but due to the rapid rate of mutation of this RNA virus, the immune system eventually
becomes overwhelmed by the constant challenge and phenotypic variety of HIV-1
and ultimately becomes dysfunctional.
1.2 Viral Characteristics.
HIV is a Lentivirus (Virus Code 61.0.6.) of the family Retroviridae (Virus
Code 61.), having two copies of single stranded RNA (Goudsmit et al., 1986; Coffin
et al., 1995). The subfamily Lentiviridae comprises five serogroups, reflecting the
hosts with which they are associated (primates, horses, cats, cattle, sheep and goats).
Some serogroups have cross-reactive gag antigens (e.g., the ovine, caprine and feline
Lentiviruses are all cross reactive). Within the primate serogroup to which HIV
belongs, the Lentiviruses are distinguished by the use of CD4 protein receptors for
attachment (Coffin et al., 1995).
The genome of HIV-1 is 9,749 nucleotides (Muesing et al., 1985). This small
genome encodes for nine proteins that are located in the central region of proviral
DNA, and can be divided into three classes:
1. Major structural proteins, (forming components of the virion surface and core).
2. Regulatory proteins; tat (trans activating factor) and rev (regulator of virion
expression).
3. Accessory proteins; vpu, vpr (viral proteins u and r), vif (virion infectivity
factor) and nef (negative trans-activating factor) (Fig. 1.1) (Gallo et al., 1988).
1.3 HIV-1 Virus Particles.
HIV-1 virions are comprised of a nucleoprotein core and transmembrane
(gp41) env protein surrounded by a composite of the cell lipid bilayer and the viral
surface (gp120) protein. The nucleoprotein core of the virion is composed of the
following:
1. A matrix (MA [p17]) gag protein associated with the inner leaflet of the lipid
bilayer.
2. The capsid (CA [p24]) gag protein that forms the structural conical core of the
viral nucleoprotein complex.
3. The nucleocapsid (NC [p9]) gag protein that selectively binds two copies of
viral genomic RNA and associated tRNA molecules (Gottlinger et al., 1989;
Bryant and Ratner, 1990).
4. The pol enzymes in the nucleoprotein core are, HIV-1 protease (HIV-1 PR),
reverse transcriptase and integrase. The HIV-1 particle also contains the viral
proteins vpr and vif (Paxton et al., 1993; Karczewski and Strebel, 1996). It
has been proposed that the viral nef and tat proteins are incorporated into
HIV-1 particles (Welker et al., 1996, Harrich et al., 1996). The host-cell
protein, cyclophilin A, is also incorporated into HIV-1 particles by an
interaction with the capsid protein and is important for viral replication (Thali
et al., 1994; Franke et al., 1994; Braaten et al., 1996).
vpu
4
1.4 Replication of HIV. The HIV-1 replicative cycle begins with the attachment of the viral particle to
CD4 and related receptors on the surface of a cell (Maddon et al., 1986; Turville et
al., 2002). Other possible modes of viral binding to cells have been hypothesised.
These involve antibody-coated virions (Homsy et al., 1989), and the glycolipid
galactosyl ceramide (Harouse et al., 1991; Bhat et al., 1993; Long et al., 1994)].
The post-binding events leading to fusion with the cell membrane have
become popular for inhibition (Eckert and Kim, 2001; Jiang et al., 2002). HIV-1, like
gag pol
vif tat
rev
env nef
vpr
Figure 1.1: Structure of HIV-1 Genome and Important RNAs Produced. The primary HIV-1
transcript contains multiple splice donors (5' splice sites), and splice acceptors (3' splice sites) within
the principle genes gag, pol and env, which can be processed to yield more than 30 alternative
mRNAs (Schwartz et al., 1990a). Many of these RNAs are polycistronic, containing the open reading
frame of more than one protein, eg a fused gag-pol precursor termed Pr55gag terminates within vif and
so contains all the proteins of gag, pol and allows the production of vif. The polycistronic mRNAs
typically express a single gene product that is translated to a poly protein or incorporated into the
virion as genomic RNA. The efficiency of the initiation codon and the proximity of the initiation
codon to the 5’ end of the mRNA govern open reading frame choice (Schwartz et al., 1992).
5
most other retroviruses infect cells in a pH-independent manner (McClure et al.,
1990). The fusion event itself has been mapped to a mutable region on the viral
surface known as the V3 loop of the viral coat glycoprotein gp120 (Skinner et al.,
1988; Page et al., 1992; Ghiara et al., 1997). Fusion occurs to chemokine receptors on
the host cell surface, and the selection of receptor is determined by the specific
mutations of the V3 loop. Receptors selected include fusin, CKCR4 (T-cell tropic
strains) (Feng et al., 1996), CKR5, RANTES, MIP-1 and MIP-2 (macrophage tropic
strains) (Alkhatib et al., 1996; Deng et al., 1996; Dragic et al., 1996; Choe et al.,
1996; Doranz et al., 1996). Some chemokine receptors, e.g. CKR3 and CKR-2b have
also been identified as accessory proteins (Choe et al., 1996, Doranz et al., 1996).
Once the viral core gains entry to the cell, a protective envelope about the core
is degraded by the packaged cellular enzyme cyclophilin A (Luban 1996, Gamble et
al., 1996). This allows the release into the cytoplasm of a pre-integration complex
(PIC), which is actively imported into the nucleus in a process mediated by the HIV-1
matrix protein and a host virion-associated matrix-interacting protein termed VAN
(virion associated nuclear protein) (Gupta et al., 2000). The PIC comprises all the
viral and purloined host proteins necessary for reverse transcription of the RNA viral
sequence and integration into the host genome (Farnet and Bushman, 1997; Gupta et
al., 2000). Once the provirus is integrated, the expression of viral genes requires the
collaborative activities of the host-cell transcription machinery (RNA polymerase and
transcription factors Sp1 and NF-κB) and viral regulatory proteins (tat and rev) (Al-
Harthi and Roebuck, 1998).
Replication can be separated into early and late stages. Since HIV-1 produces
a single transcript of structural and enzymatic mRNA (termed Pr160gag-pol), there is a
need to differentially process the RNA to express all proteins necessary to form
virions. During early replication, full length Pr160gag-pol is produced and
subsequently spliced by the cellular spliceosomal machinery yielding the "early" 2-kb
mRNAs. These transcripts give rise to the proteins tat, rev, nef, env and vpu.
Subsequent bursts of proviral mRNA production now become directly
controlled by tat. Tat, a transcriptional transactivator, binds RNA (Rosen et al., 1985).
Tat binds to a short stem loop structure known as the transactivation response element
(tar), that is located at the 5' terminus of all HIV-1 RNAs. The binding of tat to tar
increases the activation of transcription of the HIV long terminal repeats (LTRs) by at
6
least 1000-fold (Southgate and Green, 1991). It appears that tat acts principally to
promote the elongation phase of HIV-1 transcription, so that full-length transcripts
can be produced (Kao et al., 1987, Feinberg et al., 1991). In the absence of tat, HIV
generates primarily short transcripts (deficient in continuing RNA transcription once
initiated) and is unable to transcribe from the DNA template beyond a few hundred
nucleotides (Cullen, 1990).
Rev functions post-transcriptionally to selectively induce the expression of
unspliced and singly spliced mRNAs (Feinberg et al., 1986; Sodroski et al., 1986;
Malim et al., 1989b). Rev facilitates the export from the nucleus of unspliced
(Pr160gag-pol [~9.7-kb]) and incompletely spliced (Pr55gag [~4-kb]) mRNA,
effectively switching viral replication from an early to a late phase. The Pr160gag-pol
mRNAs encode the structural and functional proteins (as well being genomic RNA
which is packaged into virions), while the Pr55gag mRNA encodes nucleoproteins
(Daefler et al., 1990). Thus rev controls the differential expression of structural and
regulatory proteins.
Pr160gag-pol migrates to the outer surface of the cell wall and accumulates at
the membrane, attaching via a myristoylation tag (Dreyer et al., 1989). The Pr55gag
precursor is glycosylated and also migrates to the site of budding, assisted by the viral
matrix and host VAN proteins (Perrin et al., 1998; Gupta et al., 2000). The growing
bud incorporates genomic RNA, and viral accessory proteins as well as appropriated
cellular enzymes such as cyclophilin A (Luban, 1996), thioltransferase (Davis et al.,
1997) and VAN (Gupta et al., 2000). Indeed the promiscuous nature of lentiviral
packaging of both genetic material and proteins makes it possible to incorporate
different viral RNA strands and host proteins (Costa, et al., 2000; Gupta et al., 2000).
HIV-1 PR is a proteolytic enzyme that is active before or shortly after virion
release (Tessmer and Kraisslich, 1998; Park and Morrow, 1993), auto-catalytically
cleaving itself from the centre of the zymogenic polyprotein PR160gag-pol. Then,
essentially acting like molecular scissors, HIV-1 PR cleaves the remainder of the
precursory polyproteins (Pr160gag-pol and Pr55gag) into the structural and functional
forms. The enzymes and proteins of HIV-1 then fold and self-assemble to form a
fully mature and infective virion (Meek et al., 1989, Barklis et al., 1997).
One of two things may occur once HIV-1 proteins have matured:
1. The cell may start to form syncytia. This happens when the viral envelope
protein gp41 contacts another cell followed by fusion, releasing infective
material into more cells. This can lead to giant cells or to a mesh of cells in
one mass (Eron, 1997).
2. Budding may occur, causing the release of new virions.
The infected cells may be destroyed in several ways, either by formation of
syncytia, by cell lysis caused by osmotic leeching, or by antibodies/killer T-cells
targeting viral proteins integral to the cellular membranes and “flagging” the cell for
destruction (Ciriolo, 1997).
ATTACHMENT BUDDING
FUSION
PROTEINS
PIC
GENOME RT
IN/SP
Figure 1.2: The Major Steps of HIV-1 Replication. HIV-1 attaches to cellular CD4 receptors,
followed by fusion to chemokine receptors. Once the capsid gains entry to the cell, the PIC
(preintegration complex) uncoats and begins reverse transcription (RT), followed by integration of the
provirus, DNA transcription to RNA, RNA splicing and export (IN/SP). RNA is then translated to
Proteins or incorporated into the growing virion as genomic RNA, finally the new virion buds from the
cell and is released to mature and infect a new cell. (Figure modified from Knightley, 1989).
7
8
1.5 HIV Accessory Proteins.
HIV-1 has four accessory proteins, nef, vif, vpr and vpu. These accessory
proteins are not absolutely required for viral replication in vitro, but are critical
virulence factors in vivo. Nef is expressed from a multiply spliced mRNA and hence
is rev independent. In contrast, vpr, vpu, and vif are the product of incompletely
spliced mRNA, and thus are expressed during late replication. Most of these
accessory proteins of HIV-1 have multiple functions including down regulation of
receptors, heightening infectivity, reducing T-cell activation and blocking cell
division. However, their direct roles in viral replication appear to be their most
important function.
Nef acts post-transcriptionally to decrease the cell-surface expression and
increases the rate of endocytosis and lysosomal degradation of CD4 receptors
(Garcia and Miller, 1992; Aiken et al., 1994; Schwartz et al., 1996). A consequence
of this is a decrease in the continued infection of a cell (which already contains a
provirus) and autoinfection from progeny virions. Hence the efficiency of infecting
new cells within the host is increased.
Vpr plays a role in the ability of HIV to infect non-dividing cells and prevents
cell proliferation during chronic infection (Rogel et al., 1995; Jowett et al., 1995;
Braaten et al., 1995). Vpr also facilitates the nuclear localisation of the pre-
integration complex (Heinzinger et al., 1994, Gallay et al., 1996). The vpu
polypeptide is an integral membrane phosphoprotein that is localised in the internal
membranes of the cell (Sato et al., 1990). Vpu is translated from within the coding
sequence env at a level tenfold lower than that of env itself as the vpu translation
initiation codon is not efficient (Schwartz et al., 1990b). Within HIV-infected cells,
complexes between the viral receptor, CD4, and the viral envelope protein form in the
endoplasmic reticulum causing the trapping of both proteins within this compartment.
The formation of intracellular env-CD4 complexes thus interferes with virion
assembly (Willey et al., 1992). Vpu liberates the viral envelope by triggering the
degradation of CD4 molecules complexed with env (Willey et al., 1992). Aside from
this function, vpu also increases the release of HIV from the surface of an infected
cell (Schubert et al., 1996, Klimkait et al., 1990).
Vif is a 23-kD polypeptide that is essential for the replication of HIV in
peripheral blood lymphocytes, macrophages, and certain cell lines (Strebel et al.,
9
1987). In most cell lines, vif is not required, suggesting that these cells may express a
protein that can substitute for vif function (Liu et al., 1995). Virions generated in
permissive cells can infect non-permissive cells but the virus subsequently produced
is noninfectious (Camaur and Trono, 1996). Vif mutant virions have improperly
packed nucleoprotein cores as revealed by electron microscopy (Hoglund et al.,
1994).
1.6 Therapeutic Problems. Due to their requirement for viral replication, many of the proteins described
above have potential as therapeutic targets for drugs that can treat HIV-1 infection.
The main difficulties in generating effective drugs for any given target can be
attributed to any one or more of the following;
1. Unavailability of sufficient information on the particular target being
addressed.
2. Selectivity of compounds, where potent and specific inhibitors for the HIV-1
protein being targeted may also bind to similar host cell proteins.
3. Problem of creating cell-permeable antiviral agents that are orally-
bioavailable.
4. Other pharmacological problems that are difficult to predict and incorporate
into design such as degradation of the agent, toxicity, poor solubility, ability
to enter all body systems (eg crossing the blood brain barrier) and metabolism
of the agent.
5. Development of drug resistance through natural selection, resulting in a
weaker binding of the drug to its target.
A significant problem encountered with antiviral treatments to date has been
the emergence of viral resistance to drugs (Menendez-Arias, 2002). Resistance arises
through mutations, which decrease the affinity of an inhibitor for its target or alter the
substrate processing rates. Mutations may be lethal to the virus so that they cannot
form, infect or integrate, or they may be silent so that no noticeable difference can be
detected. Some mutations may reduce infectivity but still allow function or confer a
selective advantage to the mutant protein that leads to its dominance within the host.
The mutations occurring within HIV-1 have impeded treatment, especially in the long
10
term, as antibodies or inhibitors binding to a particular sequence in viral proteins
become ineffective once these sequences become mutated (Mayers, 1997). Currently
the best treatments for HIV-1 infection is a combination of inhibitors of HIV-1 PR,
also administered with inhibitors of other HIV-1 proteins such as reverse transcription
inhibitors (combination therapy).
An RNA virus such as HIV-1, has three key mechanisms of mutation:
1. Non-homologous recombination of the RNA between 5' and 3' fragments,
enhanced by the diploid nature of lentiviral genomes where co-packaging
allows template switching between the two RNA molecules (Hu and Temin,
1990a, Hu and Temin, 1990b). In general, first round replication analysis has
shown that an average of three changeovers occur each time a virion is
produced and switching often occurs about the long terminal repeats (LTRs),
highlighting the importance of recombination events in the HIV replicative
cycle (Jetzt et al., 2000; An and Telesnitsky 2002).
2. There exist mechanisms for RNA to mutate at second sites due to optimisation
of alternate RNA base pairings, if the first mutation disrupts localised RNA
structure (Klovins et al., 1997). For example, a single base mutation may
disrupt RNA secondary structure that facilitates subsequent mutations to occur
in order to correct - as close as possible - the structural aberration of the first
single mutation. Such errors are incorporated into the provirus, which the
progeny receive and continue to propagate (Klovins et al., 1997).
3. Being an RNA virus, HIV-1 requires an RNA dependent DNA polymerase
(reverse transcription enzyme). HIV-1 reverse transcriptase is a hypermutable
enzyme and does not faithfully copy the RNA template to DNA since it lacks
3'-5' exonuclease proofreading capabilities (Chary et al., 1997). The mutations
due to an HIV-1 reverse transcriptase error allow the virus to evolve at a rate
of approximately 1/1700 nucleotides inserted, by 1/+1 frameshifts, deletions
and additions (Chary et al., 1997). Such hypermutations lead to alteration of
cell-type tropism, drug resistance and perhaps escape of neutralisation by the
host immune system.
Resistance can arise from administration of an inhibitor. Some mutant virions are
able to escape the effects of the inhibitor, and due to removal of susceptible wild type
virions, the mutant strains become the dominant population (natural selection). The
11
strain of HIV-1, timing of initiation of a given therapy (or combination of therapies),
and the stage of infection has all been observed to influence resistance. (Menendez-
Arias 2002; Holguin and Soriano, 2002; Joly et al., 2002).
1.7 Hypothesis and Aims of Thesis.
Reverse transcriptase inhibitors were the first to enter the clinic but they have
shown low efficacy. HIV-1 PR inhibitors have now been available for about five
years and they have significantly extended the lives of HIV-infected people when
given as a cocktail with each other or with RT inhibitors. These inhibitors do
however suffer from the development of viral resistance. Specific difficulties
encountered with RT inhibitors have been:
1. The rapid mutation rate of HIV-1 (Struble et al., 1997).
2. The timeframe for the process of reverse transcription that can produce viral
DNA within 6 hours after entry (Zack et al., 1990). This gives a reasonable
window for therapeutic intervention with nucleoside analogues (NAs) [(NAs
are incorporated into DNA causing chain termination)], but requires highly
toxic dosing levels to maintain a therapeutic index with non-nucleoside
analogues (NNRTIs) [(inhibits the reverse transcriptase directly)].
3. Many NAs require phosphorylation by cellular kinases before they can be
incorporated into DNA synthesis. This can be an inefficient process in vivo
depending on the cell cycle, decreasing the NAs effectiveness (Peter and
Gambertoglio, 1998).
4. The process of reverse transcription is remote from the HIV-1 PR step, with
HIV-1 PR being inhibited after budding and reverse transcription inhibited
only after viral fusion. There are two consequences of this: (a) all cellular
mediated steps (between reverse transcription and protease activation) are not
addressed. Thus cells expressing virions can only be removed by the
compromised host immune system. (b) The cells that are producing virions
are providing a "sink" for combination therapy as many may have defective
proteins or genes involved in intermediate stages (eg integration or rev
function), but also active protease and reverse transcriptase.
5. The combination therapy is reported to be synergistic largely by cross
removal of mutants, but cross-resistant viruses arise under this selective
12
pressure, creating a population resistant to both inhibitor classes (Condra et
al., 1995).
6. There are no synergistic effects that would allow reduced dosage of either
drug except for bioavailability reasons for PIs (e.g. ritonavir strongly inhibits
cytochrome P450, increasing other PI plasma concentrations) and salvage
therapy (e.g. NA-pretreated patients develop mutations that can increase the
susceptibility of the viruses to the NNRTI) (Struble et al., 1997; Joly et al.,
2002). In both cases future treatment options can be compromised especially
in patients where combinations and toxicities of each drug are poorly
tolerated (Joly et al., 2002).
A better strategy than attacking polar stages of replication would be to attack
genuinely synergistic stepwise processes of viral replication. Because of the
advanced development and great success as a therapeutic of HIV-1 PR inhibitors
(Deeks et al., 1997; Wlodawer and Vondrasek 1998; Van Heeswijk et al., 2001), a
good synergisitic approach would be to target HIV-1 PR and another viral specific
protein that acts at a similar late stage of viral replication.
A challenge confronting a virus is to be as efficient as possible by encoding
the minimum number of genes in the smallest possible genome size. The most
efficient viruses overcome this problem by encoding genes within genes. The
activation of the alternate gene is manipulated through strategies such as amber stop
sequences or shifting the reading frame. HIV is a very subtle virus with respect to the
problem of size-efficiency. The long HIV-1 mRNA transcript is initially subject to
the full spliceosome processing of the cell. This is crucial to the virus as full splicing
leads to 2-kb RNAs encoding the early proteins. Rev facilitates the release of
incompletely spliced mRNAs into the cytoplasm that are translated into the
polyproteins Pr160gag-pol and Pr55gag, which then fall subject to HIV-1 PR
cleavage. This allows the appearance of all viral proteins necessary for parasitism,
with rev regulating the transition from early to late proteins (regulatory to crude
structural proteins) and HIV-1 PR differentially producing active structural and
enzymatic proteins. Indeed, no other processes of HIV-1 replication share the
intimate link that HIV-1 Rev and HIV-1 PR do, as HIV-1 PR is encoded in the
transcript which rev transports and proteolytic activation is the immediate viral-
13
specific step following rev export. Thus HIV-1 Rev and HIV-1 PR represent
contiguous stages that, if both were inhibited, could synergistically block the
replication of HIV-1.
Based upon this conclusion, it was decided to focus on two proteins (HIV-1
PR and HIV-1 Rev), which are required for viral replication and to work towards
designing new inhibitors of these proteins’ functions. A particular goal of the project
was the structural analysis of protein-ligand complexes, and design of potent
inhibitors of HIV-1.
Specific aims were:
1. To acquire and analyse high-resolution crystal structures for complexes of
HIV-1 PR with substrate, and to compare structures for substrate-, inhibitor-,
and product-bound HIV-1 PR complexes (Chapter 2).
2. To design, and synthesize novel HIV-1 PR inhibitors (Chapter 3).
3. To develop a reliable RNA-Rev binding assay and, to use an established
fluorometric HIV-1 PR assay to test prospective Rev and HIV-1 PR
antagonists in vitro (Chapters 3, 4 and 5).
My experimental work involved:
1. Acquisition and analysis of high-resolution data from HIV-1 PR:substrate
complexes of HIV-1 PR. Improved high resolution structures for HIV-1 PR
bound to a pre-organised β-strand substrate and its hydrolysed products,
determined using cryo-crystallographic techniques, provided new information
about substrate recognition and the mechanism of catalysis.
2. Design and synthesis of novel HIV-1 PR inhibitors. Computer-assisted design
of potent HIV-1 PR inhibitors and their synthesis, using a combinatorial
approach from cyclic peptide templates, provided compounds for testing in an
in vitro enzyme assay using a fluorogenic substrate (Bergman et al., 1995;
Abbenante et al., 1995).
3. Development of an RNA-binding assay for Rev and small molecule
analogues. Expression, labelling and purification of recombinant Rev protein
and RNA was undertaken and the material used to develop a BIAcore-based
assay to compare HIV-1 RRE binding to Rev and various peptide mimetics of
14
Rev including a helix-stabilised Rev34-50 peptide to test if helix induction
increases affinity to a biological target, as well as a competitive assay for
small molecule antagonists of Rev-RRE binding as a prelude to screening of
compounds in more advanced drug design.
15
Chapter 2: Crystal Structures of HIV-1 Protease Complexed
with Substrate and Products.
2.1 Introduction 2.1.1 HIV-1 PR Structure
2.1.2 HIV-1 PR Function
2.1.3 Mechanism of Catalysis
2.1.4 Protease Activation Assessed by Cryo-crystallography
2.1.5 Preliminary Crystallography
2.1.6 Aims
2.2 Results 2.2.1 Crystal Structure of Enzyme-Substrate Complex
2.2.2 Crystal Structure of Enzyme-Product Complex
2.3 Discussion 2.3.1 Insight from Enzyme Kinetics
2.3.2 Comparison of Substrate, Product and Analogue
2.3.3 Mechanism of Catalysis of HIV-1 PR
2.4 Experimental 2.4.1 Origin of Protein and Substrate
2.4.2 Crystallisation Methods
2.4.3 Structure Determination
2.4.4 Analysis and Alignment
16
2.1 Introduction 2.1.1 HIV-1 PR Structure
It was first proposed by Kramer et al., (1986) that inhibition of HIV-1 PR
might result in a possible treatment for HIV-1 infection. Kohl et al., (1988) and Loeb
et al., (1989) supported this hypothesis by demonstrating that mutation of key
residues of HIV-1 PR required for function prevented full maturation of virus
particles which were consequently non-infective. These early investigations and
subsequent X-ray crystal structure studies (Navia et al., 1989; Wlodawer et al., 1989;
Miller et al., 1989) enabled the design and development of a range of efficacious
inhibitors of HIV-1 PR, a handful of which are now successfully used in man for the
treatment of AIDS.
HIV-1 PR (Protein Accession; A02.001 [MEROPS], EC 3.4.34.16 [NC-
IUBMB], 144114-21-6 [CAS]) is classified as an aspartic protease, having two
catalytic aspartates in the active site. HIV-1 PR is inhibited by pepstatin (a common
hexapeptide inhibitor of aspartic proteases from Streptomyces sp), and shares both
structural and biochemical similarities with other aspartic proteases (Pearl and Taylor
1987; Lapatto et al., 1989). Aspartic proteases perform diverse roles from regulating
blood pressure (renin) and digestion (pepsin and chymosin) to industrial food
processing (cardosin A and mucor renin in cheese production). Aspartic proteases
usually process polyprotein precursors (Costa et al., 1997) and are also found in
viruses, bacteria, and fungi (e.g. penicillopepsin, rhizopuspepsin, endothiapepsin)
(Tang et al., 1978). They perform crucial roles in pathogenic organisms such as
Candida tropicalis, the blood fluke Schistosoma japonicum and the malaria parasite
Plasmodium falciparium and are important in higher plants, such as barley aspartic
protease from Hordeum vulgare L and cardosin A from Cynara cardunculus L (Costa
et al., 1997).
Members of the aspartic protease family (examples shown in Fig. 2.1) have
similar three-dimensional structures that are predominantly monomeric and feature
multiple β-sheets (Miller et al., 1997). The major differences between members of
this class are in surface loops, in particular those that surround the entrance to the
active site and substrate-binding cleft. Such loops vary in position by up to 8 - 10Å
(Sielecki et al., 1989). Minor variability in the polypeptide chain length and pH
optimum for enzymatic activity (which is generally acidic) has been observed
(Sielecki et al., 1989). Plant aspartic proteases have an additional sequence of ~100
amino acids that makes them similar to sphingolipid activating proteins (saposins)
(Costa et al., 1997). Most aspartic proteases are monomeric enzymes with two closely
approaching aspartate residues that are responsible for catalytic activity.
HIV-1 PR differs from most other aspartic proteases by being a homodimer,
with each of the two identical halves forming a unique, single and symmetrical active
site. The active site aspartic acid residues form aspartyls by sharing a single proton
between them at mildly acidic pH (Pearl and Taylor, 1987; Wlodawer, 1994). HIV-1
PR contains four short β-strands rather than the six large β-strands of other aspartic
proteases, and it is also small, being less than 70% of the size of pepsin, with an
11kDa monomer comprising 99 amino acids in its active form (Meek et al., 1989)
(Fig. 2.1).
B
A
C
E D
Figure 2.1: Crystal Structures of Aspartic Proteases. Representative members of the family of
aspartic proteases are A, Penicillopepsin (pdb accession 1apt); B, Rhizopuspepsin (pdb accession
2apr); C, Human Cathepsin D (pdb accession 1lyb); D, HIV-1 PR (pdb accession 5hvp); E, Pepsin
(pdb accession 1pso). Shown are co-crystal structures with the family inhibitor pepstatin (not shown) at
<2Å resolution except Cathepsin D (2.5Å). It can be seen there is a predominance of monomeric forms
and multiple β-sheets, in contrast to HIV-1 PR that is smaller and dimeric.
17
The structure of HIV-1 PR (Fig. 2.2) features extended β-hairpin chains that
form "flaps". These are not well ordered until they close upon inhibitors/substrates
(Wlodawer and Erickson 1993; Wlodawer et al., 1989), causing a major structural
change that creates the active site and permits numerous van der Waal and hydrogen-
bonding interactions with the ligand. The flaps are key distinguishing features of this
enzyme (Shao et al., 1997). The binding cleft that accommodates the substrate or
inhibitor is notably hydrophobic but a water molecule is used to bridge the enzyme
flaps to inhibitors and possibly also substrates (Navia et al., 1989). This is not thought
to be the catalytic water molecule needed to process substrates by hydrolysis, but
rather plays a structural role in facilitating the ligand fit to the active site, and can be
displaced in many inhibitor complexes (Wlodawer and Erickson, 1993).
FLAPS STRUCTURAL WATER
80’S LOOPS
SUBSTRATE – BINDING CLEFT
30’S LOOPS
ACTIVE SITE
Figure 2.2: Structural Features of HIV-1 PR in the inhibitor-bound “closed” form (inhibitor is
pepstatin (not shown) - pdb accession 5hvp). Labelled are the key structural features including; the
active site, substrate binding cleft, structural water, flaps and the 30’s and 80’s loops that are important
to drug resistance.
The active site contains the sequence Asp25-Thr26-Gly27 and since the
enzyme is homodimeric, this sequence is duplicated in the other half of the active site
(Asp125-Thr126-Gly127). The C- and N- termini as well as the threonine residue
from each monomer interact to hold the two halves of the active site together in a
dimeric state (Pearl and Taylor, 1987) reported as a "fireman's knot or grip”
(Wlodawer et al., 1989, Wlodawer and Vondrasek 1998; Strisovsky et al., 2000).
18
19
2.1.2 HIV-1 PR Function
HIV-1 PR processes the translated HIV-1 polyprotein Pr160gag-pol substrate
in a ratio of 1 Pr160gag-pol to 20 Pr55gag polyproteins (Roberts et al., 1992). Catalytic
activity for HIV-1 PR and most aspartic proteases (Tang et al., 1978) is optimal at
high ionic strength and low pH (pH 5.5 for HIV-1 PR) (Bergman et al., 1995). This is
the pH at which HIV-1 PR is most commonly assayed and crystallised (Hui et al.,
1993).
One controversial issue, that is important to therapeutic intervention, concerns
the precise stage during viral replication that HIV-1 PR exerts its activity. It is now
reasonably certain that HIV-1 PR becomes active following budding as opposed to
intracellularly prior to budding. Within the juvenile virion, the concentration of
proteins at the site of budding decreases the pH in a localised area approaching the
optimum pH for activity of HIV-1 PR. This has also been suggested for Rous
Sarcoma Viral protease (Jaskolski et al., 1990).
The initial step in processing the gag-pol precursor polypeptide is correct
folding and dimerisation of HIV-1 PR, requiring the loss of the zymogenic N-
terminal portion of the precursor (56 residues). Studies conducted on the p6pol-PR
cleavage site support proteolytic activity following assembly of structural proteins
(Tessmer and Krausslich 1998; Yu et al., 1998; Louis et al., 1999), since the p6pol-PR
cleavage site (N-terminal side of the protease, releasing the protease from the
zymogenic 17kDa form) reportedly acts as a negative regulatory element for protein
folding and dimerisation. At elevated pH, the regulatory element destabilises the
dimer, minimising or regulating enzyme activity during formation of the viral
particle. Once the pH decreases, the removal of p6pol-PR proceeds more rapidly,
leading to higher concentrations of active enzyme and processing of the gag-pol
precursor.
It has been convincingly demonstrated that HIV-1 PR inhibitors prevent full
maturation of virus particles (Pollard, 1994; Carpenter et al., 1998; Tavel, 2000), and
thus the enzyme must be active following budding of immature virus particles from
infected host cells. Protease inhibition during this late activation phase conceivably
presents a larger problem than during early activation, since a therapeutic agent
would need to be packaged into a budding virion to be effective. In such a scenario,
the effectiveness of a therapeutic agent would be dependent on achieving a
concentration within the virion sufficiently high to suppress catalysis by HIV-1 PR.
The enzyme recognises eight cleavage sites in gag-pol and gag polypeptides
as shown in Figure 2.3 and Table 2.1 with a preference for hydrophobic residues.
Remarkably, the enzyme is able to cleave a peptide bond between a hydrophobic
residue and a proline, something previously unreported for mammalian proteases, and
this feature formed an early basis for development of inhibitors with selectivity for
this viral enzyme over host proteases. The substrate-binding cleft is well defined
within the enzyme and can accommodate a seven amino acid polypeptide within its
cavity (Fig. 2.4). The binding mode is dominated by hydrophobic interactions with
the majority of H-bonding to the peptide backbone itself.
20
gag
pol
NC
PR
RT
RH
IN
MA = Matrix CA = Capsid NC = Nucleocapsid SP = Splice Point of NC PR = Protease RT = Reverse Transcriptase RH = RnaseH IN = Integrase
gp120 gp41
env
Pr55gag
Pr160gag-pol
CA SP MA
p17 MA p24 NC p7 CA
p66/51 RT/RH p32 IN p11 PR
pol
gag
Figure 2.3: The Processing Profile of HIV-1 PR is shown and the arrangement of the proteins within
the virion upon maturation (figure modified from Edelman 2001).
S2'S1
S1'
S3'
S3
S2
NH
HN
NH
HN
NH
HN
O
O
O
O
OP1'
P2'
P3'
P1
P2
P3
P4
R'H3N
O
S4
O O
O O
Figure 2.4: The Nomenclature of Protease Active Sites. A hexapeptide is shown in the active site of
HIV-1 PR labelled with the standard nomenclature of Schetcher and Berger, (1967) where S4-S3' are
regions of the enzyme binding pocket and P4-P3' are the corresponding substrate residues. The scissile
amide bond is between P1 and P1', corresponding to the active site between S1 and S1'.
Cleavage sites P5 P4 P3 P2 P1 P1' P2' P3' P4' P5’
p17 - p24 (MA) Val Ser Gln Asn Tyr Pro Ile Val Gln Asn
p24 – Non (CA) Lys Ala Arg Val Leu Ala Glu Ala Met Ser
Non - p7 (NC) Ser Ala Thr Ile Met Met Gln Arg Gly Asn
p7 - p6 (SP) Arg Pro Gly Asn Phe Leu Gln Ser Arg Pro
Non - p11 (PR) Val Ser Phe Asn Phe Leu Gln Ile Thr Leu
p11 - p51 (RT) Cys Thr Leu Asn Phe Pro Ile Ser Pro Ile
p51 - p15 (RH) Ile Arg Lys Ile Leu Phe Leu Asp Gly Ile
p15 - p34 (IN) Gly Ala Glu Thr Phe Tyr Val Asp Gly Lys
Table 2.1: Native Cleavage Sites for HIV-1 PR. (Table 2 was developed from Wlodawer, 1994;
March and Fairlie, 1996; Abdel-Meguid, 1993. Non = a non-coding region).
2.1.3 HIV-1 PR Mechanism of Catalysis
It has been hypothesised that substrates must adopt an extended conformation,
as observed for other aspartic proteases (Parris et al., 1992), in order to enter the
active site of HIV-1 PR. Certainly inhibitors and substrate analogues are found in
general to bind as an extended (β-strand) conformation within the active sites of
proteases, including HIV-1 PR (Tyndall and Fairlie, 1999; Fairlie et al., 2000),
although this does not mean they necessarily have to adopt such a conformation prior
21
to binding. It does suggest though that a substrate that is conformationally restricted
to an extended conformation prior to binding with HIV-1 PR, may be entropically
favoured to bind due to its pre-organisation. This had been hypothesised by Swain et
al., (1990); that a substrate or inhibitor already in an extended conformation might
have an advantage accessing the substrate-binding cleft, termed "selection hypothesis"
(Reid and Fairlie, 1997). However, surprisingly few substrates or inhibitors have been
designed with the advantage of being fixed or pre-organised in an extended
conformation for interaction with any protease. In fact almost all known HIV-1
protease inhibitors are conformationally flexible and must therefore pay an entropic
penalty for rearranging into the protease-binding conformation.
Though the steps of catalysis are known for proteolytic enzymes (Fig. 2.5),
there have been many proposed hypotheses for the mechanism of catalysis of HIV-1
PR. Catalysis is usually considered as occuring via either a general acid-base
mechanism (Polgar 1989) or via a covalently bound intermediate (Hofmann et al.,
1984; Hofmann et al., 1988). X-ray crystallographic reports for enzyme-inhibitor
(EI*) complexes (Navia et al., 1989; Wlodawer et al., 1989; Lapatto et al., 1989;
Swain et al., 1990; Jaskolski et al., 1991) have been important in understanding the
catalytic mechanism of HIV-1 PR, but those studies fail to accurately portray early or
late events in catalysis as they are not based on actual data of substrate or product
complexes. The other main approach taken to investigate catalysis has been
biochemical mechanistic studies (Ido et al., 1991; Hyland et al., 1991a; Hyland et al.,
1991b; Rodriguez et al., 1993; Meek et al., 1994). From these studies have emerged
three general proposals for the mechanism of catalysis by HIV-1 PR, as highlighted in
Figure 2.6(a) – (c).
E + S ES ES* ET*/EI* EP* EP E + P
Figure 2.5 The Steps of HIV-1 PR Catalysis: E+S represents the enzyme (E) and substrate (S) before
binding. ES is the bound complex in the ground state (no strain). The ES* is a higher energy state due
to straining of the complex towards ET* (the transition state intermediate). EI* is the inhibitor complex
believed to mimic ET*. EP* and EP are product complexes strained or unstrained in which cleavage of
the substrate has occurred. E+P is the final state where complete dissociation of the products from
enzyme has occurred (modified from Hofmann et al., 1984; Hofmann et al., 1988). The section
contained in the boxed area is mechanistically described in Figure 2.6(a) – (c).
22
HN
P1'
P1
D25
OO
D125
OO
HN
P1'
P1
D25 O
D125O
OO
O
P1
OH2N
P1'
P1
D25 O
D125
O
O
O
H2N
P1'
P1
D25 O
D125
O
OH
OHO
O
OH
D25
OO
O
OD125
OHO
OH H
H2N
P1'
H H
3 1 2
Figure 2.6: The Three Proposed Mechanisms of Catalysis by HIV-1 PR (a) – (c). Shown in the Figure 2.6(a) – 2.6(c) are the three proposed mechanisms of catalysis for HIV-1 PR.
4 5
Figure 2.6(a) Mechanism 1 (above) depicts: (1) D25 acting as a nucleophile to attack the scissile carbonyl forming the covalent E-S complex (2), which rapidly proceeds to (3) via activation of the peptide bond. In (3) a water molecule enters and is attacked by the Lewis base D125, leading to (4), a covalently short lived E-P complex that auto-electronically proceeds to the free products (5).
H2N
O
OHHN
P1'
P1
D25
OO
D125
O
O
H
HN
P1'
P1
D25
OO
D125O
O OHH
P1'
P1
HO
H
O
OH
D125
O
O
D25
OO
H
1 2 3 Figure 2.6(b) Mechanism 2: D125 activates a nucleophilic water leading to the reaction state (2), that concomitantly proceeds by reversal of this acid/base reaction (such that D25 attacks the substrate) to the formation of products (3).
H2N
OH
OHN
P1'
P1
D25
OO
D125
O
O
H2N
P1'
P1
D25
OO
D125O
HO OP1'
P1
O
O
D125
O
O
D251 2 3
OO
HOH H
H
H
23
Figure 2.6(c) Mechanism 3: The catalytic water molecule is activated by D25, attacking the scissile carbonyl, thereby electronically activating the scissile amide nitrogen to attack the hydroxyl of D125, so that (2), a zwitterionic gem-diol intermediate is formed which then proceeds to products (3), rapidly through internal electronic rearrangement.
24
Mechanism 1 is historically based and suggests hydrolysis proceeds via a
covalent E-S complex to a covalent E-P complex, before dissociation to free products.
This mechanism has been largely discredited now (Hofmann et al., 1984; Hofmann et
al., 1988), because no evidence for covalent complexes has been found. Mechanism 2
has been popularized by Wlodawer and co-workers and is a general acid/base
concerted reaction that forms a neutral intermediate. Mechanism 3 was originally
proposed by Meek and co-workers and involves a similar general acid/base reaction
forming a zwitterionic gem-diol intermediate that proceeds to products.
For all three proposals, the nucleophilic behaviour of the active site water may
be facilitated by substrate interactions with HIV-1 PR at either side of the scissile
amide bond (cleavage site). In this respect, fitting of the peptide inhibitor/substrate
side chains into the indentations that line the substrate-binding groove may help to
stretch the scissile amide bond towards the transition state for hydrolysis. The
structural water molecule that bridges to the flaps may also stretch, strain or distort
the amide-substrate backbone through the carbonyls either side of the scissile peptide
bond. It is known that once an inhibitor enters the hydrophobic cleft, the flap regions
close in, forming a "roof" over the complex, associating through H-bonding
interactions with the structural water without strain and forming a stable, non-
covalent complex with one aspartate protonated and the other unprotonated.
2.1.4 Protease Activation Assessed by Cryo-crystallography
To distinguish between these various mechanistic proposals for protease-
activated hydrolysis, we were interested in directly investigating the interactions
made between the substrate and protease by acquisition and analysis of high-
resolution X-ray crystal structure data from crystals of HIV-1 PR complexed with
substrate and/or product(s). There is currently little such information available for any
proteolytic enzyme with respect to structural changes occurring upon substrate
binding and product formation.
The reason for this is that catalysis is rapid and there are few techniques
available for structurally investigating a protease-catalysed hydrolysis of a peptide
substrate before, during and after substrate processing. There were only two practical
crystallographic methods available prior to commencement of this Ph.D.
25
1. Ultrafast collection methods that rely on a synchrotron X-ray source and a
large pool of crystals. This technique suffers from synchronisation of the
reaction so that all molecules are in the same state of catalysis at the same
time within the crystal lattice (Farber 1999).
2. Thermodynamic-based flow cell/equilibrium experiments, where a
crystallised enzyme is exposed to a large amount of substrate and allowed to
reach equilibrium, or maintained as an “open” system to release products
before mounting and data collection. This method also requires a pool of
crystals for optimisation and also needs a large enough lattice to allow
diffusion (of substrates and products) and conformational changes of the
enzyme (Farber 1999).
In each case the collection and refinement is difficult and often has poor
statistics (due to the difficulties of the systems) and therefore the interpretation can be
difficult and often subjective (Farber 1999, Farber 1989; Gupta et al., 1993;
Vershuren et al., 1993; Bolduc et al., 1995; Mosi et al., 1997). Because of these
difficulties there are only a few precedents in which protease-substrate complexes
have been crystallised and their structures determined, three of which are for
retroviral aspartyl proteases (Prabu-Jeyabalan et al., 2000; Laco et al., 1997; Rose et
al., 1996), and a fourth model-based study (Popov et al., 1999).
The earliest study by Rose et al., (1996), reported trapping one of the two
possible cleaved products in the active site of HIV-1 PR. Another structure they
solved showed only the second product in the active site of SIV protease. These two
moderate resolution crystal structures were modelled so that the two halves of the
disparate structures could be combined in silico in order to infer the structure of the
protease-substrate complex following peptide hydrolysis, and to comment upon
events leading to catalysis. The study and reported findings suffer from the fact that
an intact substrate or both products could not be found simultaneously in a single
protease active site, and because structural changes to HIV-1 PR that occur on
substrate binding are likely to be specific for the particular substrate. Without
information on an intact substrate-protease complex, it is difficult to accurately
predict the initial molecular events that define catalysis.
26
A second study (Laco et al., 1997) reported on the effectiveness of mutating
an active site residue (D30N) in FIV protease, thereby preventing substrate cleavage
without interfering with protease structure. These workers co-crystallised FIV
protease with an inhibitor and compared this structure with an identical wild-type
complex. Since native substrates could not be crystallised with FIV protease, the
researchers had to compromise by crystallising FIV protease with a peptide sequence
corresponding to the statine-based inhibitor that also resembles the capsid p24 – non
coding sequence of HIV-1 PR (being Ac-NA-Val-Leu-Ala-Glu-NA-NH2 where NA
is naphthylalanine). This peptide was only a poor substrate for FIV protease, being
turned over slowly. Furthermore the assignment of the active site mutated residues
was ambiguous, as too the conformation of the substrate, making their observations
about early stages of catalysis questionable.
A third study published in Russian (Popov et al., 1999), was a comparison of
the crystal structure of HIV-1 PR bound to the peptide inhibitor JG365 with its
computer model to develop accurate parameters for modelling a substrate (Ser-Gln-
Asn-Tyr-Pro-Ile-Val). This model was extrapolated to infer a mechanism for
catalysis. The RMS deviation alone for the inhibitor JG365 model to the crystal
structure was 0.87Å and, even assuming that this error did not increase in the
substrate model, this deviation is too high to accurately comment upon any proposed
mechanisms of catalysis.
A fourth and more recent study (Prabu-Jeyabalan et al., 2000), used a native
HIV-1 PR substrate of the capsid (p24 – non coding region being Lys-Ala-Arg-Val-
Leu-Ala-Glu-Ala-Met-Ser). However, this was not a very rigorous study and does not
add any new information to the changes HIV-1 PR undergoes through catalysis or
offer a plausible mechanism for catalysis.
2.1.5 Preliminary Crystallography
During an honours degree by this candidate (Pattenden et al., 1998), X-ray
diffraction data was collected at room temperature (290K) on crystals of a novel
substrate 1 (Fig. 2.7) bound to HIV-1 PR and a D25N mutant HIV-1 PR. The D25N
HIV-1 PR is a mutant protease in which the catalytic aspartate (position 25 in the
monomer) has been replaced with an isoelectronic and isostructural asparagine, thus
facilitating comparable crystal packing but removing catalytic activity.
That study was problematic as there was ambiguity in assigning specific
conformations at room temperature to both the ligand(s) and important regions of the
protein. The aim was to compare the binding of the bicyclic substrate 1 to both wild
type and mutant (D25N) HIV-1 PR. This substrate was highly novel, being the
equivalent of a hexapeptide substrate pre-organised in an extended conformation
through cyclic restraints. The mutant enzyme, being unable to process substrate, was
expected to trap the bicyclic substrate in the active site, thereby furnishing detailed
structural information on the initial stabilised E-S complex. On the other hand the
wild type enzyme, possessing the catalytic residues (D25, D125), was expected to
bind to the bicyclic substrate and cleave the Tyr-Tyr bond, thereby allowing insight to
either the transition state (if cleavage was slow) or a product-bound form (if product
dissociation was slow).
202
203
205
204
NH
HN
O
O
O HN
NH
O
O
O
NH
O
204
205201
206
1
Figure 2.7: The Substrate used in Crystallography Experiments. Labelled on the substrate (1) is
the crystallographic nomenclature of “residue id” used for residue and atom identification.
Electron density was observed for the bicycle 1 in the crystal structure with
the proteolytically inactive D25N mutant HIV-1 PR, confirming that it had been
trapped intact in the substrate-binding site. However the complex contained several
alternate conformations that are in important regions of the protease (near residues
N25, N125, I50, G51). Residues I50 and G51 are located in the key flap region, and
I50, I150 co-ordinate through their amide bonds to the conserved water molecule
which in turn bridges to the carbonyl oxygen atoms of the bicyclic molecule that
flank either side of the ‘scissile’ amide bond. Ambiguities in this region of the
structure negatively impacted on the final analysis of the structures and on
mechanistic interpretations.
27
It was particularly unfortunate that the compromised active-site residues N25,
N125 were in two distinct conformations. Modelling of these residues indicated that
28
they were related by a rotation around the amide bond of 180°, apparently due to
unusual 2Fo-Fc electron density in the substrate carbonyl directed towards N25. The
initial state of catalysis could therefore not be accurately ascribed. The extraneous
electron density for the putative bicyclic substrate was alternatively modelled as a
gem-diol transition state intermediate, in the reverse binding orientation for the
ligand, and as an amide tautomer. None of these possibilities satisfactorily accounted
for the extra density or improved the crystallographic R factors. Rather, a second
conformation was constructed for the entire substrate, which allowed the difficult
2Fo-2Fc-electron density map to be resolved with a slight improvement in the Rfactor.
Though this allowed the putative scissile carbonyl to enter the unusually placed
electron density, it meant that a convincing specific assignment of the position of 1
could not be made, and so the binding mode was ambiguous, as for the structure by
Laco et al., (1997).
The crystal structure obtained for the bicyclic structure 1 complexed with wild
type (D25) HIV-1 PR, had lower quality electron density than the D25N mutant
complex. Cleavage of the putative substrate could be detected, since the two expected
products were visible and slightly separated. Thus it was established that 1 is indeed a
substrate for HIV-1 PR. However, there was some ambiguity in the C-terminal
product due to a lack of 2Fo-Fc-electron density, making it possible to model the
reverse orientation of this ligand without affecting the R factors. It was not possible to
comment informatively on possible flipping of the residues about I50/I150 due to the
low resolution of the data.
2.1.6 Aims
Because of the ambiguities in important regions of the structures described
above, it was decided to reinvestigate new crystals by X-ray structure techniques
under cryostatic conditions where molecular motion would be greatly reduced. The
same synthetic, catalytically active, wild-type HIV-1 protease and the same inactive
(D25N) mutant were again separately crystallised with the bicyclic peptide substrate
1 and an analogue of the hexapeptide Nle/Leu-Val-Phe-Phe-Ile-Nle/Leu, with the
scissile bond located between the two phenylalanines.
The purpose of these new structural refinements was to potentially proffer
new detailed information on the mechanism of HIV-1 PR catalysis by determining the
structure of a substrate with catalytically inactive D25N-enzyme as well as
catalytically active D25-enzyme under cryostatic conditions. We expected to be able
to compare three-dimensional structures of the “E-S” and “E-P” complexes. The
unusual bicyclic substrate 1 was selected for this study because:
1. It would allow direct structural comparison of the E-S and E-P complexes,
containing 1 and its cleavage products respectively, with the structure known
for HIV-1 PR bound to an analogue of 1, namely 4 (Fig. 2.8), that was
designed to mimic the transition state “E-T” (Martin et al., 1999) by
incorporating an hydroxyethylamine ‘transition state isostere’ replacement for
the scissile amide bond. Thus in principle it would be possible to compare
structures for “E-S”, “E-T”, and “E-P”, thereby affording new insight to the
catalytic process.
2. Previous work conducted in the Fairlie laboratory (University of Queensland)
had shown that all proteolytic enzymes bind inhibitors and substrate-
analogues in an extended (β-strand) conformation (Fairlie et al., 2000).
Hence, the chance of successfully trapping the substrate should be greatly
improved if the substrate is constrained in an extended conformation (Martin
et al., 1999). Indeed, bicyclic substrate 1 is a highly constrained mimic of a
hexapeptide locked into a β-strand extended conformation (Fairlie et al.,
2000), as suggested by earlier work on a bicyclic inhibitor analogue (Reid et
al., 1996).
3. The bicyclic inhibitor 4 and each of its two cyclic components (2,3) have
served as templates for either combinatorial synthesis or as the basis of
inhibitor design. Therefore the information gained from this study may
provide some insight to inhibitor design – particularly for inhibitors closely
analogous to substrates (Reid and Fairlie 1997; Fairlie et al., 2000).
4. The individual component cycles (e.g. 2 and 3) of bicyclic 1 are known to be
much better inhibitors (Ki 8µM and 15µM respectively) of HIV-1 PR than the
tripeptides they mimic (Ki 60µM) (Reid and Fairlie 1997). It was therefore
expected that these products of hydrolysis of 1 would be retained in the active
site longer than normal peptide cleavage products, possibly long enough to be
co-crystallised with enzyme.
29
2 3
4
NH
HN
O
O
O HN
NH
O
O
O
NH
HN
O
O
O HN
NH
O
O
O
NH
O
NH
HN
O
O
O
HN
NH
O
O
O
H3N
O
O
HNOH
1
Figure 2.8: Structure of Novel Substrate (1), Products (2, 3) and Inhibitor (4) used in this Study.
2.2 Results 2.2.1 Crystal Structure of the Enzyme-Substrate Complex
The crystal structure of a constrained substrate complex of HIV-1 PR was
determined using a crystal of catalytically inactive D25N HIV-1 PR complexed with
substrate 1. The crystal formed at 20˚C in a drop comprising 5µL of protein-substrate
mixture (0.1M acetate pH 5.5, 5mM DMSO), over a reservoir containing 0.1M
acetate buffer (pH 5.5) and 35% saturated ammonium sulfate as the precipitant.
Crystals appeared within one week and grew over 4 weeks to their final size before
collection six months later by Dr Shu-Hong Hu of the IMB. The crystals formed in
the space group P212121, with a single protein and substrate ligand in the asymmetric
unit cell. Data for the substrate complex was measured from a single crystal of
dimensions 0.4 x 0.1 x 0.1mm3 that diffracted to 1.6Å resolution using cryo- 30
31
crystallographic conditions as described in the experimental section and collected by
Dr Shu-Hong Hu. Diffraction data was integrated and scaled using DENZO
(Otwinowski 1993) and SCALEPACK (Minor 1993) and the phase was obtained by
difference Fourier methods utilising the previous room temperature substrate complex
(Pattenden et al., 1998). The crystallographic collection and refinement statistics for
both the E-S and E-P complex are presented in Table 2.2.
The substrate complex was found to have unit cell parameters a = 50.927 b =
58.054 c = 61.566Å and crystallographic R factors of 39.6 (Rfactor) and 38.8 (Rfree)
that refined to the final values of 17.3 (Rfactor) and 20.6 (Rfree). The final model of the
refined crystal structure of the substrate complex required several residues and
molecules to be modelled with alternate conformations (R14, K45, I47, R57, Q61,
V82, Q107, R114, E121, N137, M146, I150, V182, wat329, wat337, wat362 and
wat382).
The final structure was analysed using PROCHECK (Laskowski et al., 1993)
and WHATCHECK (Hooft et al., 1996) showing excellent refinement statistics. No
residues were in generously allowed or disallowed regions of the Ramachandran plot,
and only 7.7% were in additionally allowed regions and 92.3% were in most favoured
regions (Fig. 2.9). The stereochemical parameters of the plot statistics were all inside,
or better than the reference set of 118 structures all of which were at least 2Å
resolution.
The enzyme component of the substrate complex differed from inhibitor
complexes by only the single active site mutation (D25N), and was found to have all
the structural features described for HIV-1 PR in other complexes, such as the
pepstatin peptide inhibitor complex (Fig. 2.10, pdb accession code 5hvp, Fitzgerald et
al., 1990).
The substrate 1 was found to fill the normal substrate-binding cleft from S3-
S3’, with tyrosinyl side chains in S1 and S1’, the valine and isoleucine side chains in
S2 and S2’ respectively, and the aliphatic alkyl linkers occupying S3 and S3’ as
shown in Figure 2.11. The flaps were found to enclose the substrate 1 as reported for
inhibitor structures, and the conserved structural water molecule was also present,
forming the usual bridging interaction with the carbonyls of the substrate and the
amides of the flap residues I50/150, and superimposing well with the pepstatin
structure (Figure 2.10).
32
Figure 2.9: Ramachandran Plot of the E-S Complex for 1 Bound to HIV-1 PR (D25N). Regions
represented by various shades are: residues in most favoured regions [A,B,L], residues in additionally
allowed regions [a,b,l,p], residues in generously allowed regions [~a,~b,~l,~p]. Plot is based on an
analysis of 118 structures of resolution of at least 2.0Å and Rfactor no greater than 20%. A good quality
model would be expected to have over 90% in the most favoured region (Laskowski et al., 1993; Hooft
et al., 1996).
Figure 2.10 Superimposition of the E-S complex with Pepstatin Inhibitor Complex. The E-S complex (green) was superimposed onto the pepstatin-bound structure (purple) (ligands omitted) with an RMSD of 0.65Å. It can be seen the substrate complex possesses the overall morphological features of the pepstatin-bound HIV-1 PR structure 5hvp.
S2 S1’
S4 S3’ Figure 2.11: Connolly Surface of the Active Site of the E-S Complex is shown with the bicyclic hexapeptide ligand 1 captured in the substrate binding cleft. The standard labelling nomenclature of Schetcher and Berger (1967) is included.
S3 S1 S2’
The crystal structure shows that the substrate 1 makes a large number of
favourable contacts with the enzyme (Fig. 2.12), the majority of which are
hydrophobic (64% of all contacts). There are 12 hydrogen bonds (H-bonds) and 24
VdW interactions (VdW cut-offs of 3.4 – 4.2Å, with distances <3.4Å taken as being
unfavourable, H-bonding cut-offs 2.6 – 3.4Å). Of the H-bonding contacts, 4 are to
solvent water molecules, 2 of which are to the structurally conserved water. Of the 24
VdW contacts, only one is a bad contact (G127 O and 204 CD1 at a distance of
3.4Å), however G127 also makes a H-bond to the substrate and water and so the H-
bonding may reduce the expected repulsion (Figure 2.12).
The 2Fo-Fc electron density map (Figure 2.13) showed clear evidence for
substrate 1 binding in a single orientation at the active site of the enzyme and the
substrate was unambiguously modelled and refined into this electron density. In all
respects the previous difficulties assigning residues N25, N125, I50 and G51 were
resolved and the active site was modelled in a single orientation (Figure 2.14). The
active site was found to have a slight distortion from planarity of the replacement
residues N25 and N125, but with favourable H-bonding between N125 OD1
accepting a proton from N25 ND1 at a distance of 3.1Å and N125 ND1 donating a
proton directly to the substrate carbonyl at a short distance of 2.6Å. The 4.5Å
distance from the substrate carbonyl to N25 is far too great for any direct interaction
(Figure 2.14).
33
Figure 2.12: Hydrogen Bonding Pattern of the E-S Complex. Shown is the Hydrogen bonding from
HIV-1 PR (D25N) to the bicyclic substrate 1, including the water molecules within the range of the
hydrogen bonding cut-off.
Figure 2.13 Electron Density Map of the Substrate within HIV-1 PR. The 2Fo-Fc electron density
map about the substrate 1 in a single orientation at the active site (image generated in Setor V).
34
3.1Å 2.6Å
N25
N125
Figure 2.14: The Active Site of the E-S Complex for 1 Bound to HIV-1 PR (D25N). Shown is the
H-bonding of N125 from the substrate carbonyl (2.6Å) and the distance between the active site
analogue residues of 3.1Å. The distance N25 ND1 is 4.5Å from the substrate carbonyl.
N125 T126
T26
G27 amide
G127 amide
N25
Figure 2.15: H-Bonding Across the Active Site and Dimer Interface for 1 Bound to HIV-1 PR
(D25N). The side-chain to main-chain “fireman’s grip” of the threonine residues is shown, both
measuring a distance of 3.0Å. N125 has a similar interaction and H-bonds to the amide backbone of
G127 at a distance of 2.9Å. The analogous interaction N25 - G27 is disfavoured due to the presentation
of the N25 ND1 atom to the amide of G27, however, in the wild type enzyme that presents an oxygen
atom (D25), this interaction is favourable (distance 2.8Å).
35
36
2.2.2 Crystal Structure of the Enzyme-Product Complex
HIV-1 PR was also pre-equilibrated in the presence of the substrate 1 for 16
hours at 4ºC to allow cleavage of this substrate yielding the products 2 and 3 (Fig.
2.8). The solution was co-crystallised to yield the products-bound crystal structure.
The products-bound complex crystallised in the space group P212121, with a single
protein and two captured monocyclic ligands (2 and 3) in the asymmetric unit cell.
The crystal grew in 45% saturated ammonium sulfate over a 4-week period before
harvesting 6 months later. Data for the product bound structures were measured from
a single crystal of dimensions 0.3 x 0.1 x 0.1 mm3. Diffraction data was integrated
and scaled using DENZO (Otwinowski 1993) and SCALEPACK (Minor 1993) and
the phase was obtained by difference Fourier methods from the previously solved
room temperature product complex (Pattenden et al., 1998). Data was measured to
1.6Å resolution using cryo-crystallographic conditions as for the substrate complex.
The product complex was found to have unit cell parameters a = 51.868 b =
57.646 c = 61.533Å with initial crystallographic R factors of 30.8 (Rfactor) and 34.7
(Rfree) refining to 19.3 (Rfactor) and 22.6 (Rfree). The final model of the product
complex had residues modelled with half occupancies (I33, M36, A37 and L123), as
well as some residues modelled in alternate conformations (R14, E21, I84, R114,
E134 and M146). A complete second mode of binding was modelled for the C-
terminal product 3 and rigorously tested before inclusion in the final structure as a
second conformation (vide infra).
The final structure was analysed using PROCHECK (Laskowski et al., 1993)
and WHATCHECK (Hooft et al., 1996) showing excellent crystallographic collection
and refinement statistics (presented in Table 2.2). No residues were in generously or
disallowed regions and only 5.1% were in additionally allowed regions and 94.9%
were in most favoured regions of the Ramachandran plot below (Fig. 2.16). The
stereochemical parameters of the plot statistics were all inside, or better than the
reference set of 163 structures all of at least 2Å resolution. The only real variations
seen in the structure are for the C-terminal ligand 3, which had minor deviations from
“ideal” small molecule values, but are not greater than 0.1Å, and a single CB-CA-C
bond angle for residue 204 being >10.0˚ from the “ideal” 110.1˚ at 120.2˚. In all
respects the differences are minor.
37
290K
E-S complex
290K
E-P complex
160K
E-S complex
160K
E-P complex
Maximum resolution Å 50-1.9 50-2.0 100-1.6 50-1.6
no. observations 24 094 27 987 65431 70060
no. unique reflections 11252 11 473 22094 22703
no. rejected observations 331 486 62 82
Mosaicity ° (refined value) 0.350 0.400 0.487 0.299
completeness of data (%) 73.2 (12.2) 86 (8.3) 90 (52.3) 91.7 (64.1)
Calculated I/σI (%) 36.8 (20.7) 75 (38.9) 24.5 (4.8) 22.2 (5.9)
Rmerge (%) 14.7 (4.2) 26.6 (10.6) 3.1 (14.1) 3.4 (9.3)
Resolution range 8-1.9 8-2.5 8-1.6 8-1.6
no. reflections (F>0σ F) 12786 6077 1253 21876
Rfactor 19.0 23.3 17.4 19.3
Rfree 22.8 26.4 20.8 22.6
no. non-H atoms (all) 1748 1619 1853 1847
no. waters 110 70 191 232
no. sulfates 3 2 4 3
R.M.S. deviations from ideal
bond length 0.005 0.004 0.059 0.062
bond angle 1.197 1.063 1.538 1.890
dihedral angle 28.860 26.870 26.615 26.959
Improper angle 1.261 1.071 1.828 2.170
av. B-factor (+ solvent
molecules)
18.2 9.9 10.6 15.0
av. B-factors (from Wilson plot) 29.3 23.9 25.7 18.5
Mean co-ord error 0.21 0.31 0.162 – 0.194 0.201 – 0.229
Table 2.2: Diffraction & Refinement Statistics Comparing 290K and 160K Data.
Rmerge = Σ|I-<I>|IΣ<I>. Rfactor = Σ|Fobs-Fcalc|/ΣFobs and Rfree is mathematically equivalent to Rfactor but
measured over 10% of the data as defined by Brunger (1992). RMS = Root Mean Square. RMS
deviations from ideal calculated with ligand(s) omitted for the 290K data and with ligand(s) included
for the 160K data as the ligand(s) were found to contain the greatest deviations from ideal. Mean co-
ordinate error is reported as from Luzzati plot. (%) = % in highest resolution shell. The av. B-factors is
with ligands excluded with the value from the Wilson plot including the ligands.
Figure 2.16: Ramachandran Plot of the E-P Complex for (2+3) Bound to HIV-1 PR. Regions are:
residues in most favoured regions [A,B,L], residues in additionally allowed regions [a,b,l,p], residues
in generously allowed regions [~a,~b,~l,~p]. Plot is based on an analysis of 163 structures of resolution
of at least 2.0Å and Rfactor no greater than 20%. A good quality model would be expected to have over
90% in the most favoured region.
The product ligands (in the expected orientation of the C-product, 3) make 42
contacts to HIV-1 PR, 14 are H-bonding (4 to waters, 2 are to the structurally
conserved water). There are 4 H-bonds <2.6Å and 2 additional ligand-ligand contacts
<2.6Å, which may be low barrier H-bonds. There are 28 VdW contacts, 5 of which
are bad (all bad contacts are in the C-product, 3 and <3.4Å) and 2 of these bad
contacts are additional ligand-ligand VdW contacts. The majority of the interactions
between HIV-1 PR and both ligands are hydrophobic (66% of all contacts) and the N-
product (2) accounts for significantly fewer VdW contacts than the C-product (3), but
makes the same number of H-bonding interactions (Figure 2.17).
The product complex 2Fo-Fc electron density map shows clear evidence for
cleavage of 1 and separation of the N-terminal monocyclic product ligand (N-product,
2) and C-terminal monocyclic product ligand (C-product, 3) as shown in Figure 2.18.
The electron density for the product complex is of a high quality, similar to the
38
substrate complex, except in the region of the C-product (3) which was generally of
lower quality than the 2Fo-Fc electron density map for the N-product (2).
Figure 2.17: H-bonding Between HIV-1 PR and the Product Ligands. Shown is the Hydrogen
bonding from HIV-1 PR to the products 2 and 3, including the water molecules within the range of the
hydrogen bonding cut-off.
Figure 2.18: Electron Density map of Products 2 and 3 within HIV-1 PR. The 2Fo-Fc electron
density map about the products 2 and 3 is shown with the single expected C-cycle 3 orientation (image
generated in Setor V).
39
The placement of the C-product within the electron density maps was difficult
and the lower order of the maps was also reflected in a related disorder of the protein
residues interacting with the C-product (3), that is not seen in the N-product (2). This
suggests multiple binding conformations for the C-product (3), as was initially found
for the room temperature data, however stronger 2Fo-Fc-electron density exists for
the cryo-structure studied here, making it possible to model the opposite orientation
of this ligand resulting in improved R-factors. No interconversion of the residues
about I50/150 was detected, clearly defining the atom positions for this model.
A second conformation for the C-product (3) was rigorously tested by
refinement experimentations. The individual and separate conformations of the C-
product (3) were refined individually and compared to refinement cycles where both
conformations were included with 50% occupancy. From this experiment, the Rfactor
and Rfree decreased for the dual occupancy model with respect to either of the given
conformations singly, indicating a genuine second conformation of the C-product (3).
The occupancy was also adjusted, yielding best refinement for the expected
orientation of the cycle at 40% and the unexpected orientation at 60%, with the
second conformation of the C-product (3) binding in the reverse orientation to the
first as shown in figure 2.19.
Figure 2.19: The Two Conformations of the C-Product Ligand. The expected orientation (orange)
and the unexpected orientation (red) for 3 are shown.
Two alternate modes of binding of the C-product (3) consisting of residues
204-206 were therefore included in the final crystal structure; however, despite the 40
improvement of the refinement parameters by building in the second conformation
and the vast improvement over the room temperature model, there was still a
relatively poor fit of the two conformations of the C-product in the electron density
maps of the final model.
An unusual three-way carboxyl association was noted between the N-cycle of
the product complex and the two active site residues (D25, D125), as has been
reported previously (Rose et al., 1996). The product complex reveals additional
features of this carboxyl triad, being the interactions with the amine of the cleaved
scissile bond (C-cycle). The distances of the atoms in the carboxylate triad are 2.6Å
(N-cycle - D125), 2.9Å (D125 - D25) and 2.7Å (D25 - N-cycle). None of the
carboxylates that make up the triad are co-planar, but each are rotated slightly to
maximise H-bonding. The amine of the C-cycle has the ability to make three contacts
to the carboxylate triad at distances of 2.6Å (D25), 2.4Å (203 OT) and 2.5Å (203 O)
(Figure 2.20).
D125
D25
2.6Å
2.9Å
2.6Å
204 ND2
Figure 2.20: The Carboxylate triad of the E-P Complex. View of the carboxylate triad (large
hashes) from outside the S4 pocket, also showing the close proximity of the C-product ligand to the
interaction. The carboxylate triad measures 2.6Å from both aspartates to the respective product oxygen
atoms with the longer distance between the aspartates at 2.9Å. The distance to the C-product 204 ND2
atom (small dots) is 2.6Å and the distances between the two product atoms are very close at 2.4Å and
2.5Å.
41
2.3 Discussion 2.3.1 Insight from Enzyme Kinetics
Researchers in the Fairlie group have previously determined the
affinity of HIV-1 PR for bicyclic substrate 1, monocyclic products 2 and 3, and an
acyclic analogue 5 that is mimicked by the bicycle 1 (Reid and Fairlie 1997; Fairlie et
al., 2000).
2 3
5
1
NH
HN
O
O
O HN
NH
O
O
O
NH
O NH
HN
O
O
O
HN
NH
O
O
O
H3N
O
O
NH
HN
O
O HN
NHO
O
NH
OOH
O
AcHN
1
Figure 2.21: Bicyclic Substrate, Product and Linear Substrate “Inhibitors”.
To determine the specific affinities of HIV-1 PR for the various ligands,
competition experiments were conducted with the fluorogenic substrate 2-Abz-Thr-
Ile-Nle-Phe(pNO2)-Gln-Arg-NH2 (Abz-NF*-6). This substrate contains an ortho-
aminobenzoic acid fluorophore that is intramolecularly quenched by a para-
nitrophenylalanine, until these groups are separated by substrate hydrolysis, which
liberates the fluorophore and emanates measurable fluorescence. Abz-NF*-6 was
processed in the presence of 1, 2, 3 or 5, which compete for the active site, following
the reaction profile:
k1 k2
E + Abz-NF*-6 E-(Abz-NF*-6) E + 2-Abz-Thr-Ile-Nle + Phe(pNO2)-Gln-Arg-NH2
Ki 1, 2, 3 or 5 EI
42
Thus 1, 2, 3 and 5 were used as competitive inhibitors and inhibitory potency
can be measured by alterations to the rate of formation of the fluorescing product of
Abz-NF*-6. The protease affinities for 1, 2, 3 and 5 could therefore be determined by
measuring Ki:
E + I EI Ki (k-1/k1)k1
k-1 Using this relationship, the relative affinity in terms of Ki is 0.8µM (1), 8µM
(2), 15µM (3), and 60µM (5) (Reid and Fairlie 1997; Fairlie et al., 2000). Since Ki
for a competitive inhibitor is simply a measure of its affinity for enzyme (Ki =
[EI]/[E][I]), one can conclude that the bicyclic substrate (1) has 75-fold higher
affinity for HIV-1 PR than the linear substrate (5), corresponding to ~11kJ mol-1
difference in binding energy (∆G = -RT lnK). The cycles compare with a Ki of 60µM
for an acyclic analogue AcLVF-FIL-NH2 (5), a relatively similar substrate that can
make more hydrogen bonding contacts with HIV-1 PR through its terminal amide
group at each end. Thus the measured 75 fold increase in affinity for the protease of
the cyclic analogue 1 over the linear substrate 5 is only a lower limit because 5 makes
several extra H-bond contacts between its additional termini and the enzyme. An
argument can be made that there is a distinct energetic advantage (∆G ≥ 11 kJ mol-1)
in constraining or pre-organizing the cyclic substrate in an extended conformation, an
advantage provided entirely from entropy differences between cyclic and acyclic
conformations. Although enhancing the affinity of a substrate for an enzyme is not
necessarily an advantage for catalysis, since products must dissociate readily from the
active site to enable rapid turnovers. However, for the purposes here of retaining
substrate and products in the active site, the above kinetic data do support the
significant advantages of conformational pre-organization for enzyme affinity.
In the absence of the competing fluorogenic substrate Abz-NF*-6, 1 and 5
cannot be monitored as competitive inhibitors, but their activity as substrates can be
observed. The substrate dissociation constant (Kd) is identical to the measured Ki
values described above in the following scheme:
43
E + S E-S E + P
Kd = k-1/k1 = Ki, assuming k2 << k-1
k1
k-1
k2
Thus, the bicyclic substrate (1) as well as the cyclic products of peptide
hydrolysis (2,3), are held more tightly by the enzyme than the acyclic substrate 5.
This suggested to us that we may be able to trap the cyclic substrate, transition state
intermediate and/or cyclic products in the enzyme crystal lattice following substrate
binding, but prior to product dissociation from HIV-1 PR.
[E-S E-T E-P] Products
Furthermore, the conformationally constrained cyclic substrate 1 was found to
be a poorer substrate (Km = 110µM) than acyclic 5 (Km = 20µM) (Reid and Fairlie
1997; Fairlie et al., 2000), as defined by the Michaelis-Menten constant (k2+k-1/k1).
Yet 1 is more constrained to an extended conformation and the enzyme has a higher
affinity for the substrate 1 than for 5 as judged by its Ki. This can be rationalised if
the Enzyme-Substrate (E-S) complex decomposes (k-1 + k2) more slowly for 1 than 5.
This is not a surprising result since a “good” substrate should not bind so tightly to an
enzyme as to impede turnover but rather there needs to be a compromise between
enzyme affinity and product formation and dissociation. Unfortunately, kcat (rate of
formation of E + P from E-S) was too difficult to obtain in water due to the poor
solubility of the ligands and slow turnover for such short peptides.
It is fair to conclude that due to the higher affinity of cyclic substrate/products
than acyclic substrate/products, that there were good prospects for finding the E-S, E-
T or E-P in the crystal structure following co-crystallisation of HIV-1 PR with 1, and
likely better prospects than following co-crystallisation with 5. Indeed we were
unable to crystallise 5 with enzyme, possibly due to hydrolysis and product
dissociation (disfavoured by kcat). On the other hand crystals were obtained using the
higher affinity substrate 1 with HIV-1 PR, even though products 2 and 3 were
actually seen within the active site of the catalytic enzyme from these crystals. Thus
the bicyclic substrate 1 is still a sufficiently good substrate to enable product
formation.
The trapping of both of the separated product halves 2 and 3 was believed to
have occurred because of two factors:
1. The bicyclic substrate 1 is highly constrained in the extended conformation
that is pre-organized for higher affinity binding to the protease than normal 44
acyclic substrates. Once the amide bond is cleaved, the monocyclic
components still make all the same interactions with enzyme as the bicycle
and amount to higher affinity binding than acyclic products.
2. An excess of ligand (1) to protease is used in the pre-equilibration and high
salt solution, driving the formation of products by Le Chatelier’s principle and
favouring retention of hydrophobic ligands in the active site of HIV-1 PR.
The failure in our hands of the linear substrate 5 to crystallise with the enzyme
is consistent with (a) and (b), the acyclic substrate being more likely to turnover, and
the lower affinity acyclic products more prone to dissociation, as described above.
Because of these factors, the crystals that subsequently formed would have products
of 1 bound at the active site of HIV-1 PR. If the enzyme had not cleaved the
substrate under pre-equilibration and crystallisation conditions, then electron density
at the active site would show evidence of intact substrate 1 bound to the enzyme.
However the electron density from the crystal grown in this way shows clear
evidence for products 2 and 3 bound at the active site, with defined geometry of
separation and products 2 and 3 clearly interact independently with HIV-1PR.
2.3.2 Comparison of Substrate, Product and Analogue
Three high-resolution crystal structures are now available from this series.
Substrate 1 (Results 2.2.1), products 2 + 3 (Results 2.2.2), and a previously solved
transition state analogue inhibitor 4 (Fig. 2.22) (Martin et al., 1999) have now all
been crystallised with HIV-1 PR and structures determined by the Fairlie and Martin
groups (IMB, UQ). These 3 structures can potentially provide a unique insight to
progressive structural changes that occur during catalysis by this viral enzyme.
Figure 2.22: Inhibitor Analogue of the Substrate. The inhibitor differs from a substrate by
replacement of the scissile amide bond with a hydroxylethylamine isostere linkage.
4
NH
HN
O
O
O HN
NH
O
O
O
NHOH
45
The following comparison of these three structures is focused on identifying
features that might shed light on the catalytic mechanism and structural events
through catalysis. The specific regions of interest are the flaps (including conserved
water and ligand carbonyls), the relative orientations and locations of each of the
ligands, and interactions involving the active site and scissile amide bond.
(a) Comparison of Ligand Binding Mode
The binding mode of the ligands of the E-S, E-P and E-I complex was
compared by superimposing all the protein Cα atoms (396 non-hydrogen atoms), so
the ligands align unbiased in the substrate-binding cleft. The protein chains of the E-S
and E-P complexes superimpose very well (RMS deviation 0.23Å), and the N-
terminal macrocycles superimpose almost identically within the active site, with some
variation in the linker region of 201 and the tyrosinyl group of 203 (Fig. 2.23),
suggesting the N-terminal macrocycle is essentially anchored in an almost identical
manner in the active site throughout catalysis.
204 202
206
201
203 205
Figure 2.23: Superimposition of E-S and E-P Complex. The E-S complex (yellow) was unmoved
and the E-P complex (red) superimposed onto the structure with a RMS deviation of 0.23Å. Shown is
the crystallographic labelling of residues 201 – 206.
46
The C-terminal macrocycles do not superimpose quite as well as the
analogous N- terminal macrocycles with the E-P macrocycle pivoted about the
isoleucinyl/tyrosinyl centre with respect to the corresponding substrate macrocycle
and occupying different positions within the enzyme active site. A superimposition of
34 non-hydrogen atoms (excluding the 206 linker region, isoleucinyl side chain and
different atoms about the scissile amine and product-amide), show the primary
elements of the C-terminal macrocycles have a similar orientation and shape (RMS
deviation 0.34Å over 34 non-hydrogen atoms), however the C-terminal macrocycle
has clearly moved following cleavage of the scissile amide bond and is not as
strongly maintained as the N-terminal macrocycle.
The E-S and E-I complexes also superimpose very well (RMS deviation
0.27Å over 396 Cα atoms) (Fig 2.24). In contrast to the comparison of the E-S and E-
P complexes above, the E-S and E-I ligands align almost identically at the C-terminal
macrocycles with a slight pivoting of the inhibitor about the isoleucinyl/tyrosinyl
centre of the macrocycle and differences in the linker region 206 and the isoleucine
side chain as minor deviations. The E-S and E-I N-terminal macrocycles do not
occupy the same position within the active site of the enzyme but are translated
towards the S1 – S3 pockets. A superimposition over 34 non-hydrogen atoms
(excluding the 201 linker region, valinyl group and different atoms about the scissile
peptide and product-carboxyl, RMS deviation of 0.28Å) show that the N-terminal
macrocycles adopt a similar orientation and shape, but occupy a different position in
the active site.
202 204
206
201
205 203
Figure 2.24: Superimposition of E-S and E-I Analogue Complexes. Superimposition of the E-S
complex (yellow) of and the E-I complex (green) of 4 with a RMS deviation of 0.27Å over 396 Cα
atoms. The inhibitor ligand is identical to the substrate used except for the replacement of the scissile
amide bond with a hydroxyethylamine isosteric linkage.
47
A superimposition of all three structures (Fig 2.25) highlights the difference of
all the macrocyclic ligands. The regions between the macrocycle 203Cα – 204Cα
atoms (comprising the scissile peptide, hydroxyethylamine and free carboxyl/amide
groups) represent the greatest differences between the ligands. There is significant
differences in the separation distances between the macrocycles from the 203Cα –
204Cα atoms being 3.8Å in the E-S complex, 4.8Å in the E-P complex and 4.9Å in
the E-I complex. The carboxyl/hydroxyl atoms between the macrocycles (and
attached to only the N-terminal macrocycle of the E-P complex) all occupy different
positions (0.8Å difference from the E-S to E-P, 1.0Å from the E-S to E-I and 1.6Å
from the E-P to E-I oxygen atoms).
204202
206
201
205
203
Figure 2.25: Superimposition of the E-S, E-P and E-I Analogue Complexes. The difference
between ligand locations in the active site E-S complex (yellow), E-I complex (green) and E-P
complex (red) is shown. The superimposition was conducted over 594 Cα atoms with RMS deviations
are reported in table 2.4. For the inhibitor, atoms comprising the N-terminal cycle linker 201 were
omitted in the model due to refinement difficulties (reported by Martin et al., 1999).
48
It is not surprising that there is a difference about the central linker region of
the macrocycles as the ligands are very different both physically and
stereoelectronically in this region. In the first instance the E-P complex macrocycles
differ dramatically from the substrate with the changes in hybridisation and charge
brought about by the cleavage of the scissile amide bond. The inhibitor isostere is
lengthened between the cycles with respect to the substrate by a methylene insertion
to form the hydroxyethylamine peptide analogue (-CO-NH- → -CHOH-CH2-⊕NH2-),
49
and has Sp3 hybridisation of the isostere carbon atom that is functionalised with the
hydroxyl group. The isostere is also believed to be Sp3 hybridised at the nitrogen
atom and in a protonated state within the active site (Abbenante et al., 1995). The
differences in length and hybridisation are likely the reason the isostere is displaced
towards S1-S3 with respect to the E-S and E-P complexes as seen in the
superimpositions (Fig 2.24 and 2.25), although the less restrained isostere provides
the inhibitor with greater flexibility compared to the substrate. This flexibility allows
a “puckering” of the inhibitor conformation about the isostere linkage from S1 to S1’
compared to the substrate and allows the inhibitor to maintain a very strong
resemblance to the substrate at the C-terminal region that binds from S1’ – S3’.
Superimpose N-cycles C-cycles Whole Protein
substrate – product 0.18Å 0.34Å 0.23Å
Substrate – inhibitor 0.28Å 0.16Å 0.27Å
Inhibitor – product 0.28Å 0.27Å 0.23Å
Table 2.3: Comparison of HIV-1 PR Structures and Ligands. RMS deviations of superimposing
396 atoms (whole protein Cα trace) and 34 atoms (macrocycle halves excluding the side chains,
different scissile regions and 201/206 linkers).
ligand region/atom
Specific HIV-1PR contact
Distance inhibitor
(Å)
Distance substrate
(Å)
Distance product
(Å)
Maximum difference
(Å) P3 201 N G48 O 3.1 2.9 2.8 0.3
P2 203 O D29 ND2 3.0 2.9 3.0 0.1
P2 203 O Str WAT 2.7 2.7 2.7 0
Str WAT I150 N --- 2.9 3.0 0.1
P1 203 N G27 O 2.9 2.8 2.8 0.1
P1 203OT D25/N25 N/A N/A 2.6 ---
P1 203 O D125/N125 2.6 2.6 2.6 0
P1' 204 N G127 O 3.0 3.0 3.0 0
P1' 204 O Str WAT --- 2.7 2.8 0.1
P2' 205 N G127 O 3.1 3.0 3.0 0.1
Str WAT I50 N --- 2.7 3.0 0.3
P2' 205 O D129 N 3.1 2.9 3.0 0.2
P3' 206 N G148 O 2.8 3.0 2.7 0.3
Table 2.4: Comparison of HIV-1PR H-bond Contacts to the Ligands (P3, S3 - P3', S3'). N/A is
labelled for residues not shared between HIV-1PR and the HIV-1PR D25/125N mutant enzyme and
the dashed lines indicate there is no analogous contact where the structural water molecule is
displaced. Str WAT is the structural water co-ordinated between the flaps and ligand(s).
50
With such dramatic changes about the region between the macrocyclic
ligands, it is surprising that the macrocycles overlay so closely (Table 2.3). The
hydrogen bonding between the substrate binding pockets (S3 – S3’) and the
corresponding specific contacts to the ligands (P3 – P3’) and flap/structural water
contacts were compared to observe the differences between the different ligands and
the enzyme (Table 2.4). From Table 2.4 it can be seen that despite the different
positions of the ligand atoms, especially about the scissile region between the
macrocycles, the H-bonding distances remain very similar in all three complexes,
with the major differences at the peripheries of the macrocycles, where the distances
from the 201 and 206 amide nitrogen atoms to G48/G148 carbonyl oxygen atoms,
vary by 0.3Å.
Taken together, the protein superimpositions, ligand superimpositions and
protein-ligand bonding reveal that each of the constrained macrocycles of the
substrate, product and inhibitor orders the local protein environment from S1 – S3
and S1’ – S3’ with very few and very minute observable differences in H-bonding
distance despite the differences of the scissile region between the macrocycles and
separation within the active site. The Ki of 8µM for the N-terminal product 2 and
15µM for the C-terminal product 3 (Reid and Fairlie 1997; Fairlie et al., 2000),
suggests that the C-terminal product is held more loosely by the enzyme than the N-
terminal product. This is consistent with the finding here of a second binding mode
for the C-terminal product and excellent superimposition of the N-terminal
macrocycles, suggesting that the C-terminal product has either reorganised in the
active site or left the active-site and re-entered in the opposite orientation that has
been captured by the enzyme. This could be a consequence of “salting-in” of the
ligand due to the hydrophilic nature of the crystallisation solution (Meek et al., 1994).
Other possibilities may also account for the second mode of binding of the C-terminal
product 3 such as avoiding the thermodynamically expensive reversal of catalysis
(reformation of a substrate). It is also possible that HIV-1 PR maximises the
favourable interactions of the C-terminal ammonium ion and solvents at the opening
of the substrate-binding cleft, and so accepts the constrained cycle in the reverse
orientation of what is typically expected. The reverse binding of the C-terminal cycle
3 is considered a kinetic artefact of the method of equilibration in a non-ideal system
and all further discussions focus on the expected orientation.
51
(b) Flaps, Conserved Water and Ligand Carbonyls
The flaps contain the residues I50 and G51 (and the symmetry related I150
and G151), that co-ordinate directly to the carbonyl oxygens of the substrate adjacent
to the scissile amide bond. This interaction occurs through a conserved water
molecule, and hence the flaps are believed to assist in catalysis by stretching the
scissile bond through this interaction (March and Fairlie, 1996). The superimposition
of substrate, inhibitor and products shows that the flap regions, including the bound
water, vary amongst the three structures, but the differences are once again subtle.
The water molecule moves by 0.4Å in going from the enzyme-substrate
complex to the enzyme-product complex. The distance from the water molecule to
I150 N in both the inhibitor and substrate complex does not vary (2.9Å) but is
different in the product complex (3.0Å). The analogous contact in the other flap (I50
N) does vary in all three structures significantly, with a maximum difference of 0.2Å
(substrate - inhibitor) and the inhibitor complex exhibiting the longest bond distance
(3.1Å).
The water molecule is also closer to the substrate and product carbonyls than
the inhibitor carbonyls by a maximum difference of 0.3Å (N-cycle substrate 1) and
0.2Å (C-cycle substrate 2). The relative positions of the carbonyls that interact with
the conserved water molecule are more significantly displaced in all three structures
by a maximum distance of 0.6Å (N-terminal carbonyls, substrate – inhibitor complex)
and 0.8Å (C-terminal carbonyls, substrate – product complex). The N-cycle is closer
to the water than the analogous C-cycle for both substrate and product complexes by
a very insignificant amount (slightly more than 0.1Å), but the inhibitor complex has
the C-cycle closer than either the substrate or product complex.
The overall distance from the flap nitrogen atoms to their respective ligand
carbonyls for N-cycle contacts are in the following order: product (4.0Å), inhibitor
(4.1Å) and substrate (4.3Å), whereas for C-cycle the ordering is different: substrate
(4.8Å), product (4.9Å) and inhibitor (5.3Å). The same trend is reflected in the other
flap residue G48/G148 that makes a contact through their carbonyls to the amides of
the ligand/s: the ordering for the N-cycle is product (2.8Å), substrate (2.8Å), inhibitor
(3.1Å) and for the C-cycle is product (2.6Å), inhibitor (2.8Å) and substrate (3.0Å).
The greatest difference about this region is the G149 CA atoms that are separated by
1.1Å (substrate - product). Since G149 is between the H-bond donor I150 and the H-
52
bond acceptor G148, it is not surprising there is a large shift in residue G149 between
the structures. Large shifts here would allow for the satisfaction of H-bond criteria to
the ligands between G148 and I150, as it is noteworthy that the flap H-bond donors
and acceptors themselves essentially do not move.
Taken together, the interactions about the flap regions with the conserved
water and ligand carbonyls, as well as the related H-bonding from G48/148 to ligand
amides shows HIV-1 PR alters interactions and structure about the flap region and
through the conserved water to satisfy all H-bonding interactions. The difference of
interaction between a substrate, product and inhibitor around the region of the flaps
and including the conserved water molecule and ligand carbonyls show that this
region is important in the intramolecular communication with the ligands to form an
ordered complex. When all interactions are taken into account in this region, there is
no evidence that the flaps close more tightly on this inhibitor compared to a substrate
ligand. This was unexpected as the literature suggests the flaps close tighter on
inhibitor complexes and tighter binding has been correlated with increased affinity
(Wlodawer and Erickson 1993; Martin et al., 1999; Reiling et al., 2002).
2.3.3 The Mechanism of Catalysis of HIV-1 PR
Having structural data for substrate and products in hand, together with a
previous structure of a related transition state analogue inhibitor 4, it is possible to
analyse individual steps along the catalytic pathway and to apply the results to the
different proposed mechanisms of hydrolysis of a peptide substrate. Since both the E-
S and E-P complexes do not form covalent bonds to active site residues, mechanism 1
(Figure 2.6a) is not consistent with our observations.
(a) Active Site Orientation.
From the literature there are essentially four possible orientations of the active
site as shown in Figure 2.26 below. The first orientation has both active site aspartates
deprotonated as Lewis bases, requiring a basic environment above pH 6. The second
is a shared orientation with a single proton between them in a concerted mechanism
of catalysis, the third is an uneven acid-base state of the active site and the fourth is a
di-protonated state under very acidic conditions. In general the enzyme is active over
the pH range of 2.5 – 6.5 (Smith et al., 1996).
pH >6 3 - 6 3 - 6 <3
Orientation 1 2 3 4
Figure 2.26: Possible Active Site Charged States based on NMR titration (Smith et al., 1996) and
solvent isotope experiments (Hyland et al., 1991a; Hyland et al., 1991b).
From the E-S complex crystallised at pH 5.5, the asparagine active site has
adopted a single, unequivocal orientation due to the asymmetry created by the
substrate binding. This orientation does not appear to be inherent to the asparagines,
as they could have easily adopted the opposite orientation in the absence of the
substrate. Indeed, in the room temperature structure, the aspartates formed alternate
conformations in the presence of substrates but forced the substrate carbonyl to flip in
order to satisfy the H-bonding. In this respect the cryostatic conditions have locked in
the lowest energy conformation of the E-S complex. The orientation of the
compromised active site proves the initial step in catalysis occurs with residue 125
being closest to the substrate carbonyl oxygen. The atom N125 ND2 is 2.6Å from the
substrate carbonyl and may be a similar distance in the wild-type enzyme
immediately preceding catalysis. The analogous distance from the other protein half
(residue N25 ND1) is far too great to interact directly with the carbonyl oxygen of the
substrate, being 4.5Å away as shown in Figure 2.14.
This orientation suggests that D125 plays an early role in catalysis, either
retaining or taking the shared proton away from D25 (orientation 2, Fig 2.26) and
interacting with the substrate carbonyl. If the proton is shared (orientation 2), this
would occur as the catalytic water molecule is pushed close to the active site by the
substrate docking at the pKa of 5.5 as shown in Figure 2.27.
53
1 2 O O
O O
H
O O
O O
H
O O
O O H
O O
O O
H HO
HO O
O O
H HO
H
D125
D25
D125
D25
D125
D25
D125
D25
D125
D25
O O
O
HO
H
D125
D25
OHROTATION ROTATION
Figure 2.27: Ionisation State of the Active Site showing two possible ways a shared proton between
the aspartates could assume the orientation seen at the active site of the E-S complex. 1 shows a simple
equilibrium that would allow D25 to formally hold a negative charge and 2 shows how the incoming
catalytic water molecule could elicit the change by encroaching on D25.
The orientation of the mutated active-site therefore offers support for D125
acting as a Lewis acid (orientations 2 and 3, Fig 2.26) in initial proton exchanges of
catalysis (Chatfield and Brooks 1995), and abstracting the shared proton from
between the catalytic residues as an early event of catalysis.
(b) The Catalytic Step.
Once the substrate is bound or encroaches enantiospecifically on the active
site, the catalytic water nucleophile attacks at the Re face of the amide carbonyl. The
trajectory path for this attack is very precise since only the Re face of the scissile
amide is accessible to a water molecule. In this respect the position of the water
molecule would essentially be the same position as the product hydroxyl. Modelling
the position of the product hydroxyl oxygen atom in question as the oxygen atom of
the catalytic water molecule in the E-S complex, and measuring the O-C-O angle
from the substrate carbonyl to this modelled water reveals the angle of hypothetical
attack as 107˚. This is precisely the ideal angle expected for attack by an SN2 class
nucleophile to form a tetrahedral transition state (Burgi et al., 1973) (Fig. 2.28).
54
Furthermore, modelling a hypothetical water molecule in this position in the
E-S complex shows this water would be within H-bonding distance only to D25
(2.5Å at 108° (D25 C-O…O water) and 3.0Å from OD2) and not D125 (4.2Å). This
makes it impossible for the water molecule to be activated from between the catalytic
residues as has been suggested (Laco et al., 1997). Rather this indicates that the
catalytic water is held by the enzyme atom D25 OD1, and H-bonds to 204 N (the
scissile nitrogen) during nucleophilic attack. Although a H-bonding distance of 3.1Å
from G27 and an angle (G27) C=O…O (Water) of 92º indicates a possible
involvement of this residue. If this scenario is correct it implies a water molecule
could be the nucleophile and is not necessarily an activated hydroxyl ion. This also
suggests that the transition-state contains a protonated nitrogen as shown in Figure
2.29.
N125
Figure 2.28: Model of the Active Site at the “Catalytic Event”. The E-S complex modelled with the
hydroxyl of the E-P complex as the correct position of the catalytic water H-bonded to D25 and the
possible association with G27 in a very small pocket and optimally orientated for nucleophillic attack
of the scissile amide bond at the Re face. The distance to D125 is too great at 4.2Å for interaction and
so activation between the aspartates is excluded.
NH
HN
O
O
O HN
NH
O
O
O
N
O
OH
H
NH
HN
O
O
O HN
NH
O
O
O
NH
HO O
H H
Figure 2.29: Alternate Mode of Nucleophilic Attack. If the catalytic water molecule only binds to
D25, it is possible the water molecule acts as a nucleophile with concomitant attack by the scissile
amide at 107° to form a zwitterionic intermediate.
55
56
(c) Stretching of the Amide Bond
An analysis of the E-S complex suggests that the hypothetical E-S* complex
does not exist. As the E-S complex moves towards the short lived, higher energy
hypothetical E-S* state, it is believed the substrate would encroach closer on the
active site. This would further favour nucleophilic attack and strain would be
introduced to the peptide bond, possibly through the flap residues, but could also
occur across the dimer interface (from residues G27/G127) and leading into the 30s
loop (D29/D129).
The previously shared proton held by D125, H-bonds to the scissile carbonyl,
which would help present the substrate carbon atom to nucleophilic attack. However,
no strain is discernible in the E-S complex, suggesting the E-S state and not the E-S*
state has been captured; yet all atoms are perfectly placed in terms of distances and
geometry for catalysis. Indeed the distance to N125 (the analogue of D125) is a very
small (2.6Å) and bordering on a definition as a low-barrier H-bond.
Alternatively, it is hypothesised that the flap residues, co-ordinating through a
water molecule to the substrate carbonyls either side of the scissile amide bond, may
assist in presenting the scissile carbonyl or distorting the scissile amide bond to help
facilitate catalysis. Thus the ES* state would hypothetically be achieved with precise
strain applied to the substrate molecule across the enzyme flaps and through D125.
However, there is essentially no evidence of flap movement between the E-S, E-P or
higher energy transition state analogue - E-I complex where tighter closure of the
flaps is expected and has been assumed. This suggests that there is no rate-
determining step between docking of the substrate and formation of the transition-
state, making substrate binding the rate-determining step to early catalytic events
leading to the transition-state. Furthermore, the role of the flaps can be defined as
assisting in forming the substrate binding pocket by communicating with the ligand(s)
so the local protein environment is ordered and perhaps excluding solvent from the
active site during catalysis, but there is no evidence for participation either directly or
indirectly to the catalytic step itself with the bicyclic compounds, with the geometry
of all H-bonds favouring catalysis without the need for strain of the nucleophile or
substrate.
(d) Mechanism
Based on the above information, modifications to the mechanism of catalysis
can be suggested for consideration (Fig. 2.30).
(1) D125 is in a protonated state (possibly due to the proximity of the substrate and water molecule) and the catalytic water is held close to D25 so that nucleophilic attack occurs at 107°. The stability of the N-terminal portion of the substrate would be critical for efficient electron transfer.
(2) Acid-base catalysis occurs forming the neutral gem-diol complex and rapidly moving toward products.
(3) The N-terminal product takes another proton from D125 to form an NH3
+ state. (4) The final state of the active site just
prior to release of products. For the bicyclic substrate, evidence suggests the C-terminal product leaves, creating a strong negative charge on D125 and facilitating a rotation of D25 to regenerate the active site and lead to the N-terminal cycle leaving.
(1) D125 is again in a protonated state as above and the catalytic water is held close to D25 so that nucleophilic attack occurs at 107° by the water molecule as the nucleophile.
(2) Acid-base catalysis occurs forming a charged proton species that shifts to the scissile amide and rapidly moving towards the transition-state. D125 “shares” a low-barrier proton with the substrate.
(3) The zwitterionic transition-state is formed and decays through reversal of acid-base catalysis. The C-terminal product takes the low-barrier proton from D125 to form an NH3
+ state. (4) The final state of the active site
just prior to release of products with regeneration as above.
Figure 2.30: Modified Mechanism of Cleavage of HIV-1 PR. The upper figure is mechanism proposal 1 and lower is mechanism proposal 2.
Mechanism 1
Mechanism 2
O
O
O
O
NH
O
H
HO
H O
OH
O
O
NH
O OP1'P1' H
H
O
O
O
O
N
O P1'
H
O
H
H
O
O
O
O
N
O P1'O
H
H
O
H
H H
G127
D125
D25
D125
D25
P1P1
D125
D25
P1
D125
D25
P1
3 4
1 2
O
O
O
O
NH
O
H
HO
H O
O
O
O
NH
O OP1'
P1'
H
O
O
O
O
N
O P1'O
H
O
H
H H
G127
H
H
O
O
O
O
NH
O OP1'H
H
H
O
G127
D125
D25
D125
D25
P1P1
D125
D25
P1
D125
D25
P1
1 2
57
The evidence from the active site suggests D125 is protonated and D25 binds
to the catalytic water molecule and possibly is also held by G127 O. From this point
the water molecule could be activated as a hydroxyl ion for nucleophilic attack
(mechanism proposal 1 Fig. 2.30), or may interact with the substrate through its lone
pair electrons, forming the gem-diol intermediate (mechanism proposal 2 Fig. 2.30). A neutral intermediate is favoured by a hydroxyl nucleophile and zwitterionic
intermediate by a water molecule due to the inherent lack of basicity of the scissile
nitrogen, though neither are mutually exclusive and a zwitterionic intermediate could
form from the hydroxyl nucleophile (not shown). In both cases and due to the angle
restraints and distances of the E-P complex, the final state in catalysis shows the C-
terminal cycle is likely a charged ammonium species within the active site.
The proposed mechanisms are in general agreement with other E-S and E-P
based mechanisms proposed from similar complexes (Rose et al., 1996; Laco et al.,
1997). In both cases, the N-terminal product leaves in an NH3+ state. The final state of
the active site just prior to product release has the unusual carboxylate triad
positioned with the N-cycle and two catalytic residues D25/D125 all within H-
bonding distances. For the bicyclic substrate, evidence suggests that the C-product
leaves first, but this is most likely due to the stronger binding of the N-cycle. In this
respect, the regeneration of the active site and ejection of the N-terminal cycle is
achieved by merely rotating D25 to interact with D125 and creating a carboxylate
repulsion and favouring the charged products to move into the solvent pool.
2.4 Experimental Section
2.4.1 Origin of Protein and Substrate
Aside from the D25N replacement, four typical mutations were introduced
into the synthetic monomers of both proteins (C67B, C95B, Q7K and L33I - where
B= L-α-amino-n-butyric acid). The cysteine modifications involve replacing the
reactive thiol group with a methyl group that avoids oxidation and polymerisation
whereas the Q7K and L33I mutations remove two autolysis sites from the proteins
which increases the stability of the enzyme (Bergman et al 1995; Rose et al 1993;
Mildner et al 1994).
Both the D25N and wild type HIV-1 PR (SF2 isolate - Wlodawer et al 1989)
were synthesised and purified by Dianne Alewood in the Peptide Synthesis laboratory 58
59
of Paul Alewood (Institute for Molecular Bioscience University of Queensland) using
modified Boc chemistry on an Applied Biosystems 430A Peptide Synthesiser
following protocols they have developed and described (Kent et al., 1992). These
proteins were purified by reversed-phase HPLC using a C4 Vydac column (Ø4.6mm
x 150, pore size 300 Å) and analysed on a PE-SCIEX API III triple quadrupole mass
spectrometer.
The cyclic substrate was synthesised by Dr Robert Reid in the laboratory of
Professor David Fairlie (Institute for Molecular Bioscience University of Queensland)
using procedures similar to cyclic inhibitors (Reid and Fairlie, 1997), except the N-
terminal reaction utilised Br(CH2)4COCl rather than Br(CH2)5COCl. The N- and C-
terminal cycles were coupled in the presence of benzotriazol-1-yl-oxy-tris-
dimethylamin phosphonium hexaflurophosphate and purified by RP-HPLC and
confirmed using mass spectrometry.
Dr Doug Bergman (Institute for Molecular Bioscience University of
Queensland) assessed the D25N mutant for activity and the substrate for cleavage.
Briefly, HIV-1 PR, purified from RP-HPLC fractions above, was solubilized in 6M
Gu.HCl (0.05 mg/mL), then refolded for 60min in buffer A (20mM phosphate pH 7.0,
20% v/v glycerol, 10mg/mL BSA). Time course samples of the bicyclic substrate 1 vs
HIV-1 PR were analysed by RP-HPLC and mass spectrometry to confirm recognition
of the substrate and cleavage between the cycles at the desired peptide bond. The
individual macrocyclic compound 1 was confirmed to be a genuine substrate,
releasing the cycle halves 2 and 3 (Fairlie et al., 2000).
The compounds 1 - 4 were reported as inhibitors of HIV-1 PR, competing
with the fluorogenic substrate Abz-NF*-6, for the active site (Fairlie et al., 2000).
Using protocols described previously (Toth and Marshall 1990; Bergman et al.,
1995). The buffer A/protease solution (10µL) was added to 50µM of the fluorogenic
substrate buffer B (MES pH 6.5, 37C, 100mM NaCl, 10% v/v glycerol) and varying
concentrations of the cyclic substrate. The Ki was then calculated from Dixon plots of
1/v vs [s]. Their relative potencies as competitive inhibitors are Ki : 3nM (4) > 0.8
µM (1) > 8µM (2) > 15µM (3) (Reid and Fairlie 1997; Fairlie et al., 2000). The
known fluorogenic substrate was also used in the same protocol described (Bergman
et al., 1995) to assess the catalytic ability of the D25N mutant HIV-1 PR, confirming
inactivation of the enzyme with this mutation (Kohl et al 1988).
60
2.4.2 Crystallisation Methods
Lyophilised wild-type and mutant D25N HIV-1 PR (2mg) was secondarily
purified to achieve >99.9% protein purity for controlled and uniform crystal growth
by refolding and dialysis. In each case the proteins were dissolved in 50% acetic acid
and diluted in 25 vols of buffer C (100mM acetate (pH5.5), 5% v/v ethylene glycol,
10% v/v glycerol). The refolded proteins were then buffer exchanged (period of two
hours, 10kDa cut-off YM-10 ultrafiltration vessels [Microcon]) to decrease the
amount of small carryover contaminants and undesired ions using buffer C. The
proteins were finally concentrated in YM-10 ultrafiltration vessels to a final
concentration of 5mg/mL. Proteins were then mixed with the substrate (dissolved in
dimethyl sulfoxide to a concentration of 50mM) in a ratio of 1:10 and stored for 16
hours at 4°C to allow equilibrium to be reached in all reactions for uniform crystal
growth.
Co-crystals of wild type and D25N mutant HIV-1 complexes were grown at
20°C by the hanging drop – vapour diffusion method following modified protocols of
Hui et al., (1993), using 2-5µL of protein-substrate mixture over a reservoir
containing 100mM acetate buffer (pH 5.5) and 35-60% saturated ammonium sulfate
as the precipitant. Crystals appeared within one to two weeks and grew over 4 weeks
to a maximum size of 0.4 x 0.1 x 0.1 mm3.
2.4.3 Structure Determination
For the crystal complexes conducted in this study, data was collected by Dr
Shu-Hong Hu (IMB) from single crystals using a Rigaku R-AXIS IIC imaging plate
area detector with CuKα radiation (1.5418Å) generated from a Rigaku RU-200
copper target rotating anode (46kV, 60mA) with a 0.00015” nickel filter. Cryo-
crystallographic conditions were established by Dr Hu and comprised plunging the
HIV-1 PR crystal into a solution containing 25% glycerol and 50% ammonium
sulfate and then immediately freezing the crystal in a low temperature nitrogen gas
stream (160K). Crystals used for data measurement had dimensions 0.4 x 0.1 x 0.1
mm3 (HIV-1 substrate) and 0.3 x 0.1 x 0.1 mm3 (HIV-1 product). Data was initially
collected by Dr Hu from 15min stills of both the substrate and product complex
showing diffraction to 1.6Å with no apparent radiation damage, before 45 frames of
data was collected through 180° of crystal rotation at 15min per frame.
61
Diffraction data were integrated and scaled using DENZO (Otwinowski,
1993) and SCALEPACK (Minor, 1993) and the structures were solved by difference
Fourier methods that use the phase of the incident X-ray beam and atom positions
from previously solved structures to develop the initial 3-dimension model of the
structure. The structures used for phasing were from a prior study using room
temperature data of substrate and product complexes (Pattenden et al., 1998). All
structures had 10% of the data set aside that was not refined. This data was used for
unbiased, statistical cross-validation using Rfree as a tool during the building and
refinement process to assess and maintain the validity and quality of the structures
(Brunger, 1992).
The model building was performed using the program O (Jones et al., 1991)
on a Silicon Graphics Indigo II R4400 and structure refinement was carried out on
initial models using simulated annealing in X-PLOR 3.851 (Brunger et al., 1987,
Brunger et al., 1990). Poorly ordered residues were positioned using OMIT maps
(Bhat and Cohen, 1984) where 10 - 15 residues were omitted from the refinement
processes to slowly develop an accurate model for poorly ordered residues. Simulated
annealing refined omit maps were also used on the product complex to better define
atom placements within complicated protein-protein and protein-ligand regions
(Hodel et al., 1992).
2.4.4 Analysis and Alignment
Structures were analysed using Insight IIC. Ligands were first unmerged from
the protein structures using the Biopolymer module then ligands were reassociated to
the parent protein structure. All superimpositions were then conducted on the 396 and
594 non-hydrogen atoms (Cα trace atoms 1-99 and 101-199), allowing the ligands to
be unbiased in the alignment, as they are no longer part of the protein templates that
are superimposed. All superimpositions were conducted with the substrate or
inhibitor complex as the target (unmoved).
All superimpositions were conducted of ligands within the active site of wild
type or mutated (D25N) HIV-1 PR in a similar manner for the superimpositions
described above, except atoms were selected as corresponding pairs for direct
positional placement and RMS deviation calculation in Insight II. Taking the core 18
atoms comprising the N- and C- terminal macrocycle with the substrate or inhibitor
62
complex as the target (unmoved) 34 non-hydrogen atoms were superimposed.
Excluded were those atoms, which are not part of the cyclic main chain atoms
comprising the valanyl/isoleucinyl side chains that have greater degrees of freedom.
Also excluded were the carbonyl oxygen atoms/hydroxy atoms and atom 203C as
these atoms are not uniform in all structures in terms of bonding and hybridisation.
Lastly, the carbon linker region (201 and 203) were excluded for uniformity as the
201 linker was not refined in the inhibitor complex (Martin et al., 1999).
63
Chapter 3: HIV-1 PROTEASE INHIBITORS BASED ON A
MACROCYCLIC TEMPLATE.
3.1 Introduction 3.1.1 The Problems of Current HIV-1 PR Inhibitors.
3.1.2 Cooperativity and Drug Design
3.1.3 Hypothesis, Aims and Significance
3.2 Results 3.2.1 Design of a Template for a Focussed Inhibitor Library.
3.2.2 Synthesis of a Focussed “Templated” Combinatorial
Library.
3.3 Discussion 3.3.1 Structure-Activity Relationships at P1'
3.3.2 Structure-Activity Relationships at P2'
3.3.3 Miscellaneous Compounds
3.3.4 Templating and Cooperativity
3.3.5 Bioavailability, Resistance and Cooperativity
3.3.6 Conclusions
3.4 Experimental 3.4.1 Purification of HIV-1 PR
3.4.2 Testing Inhibition of HIV-1 Protease
3.4.3 Computer Assisted Inhibitor Design
3.4.4 Inhibitor Synthesis
3.4.5 Inhibitor Purification and Characterisation
64
3.1 Introduction 3.1.1 The Problems of Current HIV-1 PR Inhibitors.
Current HIV-1 PR inhibitors (PIs) used to treat HIV infections have a number
of fundamental problems; the most important are pharmacological problems and the
emergence of viral resistance. Some of the pharmacological problems include low
bioavailability, drug-food interactions, drug-drug interactions, adverse side effects
and the need for high dosing due to drug specific intolerance, poor absorption and
poor penetration to all infected cell types, especially in the central nervous system
(Murphy, 1997). To date, bioavailability issues have been addressed by formulation
methods (eg crixivan is administered in a "soft-gel" form to facilitate absorption
across the intestinal wall). However, all PIs are reported to cause gastrointestinal
intolerance, nausea, vomiting, diarrhoea, hyperglycaemia, fat redistribution, lipid
abnormalities and increased bleeding episodes in patients with haemophilia (Van
Heeswijk et al., 2001).
The other important problem is the ease with which the virus develops clinical
drug resistance to PIs through mutations in the 99-residue amino acid sequence of
HIV-1 PR. The emergence of resistance has rendered “monotherapy” with a single
protease drug ineffective and has led to PIs being administered either in cocktails
with one another or in conjunction with other HIV-1 therapeutics (typically reverse
transcriptase inhibitors) (Deeks et al., 1997). Importantly, this combination therapy
has been spectacularly effective (especially involving crixivan) in extending by years
the lives of HIV-infected patients. Unfortunately, viral resistance still occurs, and
there is consequently an urgent need to find new ways to reduce this problem in order
to further prolong lives.
3.1.2 Cooperativity and Drug Design.
Rational design of protease inhibitors frequently begins by derivatising
peptide substrates to more pharmacologically acceptable non-peptides using
analogue-, mechanism-, and structure- based approaches to maximise inhibitor-
protease interactions (West and Fairlie 1995). Early inhibitors of HIV-1 PR were
peptidic in nature. Small peptides were largely abandoned as drug candidates due to
their poor bioavailability. They were also difficult to optimise due to cooperativity in
protease-inhibitor interactions (Abbenante et al., 1995; March et al., 1996; West and
65
Fairlie, 1995). This cooperativity is due to the induced fit of both the enzyme and the
inhibitor, which are flexible and respond to changes in one another when forming a
complex. Thus designed changes to the inhibitor initiate changes in the active site of
the enzyme during binding, resulting in loss of inhibitor potency. Similarly, mutations
in the enzyme can alter the ligand fit to the protease. The changes that occur in the
substrate-binding cleft upon ligand binding are largely unpredictable. Thus it can be
very difficult to optimise substrates/inhibitors by making systematic changes in each
of the binding pockets individually and iteratively. The enzyme orders its local
ligand-binding environment in response to changes to inhibitor structures, the
adjacent pockets or peripheral loops of the enzyme change shape resulting in “knock-
on” effects collectively termed “cooperativity” (Todd and Freire, 1999). A significant
problem in optimising enzyme inhibitors is therefore predicting how to most
effectively optimise one inhibitor region independently from another, while
minimising cooperative reshaping of the ligand-binding microenvironment of the
protein (Hofmann et al., 1988; Epps et al., 1990; Ridky et al., 1996; Majer et al.,
1997).
For these and reasons of bioavailability, the basis of drug design shifted long
ago to non-peptidic analogues. Unfortunately non-peptidic analogues are not simply
interchangeable with peptides. This is because there are no chemical compounds to
date that can accurately reproduce the geometry and bonding of a peptide (Reid and
Fairlie, 1997). Peptides are unique, having been selected by Nature as the
quintessential partner for proteins over a significant evolutionary period of time.
Peptide substrates have become the optimum molecules for forming dynamic,
complex and selective associations with enzymes. The substituting of peptides with
non-peptide analogues can alter the requirements of enzyme complementarity, ligand
binding mode, and cooperative reshaping of the enzyme's active site.
A possible compromise solution that comes close to mimicking a peptide,
while potentially avoiding their undesirable properties, is a minimalist approach to
inhibitor design involving connection of substrate side chain moieties that are close
enough to be linked in a covalent manner to form macrocyclic peptides (macrocycles)
(Figure 3.1). Such macrocycles potentially preserve amino acid components in the
same positions and orientations as a linear inhibitor bound to the enzyme (Abbenante
et al., 1995; March et al., 1996), but confer greater stability to metabolic degradation
and more closely mimic a peptide structure than any known non-peptide molecule.
There are many noteworthy advantages of the macrocycles described herein:
1. Macrocycles are fixed in the preferred "extended peptide conformation" of
substrates (Swain et al., 1990; Fairlie et al., 2000) that bind to proteolytic
enzymes. Macrocycles are thus pre-organised for binding to HIV-1 PR and
therefore have a selective advantage over acyclic peptidic inhibitors that need
to rearrange to the protease-binding shape (i.e. macrocycles out-compete
substrates). By being in a pre-organised conformation, macrocycles do not
have to overcome as large an entropy barrier to reorganise their structures to
the receptor-binding extended conformation.
2. Regional structural mimicry can be achieved with very subtle modifications of
the macrocycles and potential opportunistic associations capitalised upon, thus
optimisation of localised structural activity can be achieved. This means
inhibitors can be “tuned” as desired to fit the substrate binding cleft
(Abbenante et al., 1995).
3. The potency is found to be as good (if not better) than the best acyclic and
non-peptidic inhibitors (Reid and Fairlie, 1997).
4. The reduced conformational flexibility enhances regioselectivity and reduces
toxic side effects caused by non-specific interactions with other proteins.
5. Since the macrocycles involve minimalist deviations from the linear peptide
substrates they are designed to mimic, it is possible they encounter fewer drug
resistance problems (Tyndall et al., 2000).
NH
HN
O
O
O
HNH
N NH
O
O
O
NH
HN
O
O
HNH
N NH
O
O O
HO
O
OH
1
2
H2N
Figure 3.1: The Mimicry of Peptides by Macrocycles. (A) is a linear peptide substrate (LVF-FIV) and (B) is a macrocyclic inhibitor analogue constructed by ester linkage of the P1+P3 and P1’+P3’ moieties. Inhibition is achieved through a hydroxyethylamine replacement for the scissile amide bond.
A
B
66
3.1.3 Hypothesis, Aims and Significance
Our crystallographic studies in Chapter 2, performed with the E-S and E-P
complexes, revealed that HIV-1 PR maintains critical interactions within its substrate-
binding cleft throughout the reaction profile especially in the N-terminal cycles.
The Hypothesis to be tested is that "a macrocyclic peptide, which structurally
mimics a tripeptide component of a substrate for HIV-1 protease, can be used as an
effective template for the design and construction of potent inhibitors of HIV-1
protease".
The aims of this Chapter were restricted to:
1. Using an N-terminal macrocycle (e.g. from 2 in Chapter 2) as a template for
design and development of a small library of pre-organised inhibitors of HIV-
1 PR via combinatorial chemical synthesis.
2. Assessing the potencies of this library of compounds as inhibitors of substrate
processing by HIV-1 PR.
3. Examining the likely structural reasons for the relative observed potencies of
these enzyme inhibitors.
3.2 Results 3.2.1 Design of A Template For A Focused Inhibitor Library.
The N-terminal cyclic component of 6 (Fig. 3.2) resembles the N-terminal
cycles from Chapter 2, except for a lengthening by one carbon atom at P3, and so this
was chosen as the base template for a small and focused combinatorial library. The
major difference between the substrate 1 (from Chapter 2) and the planned inhibitors
is the replacement of the scissile amide bond by the hydroxylethylamine isosteric
linkage (as used in inhibitor 4 of Chapter 2).
NH
HN N
SO
O
O
O
OH
O
NH2
6
Figure 3.2: Potent Peptidomimetic Inhibitor 6 of HIV-1 PR. 67
A superimposition of the HIV-1 PR atoms of the E-I complex of 6 (Tyndall et
al., 2000) and the E-S (1), E-P (2, 3) and E-I (4) complexes from Chapter 2 (Fig. 3.3),
show that the proteins superimpose very well (RMS deviation 0.33Å, 0.27Å and
0.24Å for 396 non-hydrogen atoms onto the E-S, E-P and E-I (4) respectively). The
atom positions on the left hand side between P1 to P3 (Fig. 3.3) are not translated in
the same manner as the bicyclic inhibitor 4 towards S1 – S3 and comparatively align
very closely with the RMS deviation of the 34 cyclic atoms as conducted in Chapter 2
showing the product N-terminal cycle 2 superimpose best with the inhibitor cycle 6
(0.57Å, 0.14Å and 0.30Å for the N-terminal cycle of 1, 2 and 4 respectively) but all
are very alike within the active-site of HIV-1 PR. This is despite the variations in the
cycle size (5 vs 4 methylene carbons) and the diversity in the right hand side C-
terminal substituents (P1’ − P3’).
Figure 3.3: Templating Drug Design. An Overlay of the E-S complex of 1 (yellow), E-P complex of
2 and 3 (red), E-I complex 4 (green) of Chapter 2 and the E-I complex 6 (shown in purple - pdb
accession 1D4L, resolution 1.75Å). The overlay demonstrates the principle that a constrained
macrocycle can serve as a template to anchor an inhibitor to a specific region of the substrate-binding
cleft. It is evident that regional structural mimicry is achieved by the N-terminal cycles.
The N-terminal cycle and the surrounding protease from S1 – S3 appears to be
highly ordered, while the C-terminal acyclic appendages have little effect on protease
structure in the vicinity of the N-terminal cycles. This previously observed
conformational uniformity for inhibitors (Reid et al., 1996) localises the N-terminal
68
69
cycle within the substrate-binding protease cleft, fixing the positions of P1 – P3
substituents, and ordering the localised environment from S1 – S3, thereby
minimising cooperative enzyme-inhibitor interactions as all these ligands have
essentially the same binding mode. Upon this basis the N-terminal cycle of compound
6, and its N-alkylsulfonamide attachment, was considered a good template for the
following investigation. Once anchored, the N-terminal cyclic template should allow
independent fine-tuning of P1’ – P3’ appendages, represented in 6 by the N-
alkylsulfonamide (P1’ and P2’), that bind to adjacent regions of the protease.
Compound 6 contains two inhibitory components that must be considered for
library design. One is the N-terminal cycle of 6, which closely resembles the N-
terminal cycles from Chapter 2, except for a lengthening by one carbon atom at P3.
The cycle is also attached to a hydroxyethylamine isostere that replaces the cleavable
peptide amide bond of the substrate. Both this isostere and the cyclic template, which
mimics the tripeptide sequence L-V-F (and similar tripeptide analogues) and binds
strongly and predictably (Chapter 2 and Figure 3.3) anchoring the N-terminal
template, are constant in the inhibitor series described below. The second inhibitory
component is the N-alkyl sulfonamide of 6, which is similar to that in the potent
inhibitor amprenavir (VX-478), which contains an isoamyl functionality at P1’ and a
para-substituted benzylsulfonamide at P2’. This hybrid of the template cycle and
amprenavir is a potent inhibitor of HIV-1 PR, both in a competitive fluorogenic-
substrate assay for HIV-1 protease and in cells infected with HIV-1 (Tyndall et al.,
2000). Using this as a basis for similarly developing potent inhibitors with N-
alkylsulfonamide appendages, we were interested in probing and independently fine-
tuning P1’ and P2’ without affecting the binding to the S1-S3 region of the protease.
Thus our goal was to minimise cooperativity effects by dampening the induced fit
through use of a macrocycle that mimics a tripeptide segment of an
inhibitor/substrate. To evaluate substituent preferences at P1’ in 6, a series of
compounds with the N-terminal cyclic template and the P2’ para-
aminobenzylsulfonamide would be maintained, while the P1’ substituent is varied
(Fig 3.4). Similarly, the P1’-isoamyl is retained on the template, while a series of
different aromatic sulfonamides and ureas were synthesised for probing the P1’
pocket (Fig 3.4).
From the crystal structure of the E-I complex of 6, the S1’ pocket forms a
broad trough having no clear demarcation to S3’. The limit to the length from the
template-amine of 6 is ~4.5Å to make a VdW contact to Val82 (7.5Å away), and the
limit in width is an arc of 4.3Å. The available space for S1’ to S3’ is defined by the
residues; Arg8, Leu23, Pro81, Val82, Ile84, Gly127, Gly149 and Ile150 and has an
additional 4 water molecules filling the E-I complex of 6 (Fig. 3.5). Aside from Arg8
that borders the S3’ pocket, the S1’ pocket is hydrophobic. In contrast, the S2’ pocket
is more clearly defined and has a partial solvent face bordering at the bottom of this
pocket. The limit of S2’ pocket is hydrophilic, with a useable length of up to 6.5Å to
H-bond to Asp 130 (9.3Å from the template-amide) and a useable arc of 4.3Å to a
mixture of hydrophobic and hydrophilic residues. Within the E-I complex of 6, the
S2’ pocket comprises Ile50, Gly127, Ala128, Asp129, Asp130, Ile147 and Gly148
(Fig. 3.5) as well as 4 additional water molecules (waters not shown in Fig. 3.5).
Figure 3.4: The Concept of a Templated Combinatorial Library. The region of the inhibitor (P3 –
P1) is locked in a pre-organised conformation (yellow) in an attempt to order the surrounding enzyme
at S3-S1. This was anticipated to allow independent optimisation of the P1’ and P2’ inhibitor regions
(green) to specifically probe the enzyme subsites S1’ and S2’ without disturbing the S3-S1 enzyme
subsites through cooperative effects.
70
S3’ S1’
Arg8
Figure 3.5: The S1’ and S2’ Pocket Modelled from the E-I complex of 6. The model of the pockets that provides HIV-1 PR residue targets to optimise P1’ and P2’ functionalities is shown with a Connolly surface of the surronding enzyme. S2’
3.2.2 Inhibitor Design – P1’ and P2’ Variation.
The hydrophobic S1’ pocket generally favours likewise hydrophobic
substituents at P1’, such as large hydrophobic or aromatic groups found in many of
the substrates at P1, although Pro is also featured in some substrates. It was therefore
desirable to fill this pocket with large and bulky aromatic groups, short aliphatics, or
long aliphatics with flexibility to potentially adapt to changes in the pocket caused by
resistance-inducing mutations. Specifically enzyme residues Ile84, Ile50 and Val82
have been noted to cause or strongly contribute to resistance by altering the shape of
the S1’ pocket by contacting the top and side of the pocket.
There were several hypotheses for inhibitor design to match this pocket, with
the compounds proposed for synthesis shown in Figure 3.6.
1. Use of bulky aromatics that are a similar size to bulky substrate functionalities
at P1’ (compounds 9 and 10).
2. Use of short aliphatics to accommodate shrinkage of the S1’ pocket through
mutation (compounds 11 and 12).
3. Use of flexible aliphatics to adapt to possible shape changes in the S1’ pocket
(compounds 7 and 8).
71
4. Change the fit at S1’ by extending through the trough to S3’, trying to make
new interaction(s) with the highly conserved Arg8 charged side chain
(compound 7 and 9).
8
10 11 12
7 9
NH
HN
O
O
ON
OH
SO
O
NH2
NH
HN
O
O
ON
OH
SO
O
NH2
NH
HN
O
O
ON
OH
SO
O
NH2
HOO O
NH
HN
O
O
ON
OH
SO
O
NH2
NH
HN
O
O
ON
OH
SO
O
NH2
NH
HN
O
O
ON
OH
SO
O
NH2
Figure 3.6: Templated S1’ Probes of HIV-1 Protease.
Because of the clear definition of the S2’ pocket and because of the
amphipathic nature of the pocket, a greater variety of functionalisation of P2’ is
possible. This is reflected in the variability of the functionalities of the substrate, from
short aliphatic side chains as in Valine, to longer and bulkier aliphatics of
Isoleucine/Leucine, to charged residues of Glutamic acid and Glutamine. In this
respect the H-bonding and filling of this pocket is important and due to the variation
in size and charge, incorrect selection of P2’ functionalities of an inhibitor to this
pocket may influence inhibitor binding through cooperative effects.
Within the E-I complex of 6, the S2’ pocket accommodates associations with
ligands that have short charged groups, large bulky intermediate groups, or terminal
charges. For amprenavir mutations at Asp130, Val132, Met146, Ile147 and especially
Ile50 are important to inhibitor design and cooperativity. In this regard compounds 13
– 16 were made (Fig. 3.7).
72
13 14
15 16
Figure 3.7: Templated S2’ Probes of HIV-1 Protease.
NH
HN
O
O
O OHN
SO
O
NH
HN
O
O
O OHN
NH
HN
O
O
O OHN
SO
O
NO2
HN
ONH
HN
O
O
O OHN H
N
O
73
Compound 13 was designed with an opposite characteristic to amprenavir,
containing a proton acceptor rather than a proton donor so as to interact less with the
charged S2’ groups directly or perhaps with water molecules in the S2’ pocket.
Compound 14 was designed to fill the S2’ pocket with hydrophobic bulk and hence
push the limits of the templating method to overcome cooperativity. Compound 15
was anticipated to place a hydrophobic functionality with limited flexibility
(cyclohexane) on the urea linker (as opposed to a sulfonamide) into S2’, providing
information on urea substitution vs sulfonamides. Compound 16 was similar to 15,
but longer by a carbon atom and more constrained with an aromatic benzyl group to
test urea functionalisation.
3.2.3 Synthesis of a Focussed “Templated” Combinatorial Library.
The macrocyclic epoxide precursor (Fig. 3.8) was provided in large quantities
by Dr Robert Reid (IMB) who synthesized the material as previously reported (Reid
and Fairlie, 1997). The epoxide was then regioselectively opened in my chemical
reactions by primary amines designed to fit into the S1’ pocket, yielding a range of
hydroxylethylamine derivatives. These secondary amine intermediate products were
dried in vacuo to remove excess amine and solvent before I reacted them with the
appropriate sulfonyl chloride to yield the final N-alkyl sulfonamide substituted library
(7-14), or by isocyanates to contribute to a urea library (15, 16) as summarized by
Figure 3.8. The opening of the epoxide and subsequent amine substitution were
monitored using electrospray mass spectrometry, to ensure completion at the two
separate stages before purification using reversed phase HPLC. All compounds were
characterised by proton NMR spectroscopy and high-resolution mass spectrometry as
detailed in the experimental section.
Concomitant with the synthesis outlined, Dr Robert Reid synthesised
additional compounds (17-29) to increase the library and further test the templating
approach. All compounds (6 – 29) were assayed for efficacy in a competitive
fluorogenic assay against HIV-1 PR (experimental section 3.4.2) and the Ki relative
potencies as competitive inhibitors are shown in Table 3.1.
NH
HN
O
O
O OHBr N
H
HN
O
O
O OHBr N
H
HN
O
O
O O
RH2N
NH
HN
O
O
O OH H2N
R NH
HN
O
O
O OHN
R
R'
R’SO2Cl or R’NCO NaHCO3, THF/H2O
dried in vacuo
EtOH, ∆
Figure 3.8: General Synthetic Methods. The cyclic epoxide is opened using a primary amine (P1’
substituent). The intermediate secondary amine is further reacted with either a sulfonyl chloride or
isocyanate in the presence of mild base to yield the final inhibitor.
NH
HN
O
O
O OHN
SO
O
NH
HN
O
O
O OHN
HN
O
NH
HN
O
O OHN
S
NH
O
O O
O
HNHN
O
OH
O
(CH2)n
NS
O
OO
NH
HN
O
O OHN
SO
O
N
23
O
NH
HN
O
O
O OHN
HN
O
CO2H
NH
HN
O
O
O OHN
NO
OH
PhNH
HN
O
O
OO
NH
CONH2HN
O
O OHN
SO
O
20 n=3 21 n=4 22 n=5
17 18 19
O
NH
CONH2HN
O
O
O OHN
O NH
NH
CONH2HN
O
O
O OHN
O NH
HN
O
CONH2 NH
HN
O
O
O OHN
S
NH2
O
O
H2N
27 28 29
24 25 26
Figure 3.9 Inhibitors Designed and Synthesised by Dr Robert Reid. Above are the compounds
made by Dr Reid for probing the S1’ and S2’ pockets of HIV-1 PR.
74
Compound Ki (nM) Compound Ki (nM)
6 1.7 18 295
7 60 19 10
8 14 20 4
9 3 21 0.4
10 8 22 1.6
11 2 23 1800
12 3 24 20,000
13 145 25 12
14 4700 26 1
15 207 27 12
16 314 28 12
17 1 29 0.9
Thi
s Ph.
D
Table 3.1 Inhibitor Potencies vs HIV-1 PR. The outlined compounds 7 – 16 were synthesised by
Pattenden and the compounds 6 and 17 – 29 were synthesised by Dr. Reid of the IMB, UQ.
3.3 Discussion 3.3.1 Structure-Activity Relationships at P1'
Compounds 7 – 12 contain changes at P1’ that were made in order to gain
insight to inhibitor binding at the S1’ enzyme pocket. Compounds 7 – 12 maintain the
para-aminobenzylsulfonamide as used in the commercial compound amprenavir. In
this series the most potent compounds were small and bulky about the isostere
linkage (11, 12), or had extended hydrophobic bulk projecting further into the pocket
(7-10). Compounds 9-12 were similar in potency to one another and to amprenavir. In
contrast compounds 7 and 8 were significantly less active. Compound 8 has the
extended aliphatic chain at P1' and this is clearly not well accommodated at S1' in the
enzyme. Compound 7 was designed to extend P1' to stretch from S1' into the S3’
subsite (and perhaps associate with Arg8), but it is the poorest inhibitor in the series.
It is plausible that 7 makes the intended contact to Arg8 but that this association is
detrimental to binding. In this respect the role of Arg8 – a conserved residue in all
retroviral aspartyl proteases - may be specific to S3’ and its presence closer to S1’
may be due to cooperative arrangements of the pockets S1’ and S3’ when there is no,
or small functionalities in S3’. Furthermore, attempts to directly fix Arg8 into the S1’
pocket would be predicted to be detrimental.
75
76
3.3.2 Structure-Activity Relationships at P2'
The probes for the S2’ pocket can be broken into two compound classes, N-
alkyl sulfonamides and N-alkyl ureas. The N-alkyl sulfonamides comprise
compounds 7-14 and 17–22, 26 and 29, whereas the N-alkyl ureas comprise the
compounds 15, 16, 24 and 25.
In general the N-alkyl sulfonamides were the more potent inhibitors (IC50 1-
4700nM). The compounds 17, 19, 22 and 29 had inhibition <12nM, restricting
substitution of the P2’ position to the unsubstituted or para-substituted benzyl or
napthyl sulfonamides. The best of these was the 2-substituted napthalene 17, having a
Ki of 1nM, whereas the addition of the extra amine to the template of 6 forming 29
had similar potencies to 17. An important finding was a comparison between 17 and
18, which revealed the importance of the substitution pattern. The 1-position of the
quinoline functionality (18) directed the aromatic group to clash with the bottom of
the S2’ pocket, lowering the Ki value to 295nM. In contrast the 2-position directs the
napthalene of 17 along the length of the S2’ pocket and through the S3’ pocket,
thereby maximises the filling of both pockets and conveying the highest potency of
all compounds in this series, that was slightly greater than the amprenavir substituted
template (1nM vs 1.7nM).
The N-alkyl ureas proved to be poorer inhibitors than the N-alkyl
sulfonamides with Ki values ranging from 12nM – 20,000nM, with compounds 25, 27
and 28 being equipotent (12nM). This was likely due to inflexibility of the planar
urea moiety, with the substituents being directed along the H-bonding channel that
the peptide backbone normally occupies rather than into the S2’ pocket as for the N-
alkyl sulfonamides. The use of hydrophobic bulk along the line of the peptide
backbone was detrimental to binding (15 and 16). The only exception to this was 25
(12nM), which contains a t-butyl group that occupies smaller space.
3.3.3 Miscellaneous Compounds
The size of the cyclic template was varied in the inhibitors 20 – 22, all having
a common N-(isopentyl)-benzylsulfonamide substituent at P1' and P2' and lacking
only the para-amide group of 6. Compound 22 is the closest structural analogue to 6
and has essentially identical potency (1.6nM versus 1.7nM respectively). The Ki for
77
the series 20 – 22 show decreasing potency for n = 4 (4x) > n = 5 (2x) > n = 3, with
the difference from n = 4 to n = 3 being 10 fold. The differences in potency are
possibly related to more optimal dampening of cooperative effects by the n = 4 and 5
macrocycles compared to n = 3, and this appears to have a limit as the n = 4 had a
marginal advantage over the n = 5 macrocycle.
Compound 26 is almost identical to 20, but uses an Asn side-chain as part of
the cyclic template (L-N-F), instead of the valine (L-V-F). The Asn substitution
appears to increase the potency of the template by making a H-bond in S2 (1nM for
the Asn vs 4nM for Val). It can be concluded that, despite minor differences in
potencies induced by the size and minor substitution of the templates, any of these
templates could have been used to probe S1’ and S2’ as differences between cycles
for the substrate (n = 4) and the inhibitor 6 (n = 5) do not alter the binding mode
(Figure 3.5).
Compounds 27 and 28 contain a P2 Val to Asn change in the cyclic template
as for 26, changes from an N-alkyl substitutent at P1' to either Pro or Pipicolate, and
the benzylsulfonamide at P2' in 26 is either Ile or tert-butyl. These compounds have
the same potency (12nM) despite the differences in size and functionality. Their
binding in S1’ will be similar to compounds 11 and 12, but their S2’ (and S3’)
binding will be quite different due to differences in their main-chain H-bonding
arrangement that is more substrate-like. Compounds 27 and 28 do, however, provide
a yard-stick to measure the other inhibitors, as the consensus sequence of 27 is close
to the p17 – p24 and p11 – p51 polypeptide substrate cleavage sites. Compound 28,
having essentially the same potency as 27, shows that the S3’ pocket does not
contribute substantially, or at least isn't required, for high potency. Therefore
compounds with Ki values greater than 27 and 28 (12nM) bind more poorly than
substrate-analogues due either to unfavourable interactions or cooperative effects, and
thereby substitutions incurring Ki values of >12nM can be taken as non-ideal for this
compound series.
The urea 24 and the peptide analogue 23 were found to have unusually high
Ki values of 20,000nM and 1800nM respectively, much higher than any other
compound in the N-alkyl urea series. It is obviously important to know what binds
tightly for drug design, but in terms of cooperativity it is equally important to know
what constitutes a poor inhibitor to set limitations to drug design and learn more
78
about the enzyme target. For compound 23 there are several possible reasons for its
poor binding. The loss of the peptide planarity by omission of a nitrogen, together
with the bulky aromatic group occupying space where a peptide backbone sits for
S3’, is detrimental in terms of cooperativity. It is also possible that a rearrangement
has occurred as shown for 23 in Figure 3.10 which could cause crowding in S1’,
allowing the isoamyl group to switch into S2’ and change the binding mode
completely. In a similar scenario, 24 may alter it’s binding mode away from the
aspartic acid residues in the S2’ pocket by placing the P1’ isoamyl group into the S2’
pocket and the urea-substituted valaninyl into the S1’ pocket. In such a scenario the
inhibitor would suffer from the rigidity of the urea and the negative charge that is of a
sufficient length to interact with Arg8 at S3’.
3.3.4 Templating and Cooperativity.
An overlay of the crystal structure of 6 with the substrate and product
complexes of Chapter 2 established that the template binds to HIV-1 protease in the
same conformation and mode as the cyclic substrate (1) and product ligands (2 and
3), despite the size of the template and functionality (Fig. 3.3). The macrocycle
appears to organise both itself and its surrounding protease environment, while the C-
terminal acyclic appendages have little effect on protease structure in the vicinity of
the macrocycle. In agreement with this structural finding, the structure-activity
relationship (SAR) investigation of P1’ and P2’ using the templating method found
that the template locked P1 – P3 in an ordered manner for probing S1’ (Fig. 3.3). The
success of this strategy is attributed to the template occupying three contiguous
subsites S3 – S1 in the protease, thereby maintaining all the hydrogen bonding and
pocket filling interactions with the protease and fixing the surrounding protease
environment in the vicinity of the macrocycle at P3 – P1.
From the SAR investigation of P1’, it was found that the benzylsulfonamide
component of amprenavir was good for maintaining P2’ without rotation about the
nitrogen linker, allowing investigation of P1’ using a variety of inhibitory strategies.
The best functionalities were found not to contain charge or flexible alkyl chains.
Rather local structural order of P1’ functionalities towards the top of the S1’ pocket
or very low to the pocket were found to be essentially equal in potency and favoured
by the enzyme, and in contrast, the SAR investigation of the S2’ pocket found that N-
alkyl ureas bind similarly to peptides, but their potency was reduced by substitution
towards the S3’ pocket. It may be possible to further tune the N-alkyl ureas on the
cyclic template by secondary substitution towards the S2’ pocket as shown in Figure
3.11. By maintaining the planarity and H-bonding of the peptide-like backbone of the
inhibitor and probing the trough of S2’ it may be possible to take advantage of the
charged aspartic acids in the bottom of this pocket with hydrogen donors such as a
secondary amine. This method would be favourable if a more peptide-like compound
from S1’ – S3’ is desired, however, the N-alkyl sulfonamides have an advantage to
other inhibitors as they do not follow the peptide backbone, but directly penetrate into
S2’ from the nitrogen linkage. This novel mode of binding takes advantage of the
amphipathic nature of the S2’ pocket while allowing filling of the extended trough.
From the SAR investigation of P2’ the 2-substituted napthylsulfonamide (17)
conferred the most potent inhibition in this series followed by the benzylsulfonamide
(22) being essentially equivalent in potency to the benzylsulfonamide of 6 and the
template modified analogue of 6 (29), providing suitable alternatives to the
amprenavir P2’ substituent.
NH
HN
O
O
O OHN
O R1
R2
O
Figure 3.10: Possible Optimisation of the N-alkyl Ureas by retention of the planarity around R2
while utilising the space in S2’ via substitution at R1.
3.3.5 Bioavailability, Resistance and Cooperativity.
For successful drug development, all parameters that can influence the
efficacy of a compound must be considered at the design stage. Moving from native
peptide substrates to more bioavailable synthetic analogues can cause changes in
ligand recognition by the enzyme. Similarly, the emergence of resistance to these
analogues could be due to a change in the fit of an inhibitor to the substrate-binding
cleft through enzyme mutation. Resistance to current PIs often arises from mutations
79
occurring within the substrate-binding groove of the enzyme, thereby changing its
shape and directly influencing the fit of the inhibitor and its consequent affinity for
the protease (Winslow and Otto, 1995; Condra et al., 1995; Markowitz et al., 1995;
Partaledis et al., 1995; Baldwin et al., 1995; Chen et al., 1995; Gulnik et al., 1995;
Mahalingam et al., 1999; Olsen et al., 1999). Alternatively, important mutations can
also occur remote to the active site causing resistance either by structural packing or
knock-on effects that influence the active site shape or by altering the kinetics of
substrate processing.
HN
ON N
O
A
NH
N
O
HN B
OHO
O
H
O
H
H
HN H
HN
ON N
O
A
NH
N
O
HN B
OHO
O
H
O
H
H
HN H
P2'
S2' S2'
P2'
1 2
i ii
Figure 3.11: Schematic of Interactions in HIV-1 PR and Re-shaping of the Substrate-binding
Cleft. Chains A and B represent a β-sheet of HIV-1 PR in which A is remote from the substrate
binding cleft S2’ and B has an aspartic acid group extending into the pocket and interacting with an
inhibitor moiety P2’. Scheme (i) shows a VDW association between a valine on A and an isoleucine on
B. In scheme (ii) the valine is mutated to a leucine causing shifts in B to avoid the steric effects of
VDW repulsions. This subsequently has a “domino effect” shifting the aspartic acid in S2’, making
interactions with the inhibitor moiety in P2’ sterically disfavoured, causing an adverse change to the Ki
(perturbation of affinity) and rendering the drug ineffective in vivo.
The effects of mutations more remote to the substrate-binding cleft are not
always obvious and can alternatively influence dimer stability, inhibitor-binding or
substrate-binding kinetics, or active site re-shaping through long-range structural
perturbations (Boden and Markowitz, 1998; Erickson et al., 1999). Mutations that re-
shape through long-range interactions can expand or contract the size/shape of the
hydrophobic binding pockets so that inhibitors either cannot make adequate H-
80
81
bonding contacts or are influenced by van der Waal’s steric clashes as shown in
Figure 3.11 above (Olsen et al., 1999). Mutations can thus decrease the affinity of an
inhibitor for an enzyme, or they can increase the rate of processing of a substrate,
potentially with the same result, namely reduced inhibitor affinity and potency.
Sequence analysis of drug resistant viruses has shown that mutations can also
occur within polypeptide substrates (Doyon et al., 1996; Zhang et al., 1997,
Mammano et al., 1998). These mutations appear to be compensatory rather than
primary. Patterns of resistance have been specifically noted for different regions of
HIV-1 PR, especially in the so-called 30's and 80's loops (Towler et al., 1997). Table
3.3 below is a summary of known modes of resistance for current PIs.
MUTATION INDINAVIR RITONAVIR SAQUINAVIR NELFINAVIR AMPRENAVIR SITE CRIXIVAN NORVIR INVIRASE VIRACEPT AGENERASE Leu10 Accessory Accessory Accessory Accessory Accessory Lys20 Accessory Accessory Accessory Accessory Accessory Leu24 Contributes -- -- -- -- Asp30 -- -- -- Causes -- Val32 substrate Contributes Contributes -- -- Contributes Met36 Accessory Accessory Accessory Accessory Accessory Met46 flap Contributes Contributes -- Contributes Contributes Ile47 flap Contributes Contributes -- -- Contributes Gly48 flap-subs Contributes Contributes Contributes Causes Contributes Ile50 flap-subs -- Contributes -- -- Causes 53 flap Contributes Contributes Contributes Unknown Unknown Ile54 flap Contributes Contributes Contributes Contributes Contributes Leu63 Accessory Accessory Accessory Accessory Accessory Ala71 Accessory Accessory Accessory Accessory Accessory Gly73 Contributes Contributes Unknown Contributes Unknown Val77 Accessory Accessory Accessory Accessory Accessory Val82 substrate Causes Causes Contributes Contributes Contributes Ile84 substrate Causes Causes Causes Causes Causes Asn88 Contributes Unknown Unknown Contributes Hypersensitivity Leu90 Contributes Contributes Causes Causes Contributes Asn93 Accessory Accessory Accessory Accessory Accessory Table 3.2: HIV-1 PR Inhibitors and their Known Mutations. Substrate = region of substrate-
binding cleft; flap = region of the flaps; flap-subs = region of the flap and substrate-binding cleft; -- =
no selection from the drug; Unknown = unknown if effects; Accessory = found as polymorphic
variation in untreated infected patients and contributes to resistance in the presence of other mutations
(does not directly cause); Contributes = directly contributes to phenotypic resistance in association
with other mutations; Causes = directly causes phenotypic resistance and viral persistence;
Hypersensitivity = causes hypersensitivity to the drug. (adapted from Condra et al., 1996; Molla et al.,
1996; Schapiro et al., 1996; Murphy 1997; Boden and Markowitz 1998; Craig et al., 1998, Patick et
al., 1998; Shafer et al., 1999; Atkinson et al., 2000). To date there is only limited work conducted on
resistance for lopinavir (Carrillo et al., 1998).
82
It can be seen that some commonality exists in resistance patterns for all PIs
and some close similarities between specific PIs (crixivan and norvir, saquinavir and
nelfinavir), although amprenavir is different. This is most likely due to similarities of
structural size occupying the regions Ile54-Leu90 and the similarity in binding sites
(S3-S3') for the similar drugs, whereas in the case of amprenavir, the binding sites are
from S2-S2' (West and Fairlie, 1995). Specific problems have been found for
saquinavir from G48V and L90M mutants (Ohta and Shinkai, 1997) and, for
amprenavir, selection of M46I, I47V and I50V (Partaledis et al., 1995).
It is likely that resistance and cooperativity are related. When HIV-1 PR
mutates a residue, there are changes however subtle in the packing, folding or kinetic
processing associated with the mutant enzyme. Frequently the cooperative or knock
on effects will influence ligand binding in the active site. To maintain a viable virion,
the mutant HIV-1 protease can slow, but must not prevent, substrate processing. Thus
templated drug design of inhibitors that closely mimic a protease substrate could
potentially overcome the problems of resistance, since such inhibitors will only be
impeded from exerting their affects by competitive pressure from unprocessed
substrate. By developing inhibitors that are more substrate-like, it should be possible
to (a) limit viable mutations to those that stop or drastically slow processing, and (b)
dampen the cooperative effects of resistance mutations that normally inactivate
inhibitors.
3.3.6 Conclusions.
The cooperativity of enzyme-ligand interactions is a significant impedance to
effective design of potent and long-acting inhibitors. To overcome this problem we
have successfully employed a constrained cyclic tripeptide as a pre-organised beta
strand template, which orders the surrounding S1-S3 enzyme environment and thus
can dampen any cooperativity effects due to alterations elsewhere in the enzyme. The
conformational rigidity of the template localises the cycles within the S1-S3 section
of the substrate-binding groove of the protease, while allowing independent
regioselective optimisation of inhibitor segments elsewhere (e.g. S1'-S3') via focussed
combinatorial libraries.
An analysis of hundreds of crystal structures of aspartic, serine, metallo and
cysteine proteases has established that proteases exclusively recognise the β-strand
83
conformation of inhibitors/substrates (Tyndall and Fairlie 1999; Fairlie et al., 2000).
While most reported protease inhibitors are acyclic and therefore conformationally
flexible, the constrained macrocyclic template used herein was found to structurally
and functionally mimic the P1 – P3 tripeptide segment of a substrate for HIV-1 PR.
This templating approach could be extended to other classes of proteases and other
classes of inhibitors as a means of developing potent substrate-based inhibitors,
including those for which structural information is unavailable but substrate
sequences are known.
It is possible with this approach to tune an inhibitor to fit one half of a
protease active site (e.g. S1’ and S2’), and then use a template to tune it to the other
half (e.g. S1 – S3). But there are some difficulties in simply combining the optimised
acyclic segments together, because the acyclic portions do not themselves pre-
organise the enzyme. In comparison to acyclic inhibitors and peptides, the cyclic
template is more effective because (a) the macrocycle is pre-organised for binding to
HIV-1 PR and therefore has a selective entropic advantage over acyclic peptidic
inhibitors (Reid and Fairlie, 1997); (b) a macrocycle is able to maintain the binding
mode from S1 – S3, despite a diversity of functionalisation in P1’ and P2’ and thus
overcome cooperativity and allow optimisation of inhibitor components at P1’ and
P2’; and (c) regional structural mimicry can be achieved with very subtle
modifications of the macrocyclic template or novel non-peptidic appendages,
independently “tuned” to optimally fit the substrate binding cleft (Abbenante et al.,
1995).
It is known that the substrate-binding cleft must ‘breathe’ (expand and
contract) to process differing substrates, and resistant HIV-1 PRs have truncations
and elongations within the substrate-binding cleft that reduce inhibitor specificity
(Towler et al., 1997). However, the substrate must also be recognised in spite of
mutations in the protease, or the enzyme will not process it. A macrocyclic inhibitor
offers a possible way to overcome resistance, since it is very substrate-like while
maintaining potency. Within the macrocycles it is possible to reduce the size of the
side chains responsible for selectivity and present a structurally similar compound to
a substrate. It is often found that more potent inhibitors are highly functionalised to
fully occupy enzyme pockets. On this basis it is predicted that reducing the inhibitor
size will also lead to a reduction in the Ki due to decreasing contacts in the substrate
84
binding cleft. However, the SAR findings for P1’ and P2’ described above are very
exciting, as reduction from the isoamyl to proline or 2-substituted butyl groups at P1’
(such as 11 and 12), or removal of the amine substitution in P2’ with the less
substituted benzyl sulfonamide (22) or control of the directionality along the substrate
trough (as for 17), were found to be equipotent to 6. In this respect the macrocycle
template might be employed to overcome resistance without significant loss of
potency.
Alternatively, the template may provide a platform to support flexible
substituents that adopt the receptor-binding shape recognised by this protease. Thus
the inhibitor is pre-organised for receptor binding, but has sufficient in-built
flexibility to ‘breathe’ and respond to changes in the substrate-binding cleft of the
protease while maintaining inhibitor affinity for the protease (March and Fairlie,
1996). The macrocycles described here can potentially solve this problem if used as
templates with flexible aliphatic chains, enabling the otherwise constrained cycles to
“breathe” in response to mutation-induced changes in the protease. This could allow
the maintenance of the essential threshold of H-bonds and VDW contacts of the
inhibitor with HIV-1 PR.
3.4 Experimental 3.4.1 Purification of HIV-1 PR.
In order to assess the inhibitor potency, crude lyophilised HIV-1 PR was
obtained from Diane Alewood (Centre for Drug Design and Development, University
of Queensland) and dissolved in a minimum volume (4mL 15% solution) of buffer B
(90% acetonitrile, 9.9% ddH2O, 0.1% TFA). The solubilised material was applied to
a Vydac C-4 semi-preparative column (Ø4.6mm x 150, pore size 300Å) at a gradient
of buffer B of 1mL/min (buffer A = 90% ddH2O, 9.9% acetonitrile, 0.1% TFA). The
protein was found to elute in 43 - 45% B, and 500µL cuts were taken over this range.
Fractions were injected into a PE-SCIEX API III triple quadrupole mass spectrometer.
Fractions were analysed over 25 scans (dwell 0.3msec) from a mass range of 700.0 to
2100.0amu in increments of 0.2amu. The peaks were reconstructed from the XIC
(excitation ionisation chromatogram) of +Q1 data (I, I+1 patterns) showing a
characteristic HIV-1 PR series of peaks of 1076.8 [M + 10H]10+, 1195.9 [M + 9H]9+,
1345.4 [M + 8H]8+, 1537.4 [M + 7H]7+, 1793.6 [M + 6H]6+ reconstructing to a
singular molecular mass of 10755.7amu (+/- 1.4amu) for pure fractions that compares
closely to a calculated mass of 10754.7amu. The three most pure fractions were then
pooled and lyophilised before dissolution in 6M Gu.HCl (0.05mg/mL) for storage (-
20°C).
2.61e4 cps+Q1: 18.79 min (33 scans) from HIV-1PR191198, smoothed
1 1 9 5 .9
1 3 4 5 .4
1 5 3 7 .4
1 7 9 3 .61 0 7 6 .8
800 900 1000 1100 1200 1300 1400 1500 1600 1700 1800 1900 2000m/z, amu
5000
10000
15000
20000
25000
Inte
nsity
, cps
5.46e3 cpsBioSpec Reconstruct for +Q1: 18.79 min (33 scans) from HIV-1PR191198, completed iteration: 7, smoothed
1 0 7 5 5 .7
9600 9800 10000 10200 10400 10600 10800 11000 11200 11400 11600 11800Mass, amu
500
1000
1500
2000
2500
3000
3500
4000
4500
5000
Inte
nsity
, cps
Figure 3.12 Mass Spectrometry of Purified HIV-1 PR. Shown is the +Q1 and reconstruct of
molecular ions from the XIC, forming a singular molecular mass of 10755.7amu (+/- 1.4amu) that
compares closely to a calculated mass of 10754.7amu.
3.4.2 Inhibition of HIV-1 Protease
In order to assess inhibitor potency, HIV-1 PR was refolded for 60mins in
buffer A (20mM phosphate pH 7.0, 16% v/v glycerol, 2mg/mL BSA). The buffer
A/protease solution (10µL) was then added to buffer B (MES pH 6.5, 37ºC, 100mM
NaCl, 10% v/v glycerol). The final assay condition was buffer B and a final
concentration of 0.1% DMSO that was used in the assay for solubilisation and
dilution of the substrates or inhibitors. The substrate used to calculate Km and Ki was
Abz-NF*-6 which has the following sequence: Abz-Thr-Ile-Nle-phe(NO2)-Gln-Arg-
NH2 and is based on a non-native substrate Ac-Thr-Ile-Nle-Nle-Gln-Arg-NH2, as
used with the early inhibitor MVT101 (Toth et al., 1990). When calculating the Km
85
from Hanes plots (Fig. 3.13), the concentration of Abz-NF*-6 was varied. The Km
was found to be 35.33µM. Hanes plots were also used as a control to confirm enzyme
consistency and activity following folding.
Hanes Plot HIV-1 Protease
y = 33003x + 1.166R2 = 0.987
-0.2
0
0.2
0.4
0.6
0.8
1
1.2
1.4
1.6
1.8
2
-4.00E-05 -3.00E-05 -2.00E-05 -1.00E-05 0.00E+00 1.00E-05 2.00E-05
[S] Abz-NF*-6
[S]/v
Figure 3.13: Hanes Plot of [S]/v vs [S]. Shown is a Hanes plot of HIV-1 PR activity using variable
substrate concentrations. The Km (x-intercept) was found to be 35.33µM and the equation of the line
and R2 value is embedded in the figure.
[S]/v
Y= 33003X + 1.166 R2 = 0.99
[S] Abz-NF*-6
Inhibitors were dissolved in 100% DMSO and serially diluted into 10%
DMSO before addition to assay solutions (Abz-NF*-6 was immediately dissolved in
10% DMSO due to it’s solubility). Certain compounds were very hydrophobic and
needed to be serially diluted in 100% DMSO to remain in solution. In such cases less
volume of the inhibitor was added to the assay, so at all times the final DMSO
concentration in the assay did not exceed 0.1%. A fixed concentration of the substrate
(50µM) was used for calculating the inhibition values (IC50 and Ki) with a varied
concentration of the inhibitor. Dixon plots were employed for calculating the IC50
(plotting 1/v vs [I]), where the x-intercept = -IC50 (Fig 3.14).
In the case of tight binding inhibitors or where [I] approaches [E], Henderson
plots (Fig 3.15) were employed to determine the IC50 values (plotting [I]/{1-(vi/vo)}
vs vo/vi), where vo is the rate of turnover of substrate and vi is the rate in the
presence of the inhibitor and the slope = IC50. The Km for the purified enzyme was
then used for converting IC50 values to Ki values according to the formula Ki =
IC50/(1+[S]/Km). When calculating inhibitor Ki, compound 6 (Ki 1.7 nM) was used as
86
a control to ensure overall accuracy and consistency of the assay from previous
assays conducted and when using different batches of enzyme.
Dixon Plot Compound 10
y = 5E+06x + 0.0879R2 = 0.9919
0
0.1
0.2
0.3
0.4
0.5
0.6
0.0E+00 2.0E-08 4.0E-08 6.0E-08 8.0E-08 1.0E-07 1.2E-07
[I]
1/V
X-intercept –17nM (IC50) R2 0.99 Sy.x .018
[I]
Fig 3.14: Dixon Plot Compound 10. Shown the embedded analysis of the data from GraphPad Prism
over the Dixon plot generated from Microsoft Excel for compound 10. The X-intercept yields the IC50
value that is then converted to a Ki.
Henderson Plot Compound 10
y = 17.382x - 2.6552R2 = 0.9757
20
30
40
50
60
70
80
90
100
110
120
2 2.5 3 3.5 4 4.5 5 5.5 6 6.5 7
Vo/Vi
[I]/{1
-(Vi/V
o)}
[I]/{
1-(v
i/vo)
}
Slope (IC50) 17 +/- 2nM R2 0.98 Sy.x 7.17
vo/vi Fig 3.15 Henderson Plot of Compound 10. Shown the embedded analysis of the data from GraphPad
Prism over the Henderson plot generated from Microsoft Excel for compound 10. The X-intercept
yields the IC50 value that is then converted to a Ki.
87
88
3.4.3 Computer Assisted Inhibitor Design
The X-ray crystal structure coordinates of compound 6 were imported into
Insight II and potential inhibitors designed from this structure. In this approach to
drug design, it was assumed that the template and functionalities which closely
resemble 6 would occupy the same 3D space as in the crystal structure of 6 bound to
HIVPR, and this model would be more accurate than independent energy minimised
models. Therefore, using the Builder module of Insight II inhibitors were constructed
by modifications of 6 within the restraints of the S1’ and S2’ pocket. It was found
from the crystal structure of the E-I complex of 6, that distance and angle restraints
could be developed to allow fitting to the trough S1’ – S3’ about residues Arg8,
Leu23, Pro81, Val82, Ile84, Gly127, Gly149 and Ile150 and including the 4 water
molecules occupying these pockets. Within the S2’ pocket up to 6.5Å x 4.3Å space
was available, comprising Ile50, Gly127, Ala128, Asp129, Asp130, Ile147 and
Gly148 as well as 4 additional water molecules.
3.4.4 Inhibitor Synthesis
A solution of the macrocyclic epoxide (Fig. 3.8) (10mg, 28µmol) in ethanol
(2mL) was stirred and refluxed with 10 - 20 equivalents of the desired amine (12h).
The solution was evaporated to dryness under high vacuum to remove as much of the
excess amine as possible. The residue was dissolved in THF (3mL) and saturated
aqueous NaHCO3 (100µL), then the desired sulfonyl chloride or isocyanate (2-5
equivalents) was added with stirring at room temperature. Acylation proceeded
rapidly, (less than 30min), and was monitored for complete consumption of the
intermediate secondary amine (Fig 3.8) by mass spectroscopy. Inhibitors that required
no further elaboration were evaporated and re-dissolved in MeCN/H2O and purified
by reverse phase HPLC. Fractions containing products were lyophilised to give white
powders (typically 2 - 8mg).
Inhibitors 7-12 were prepared using 4-acetamidobenzenesulfonyl chloride and
therefore required an additional hydrolysis step to deprotect the para-amino group.
Methanol (3mL) and 2M HCl (1mL) were added and the solution was stirred and
heated at 60°C for 1 - 3h. Cleavage of the acetyl group was conveniently followed by
mass spectroscopy and no undesirable degradation of the macrocycle amide bonds
89
was observed. The solution was evaporated and the residue was purified by reversed
phase HPLC as above.
The urea derivative inhibitor 15 was prepared in two steps from the amine
(Fig. 3.8) and valine methyl ester isocyanate in THF, followed by hydrolysis of the
methyl ester with NaOH. The solution was acidified with TFA and purified by RP-
HPLC 50:50% MeCN:H2O + TFA 0.1% isocratic, retention time 6min, giving a white
powder after lyophilisation.
3.4.5 Inhibitor Purification and Characterisation
Proton and carbon NMR spectra were recorded on a Varian Gemini 300 or
Bruker ARX-500 spectrometer at 298K and were referenced internally to the residual
solvent peak: CD3OH δH 3.31, δC 49.0; DMSO-d6 δH 2.50, δC 39.7; CD3CN δH 1.94,
δC 1.39; CDCl3 δH 7.27, δC 77.0 ppm. Routine Mass spectra and ESI-MS were
measured on a PE-SCIEX API-III or Perceptive Biosystems Mariner API-qTOF
instrument equipped with an LC pump and Rheodyne injector. Reaction mixtures
were sampled and diluted with 70% MeCN/30% H2O and were introduced into the
mass spectrometer at 30 - 50µL/min. As a result of the atmospheric pressure
ionisation process (electrospray), only molecular ions MH+ were observed for each
component in the sample. High Resolution Mass Spectra of purified products (except
where stated as ESI-MS) were measured on a Finnigan 2000 Fourier Transform Mass
Spectrometer at 3 Tesla with resolving power of greater than 20,000 by Graham
MacFarlane (Dept. of Chemistry, UQ). Preparative reverse phase HPLC was
conducted on a Waters 600 system equipped with a Rheodyne preparative injector
with 5 mL loop volume on a Vydac C-18 90Å 10 µM column 250 x 22 mm at 20
mL/min using gradient elution (solvent A: water + 0.1% TFA, solvent B: 90% MeCN
10% water + 0.1% TFA) and UV detection at 280 nm.
6-{(4-Amino-benzenesulfonyl)-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-
diaza-bicyclo[13.2.2]nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-amino}-hexanoic
acid 7. 1H NMR (500 MHz, CD3OH) δ 7.59 (d, J = 8.8 Hz, 1H), 7.50 (d, J = 8.8 Hz, 2H),
7.08 (d, J = 8.0 Hz, 2H), 6.85 (d, J = 8.8 Hz, 1H), 6.78 (d, J = 8.0 Hz, 2H), 6.70 (d, J
= 8.8 Hz, 2H), 4.23 (m, 1H), 4.18-4.03 (m, 3H), 3.76 (m, 1H), 3.44-3.14 (m, solvent
90
obscured), 3.08 (m, 1H), 2.94 (m, 1H), 2.42 (t, J = 13.2 Hz, 1H), 2.30-2.23 (m, 2H),
2.19-2.11 (m, 2H), 1.87 (m, 1H), 1.75-1.64 (m, 2H), 1.64-1.40 (m, 6H), 1.40-1.06 (m,
7H), 0.88 (d, J = 6.7 Hz, 3H), 0.77 (d, J = 6.8 Hz, 3H). HRMS 683.3079 MNa+, calc.
for C33H48N4O8SNa 683.3085.
4-Amino-N-hexyl-N-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diaza-
bicyclo-[13.2.2]nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-benzenesulfonamide
8. 1H NMR (500 MHz, CD3OH) δ 7.94 (d, J = 9.6 Hz, 1H), 7.51 (d, J = 8.6 Hz, 2H),
7.17 (d, J = 8.8 Hz, 1H), 7.07 (d, J = 8.0 Hz, 2H), 6.77 (d, J = 8.0 Hz, 2H), 6.68 (d, J
= 8.6 Hz, 2H), 4.23 (m, 1H), 4.17-4.04 (m, 4H), 3.90 (m, 1H), 3.36-3.19 (m, solvent
obscured), 2.86 (dd, J = 14.6, 2.9 Hz, 1H), 2.41 (t, J = 12.3 Hz, 1H), 1.93-1.84 (m,
2H), 1.81-1.09 (m, 15H), 0.89 (d, J = 6.7 Hz, 3H), 0.75 (d, J = 6.7 Hz, 3H). HRMS
653.3335 MNa+, calc. for C33H50N4O6SNa 653.3343.
4-Amino-N-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diaza-bicyclo-
[13.2.2]-nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-N-[2-(4-methoxy-phenyl)-
ethyl]-benzenesulfonamide 9. 1H NMR (500 MHz, DMSO-d6) δ 7.74 (d, J = 9.6 Hz, 1H), 7.12 (d, J = 8.9 Hz, 1H),
7.70 (d, J = 8.8 Hz, 2H), 7.06, (d, J = 8.6 Hz, 2H), 7.03, (d, J = 8.6 Hz, 2H), 6.84, (d,
J = 8.6 Hz, 2H), 6.72, (d, J = 8.4 Hz, 2H), 6.60, (d, J = 8.7 Hz, 2H), 4.17 (m, 1H),
4.08 (m, 1H), 4.03 (dd, J = 9.0, 4.0 Hz 1H), 3.99 (m, 1H), 3.71 (s, 3H), 3.63 (m, 1H),
3.36 (m, 1H), 3.31 (m, 1H), 3.11 (m, 1H), 3.07 (m, 1H), 2.82 (dd, J = 14.1, 8.0 Hz,
1H), 2.69 (m, 2H), 2.33 (m, 1H), 2.16 (m, 1H), 1.91 (m, 1H), 1.76 (m, 1H), 1.64-1.49
(m, 2H), 1.35 (m, 1H), 1.27-0.96 (m, 3H), 0.77 (d, J = 6.7 Hz, 3H), 0.69 (d, J = 6.7
Hz, 3H). ESI-MS: 681.3 MH+.
4-Amino-N-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diaza-bicyclo-
[13.2.2]-nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-N-phenethyl-
benzenesulfonamide 10. 1H NMR (500 MHz, CD3OH) δ 7.59 (d, J = 8.8 Hz, 1H), 7.50 (d, J = 8.5 Hz, 2H),
7.31-7.23 (m, 2H), 7.22-7.12 (m, 3H), 7.09 (d, J = 8.0 Hz, 2H), 6.84 (d, J = 8.5 Hz,
1H), 6.79 (d, J = 8.0 Hz, 2H), 6.71 (d, J = 8.5 Hz, 2H), 4.27 (m, 1H), 4.19-4.06 (m,
91
3H), 3.81 (m, 1H), 3.53-3.43 (m, 2H), 3.19 (m, 1H), 2.99 (m, 1H), 2.94-2.79 (m, 2H),
2.43 (m, 1H), 2.21-2.11 (m, 2H), 2.03 (m, 2H), 1.88 (m, 1H), 1.77-1.44 (m, 4H),
1.40-1.07 (m, 5H), 0.88 ((d, J = 6.8 Hz, 3H), 0.77 (d, J = 6.7 Hz, 3H). HRMS
673.3028 MNa+, calc. for C35H46N4O6SNa 673.3030.
4-Amino-N-sec-butyl-N-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diaza-
bicyclo-[13.2.2]nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-benzenesulfonamide
11. 1H NMR (500 MHz, CD3OH) δ 7.88 (d, J = 9.8 Hz, 1H), 7.49 (d, J = 8.7 Hz, 2H),
7.15 (d, J = 9.2 Hz, 1H), 7.06 (d, J = 8.0 Hz, 2H), 6.76 (d, J = 8.3 Hz, 2H), 6.69 (d, J
= 8.7 Hz, 2H), 4.22 (m, 1H, H-3), 4.17-4.05 (m, 2H, H-3, H-13), 4.06 (dd, J = 9.0, 6.0
Hz, 1H, H-10), 3.76 (m, 1H, CHOH), 3.4-3.2 (m, solvent obscured), 2.98 (dd, J =
13.5, 8.0 Hz, 1H), 2.91 (dd, J = 14.7, 8.5 Hz, 1H), 2.84 (dd, J = 13.5, 6.8 Hz, 1H),
2.39 (t, J = 12.9 Hz, 1H), 2.21-2.10 (m, 2H), 1.97 (m, 1H), 1.84 (m, 1H), 1.75-1.63
(m, 2H), 1.52-1.40 (m, 1H), 1.38-1.10 (m, 3H), 0.91 (d, J = 6.6 Hz, 3H), 0.89-0.84
(m, 6H), 0.74 (d, J = 6.8 Hz, 3H). HRMS m/e 603.3193 MH+, calc. for C31H47N4O6S
603.3211.
4-Amino-N-cyclopentyl-N-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-
diaza-bicyclo[13.2.2]nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-
benzenesulfonamide 12. 1H NMR (500 MHz, CD3OH) δ 7.90 (d, J = 9.9 Hz, 1H), 7.51 (d, J = 8.7 Hz, 2H),
7.18 (d, J = 9.2 Hz, 1H), 7.07 (d, J = 8.0 Hz, 2H), 6.77 (d, J = 8.0 Hz, 2H), 6.72 (d, J
= 8.7 Hz, 2H), 4.22 (m, 1H), 4.16-4.04 (m, 2H), 4.07 (dd, J = 9.1, 6.0 Hz, 1H, Val-
αH), 3.75 (m, 1H, CHOH), 3.39 (dd, J = 14.5, 3.9 Hz, 1H), 3.25 (m, 1H), 3.18 (dd, J
= 13.6, 3.4 Hz, 1H), 3.07 (m, 1H), 2.94 (dd, J = 14.5, 8.6 Hz, 1H), 2.40 (t, J = 12.9
Hz, 1H), 2.21-2.09 (m, 2H), 1.85 (m, 1H), 1.74-1.63 (m, 2H), 1.60-1.40 (m, 3H),
1.38-1.10 (m, 11H), 0.90-0.84 (m, 6H), 0.76 (d, J = 6.8 Hz, 3H). HRMS 615.3182
MH+, calc. for C32H47N4O6S 615.3211.
3-Nitro-N-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diaza-bicyclo-
[13.2.2]-nonadeca-1(18),15,(19),16-trien-13S-yl)-ethyl]-N-(3-methyl-butyl)-
benzenesulfonamide 13.
92
1H NMR (500 MHz, CD3OD) δ 8.62 (t, J = 2.0 Hz, 1H), 8.50 (ddd, J = 8.0, 2.2, 1.0
Hz, 1H), 8.24 (d, J = 8.0 Hz, 1H), 7.87 (t, J = 8.0 Hz, 1H), 7.07 (d, J = 8.5 Hz, 2H),
6.79 (d, J = 8.5 Hz, 2H), 4.24 (m, 1H), 4.14 (m, 1H), 4.10-4.04 (m, 2H), 3.72 (m,
1H), 3.53-3.45 (m, 2H), 3.31-3.24 (m, solvent obscured), 3.21-3.14 (m, 2H), 2.40 (t, J
= 12.9 Hz, 1H), 2.21-2.10 (m, 2H), 1.87 (m, 1H), 1.75-1.64 (m, 2H), 1.62-1.08 (m,
8H), 0.90 (d, J = 6.5 Hz, 6H), 0.87 (d, J = 6.8 Hz, 3H), 0.77 (d, J = 6.8 Hz, 3H).
HRMS m/e 647.3066 MH+ calc. for C32H47N4O8S 647.3115.
3-Benzyl-1-[2R-hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diazabicyclo-
[13.2.2]-nonadeca-1(18),15(19),16-trien-13S-yl)-ethyl]-1-(3-methyl-butyl)-urea
14. 1H NMR (500 MHz, CD3OD) δ 7.34-7.19 (m, ), 7.05 (d, J = 8.2 Hz, 2H), 6.77 (d, J =
8.5 Hz, 2H), 4.37 and 4.36 (AB system JAB = 15.3 Hz, 2H), 4.23 (m, 1H), 4.14 (m,
1H), 4.08 (d, J = 6.2 Hz, 1H), 4.06 (m, 1H), 3.72 (m, 1H), 3.44-3.25 (m, solvent
obscured), 3.15 (dd, J = 13.7, 3.7 Hz, 1H), 2.41 (t, J = 13.2 Hz, 1H), 2.21-2.11 (m,
2H), 1.88 (m, 1H), 1.76-1.64 (m, 2H), 1.59 (m, 1H), 1.54-1.08 (m, 8H), 0.94 (d, J =
6.6 Hz, 3H), 0.89 (d, J = 6.8 Hz, 3H), 0.81 (d, J = 6.8 Hz, 3H). HRMS m/e 595.3804
MH+, calc. for C34H51N4O5 595.3859.
2-[3-[2R-Hydroxy-2-(10S-isopropyl-8,11-dioxo-2-oxa-9,12-diaza-
bicyclo[13.2.2]nonadeca-1(18),15(19),16-trien-13-yl)-ethyl]-3-(3-methyl-butyl)-
ureido]-3-methyl-butyric acid 15. 1H NMR (300 MHz, CD3CN) δ AA′XX′ system: 7.06 (m, 2H, JAX+JAX′ = 8.5 Hz,
ortho to CH2), 6.77 (m, 2H, JAX+JAX′ = 8.5 Hz, ortho to CH2), 6.45 (d, J = 9.7 Hz,
1H, H-12), 6.19 (d, J = 8.7 Hz, 1H, H-9), 6.13 (br, 1H, urea NH), 4.24 (ddd, J = 12.0,
6.0, 4.0 Hz, 1H, H-3), 4.15-3.98 (m, 3H, H-3, H-10, H-13), 3.69 (m, 1H, CHOH),
3.44-3.13 (m, 3H), 3.08 (dd, J = 13.7, 3.9 Hz, 1H), 2.45-2.00 (m, also H2O peak), 1.9-
1.78 (m, 1H), 1.71-1.00 (m), 0.98 (d, J = 6.9 Hz, 3H), 0.95 (d, J = 6.9 Hz, 3H), 0.92
(d, J = 6.5 Hz, 6H), 0.84 (d, J = 6.8 Hz, 3H), 0.76 (d, J = 6.8 Hz, 3H). 13C NMR
(CD3CN) δ 174.6, 172.5, 171.6, 161.1, 157.8, 131.6, 131.3, 117.0, 74.9, 68.4, 60.4,
57.7, 54.6, 52.3, 47.4, 37.6, 36.2, 35.7, 33.2, 30.7, 29.9, 26.8, 25.6, 25.6, 22.9, 22.8,
19.8, 19.7, 18.4, 18.1. ESI-MS m/e 605.4 MH+ calc. for C32H53N4O7 605.4.
93
2-(11S-{2-[Benzenesulfonyl-(3-methyl-butyl)-amino]-1R-hydroxy-ethyl}-6,9-dioxo-2-
oxa-7,10-diaza-bicyclo[11.2.2]heptadeca-1(16),13(17),14-trien-8S-yl)-acetamide
16. 1H NMR (500MHz, CDCl3) δ 6.55-7.79 (m, 13H, ArH, Asn-NH2, Asn-NH, Tyr-NH),
4.23 (m, 1H, OCH), 4.12-4.19 (m, 2H, OCH, Asn-αCH), 4.04 (m, 1H, Tyr-αCH),
3.67 (m, 1H, CHOH), 3.40 (m, 1H, CHN), 3.02-3.25 (m, 3H, NCH2, Tyr-βCH), 2.98
(m, 1H, CHN), 2.20-2.45 (m, 4H, Tyr-βCH, Asn-βCH2, CH(CO)), 1.82-2.08 (m, 3H,
CHC(O), CH2), 1.27-1.49 (m, 3H, CH, CH2), 0.79 (d, J = 7.7 Hz, 6H, (CH3)2). ESI-
MS 589 (M+H)+.
94
Chapter 4: Expression, Purification and Labelling of
Recombinant HIV-1 Rev and RBE3 RNA.
4.1 Introduction 4.1.1 HIV-1 Rev – Background.
4.1.2 The Functional Domains and Function of Rev.
4.1.3 Aims and Hypothesis.
4.1.4 The Problems of Labelling and Purifying RNA.
4.2 Results and Discussion 4.2.1 Cloning, Expression and Purification of HIV-1 Rev.
4.2.2 Refolding Experiments of HIV-1 Rev.
4.2.3 In Vitro Transcription, labelling and Purification of
RBE3 RNA.
4.3 Experimental 4.3.1 Constructs used and Transformation into JM109 Cells.
4.3.2 Restriction Digests.
4.3.3 Preparation of Electrocompetent E.coli for Protein
expression.
4.3.4 LB Antibiotic Media.
4.3.5 SDS-PAGE and Urea-PAGE Protocols.
4.3.6 Transfer Protocol for Western Blotting and Biotin
Detection.
4.3.7 Calculation of Protein Concentrations.
4.3.8 RP-HPLC and Mass Spectrometry.
95
4.1 Introduction. 4.1.1 HIV-1 Rev - Background.
Human Immunodeficiency Virus Type 1 Regulator of Virion Expression
(Rev) previously termed art or trs (Feinberg et al., 1986; Sodroski et al., 1986;
Perkins et al., 1989) is transcribed from two exons to form a variable sized (14-
20kDa) phosphorylated nuclear/nucleoli protein (Cullen et al., 1988; Goh et al., 1987;
Perkins et al., 1989; Hauber et al., 1988; Daefler et al., 1990). The Rev gene can also
be fused with tat (from the first exon), leading to a bifunctional protein called tev
without noticeable adverse effects on replication (Rosen and Pavlakis, 1990).
Rev was shown to be essential for viral replication since: (i) a provirus that
lacks Rev function is transcriptionally active but does not express viral late genes and
thus does not produce virions, suggesting Rev is essential for viral replication
(Dayton et al., 1986; Terwilliger et al., 1988; Sadaie et al., 1988; Malim et al., 1989a)
and (ii) a decrease in the rate of splicing of viral RNA down regulates Rev expression
levels and it seems the two processes are tightly regulated. Thus, inhibition of Rev
expression causes both an antagonism of Rev function (export) and a decrease in viral
production in vitro (Felber et al., 1990).
Rev is a basic protein, active over a large range of salt concentrations from
20mM to 500mM (Daefler et al., 1990), and interacts with a highly structured RNA
element known as the Rev response element (RRE) and was the first protein shown to
regulate nuclear export in a sequence-specific manner (Malim et al., 1989b). The
RRE is present in the unspliced (Pr160gag-pol), as well as the singly spliced (PR55gag)
RNA transcripts, but not in completely spliced transcripts containing tat, rev and nef.
Differential expression is then achieved when a mean threshold of Rev builds to
export late RNA transcript products containing the RRE (Emerman et al., 1989;
Felber et al., 1989). The RRE comprises 234 to 251 nucleotides of the env-coding
region (Malim et al., 1989b; Nakaya et al., 1997). Within the RRE a 37 nucleotide
sequence has been found to be the minimum region required to interact with Rev
(Cook et al., 1991). This region termed "the Rev high affinity binding site,” RBEIII
or RBE3 (contained within stem loop II of the RRE), is a double-stranded RNA helix
containing a "bubble" formed by non-Watson and Crick or Hoogsteen basepairing
(G:A and G:G) (Charpentier et al., 1997; Bartel et al., 1991).
There have been numerous structural studies involving Rev or the RRE,
including native protein studies of Rev (Scanlon et al., 1995; Watts et al., 1998;
Blanco et al., 2001) and native studies of the RRE/RBE3 (Peterson et al., 1994;
Ippolito and Steitz 2000; Hung et al., 2000). However, the most important structural
studies utilise a fragment of Rev (Rev34-50) complexed to the RBE3 (Battiste et al.,
1995; Ye et al., 1996; Charpentier et al., 1997; Grate and Wilson, 1997). These
structural studies show the deep penetration of Rev through the RRE bubble,
neutralising charge on both Rev and the RRE and disrupting the very specific
Hoogsteen interactions to distort the RNA major groove to form the Rev-RRE
complex as shown in Figure 4.1. More recent structural work utilises Rev34-50 derived
peptides, mutated to increase helicity and form a stronger interaction with the RBE3
RNA, but does not elucidate further information about the native peptide interaction
to the RRE (Zhang et al., 2001).
Figure 4.1: Structure of Rev34-50 interacting with Stem Loop II of the RRE. The α-helical rev34-50
(centre yellow) can be seen interacting with a “bubble” in stem loop II of the RRE. It can be seen that
the localised RNA structure is perturbed about the “bubble” due partially to G:A and G:G pairing, but
also binding to Rev34-50 (Image created using Accelry ViewerLite (co-ordinates Battiste et al., 1995
pdb 1ETF)).
96
97
4.1.2 The Functional Domains and Function of Rev.
Rev has been shown to contain at least four functional domains. (1) An
arginine-rich RNA binding domain (Malim et al., 1989a). (2) A nucleoli localisation
domain within the arginine-rich binding domain (Fischer et al., 1995). (3) A
multimerisation domain - Rev is believed to form a homotetramer in solution (Zapp et
al., 1991; Hope et al., 1990). (4) An effector domain - that is a specific nuclear export
signal (NES). The N-terminal region of Rev contains the first three domains (RNA
binding domain, multimerisation domain and the nucleoli localisation domain) as
well as a phosphorylation site (Malim et al., 1989a; Malim and Cullen, 1991; Berger
et al., 1991; Fischer et al., 1994). The C-terminal domain, termed the activation or
effector domain, has been suggested to interact with many cellular factors (Fischer et
al., 1994; Fischer et al., 1995; Askjaer et al., 1998).
There are four major variants of Rev that are studied; a consensus sequence
and sequence of the Rev construct used in this study are shown in figure 4.2. The
phosphorylation of Rev on Serine 5 and 8 (numbering taken from the consensus
sequence) was initially reported not to be required for Rev function (Malim et al.,
1989a), though a subsequent study has shown phosphorylated Rev has a greater
interaction with the RRE, possibly due to an increase in the helical content of the
RNA binding site (Fouts et al., 1997).
The exact means by which Rev functions to transport the RRE and what
cellular proteins/pathways Rev exploits have not been adequately established and are
contentious in the literature. It is also not understood why such complex Rev-
mediated regulation exists, though a suspected role in latency may assist in immune
escape and contribute to the complicated pathology of infection (Pomerantz et al.,
1990; Michael et al., 1991; Garcia and Cullen 1991). Rev appears to perform a dual
role in the HIV-1 replicative cycle, first through binding to the RRE, Rev decreases
the affinity between spliceosomal proteins for inhibitory sequences on the RRE
(Mikaelian et al., 1996), allowing stabilisation of the RRE within the nucleoli
(Schneider et al., 1997). Second the NES allows the Rev:RRE complex to be a
substrate for nuclear export, and once in the cytoplasm the RNA-Rev-transport
protein complex then dissociates, allowing the RNA to be translated into proteins, and
the transport protein and Rev to re-enter the nucleus. This consensus is summarised in
figure 4.3.
MAGRSGDSDE10NLLKAIRLIK20FLYQSSPPSP30EGTRQARRNR40RRRWRARQRQ50IHSIGERII
C60TFLGRPEEPV70PLQLPPLERL80NLNCSEDCGT90SGTQGVGSPQ100IPVEPPAVLE110SGTEE
MRRASVAGRS10GDSDEELIR20TVRLIKLLYQ30SNPPPNPEGT40RQARRNRRRR50WRERQRQI
HS60ISERILSTYL70GRSAEPVPLQ80LPPLERLTLD90CNEDCGTSGT100QGVGSPQILV110ESPTVLE
SGA120KEHHHHHH128
Figure 4.2: The Sequence of HIV-1 Rev. The top sequence is constructed as a consensus sequence based on known gene isolates from Genbank and follows the standard numbering nomenclature used within the literature. The lower sequence is that used in this study (Cochrane et al., 1989; Jensen et al., 1997; Jensen et al., 1995) and corresponds to the consensus sequence from residue 7 and excluding the poly-His tag from residue 123. The consensus sequence numbering is adopted as the nomenclature throughout the remainder of the thesis. The Light blue colouring indicates the nucleoli localisation sequence (17mer), which also contains the bold blue RRE-binding domain. The red is the trans-dominant nuclear export signal region and the green region has been found to be functionally redundant. The minimal functional unit of Rev has been shown to be the first 92 nucleotides of the consensus sequence, which is sufficient to confer interactions with the RRE and export viral mRNA. The pink residues are a known splicing point (Perkins et al., 1989). The orange and yellow sequences are a Heart Muscle Kinase site and poly-His tag for P32 labelling and protein purification respectively.
Cytoplasm RRE
Nucleus
gag-pol/pol
Nucleolus dissociation of RNA
Figure 4.3: The In Vitro Cycling of Rev. Within the nucleolus, Rev binds to a secondary structure in the viral pre-mRNA (RRE) rescuing the RRE from splicing. Rev further multimerises on the RRE facilitating the binding of an mRNA nuclear export protein (a debated cellular factor) to initiate export of the Rev:RRE complex (possibly through 5S rRNA and snRNA pathways) (Woodrich and Krausslich, 2001; Askjaer et al., 1998; Fridell et al., 1997; Fischer et al., 1995). The exported complex then dissociates and the RRE is translated to the gag-pol and pol proteins to be further processed by HIV-1 PR (not shown). The cellular factor and Rev re-enter the nucleus to reinitiate the cycle. Discrete functional domains within Rev mediate the interactions with cellular proteins and viral RNA for nuclear localization, RNA binding, multimerisation and nuclear export. The late, structural phase of gene expression is dependent on the presence of Rev. In the absence of Rev there is an accumulation of small, fully spliced mRNAs and no production of infectious virions.
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RRE binding domain and Nucleoli Localisation Domain. The sequence that directs
Rev to the nucleus (Nuclear Localisation Signal or NLS) is contained within the
RRE-binding domain (17mer, Rev34-50). All NMR studies of Rev:RBE3 complexes
have been conducted using this 17mer or a derivative thereof and not the complete
Rev protein (Battiste et al., 1995; Ye et al., 1996; Zhang et al., 2001). A helical
content between 11-57% has been reported for this 17mer alone based on circular
dichroism studies, with an apparent Kd for RRE of 10-20nM compared to 1-10nM
reported for the complete Rev protein (Kjems et al., 1992). The secondary structure
of the RRE is more important for recognition by this portion of Rev than the RRE
nucleotide sequence itself since mutations which do not disrupt RNA secondary
structure do not affect activity (Bartel et al., 1991; Heapy et al., 1990; Dayton et al.,
1989). The sequence of the 17mer predominantly localises Rev to the nucleus, and
more specifically, the nucleoli (Cullen et al., 1988; Rosen et al., 1988; Felber et al.,
1989; Malim et al., 1989a). This signal sequence contains an arginine rich motif -
NRRRRW (Perkins et al., 1989) - that forms part of the high affinity RNA binding
domain, and is homologous to known NLSs (Dingwall and Laskey, 1986; Malim et
al., 1989b).
Although it has been demonstrated that the NLS allowed a fusion protein of
rev40-45-β-galactosidase to accumulate in the nucleus (Perkins et al., 1989), additional
sequences or even Rev secondary structure about the NLS are probably important for
the function of nucleolar localisation. Furthermore, the removal of the arginine-rich
sequence within the NLS alone causes retention in the cytoplasm (Perkins et al.,
1989). The NLS also comprises a significant portion of the RRE-binding domain.
NMR structural studies indicate that the NLS is "shielded” by formation of the
complex with the RRE, enabling export of this complex (Battiste et al., 1995; Ye et
al., 1996; Charpentier et al., 1997; Grate and Wilson, 1997).
Multimerisation Domain. Rev multimerises on the RRE, forming aggregates
believed to be 3 on stem loop II (the Rev high affinity binding site) and 8 over the
entire RRE structure. Multimerisation is important to Rev function, and a mean
threshold of Rev concentration has been suggested to be necessary for efficient export
(Cook et al., 1991; Kjems et al., 1991).
The NES Domain. The C-terminus contains a 35 amino acid sequence
(comprised of a highly basic, leucine rich sequence), which functions as a trans-
100
acting gene product and is an NES (Fischer et al., 1995). This region is believed to
bind cellular cofactors that increase the rate of export into the cytoplasm, and possibly
include the debated cellular export protein (Rosen, 1992; Tronto and Baltimore,
1990). Mutations within the NES domain create a trans-dominant negative
phenotype, inhibiting Rev function by disabling transport to the cytoplasm (Malim et
al., 1989a; Malim et al., 1991; Fischer et al., 1994). However, the minimal functional
unit of Rev has been shown to be the first 92 nucleotides, which is sufficient to confer
interactions with the RRE and export viral mRNA (Perkins et al., 1989).
4.1.3 Aims and Hypothesis.
The 3rd aim of the project was the development of an RNA-binding assay to
test prospective Rev antagonists in vitro. It was hypothesised that recombinant
protein and RNA could be used to develop a new surface plasmon resonance (SPR)
assay to screen potentially useful drugs. This chapter describes the expression,
purification and labelling of material suitable to develop such an assay.
4.1.4 The Problems of Labelling and Purifying RNA.
The major problem with developing RNA-based assays for large libraries is
the problems of economically labelling and purifying RNA at scale (Wyatt et al.,
1991). Generally the method of choice for labelling RNA is random incorporation
through in vitro transcription using radioactive labels, or specifically during chemical
synthesis using nucleotide tags (ie a nucleotide modified with biotin, a chromogen or
other common detectable derivatives). Following labelling, the RNA is purified to
remove unincorporated nucleotides and labels by precipitation and/or
chromatography (Cunningham and Lu, 1996). For a BIAcore-based assay relying on
the capture of RNA, the most suitable assay system with respect to immobilisation
chemistry would utilise Streptavidin as the ligand and capturing the RNA to the
surface by a biotin label. For this project it was not economically viable to purchase
chemically synthesised and labelled RNA and so it was necessary to label RNA
produced from in vitro transcription kits.
The other option to labelling RNA is by enzymatic incorporation of the label,
but unlike DNA, that is simple to synthesise and label using end-labelling enzymes
such as DNA ligase or terminal deoxynucleotidyl transferase, RNA is difficult to end-
101
label, is more susceptible to degradation and difficult to purify at scale (Rosemeyer et
al., 1994). These difficulties can be largely attributed to the different physical
properties of RNA that allow RNA to adopt a greater number of secondary and
tertiary structures and even form different functional units in some cases (Conn and
Draper, 1998; Tinoco and Bustamante, 1999). Because of these differences, general
methods need to be adjusted in individual RNA purifications to optimise the purity
and yield. The present study required the development of more consistent and reliable
means of purifying labelled RBE3 RNA product from large scale in vitro
transcription reactions.
Generally most RNA purifications utilise the physical and chemical properties
of RNA and rely on a three-step strategy consisting of, (1) The inactivation of
RNases, (2) the solution-phase partitioning of RNA from contaminants and (3) the
selective precipitation of either the contaminants or RNA.
The inactivation of RNases is difficult since, unlike DNases, RNases do not
require metal ions for activity (therefore chelating agents such as EDTA are
ineffectual) (Jarrous 2002). RNA also has the disadvantage of nucleophilic self and
cross cleavage caused by the 2’ hydroxyl group adjacent to the phosphodiester
linkages acting as a reactive species in solution (Chetverin et al., 1997). Safest
methods of inactivation are those that use guanidinium ions and have the added
advantage of denaturing all proteins in the solution. Guanidinium was first used in
buffers to isolate RNA by Cox (1968), and subsequently by Nozaki and Tanford
(1970) and Gordon (1972), who demonstrated guanidinium’s excellent protein
denaturing ability. Indeed guanidinium chloride is 2-3 fold/mol better at denaturing
proteins than other denaturants such as urea (Monera et al., 1994), and thus
guanidinium replaced phenol extractions as a first step for RNA purification.
The next step in RNA purifications is partitioning, which separates the RNA
from all other species based on the differing physical or chemical properties. Since
DNA is typically soluble above ~pH 6.7, buffers that are more acidic than this can be
used for partitioning high amounts of DNA and RNA and it has been reported that as
much as 5-10mg of nucleic acid can be present in large scale transcription reactions
(Promega, 1999). For partitioning denatured proteins from the RNA, Feramisco et al.,
(1982) revived the use of phenol in developing the first rapid procedure for RNA
purification. Their method involved combining a guanidinium and hot phenol step
102
that was systematically optimised by others (Chomczynski and Sacchi 1987; Meltzer,
1990) to develop a single-step method of RNA extraction, which method evolved to a
kit form that is marketed by major laboratory supply companies. Such single-step
methods often include isoamyl alcohol/chloroform. The isoamyl alcohol acts as a
defoaming agent, which prevents nucleic acid shearing and improves solution
separation at the interphase from the phenol and chloroform organic solvents
(Chomczynski and Sacchi, 1987).
The final step is then the precipitation of materials from the aqueous phase.
Biological precipitation is dependent on pH, dehydrant used, and the type and
concentration of the salt used to facilitate supersaturation. Typically, ammonium,
sodium or lithium ions are chosen depending on how tight a pellet of RNA is desired
to facilitate subsequent washing steps with 70% ethanol (Cathala et al., 1983; Lev
1987; Osterburg et al., 1975, Chang et al., 1990), but there is no absolute agreement
on the monovalent salt concentration and can range from 0.1 - 0.5M depending on the
size of the RNA molecule(s) (Wilkinson, 1991).
RNA precipitation is often facilitated using dehydrants such as ethanol or
isopropanol. All precipitations (of any chemical/biological material) should be
conducted progressively over time to maximise yield and efficiency, since as much as
40% of the yield can be lost from insufficient precipitation times and conditions
(Wilkinson, 1991). A “seed” should be created at the solvent interface and gently
mixed into the solution, dispersing the solvent and facilitating precipitation in a slow
and ordered manner under temperature control until supersaturation is approached.
Once supersaturation is achieved precipitation should then proceed rapidly for an
extended period (dependent on the volume and size of the RNA). Precipitation should
never be conducted at –80°C as the yield is poor since the solution freezes before
precipitation occurs (Wilkinson, 1991).
4.2 Results and Discussion 4.2.1 Cloning, Expression and Purification of HIV-1 Rev.
Plasmids of Rev and RRE-containing constructs were obtained from Dr
Jørgen Kjems (University of Aarhus, Denmark) and were propagated in JM109 cells
(Experimental section 4.3.1). The protein-encoding plasmid constructs provided are
designated pET-His-Rev-HMK and pET-HMK-Rev-His, generating N- or C-
103
terminally labelled poly-His tagged Rev respectively. In order to maximise protein
yield the two constructs were introduced in parallel transformations into two separate
cell lines (two transformations per cell line, or a total of four transformations)
(Experimental section 4.3.1 – 4.3.5). The expression vectors chosen were
BL21(DE3)pLysS (Promega) and AD494(DE3) (Novagen), and both small and large
scale cultures were grown of each plasmid in both cell lines to determine the best
construct and conditions for expression and purification of Rev.
For small-scale experiments, 2mL of overnight cultures in LB-selection media
(37°C, 250rpm) were pelleted (12,000 x g, 5min, 4°C) and inoculated into 200mL
LB-selection media (37°C, 200rpm). The expression cultures were grown to an OD600
greater than 0.67 and induced using either 1mM or 2mM IPTG and were harvested 4
– 6h post induction in a Savant centrifuge (3500rpm, 15min, 4°C). Pellets were stored
frozen (12 – 16h) at –20°C. The frozen pellets were thawed on ice and resuspended in
5mLs denaturation lysis buffer (100mM sodium phosphate, 10mM Tris pH 8.0, 6M
Gu·HCl) and lysed by vortexing.
The viscosity of the lysates was reduced with sonication using a Coleman
Palmer Torbéo ultrasonic cell disruptor 36810-series (7w, 20sec), then cleared cell
lysates prepared (10,000 x g, 30min 22°C) and incubated on an agitating table in the
presence of 100µL of Ni-NTA Superflow (Qiagen) for 60min. The lysate-Ni-NTA
slurry was loaded onto Biorad narrow bore disposable 5mL columns and washed (50
column vols for each wash) with pH changes of buffer B (100mM Na·H2PO4, 10mM
Tris, 8M urea) through the series pH 8, pH 6.3 and pH 5.8. His-tagged protein eluted
with buffer B at pH 4.5 and pH 2. Fractions were collected and analysed
spectrophotometrically for the presence of protein, yielding highest total protein
concentrations from the pH 4.5 elutions using either the BL21(DE3) or
AD494(DE3)pLysS. Both low pH samples were neutralised with 10 volumes of Tris
buffered saline (TBS – 10mM Tris (pH 8), 150mM NaCl) and applied to a Western
blot (Experimental section 4.3.5 and 4.3.6). A significant single band was visible for
the proteins on the Western blot of the C-terminal constructs, whereas, the N-terminal
constructs produced only a very faint band (Fig. 4.4). In comparing the best cell line
and construct, the AD494(DE3) cells were found to produce the greatest amount of
recombinant protein of both N- and C-terminally labelled protein, but more total
protein was produced using either cell line and the C-terminal construct, with little
104
discernible difference between 1mM or 2mM IPTG and therefore the AD494(DE3)-
pET-HMK-Rev-His transformant was used for protein expression.
For large scale production, a 5mL overnight culture of AD494(DE3)-pET-
HMK-Rev-His was grown in LB-selection media (37°C, 225rpm) and the cell pellet
(14,000 x g, 5min, 4°C) was resuspended in 2 x 400mLs selection media and grown
to an OD600 of 0.74 (37°C, 160rpm) and induced with 1mM IPTG. The expression
was harvested 6h post induction (3500rpm, 15min, 4°C) and the pellets stored
overnight (-20°C). The cells were lysed and sonicated as for small-scale production
described above in 50mLs of denaturation lysis buffer. A clear cell lysate was
prepared (12,000 x g, 30min, 18°C) and incubated on a table agitator for 1h in the
presence of 3.2mL Ni-NTA resin (QIAGEN Superflow).
The buffer B wash and elution buffer was as described, except the washing
and elution conditions were pH 6.38, pH 5.95 (wash) and pH 4.48, pH 2.0 (elution
~11mLs in total). Both washing and elutions were conducted while monitoring the
absorbance changes at 280nm and 320nm until no more protein could be seen
spectrophotometrically (>10 column volumes for washes ~20 mLs/wash), with the
strongest protein profile found in the pH 4.48 samples (Fig. 4.5). Purifications yielded
as much as ~22mgs of Rev from the large scale cultures (800mL) in the pH 4.48 and
pH 2.0 samples. It is interesting that Rev can be seen eluting early in the pH 6.38 and
pH 5.91 samples, suggesting the resin may be saturated (maximum binding capacity
of the Ni-NTA resin is 8mg/mL), if this is the case, the expression level for Rev could
be greater than 32mg/L.
The pH 4.48 material from primary Ni-NTA purifications was assessed using
image quantitation software (ImageQuant™ – Amersham Biosciences) where the
pixels from a scanned image of the protein band of interest are quantitated in relation
to the total protein in the well. Using this method the purity was shown to be greater
than 90% pure (Figure 4.6). To confirm this a RP-HPLC purification was conducted
using a C-8 Vydac column (Ø20mm x 250mm) (Experimental Section 4.3.8) showing
a major species at 31.5min and several minor species present (Fig 4.7). The peak
integration confirmed the image quantitation assessment that primary isolation was
~90% pure.
The Rev gene used to derive the Kjems construct was provided to the Kjems
lab from an external source and has been reported to produce recombinant His-tagged
Rev that migrated at slightly more than its 14kDa mass (~21.5kDa) (Cochrane et al.,
1989), as observed here (Fig 4.5).
Figure 4.4: Western Blot Comparison of different Constructs in different cells. The N-terminal constructs in AD494(DE3) cells (lanes 1 – 4) can be seen producing far less protein (barely visible) than the C-terminal constructs in BL21(DE3)pLysS cells (lanes 5 – 10), with little difference between IPTG concentrations (1mM lanes 1, 2, 5, 6, 9 and 2mM lanes 3, 4, 7, 8, 10). The AD494(DE3) cells produced far more protein from both N- and C- terminal constructs (not shown in figure).
1 2 3 4 5 6 7 8 9 10
MW pH 6.38 5.91 4.48 2.0 MW pH 4.48
Figure 4.5: SDS-PAGE and Western Transfer of the Ni-NTA purification. The lanes of a 12.8%
SDS-PAGE (left) show wash and elutions from a scaled-up purification of Rev expressed from the
pET-HMK-Rev-His (C-terminal) construct, grown in AD494(DE3) cells. Rev began eluting from pH
6.38, with essentially no contaminants visible until the pH 4.48 elution. The Western transfer (right)
used a primary Mouse-anti-His antibody that was detected with a secondary anti-mouse-alkaline
phosphatase antibody (Experimental section 4.3.6), confirming the purified protein is His-tagged. The
prestained MW marker (Bio-rad) comprised; Phosphorylase B (107kDa), BSA (76kDa), Ovalbumin
(52kDa), Carbonic Anhydrase (36.8kDa), Soybean Trypsin Inhibitor (27.2kDa) and Lysozyme
(19kDa). The recombinant Rev migrated with an apparent MW of 20 – 22kDa.
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Figure 4.6: Image Quantitation Analysis. Samples of Rev from the pH 4.48 elution loaded on an SDS-PAGE and stained with Coomassie Blue. The boxes 1, 4 and 6 (the Rev protein) were quantitated against the remainder of the wells (2, 5, 7) and corrected for the background (3). Purity was found to be 92% for the lane labelled 2, 93% for 5 and 96% for 7.
5 7
The protein was exposed to stronger denaturing and reducing conditions but
would not migrate at less than the 19kDa marker. In order to resolve this mass
anomaly, LC-MS was conducted with two species found close to the expected mass
of 14467.3Da (14486.6 and 14931.4) (Fig. 4.7), but no species was present above
these masses contrary to SDS-PAGE. It is possible the first peak (14486.6amu) is
possibly the mass +1 sodium ion as Rev was loaded onto the column in a high salt
buffer. The second differs possibly due to TFA ions (+4 TFA ions) at 14931.4amu
from the expected mass, though it is not possible with this data to draw definitive
conclusions and it is possible a combination of many ions could account for the
difference. By mass spectrometry, no species was present above these masses as
suggested by SDS-PAGE (~21.5kDa) and therefore it is plausible that the Rev protein
used here has an intrinsic anomalous migration in SDS-PAGE gels. In order to
unequivocally rule out protein sequence differences, possible modification and
confirm the protein sequence, protease footprinting by mass spectrometry or DNA
sequencing would be advisable for future work.
Figure 4.7: HPLC profile and Mass spectrometry of crude Rev. The RP-HPLC profile of the pH
4.48 elution following Ni-NTA affinity purification and dialysis (left) with the major protein peak at
31.5min. The deconvoluted mass spectrum (right) of proteins from the 31.5min fraction shows a major
peak of 14931.4amu as well as a minor peak of 14486.6amu.
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4.2.2 Refolding Experiments of HIV-1 Rev.
Denatured Rev has previously been refolded experimentally from denatured
and lyophilised states (Tange et al., 1996; Cochrane et al., 1989). A variety of
different refolding methods, buffers and conditions were tested here as either a
refolding experiment or in order to change conditions by ion exchange
chromatography to try to further purify the protein, however no improvement to
purity could be made except by RP-HPLC. Ideal refolding conditions were found to
follow serial dialysis and temperature change through the following pathway:
1. Room temperature dialysis into 50mM Hepes (pH 7.4), 6M Gu.HCl, 1mM
EDTA for 1h to standardise the pH of the solutions and exchange the urea for
guanidinium to avoid urea precipitating at 4°C in subsequent steps.
2. Dialysis into 50mM Hepes (pH 7.4), 4M Gu.HCl, 500mM NaCl, 0.5mM
EDTA, 10% Glycerol for a minimum of 1h at room temperature before
centrifugation of any precipitated material (10,000 x g, R.T. 10min).
3. Dialysis into 50mM Hepes (pH 7.4), 2M Gu.HCl, 500mM NaCl, 0.5mM
EDTA, 10% Glycerol for 2h at 4°C.
4. Dialysis for a minimum of 4h at 4°C in 50mM Hepes (pH 7.4), 150mM
Gu.HCl, 500mM NaCl, 0.5mM EDTA, 10% Glycerol and recentrifugation of
this material (10,000 x g, 10min, 4°C).
5. Final dialysis in 50mM Hepes (pH 7.4), 500mM NaCl, 10% Glycerol O/N at
4°C for use the following day or storage for less than 48h.
It was found that dialysis was more successful if the protein concentration was
less than 1mg/mL. If salt concentrations below 500mM were desired, such conditions
had to be approached by dialysis through decreasing salt concentrations from the final
500mM buffer and it was necessary to use <0.5mg/mL protein in order to reduce
precipitation at lower salt concentrations, with 300mM being generally the lowest
limit. Rapid refolding (where a small protein volume is diluted into a larger buffer)
was tested through dilution from 50mM Hepes (pH 7.4), 2M Gu.HCl, 500mM NaCl,
0.5mM EDTA, 10% Glycerol at 4°C with no less than a 10-fold factor and then
concentration and rapid buffer exchange using Microcon YM-10 concentrators. It was
found that rapid methods were not as good as dialysis and protein concentrations had
to be kept below 0.1mg/mL for sufficient recovery starting from protein
108
concentrations of 0.5mg/mL. Typical conditions were 10 - 25-fold dilution initially
and then further dialysis or dilution (later BIAcore experiments in Chapter 5 showed
up to 1/5000 dilution was acceptable from 0.5mg/mL stock and a further 1/10 dilution
of this sample was detectable in the BIAcore). In some experiments concentration of
the protein was effected using YM-10 concentrators and protein would consistently
precipitate when exceeding 2mg/mL.
Long term protein storage was in 50mM Hepes (pH 7.4), 6M Gu.HCl (4°C) or
50mM Hepes (pH 7.4), 4M Gu.HCl, 500mM NaCl, 0.5mM EDTA, 10% v/v glycerol
(4°C) for no greater than 3 months. Experiments were planned well in advance and
protein was stored in 50mM Hepes (pH 7.4), 150mM Gu.HCl, 500mM NaCl, 0.5mM
EDTA, 10% v/v glycerol (4°C) for up to one week. The final dialysis buffer
conditions (50mM Hepes (pH 7.4), 500mM NaCl, 10% v/v glycerol, 4°C) were not
used beyond 48 hours as material would begin to precipitate beyond this time.
Storage conditions tested at –20°C consistently led to precipitation and only
lyophilised protein from the analytical RP-HPLC purifications was stored at -80°C.
In conclusion, the expression and purification of HIV-1 Rev from the C-
terminal plasmid construct of pET-HMK-Rev-His grown in AD494(DE3) cells was
the most efficient expression system, producing in large quantities (~22mg/800mL of
culture) from induction of cultures with 1mM IPTG at an OD600 of 0.76. The cells
were lysed under standard denaturing conditions for affinity purification using Ni-
NTA Superflow resin and primary purifications yielded protein of ~90% purity.
Denatured purification worked well, recovering large quantities of protein, of high
purity, and after dialysis provided simplified storage conditions.
4.2.3 In Vitro Transcription, labelling and Purification of RBE3 RNA.
The RBE3 plasmid construct provided by the Kjems lab was propagated in the
JM109 cells (Experimental section 4.3.1) and grown to saturation levels (OD600 ~2.0)
before purification using QIAGEN MAXIprep DNA kits as per suppliers protocol.
The purified DNA (A260/280 not <2.0) was digested using the restriction enzyme Dra I
to produce linearised, blunt ended DNA (blunt ended DNA is necessary to avoid a
form of rolling circle transcription resulting in very large transcripts or T7 RNA
polymerase incomplete termination and dissociation in the presence of the –ve sense
strand overhangs) and this material was used as the template to synthesise RNA under
control of the T7 promoter (Fig 4.8).
Small-scale production of RNA was carried out using standard protocols
supplied with the kits (Promega RiboPROBE) under the following conditions; 40mM
Tris-HCl (pH 7.5), 6mM MgCl2, 2mM spermidine, 10mM NaCl, 10mM DTT, 0.5mM
each of the rNTPs, 1,000U/ml Promega RNasin® Ribonuclease Inhibitor, 400U/ml T7
RNA polymerase, 50-100µg/ml DNA template. Synthesis was conducted for 2 hours
at 37°C and typically yielded purified RNA no greater than the DNA template
concentrations; further addition of template did not increase the yields in contrast to
recommendations (Dasso and Jackson, 1989). At the time the work was conducted,
kit advice for purification was a DNase digestion step followed by ethanol/propanol
precipitation and washing, though it is now advised to conduct anion column
purification (Promega, 1999). Initially purification of small RNA transcription
reactions was conducted as per kit instructions using DNase digestion, precipitation
and later modified with anion chromatography. These methods failed to produce
suitably pure RNA and purification was improved using an extraction solution
(500µL citrate buffered phenol (pH 4.0), 85µL 3M sodium acetate (pH 4.6) and
500µL 1-bromo-2-chloropropane at 4°C), followed by precipitation of the upper
aqueous phase with 1.2 volumes propan-2-ol. This protocol was optimal for solution
volumes no greater than 500µL for the RiboPROBE kit, resulting in A260/A280 ratios
>2.0 and A260/A230 ratios within the range of 1.5 – 2.
GTATGGGCGCAGCGTCAATGACGCTGACGGTACAGGCCCCTTTTTAAA
CATACCCCCGTCGCAGTTACTGCGACTGCCATGTCCGGGGAAAAATTT
Dra I site T7 RNApol Transcription
T7 PROMOTER
Figure 4.8: The in vitro transcription reaction. The DNA template is cleaved with the restriction
enzyme Dra I at the palindromic sequence TTT∨AAA, resulting in blunt-ended DNA. T7 RNA
polymerase transcribes the RNA yielding the sequence from the promoter to the blunt end. For the
RBE3 fragment the sequence was 45 nucleotides;
5’-GUAUGGGCGCAGCGUCAAUGACGCUGACGGUACAGGCCCCUUUUU-3’.
Labelling the 3’-ends of RNA strands was attempted using T4-RNA ligase.
Briefly, the 5’-ends of the RNA strands were modified using calf intestinal alkaline
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phosphatase (Boehringer Mannheim) in order to remove the phosphates present on
the 5’ ends and prevent 5’ – 3’ ligation of template strands. The alkaline phosphatase
reactions were conducted using 120 – 6000pmol RNA and 0.01U enzyme/pmol of
RNA (600µL volume reactions in 50mM Tris.HCl (pH 9.3), 1mM MgCl2, 0.1mM
ZnCl2, 1mM spermidine). The Calf intestinal alkaline phosphatase was inactivated
with 6 volumes of stop buffer (10mM sodium citrate (pH 4.0), 5mM EDTA, 0.5%
SDS) and heating the solution to 65°C. The RNA was extracted with the extraction
solution described above. The modified RBE3 RNA was resuspended in the T4-RNA
ligase reaction buffer (Promega) (50mM Tris (pH 7.8), 10mM mgCl2, 5mM DTT and
1mM ATP) to 10µmol concentration. Biotin-16-UTP was added at 1/10 the
concentration of the RNA target with 0.16U of the enzyme/µL reaction volume
(120µL reaction). The labelled RNA was purified as described above and visualised
by first running the material on 18% urea-PAGE gels before transferring to
nitrocellulose membranes and detection using CDP-StarTM and Streptavidin-alkaline
phosphatase antibodies as described in the Experimental section 4.3.5 and 4.3.6 (Fig
4.9). As can be seen in figure 4.9, the labelling by RNA ligation was inefficient, with
larger sized labelled RNA present than was expected. It is possible this method of
labelling failed due to incomplete digestion of 5’ phosphate ends by calf intestinal
alkaline phosphatase leading to 5’ – 3’ ligation of template strands and hence the
larger biotin-labelled products seen (self ligation to form circular structures has also
likely occurred but is not detectable on the transfer as the 3’ ends would have been no
longer available for ligation to biotin-16-UTP).
higher mw RNA than expected expected position for labelled RNA electrophoretic front containing unincorporated label.
Figure 4.9: The Endlabelling of RNA using T4 RNA Ligase. The RNA was ran on an 18% urea gel and transferred to a nitrocellulose membrane for detection using a Streptavidin-alkaline phosphatase conjugate as described (Experimental section 4.3.5 and 4.3.6). Labelled material is present just below the well migratory point or in the electrophoretic front (unincorporated label). No labelled RNA of the correct size is detected. The mobility was estimated using a biotin-labelled DNA primer provided by Dr Marcus Hastie (QUT) of 35 nucleotides in length (not shown).
Labelling 3’ ends of the RNA strands was also attempted using terminal
deoxynucleotidyltransferase (Boehringer Mannheim) following published methods
110
(Rosemeyer et al., 1994), but yielded insufficient labelled RNA (<0.01% yield, data
not shown). A more general method of labelling incorporated the biotin label into the
in vitro reaction using RiboMAX in vitro transcription kits (Promega), RiboMAX kits
produced far more amounts of transcribed RNA than added DNA template (in
contrast to the RiboPROBE kit) due to the improvements to the buffer and buffer
additives of the reactions based on the work of Gurevich et al., (1991) and
Cunningham and Ofengand (1990).
The reactions were conducted as follows: 80mM HEPES-KOH (pH 7.5),
12mM MgCl2, 2mM spermidine, 40mM DTT, 25mM each of the rNTPs (with UTP
constituted by biotin-16-UTP and UTP at a ratio of 1:11), 1,000U/ml RNasin®
Ribonuclease Inhibitor, 5U/ml yeast inorganic pyrophosphatase, 1,800U/ml T7 RNA
polymerase, 100µg/ml template DNA. Transcription was conducted over a 4-hour
period (2 hours at 37°C then a further 1,800U/ml of T7 RNA polymerase was added
and incubation continued for 2 more hours). The RNA was extracted using the
extraction solution as for small-scale productions and though this worked very well
on a small scale, on a large scale the preparations were found to contain significant
carryover of biotin-16-UTP and possibly incompletely transcribed biotin labelled
RNA as a contaminant (Fig 4.10). An attempt was made to remove lower molecular
weight contaminants using YM-10 concentrators, but this was unsuccessful.
Figure 4.10: Extracted and Precipitated Labelled RNA. The RNA
was run on an 18% urea gel and transferred to a nitrocellulose
membrane for detection using a Streptavidin-alkaline phosphatase
conjugate as described in Experimental section 4.3.5 and 4.3.6. The
labelled RNA (upper bands of the gel) can be seen contaminated with
either labelled but incompletely transcribed RNA, unincorporated
Biotin label or both (lower well positions).
Of all the methods attempted, the labelling of RNA was most effective using
the random technique above where biotin-16-UTP was mixed in a ratio of 1:11 (Fig
4.10), which resulted in large amounts of labelled material in a robust reaction
(milligram quantities of RNA were produced compared to microgram quantities by
end labelling techniques). The ratio was determined based on the sequence of RBE3
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itself that contains a total of 11 Us including a run of 5 poly-Us at the 3’ end of the
construct (Fig 4.8). By using the biotin-16-UTP as a limiting resource in the mixture,
conservative random labelling of the RNA could occur so ~1 biotin-16-UTP was
inserted/strand of RNA. There was also a greater chance of the label being
incorporated at the 3’ end and away from the RBE3 coding sequence as there is a run
of 5 poly-Us at the 3’ end of the RBE3 sequence.
Several attempts were made to purify the labelled RNA, but failed, including
FPLC using a Rev34-50 affinity column as described by Englebretsen et al., (1996),
RP-HPLC and anion resins, and so an elaboration of the small-scale method was
applied. This method is in essence a stepwise application of the individual reagents
present in commercial RNA extraction kits and reagents like Tri-reagent (Sigma-
Aldrich), derived from the methods of Feramisco et al., (1982), Chomczynski and
Sacchi (1987), Meltzer (1990) and Maxam and Gilbert (1977).
Upon completion of the in vitro transcription reaction the pH of the solution
was changed by adding a denaturant buffer of 25mM sodium citrate (pH 4.0), 4M
guanidinium thiocyanate with the volume adjusted based on crude A260 readings, so a
minimum of 1mL buffer was added per 2mg of total nucleic acid (RNA and DNA).
The reducing agent β-mercaptoethanol (0.5% v/v) was added and shaken vigorously
before incubating on ice for 15min and clarification (10,000 x g, 4°C, 20min).
The aqueous phase was retained and 3M sodium acetate (1/10th volume, pH
4.0) was added to the solution and shaken gently for 5min. Room temperature citrate-
buffered Phenol (pH 4.3, 0.5 vols) was added to the solution and mixed gently. This
solution was left to sit for 10min to help dissociate nucleic acid-bound protein
complexes that may be present from the in vitro transcription reaction. Then 1-
bromo-3-chloropropane (0.5 vols) was added and the solution was mixed vigorously
to form an emulsion, then cooled on ice for 15min. Samples were finally clarified
(10,000 x g, 4°C, 20min), separating the RNA into the upper, clear solution phase.
Visualisation of the separate phases was not difficult as the compound 1-bromo-3-
chloropropane formed a very strong demarcation to the separate phases.
The clarified RNA was decanted, then precipitated by adding 1.2 volumes of
propan-2-ol to the top of the solution. The solution was left without agitation at room
temperature for 10min before mixing and placing on ice (4°C for at least 1h) and final
precipitation (-20°C overnight). Precipitated material was centrifuged (10,000 x g,
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4°C, 20min) and resuspended in DEP-treated ddH20, before mixing with a 2x
formamide loading buffer (95% v/v formamide, 10mM NaOH, 1mM EDTA, 0.3%
w/v bromophenol blue), in volumes such that RNA concentrations do not exceed
2mg/mL. Solutions were heated to 65°C for 15min, then chilled on ice before loading
onto the gels.
Final purification was conducted using 18% urea polyacrylamide gels and the
RNA was extracted using modified “crush and soak” methods of Maxam and Gilbert
(1977). Two volumes of gel extraction buffer (500mM ammonium acetate (pH 8.0),
10mM Mg2+ acetate, 1mM EDTA, 0.1% SDS) was incubated at 4ºC overnight with
the sliced gels before centrifugation (10,000 x g, 4ºC). This was followed by a second
addition of (2 volumes) extraction buffer with vigorous vortexing before
centrifugation (10,000 x g, 4ºC). The RNA was precipitated with 1.2 volumes of
propan-2-ol as described above and the pelleted RNA was washed with 70% ethanol
and resuspended in 40% formamide at up to 4 mg/mL (–80ºC).
The purity of RNA was estimated from the relative absorbances at 230, 260
and 280nm with A260/A280 ratios typically >2.0. and extraction steps were repeated if
the ratio was less than 1.7. The A260/A230 ratio was typically within the range of 1.5-2.
The approximate yield of total RNA obtained was determined spectrophotometrically
at 260nm, where 1 absorbance unit (A260) = 40µg of single-stranded RNA/mL with
between 5 – 10mg purified and labelled RNA produced per RiboMAX in vitro
transcription kit.
When compared to a commercial anionic resin purification method (QIAGEN
Rneasy Protect Maxi kit), the RBE3 RNA was found to bind poorly to the resins
resulting in only a 2% recovery of the RNA, with the majority of RNA flowing
through the resin or eluting during washing. QIAGEN advised a modified protocol
for small and highly structured RNA and supplied new buffers (contents unknown as
QIAGEN do not describe their buffer contents) and suggested applying a heated
solution of the RNA to the resin (65˚C). The QIAGEN resin was able to remove the
unincorporated biotin label from the correct size and labelled RNA but the yield of
labelled RNA produced was far less than using the described extraction method as
shown in the QIAGEN resin purification labelled 3 in Fig 4.10. This purification
contains ~80% less material based on A260 values than the derived method (1 and 2
of Fig 4.11), and the purity is far less with the QIAGEN material having an A260/A280
ratio of 1.3 compared to A260/A280 for the derived method of 2.1 and 2.2 for (1) and
(2) respectively.
Upon this basis, the production of labelled RBE3 RNA was made using the
RBE3 construct provided by Jørgen Kjems through a random incorporation of biotin-
16-UTP. Purification was effected following in vitro transcription under acidic
conditions and using a sequential application of guanidinium, phenol and 1-bromo-3-
chloropropane before final purification and extraction from 18% urea-PAGE gels.
7 6 1 2 3 4 5
Figure 4.11: Extraction of RNA from urea-PAGE gels. The sliced gels (left) are shown before
extraction and illuminated RNA is clearly seen using UV light and staining with SYBRgold (as per
Experimental section 4.3.5). The final purified material as shown on a transfer (right), The wells
labelled are as follows; (1) RBE3 purified by the method as described (2) as for (1) but with a longer
precipitation time (overnight), (3) is the same material purified from a QIAGEN anion resin, (4 and 5)
are QIAGEN washes (flow through) of the resins that indicated possible RNA content due to an A260
profile. The wells labelled 6 and 7 are repeated extraction of the resin by centrifugation (in the case of
6) and re-extraction (in the case of 7) using the same methods described for well 3.
4.3 Experimental 4.3.1 Constructs used and Transformation into JM109 Cells.
Constructs for HIV-1 Rev and different length RREs (full length RRE
sequence of 295 nucleotides, a 260 nucleotide construct and the RBE3) were
provided from the lab of Dr Jørgen Kjems of the University of Aarhus, Denmark. Dr
Kjems provided two Rev-containing plasmid constructs designated pET-His-Rev-
HMK and the second pET-HMK-Rev-His as described in Jensen et al., (1997) and
Jensen et al., (1995). Both constructs were modified pET vectors (containing a T7
promoter and an ampicillin resistance gene). The E.coli codon-optimised Rev gene
was likewise provided to the Kjems lab by others (Cochrane et al., 1989). The lab of
Dr Kjems modified this construct to include a poly-His coding sequence and a heart 114
muscle kinase site, which allow easy affinity purification of the recombinant protein
and 32P labelling, respectively, of the recombinant protein for use in gel mobility-shift
assays (Jensen et al., 1995).
The plasmids were provided as precipitated mini-preps of unknown quantity.
These plasmids were transformed into JM109 cells (Promega) to propagate large
quantities of stock plasmid DNA and minpreps and DNA digests of the transformants
were conducted to confirm plasmid sizes as described in Experimental section 4.3.2
(Fig. 4.12).
Figure 4.12: Restriction Digest of Constructs Obtained. A 1.2% TBE agarose gel visualised with ethidium bromide and UV light is shown where the FLRRE is the full length RRE sequence digested with Eco RI and 260RRE is a 260 nucleotide RRE construct digested with Eco RI. Single plasmids were obtained for all except the RBE3 containing plasmid that contains multiple restriction sites for Dra I and produces five fragments. Sizes of ~5kb were obtained for the Rev constructs (Bam HI digests of the C-terminal His tagged Rev plasmids and Eco RI digests for the N-terminal His tagged Rev constructs) and ~4kb for the RRE plasmids compared to DNA molecular weight markers (not shown).
RBE3 FLRRE 260RRE C-term N-term
JM109 cells were prepared according to a modified protocol of Hanahan
(1985). Frozen competent cells (100µL) were thawed on ice, 1µL of the plasmid
suspension was added and the cell/plasmid transformation was incubated on ice for
10min. Cells were then heat-shocked for 45 seconds in a water bath (42°C) and
placed back onto ice for 2min. After closing of the cells, SOC media (900µL, 4°C)
[SOC media – 2g Bacto-Tryptone, 0.5g Bacto-Yeast Extract, 1mL 1M NaCl, 0.25mL
1M KCl, 1mL 2M Mg2+ stock [101.5g MgCl2·6H2O, 123.3g MgSO4·7H2O in 500mL
ddH2O, 0.22µm filtered], 1mL 2M glucose (sterile filtered)] was added and the
suspension was incubated for an hour (37°C, 250rpm) and selected using LB-
ampicillin plates overnight (10µL and 100µL of the cells were plated onto the
selection media) [LB 1L – 10g Bacto-Tryptone, 5g Bacto-Yeast Extract, 5g NaCl, pH
7.5, 15g agar for plates and 100µg/mL ampicillin]. Transformations were highly
efficient yielding hundreds of colonies containing at least one plasmid insert.
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A single colony of the transformed cells were selected and grown overnight in
LB-ampicillin media (3mLs culture, 37°C, 280rpm). The culture was split into two
(1.5mL each) and half was stored as a stock (-80°C, 50% v/v with 80% v/v glycerol).
The other half of the culture was pelleted (12,000 x g, 5min, 4°C), resuspended in
100µL TE-lysis buffer (25mM Tris·HCl pH 8.0, 10mM EDTA, 50mM glucose) and
lysed in the presence of 200µL alkaline lysis solution (0.2M NaOH, 1% SDS).
To the lysed culture was added 150µL acidifying buffer (3M K·CH3COO-, 5M
CH3COOH pH 4.8, 4°C) and insoluble debris pelleted (12,000 x g, 5min, 4°C). The
supernatants were layered with 2.5 vols EtOH and incubated on ice 5min before
pelleting the precipitated DNA (12,000 x g, 5min, 4°C). The DNA pellet was washed
with 70% EtOH (-20°C), repelleted (12,000 x g, 5min, 4°C), and dried at room
temperature.
4.3.2 Restriction Digests.
Restriction digests (Fig. 4.12) were conducted using standard protocols
supplied with reagents (Promega) except in the case of large scale digests (up to 32µg
DNA template) using Dra I where an excess of reagents were used to ensure digest
completion. For Eco RI and Bam HI digests of Rev constructs, the reactions were
conducted at 37°C overnight in 20µL reactions comprising 1µg DNA template,
0.1mg/mL BSA, Multi-core buffer (25mM Tris acetate (pH 7.8), 10mM Magnesium
acetate, 100mM Potassium acetate, 1mM DTT), with 10 units each of enzyme in
storage buffer. The final dilution of storage buffer was 1:20 in reactions with the Eco
RI storage buffer comprising 10mM Tris-Cl (pH 7.4), 400mM NaCl, 0.1mM EDTA,
1mM DTT, 0.15% Triton X-100, 0.5mg/mL BSA, 50% v/v glycerol and the Bam HI
buffer being 10mM Tris-HCl (pH 7.4), 300mM KCl, 5mM MgCl2, 0.1mM EDTA,
1mM DTT, 0.5mg/mL BSA, 50% v/v glycerol.
The Dra I digestions of small and large scale preps were conducted at 37°C
overnight with 1µg (small scale) or 32µg (large scale) DNA, 0.1µg/mL BSA, and
digest buffer (6mM Tris-HCl (pH 7.5), 50mM NaCl, 6mM MgCl2, 1mM DTT) in the
presence of 5 (small scale) or 60 (large scale) units of Dra I (Dra I supplied in storage
buffer that was diluted 1:20 in the digest buffer conditions (storage buffer comprised
10mM Tris-HCl (pH 7.3), 300mM NaCl, 1mM DTT, 0.1mM EDTA, 0.5mg/mL BSA
and 50% v/v glycerol).
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4.3.3 Preparation of Electrocompetent E.coli for Protein Expression.
A 5mL overnight culture was grown in SOB media (37°C, 250rpm) [SOB
800mL autoclaved, Mg2+ deficient – 16g Bacto-Tryptone, 4g Bacto-Yeast extract,
1.6mL 5M NaCl, 667µL 3M KCl], inoculated into 500mL SOB media and grown to
an OD550 of 0.8. The cultures were pelleted (4,000 rpm JA10.5, 4°C, 10min) and
washed 3 times in 10% v/v glycerol (4°C) by centrifugation and resuspension (4,000
rpm JA10.5, 4°C, 10min).
The final pellet was resuspended in glycerol (10%) to a minimal volume
yielding an OD550 = 200-300 (2mL) for storage/electroporation. Electroporation was
conducted using a Quantum EC100 electroporator with 10ng of plasmid DNA in TE
buffer (10mM Tris·HCl (pH 8.0), 5mM EDTA) and 160µL of competent cells at
2.5kV/25µFd 5 msec-1. SOC media was added to the electroporated cells (1mL, 4°C)
and the cells were grown for 1 hour (37°C, 250rpm).
4.3.4 LB Antibiotic Media
Both cell-type constructs were grown on LB-antibiotic media (AD494(DE3) –
100µg/mL ampicillin and 15µg/mL kanamycin, BL21(DE3)pLysS - 100µg/mL
ampicillin and 34µg/mL chloramphenicol) with 10µL and 100µL of transformed
cultures plated and grown at 37°C overnight. All plates yielded hundreds of selected
bacteria, of which single colonies were taken and glycerol stocks made for long-term
storage at –80°C.
4.3.5 SDS-PAGE and Urea-PAGE Protocols.
Sodium Dodecyl-Sulfate Poly-Acrylamide Gel Electrophoresis (SDS-PAGE)
was conducted following modified methods of Laemmli (1970) in the following
manner. An acrylamide (12.8% v/v) resolving gel was made and poured comprising
acrylamide:bis-acrylamide (29:1), 0.37M Tris (pH 8.8), 0.1% v/v SDS, 0.01% v/v
TEMED and 0.05% w/v ammonium persulfate. The resolving gel was poured to
approximately an inch below the smaller plate height (40mm mould) and layered with
butanol until set. Once the gel was set, the butanol was removed and an acrylamide
(5% v/v) stacking gel comprising acrylamide:bis-acrylamide (29:1), 0.126M Tris (pH
6.8), 0.1% SDS, 0.15% v/v TEMED and 0.05% w/v ammonium persulfate was
118
poured onto the top of the resolving gel and a well-forming comb was inserted into
the forming matrix and the gel allowed to set.
Protein samples were boiled for 5min in the presence of 10% v/v β-
mercaptoethanol and an equal volume of 2x loading buffer was added (1% v/v
bromophenol blue and 60% v/v glycerol). Samples were centrifuged briefly at 14,000
x g for 2min before loading onto the gel. The electrode buffer was a solution of
14.4g/L Glycine, 3.3g/L Tris, 0.1% SDS – the pH was checked but rarely adjusted
being typically close to pH 8.3. The gels were run under amperage-limited conditions
at 10mA through stacking phase then 15mA through the resolving gel.
Urea-PAGE gels were made using the same apparatus as SDS-PAGE gels, but
comprised 18% v/v acrylamide:bis-acrylamide (29:1), 0.75g/mL urea, 100mM Tris
(pH 8.0) and 0.1% w/v EDTA. The gel solution was dissolved by heating in an 800w
microwave oven for 5min (800w setting), and the solution was filtered hot through a
0.45µ filter to remove particulate matter before polymerisation (0.27% v/v TEMED
and 0.09% ammonium persulfate) and setting as per normal SDS-PAGE protocols.
Gels were pre-run for 10min in 50mM Tris (pH 8.0), 0.05% EDTA tank buffer and
samples were loaded and typically run for 1h at 300V and stained with either
SybrGreen II or SybrGold (Molecular Probes): 50mM Tris (pH 8.0), 0.05% EDTA
(1:10,000 dilution stained for 25min). Using these dyes, the RNA could be visualised
and pure RNA bands excised from the gel.
4.3.6 Transfer Protocol for Western Blotting and Biotin Detection.
The SDS-PAGE gels were placed into transfer buffer for 30min to equilibrate
[transfer buffer - 10mM CAPS pH 11 and 10% Methanol, 4°C]. Transfer filter papers
(4) were cut to size and the transfer membrane was prepared through serial
equilibration in methanol (20 seconds), ddH2O (1min) and finally the transfer buffer
(5min or until used). Transfer was conducted onto nitrocellulose film at 250mA for
2h, 300mA 1h or 500mA for 30min. Transfers were conducted in the cold by placing
the tank onto a stirring block surrounded by ice (in an esky) and filled with cold
CAPS buffer (stirred using a magnetic stirrer during the transfer).
After transferring, the blot was placed into a BSA (1%)/TBS solution (the Tris
buffered saline was made by titrating 1M HCl with a solution of 4mM Tris.HCl and
0.1M NaOH to pH 7.5) for 20min at room temperature or overnight at 4°C. A BSA-
119
blocking solution was prepared using ~5g/mL BSA in TBS, and 40µL of this
BSA/TBS solution was used to make a His-Ab solution (His-Ab was a hexa-His-
Mouse polyclonal supplied by Clonetech, Lot #6110562 and diluted 1:2000,
4µL/8mL) to inhibit binding of non-specific His-Abs that may interact with BSA and
generate background as well as reduce loss of the His-Ab by adhering to surfaces.
Membranes were placed onto nescofilm and rocked gently for 45min with the His-Ab
solution. At the end of the cycle the membranes were briefly washed with the
BSA/TBS solution and then washed with a detergent solution comprising
TBS/TWEEN (0.5mL TWEEN/L TBS buffer) (three times for ~5min).
The secondary Ab (mouse-alkaline phosphatase) was added to TBS buffer as
1:5000 dilution (1.6µL/8mLs of TBS) and 40µL BSA/TBS was again added to inhibit
binding of non-specific Ab’s and this solution was poured onto the membranes while
agitated on nescofilm. Finally the membranes were again washed with TBS/TWEEN
three times followed by TBS. Membranes were chemically activated with Western
Blue (a substrate for alkaline phosphatase) for 20min.
For detection of biotin labelled RNA, the RNA run on 18% urea-PAGE gels
(Experimental section 4.4.5), was transferred to a nitrocellulose as described above
using 0.4M NaOH as the transfer buffer and applying 500mA across the membrane
(voltage limiting conditions of 15v) for 25min.
The transfer was briefly washed in TBS (as above) and non-specific binding
blocked using the Boehringer Mannheim blocking reagent supplied with the CDP-
StarTM kit in TBS solution from 40min to 14h time periods. The transfer was then
washed four times with TBS before incubating with a Streptavidin-alkaline
phosphatase-antibody conjugate (Boehringer Mannheim) diluted 1:2500, with 1%
blocking reagent in TBS for 40min with agitation. The transfer was washed 4x in
0.3% TWEEN (in TBS) before incubating with the alkaline phosphatase
chemiluminescent substrate CDP-STARTM (1:100 dilution in 100mM Tris (pH 9.5),
100mM NaCl).
All transfers were blot dried with a Whatman 3MM paper briefly before
chemiluminescent detection by exposure to Agfa, Curix blue HC-S plus X-ray film
for 0.5 – 5min and developed using an Agfa, Curix 60 autodeveloper.
120
4.3.7 Calculation of Protein Concentrations.
The Expasy programme “ProteinParam tool” (http://www.expasy.ch/cgi-
bin/protparam) was employed to calculate a theoretical molar extinction co-efficient,
which in turn was used to calculate the protein concentration of Rev from solution
absorbances at 280nm. The specific value of ε for HIV-1 Rev was found to be 0.579g-
1 L-1 cm (assuming cysteines form complete disulfides). A secondary check of the
protein concentration was made using the Coomassie dye method of Bradford that is
known to bind aromatic and basic amino acids, especially arginine, and cause a shift
of λmax from 465 to 595nm upon protein binding (Bio-rad, 2001). Because of the
arginine rich content of Rev, this method was considered to be the most sensitive of
standard protein assays for assessing protein concentration.
A microplate micro-assay approach was used with a BSA standard (ε = 0.63g-
1 L-1 cm). Each of the standard BSA wells also contained an appropriate amount of
storage buffer or assay buffers to normalise the standards with buffer additives. Both
the spectrophotometric and Coomassie dye method agreed with the concentrations of
HIV-1 Rev protein to within 3%.
4.3.8 RP-HPLC and Mass Spectrometry.
RP-HPLC was conducted with Rev injected in final refolding buffer onto a
Vydac C-8 analytical (Ø5mm x 250mm) column pre-equilibrated with buffer A (90%
ddH2O, 9.9% acetonitrile, 0.1% TFA) at a gradient of buffer B of 1mL/min (buffer B
= 90% acetonitrile, 9.9% ddH2O, 0.1% TFA). Protein was found to elute at 12min of
mass ~ 6kDa and 31 – 32 min of ~14 – 15kDa as assessed by direct injection into the
Mariner mass spectrometer. Samples were directly injected from the HPLC column
for rapid identification, or run through an RP-HPLC column from the final refolding
buffer conditions. The column was a Zorbax 300SB column (Ø2.1 x 150mm C3 5µm)
using the same buffers as above with a flow rate of 250µL/min. The column fed
directly into a Mariner Electrospray (ES) – Time-Of-Flight (ES-TOF) mass
spectrometer (Applied Biosystems, California, USA). With the ions focused from 700
– 1295amu in increments of 0.2amu. The peaks were reconstructed from a repeating
series of associated peaks to two species of 14486.6amu and 14931.4amu (+/- 2amu)
for pure fractions that compares to a calculated mass of 14467.3amu for Rev based on
the sequence in Fig 4.2.
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Chapter 5: Application of Surface Plasmon Resonance Assays
to Rev:RRE interactions.
5.1 Introduction. 5.1.1 Assaying RNA.
5.1.2 Rev:RRE Inhibition.
5.2 Results and Discussion. 5.2.1 High Throughput Screening of Aminoglycoside
Antibiotics.
5.2.2 Assay of Helix Mimetics.
5.2.3 Does Helix Induction Increase Affinity for the RBE3
Aptamer?
122
5.1 Introduction. 5.1.1 Assaying RNA.
Although RNA presents a novel target for therapeutic intervention against
pathogens such as bacteria, yeasts and viruses, as well as disease states, especially
cancer, effective development of such therapeutics is difficult. One of the major
problems in developing RNA therapeutics is the difficulty of assaying large libraries
of compounds for their interaction with RNA. The common methods employed to
assay RNA interactions are: filter binding, gel mobility shift, fluorescence techniques
and surface plasmon resonance (SPR).
Filter binding assays rely on the strong binding of protein (but not RNA) to a
nitrocellulose filter. Following a washing step, the RNA retained on the membrane is
then quantitated allowing calculations of stoichiometry and equilibrium constants.
Gel mobility-shift assays rely on a change in the electrophoretic mobility in non-
denaturing gels caused by an association between the RNA and the analyte; the
difference in the mobility at various concentrations of analyte can be used to
determine the equilibrium constants and stoichiometry. However, filter binding and
gel mobility-shift assays suffer from several technical problems. (1) Production of
pure RNA and subsequent handling and running of the materials used in the assay
must be under strict conditions to avoid RNase contamination. (2) The labelling of
sufficient RNA or protein (or both) to accurately detect an association in the assay.
(3) The difficulties of reproducing results because a measure of technical skill and
experience is essential to ensure success (Revzin et al., 1986; Molloy, 2000; Draper et
al., 1988; Hall and Kranz, 1999). In the case of filter binding experiments, there are
problems if washing steps are incomplete especially with larger RNA molecules.
However fewer artifacts are encountered in filter binding experiments than gel
mobility shift assays, as the half-life of most RNA-protein complexes is significantly
less than the time of separation of complexed and uncomplexed RNA (Xavier et al.,
2000).
Fluorescent techniques tend to rely on tryptophans in proteins for fluorescence
or fluorophores that must be chemically attached and successfully quenched when an
association occurs. This method can suffer from similar problems to gel mobility shift
assays (described above), however the technique has greater reproducibility (Xavier
et al., 2000). Fluorescence techniques can also suffer from the problems of internal
123
quenching, a poor tryptophan signal and signal to noise problems of the particular
system employed (Kwon and Carson, 1998).
Filter binding, gel mobility shift and fluorescence assays only provide limited
information on equilibria. None can elucidate the type of binding event, such as
classical Langmuir association, conformational changes, heterogeneity or a
combination of any of these to form a complicated binding model. Furthermore, only
fluorescent techniques are routinely used for high-throughput screening of libraries
(Xavier et al., 2000), though examples of modified filter binding techniques using
Streptavidin beads are becoming popular (Chapman et al., 2002). In this regard SPR
offers the potential to overcome the problems encountered by these assay techniques,
to deliver high-throughput screening, and to provide a more thorough understanding
of the nature of analyte and ligand association.
Three specific SPR-based assays involving the RRE have been described
(West and Ramsdale, 1997; Hendrix et al., 1997; Van Ryk and Venkatesan, 1999).
The assay of West and Ramsdale (1997) was conducted using BIAcore CM5 chips
that had been chemically coupled to the Rev34-50 peptide. The Rev34-50 peptide was
succinylated and had a small spacer addition (Gly-Ser-Gly) to raise the peptide from
the chip surface and avoid crowding when the RBE3 RNA analyte formed an
association. Competitive assays were conducted by pre-equilibrating RBE3 RNA
with different concentrations of small molecule antagonists and Ki values calculated
from dose-response curves (IC50 values). This study differs from other SPR studies
involving the Rev:RRE that immobilise RNA (Hendrix et al., 1997; Van Ryk and
Venkatesan, 1999), and does not examine the full length Rev protein, but relies on the
findings of Kjems et al., (1992) that the KD for the protein and peptides are similar at
10 – 20nM for RBE3 RNA. The study by West and Ramsdale (1997) also contains a
technical difference: the regeneration solution is 6M Gu.HCl that is contraindicated
for CM5 surfaces by BIAcore information sheets that accompany the chips.
The study of Hendrix et al., (1997) established some structure-activity
relationships by screening aminoglycosides. They captured RBE3 RNA onto SA
chips or custom Streptavidin surfaces on CM5 chips. The RBE3 sequence was
generated using in vitro transcription with the label randomly incorporated using
guanosine-5’-monophosphothioate resulting in termination products requiring
repeated gel purification. The phosphothioate was converted to a biotin label by
124
reacting biotin-iodoacetamide in DMF with the phosphothioate-RNA, and the
labelled material was repeatedly gel purified. The study utilised four Rev34-50 peptides
in order to assess the folded state of RNA on the chip surface. These comprised a
native sequence, a native sequence with a poly-alanine sequence at the N- and C-
termini to induce helicity, the alanine sequence with an additional N-terminal cysteine
for future modifications, and a succinylated form of the cysteine containing sequence.
Van Ryk and Venkatesan (1999) reinvestigated Rev:RRE interactions using a
variety of truncated and full length RRE each containing a 3’ poly A sequence to bind
to a chemically synthesised biotinylated poly T capture arm that was preimmobilised
onto BIAcore Streptavidin (SA) chips. The poly A – poly T runs base paired to form
the ligand surface and full length Rev, NES mutants and truncated/hybrid Rev
sequences as well as the aminoglycosides Neomycin B, Kanamycin A and
Hygromycin B were analysed on these surfaces.
5.1.2 Rev:RRE Inhibition.
Since the initial recognition that HIV encoded Rev, antagonists of the
Rev:RRE interaction have been suggested as potential therapeutics for treatment of
HIV-1 infections (Feinberg et al., 1986; Sodroski et al., 1986). Analysis of the
Rev:RRE interaction has led to the discovery of compounds which can perturb the
Rev:RRE association, but a better understanding of this association through structure-
activity relationships will assist further antagonist design. Inhibitors to the Rev:RRE
interaction can be discussed by the two main approaches used to design them : (i)
biological strategies and (ii) chemical strategies (for a generalised review see Heguy,
1997).
5.1.2.1 Biological Strategies.
The scientific literature on Rev:RRE inhibition largely comprises biological
approaches involving either: (a) oligonucleotide/antisense strategies directed against
the RRE or (b) decoy strategies where mimics of Rev or the RRE are employed.
Biological strategies are innovative and potent due to their highly complementary
nature to either the RRE or Rev (as either antisense or trans dominant mutants), but
they have drawbacks that have prevented their use to date as therapeutics.
Oligonucleotide/antisense strategies: The majority of biological strategies are
125
focused on antisense RNA approaches employing short nucleic acid sequences
complementary to the RRE. When bound to the RRE the antisense sequence is a very
potent inhibitor of Rev binding and HIV-1 replication (Fraisier et al., 1998). Another
approach employs ribozyme elements with complementary RRE sequences to bind
and cut the RRE, leading to improperly translated proteins (Yuyama et al., 1994;
Duan et al., 1997). The best targets for antisense strategies on the RRE contain large
sections of unpaired RNA or a stem loop bulge, since the antisense molecule does not
have to compete with an in situ complementary sequence (Cload and Schepartz,
1994). The favoured target on the RRE is stem loop II (Li et al., 1993), though
targeting stem loop V is also attractive since the binding of antisense sequences to it
was found to expose stem loop III to RNase CV digestion and possibly block Rev
multimerisation on the RRE (Chin, 1992).
Decoy strategies: Decoy strategies employ inactive mimics of either Rev or
RRE, thus bringing about inhibition by perturbing the threshold of target or activator.
The most advanced and potent decoys are based on the pM10 Rev protein mutant
(NES mutant), or derivatives thereof (Malim et al., 1989a).
5.1.2.2 Chemical Strategies.
There is relatively little published work on chemical antagonists of the
Rev:RRE interaction compared to chemical antagonists of other HIV-1 targets. This
is because of incomplete knowledge of fundamental biology integral to Rev:RRE
function, as well as the difficulties associated with designing compounds for selective
interactions with small specific pieces of RNA that nevertheless involve large surface
areas. There are three separate mechanisms by which chemical strategies work to
inhibit Rev function: (a) inhibiting the export of RNA from the nucleus to the
cytoplasm, (b) inhibiting intron splicing, and (c) inhibiting the formation of Rev:RRE
complexes. The chemical strategy has largely arisen from screening compounds
expected to hydrogen bond to the RRE, but few compounds have been found to be
antagonists at even low micromolar concentrations. Of those compounds that are
active, many (such as aminoglycosides) are difficult to build synthetic libraries upon,
or are not drug-like. Thus most lead compounds have been inadequate to serve as a
pharmacophore.
(a) Inhibitors of RNA Export.
Leptomycins and Kazusamycins (Fig 5.1) act as potent (nanomolar) inhibitors
of Rev export by blocking all traffic through the 5S rRNA pathway. However,
because all activity through these pathways is inhibited, these compounds are highly
cytotoxic and are generally not considered to be drug candidates (Wolff et al., 1996).
126
HO
O
OH OR1
R2
O
O
Compound
R1
R2
IC50
(nM)
Toxicity
(nM) 72h
Leptomycin A CH3 CH3 3.5 2.6
Leptomycin B CH3 CH2CH3 0.6 0.9
Kazusamycin B CH2OH CH3 20 7.4
Kazusamycin A CH2OH CH2CH3 2.5 2.9
Figure 5.1: Representative Structures of Potent RNA Export Inhibitors. Inhibition IC50 is defined
as the concentration for 50% inhibition of transport of Rev to the cytoplasm; toxicity is defined as 50%
reduction in cell number over 72h (Wolff et al., 1996; Wang et al., 1997).
(b) Inhibitors of Intron Splicing.
Inhibitors of Group I intron splicing interfere with the conserved core unit of
intron splicing causing a general decrease in protein production and include
secondary metabolites, cyclic peptides and many aminoglycoside antibiotics. For
most known inhibitors of group I intron splicing that have antiviral activity towards
HIV-1, the antiviral action is attributed to secondary effects where no direct inhibition
of the Rev:RRE interaction is observed with inhibition ranging from 1-50µM. A
complication in understanding the mechanism of action of inhibitors to group I intron
splicing is their promiscuous nature towards different RNA sequences (often making
these inhibitors cytotoxic) (Wank et al., 1994; Schroeder et al., 2000).
(c) Direct Inhibitors to the Rev:RRE Interaction.
To date the greatest source of lead compounds and pharmacophores used for
direct inhibition of the Rev:RRE interaction are aminoglycosides (Fig. 5.2). The best
natural inhibitor was found to be Neomycin B with a Ki of 1µM (Zapp et al., 1993)
and in vitro inhibition of 70.5% of gag protein production at 200µM (Park et al.,
1996).
127
O
O
O
R1
HOR2 H2N
ONH2
NHR3
OHOOH
OHONH2H2/H2N
HOR4O
Compound R1 R2 R3 R4 Inhibition
Neamine NH2 OH H --- >100 µM
Ribostamycin NH2 OH H --- 100 µM
Butirosin NH2 OH AHBA ---
Neomycin B NH2 OH H H 1µM
Paromomycin I OH OH H H >100 µM
C
B
A
Figure 5.2: Base aminoglycoside structure. Naming nomenclature of aminoglycosides A= Neamine
B= Ribostamycin A+B= Butirosin B+C= Neobiosamine. Nomenclature used here for A+C=
Neoribostamine. By comparison, the aromatic and sugar aspects of the base aminoglycosides (A)
Neamine and (B) Butirosin are 100 fold less potent than Neomycin B (>100µM concentrations for
inhibition). Neoribostamine substituted compounds form a class of typical antibiotics with Ki values
between 10-100µM. AHBA= L-α-amino-n-hydroxybutyric acid. Figure adapted from Zapp et al
(1993).
A B C
O
H2N
HOHO H2N
ONH2
NH2OHO
N
NHO
HO
O
OHH2NO
O
H2N
O
H2N
HOHO H2N
ONH2
NH2OHO
N
NHO
HO
O
OHH2N O
OHO
O
H2N
HOHO H2N
ONH2
NH2OHO
N
NHO
HO
O
OHH2NO
OO
HO
Figure 5.3: Representative 4,5-disubstituted 2-DOS Compounds with Improved Rev:RRE
Inhibition over Neomycin B. (A) Has 81% inhibition at 200µM (this is 10.5% greater than Neomycin
B). (B) Has 91% inhibition at 200µM (this is 20.5% greater than Neomycin B). (C) Has reported 87%
inhibition at 200µM (this is 16.5% greater than Neomycin B) as found by Park et al., 1996.
New synthetic compounds based on Neoribostamine substitution
(disubstituted 2-deoxystreptamine (2-DOS) compounds – Fig. 5.2 and Fig 5.3) show
low micromolar inhibition, with the best claimed to achieve 91% inhibition of gag
protein production at 200µM, or some 20% greater inhibition than Neomycin B in
vitro (Park et al., 1996). The most potent aminoglycoside is a synthetic Neomycin B-
acridine conjugate that was designed to increase the structural rigidity of Neomycin B
and thereby its potency (Fig 5.4) (Kirk et al., 2000). The Neomycin-acridine
conjugate is reported to have a Ki of 1.5nM, close to the affinity of ~1nM of the Rev
protein for the RRE, but toxicity, bioavailability and in vitro potency are not reported.
128
O
O
O
H2N
HOHO H2N
ONH2
NH2OHO
OHONH2
HOH2N
OHO
HN
Figure 5.4: Neomycin-acridine conjugate. Kirk et
al., (2000) reported the 9-fold enhanced IC50 of the
conjugate over Neomycin B.
The aminoglycosides suffer from potency, specificity and toxicity problems;
they interfere with protein synthesis, bind to rRNA (Wank et al., 1994) and can be
modified through post-translational protein pathways, generally making these
compounds unsuitable therapeutics (Park et al., 1996).
O
O
O
OHO
OO
OO
O
O
O
O
OH
O
N
O
HOO
O
OOH
)3
(
O OHOH
OH
OH
HO O
C
B
A
Figure 5.5: Other Inhibitors of Rev:RRE Function. (A) Niuriside from the Indian medicine plant
Phyllanthus niruri, IC50= 3.3µM but did not protect infections at concentrations <260µM (Qian-
Cutrone et al., 1996). (B) Fusaricide, an N-hydroxypyridone (extracted from an unknown Fusarium sp)
IC50 = 40µg/mL (McBrien et al., 1996) and (C) Orevactaene an oxopolyene extracted from
Epicoccum nigrum IC50= 3.6mM (Shu et al., 1997).
Other inhibitors that work by unknown mechanisms or bind in a non-specific
manner have been documented and include synthetic compounds like diphenylfuran
and aromatic cations (Zapp et al., 1997; Chapman et al., 2002) as well as natural
products. None of these compounds are potent in vitro and all have moderate
129
cytotoxicity, interacting in unknown ways with other cellular components (Qian-
Cutrone et al., 1996; McBrien et al., 1996; Shu et al., 1997).
In conclusion there are currently no small molecule compounds suitable to
advance into clinical trials as antagonists of Rev:RRE function and more studies in
this area are clearly warranted.
5.1.3 Helix Induction of a Decoy Inhibitor.
Because the helical RRE-binding domain of Rev interacts in a highly specific
manner with the RRE, mimetic-inhibitors based on this recognition sequence have the
potential to be more specific and potent than inhibitors such as aminoglycosides.
Development of helical inhibitors is also highly attractive as many key recognition
motifs that mediate numerous biological processes are constrained in a helical
conformation (Gross, 2000; Groves and Barford, 1999; Burkhard et al., 2001). The
problem with using helices derived from proteins as inhibitors is the tendency to lose
helical conformation when shortened to a minimal peptide-binding unit away from
the helix-stabilised environment provided by the protein. Therefore methods of fixing
helicity in short peptides might be expected to increase affinity and facilitate design
of potent inhibitors of helix-mediated biological processes. A few molecules have
been reported to nucleate helicity in short test peptides when chemically attached to
their N-termini, but this approach has not been successfully demonstrated for
biologically important peptides (Kemp et al., 1991; Muller, et al., 1993; Austin et al.,
1997; Aurora and Rose, 1998).
For the purpose of testing the proposition that helix induction can increase
ligand affinity for a biologically important drug target, the synthetic template 1 (Fig.
5.6) was used as a helix-stabilising cap on the peptide sequence from Rev34-50 to
generate 4 and compare the affinity of 4 to 2 (the native peptide product from the
fmoc synthesis of Rev34-50), 3 (the succinylated form of 2) and the recombinant Rev
protein (Fig. 5.6). The template 1 is believed to stabilise helicity by pre-organising its
carbonyl oxygen atoms, allowing hydrogen bonding to three amide nitrogens in
attached peptide sequences.
130
NH
O
OH
O
OO
HN TRQARRNRRRRWRERQR-NH2
TRQARRNRRRRWRERQR-NH2HN
OO
TRQARRNRRRRWRERQR-NH2H3N
HN
O
OHO
RO
O
O
1
2
3 4
Figure 5.6: Peptides Used in the Study. The helix cap (1) used to induce helicity, the native Rev34-50 sequence (2), the succinylated form (3) and the cap (1) attached to the peptide (2) forming peptide 4. The helix-inducing cap has been reported to induce α-helicity at 22°C for a test peptide containing alanines (Austin et al., 1997).
HO
O
5.1.4 Aims.
There is a need for rapid screening of small molecule antagonists to develop
structure-activity relationships and refine the drug design process. Recombinant Rev
and in vitro transcribed and labelled RBE3 (Chapter 4) was produced to be used in a
BIAcore based SPR assay towards the 4th aim of the project - the development of an
RNA-binding assay to test prospective Rev antagonists in vitro. The description of
this aim needed to be broadened because of the nature of the inhibitors to the
Rev:RRE interaction and the sensitivity of the BIAcore 2000 (small molecules below
1kDa in size often do not produce a signal large enough to be detectable above
background noise). The first aim of this chapter was to develop a competitive SPR
based assay for high throughput screening of small molecule antagonists of the
Rev:RRE interaction. In contrast, molecules bigger than 1kDa that can show an
appreciable increase in plasmon resonance and so antagonists cannot be discriminated
from the Rev protein binding to the RRE. Therefore a second aim of this chapter was
to develop a non-competitive SPR based assay for specific screening of potent
biological antagonists of the Rev:RRE interaction.
131
5.2 Results and Discussion. 5.2.1 High Throughput Screening of Aminoglycoside Antibiotics.
All buffers and solutions used in the BIAcore machine were filtered and
degassed through a 0.22µ filter under house vacuum or using syringes and
microfilters (0.22µ Millipore) except for BIAcertified buffers that were used as
supplied. The method of preparing the surface for immobilisation or capture was the
same for both chips used in this chapter (Streptavidin – SA or low density
carboxymethyldextran – Pioneer B1). The chips were docked into a BIAcore 2000
machine and the surfaces conditioned in BIAcertified HBS-EP running buffer (25ºC)
at flow rates from 5 – 50µL/min for up to 30min (BIAcertified HBS-EP is a BIAcore
certified buffer comprising 10mM Hepes (pH 7.4), 150mM NaCl, 3mM EDTA,
0.005% Surfactant P20 where P20 is Polyoxyethylenesorbitan, a nonionic surfactant
recommended for inclusion in the buffers used in BIAcore systems). The surfaces
were repeatedly washed at a flow rate of 10µL/min with alternation between 1M
NaCl and 50mM NaOH (5µL injections) until the baseline was stable before
immobilisation of protein or in the case of SA chips, the capture of RNA.
For the high-throughput screen, the biotinylated-RBE3 RNA was prepared by
diluting 40µL of stock biotin-RBE3 RNA (129.8µg/mL, 40%v/v formamide) with
160µL of BIAcertified HBS-EP buffer and annealed by heating to 80ºC for 10min
and allowed to slowly return to 25°C (30 – 45min). The RNA was then further diluted
with BIAcertified HBS-EP buffer to 10.7µg/mL and 300µL of this RNA solution was
flowed onto the SA chip at 10µL/min. The RNA-captured surfaces were washed at
10µL/min with two washes of 1M NaCl (40µL each), stabilising the surface and
resulting in 1553RU of labelled RBE3 RNA captured (Fig. 5.7).
Various buffers and additives were tested to develop running and regeneration
conditions. Best conditions were found to be 50mM Hepes (pH 7.4), 300mM NaCl,
3mM EDTA. These conditions were reached by dialysis of 0.1 – 0.5mg/mL Rev in
the final refolding buffer (50mM Hepes (pH 7.4), 500mM NaCl, 10% v/v glycerol,
4°C) against 50mM Hepes (pH 7.4), 500mM NaCl, 3mM EDTA for 6 hours at 4°C,
followed by dialysis into 50mM Hepes (pH 7.4), 300mM NaCl, 3mM EDTA,
overnight at 4°C. Under these conditions Rev remained stable when serially diluted
into running buffer at 25°C, but elevated Rev concentrations above 0.5mg/mL or
lower salt concentrations below 300mM caused precipitation to occur (both at 4°C
and 25°C).
17000
18000
19000
20000
21000
22000
23000
24000
25000
-1000 0 1000 2000 3000 4000 5000 6000 7000 8000Time s
Res
pons
e
RU
2 3 4 5
1
Figure 5.7: Immobilisation of RBE3 RNA onto SA chips. 1 Baseline level, 2 RBE3 being captured,
3 dissociation, 4 stabilisation of the surface using 2 x washes of 1M NaCl and 5 short period (~30min)
of monitoring the surface for stability resulting in 1553RU of captured RNA.
Optimised conditions were found by modifications of the Van Ryk and
Venkatesan (1999) methods and comprised a running buffer of 50mM Hepes (pH
7.4), 300mM NaCl, 3mM EDTA, 25ºC, and a regeneration solution of 50mM Hepes
(pH 7.4), 450mM NaCl, 0.1% SDS, using either repeated pulses or a wash of 20µL
injected at a rate of 5µL/min (here a pulse is defined as an injection of half the
volume of the flow rate; for a 30µL/min flow rate a pulse is 15µL). Therefore these
buffer conditions were applied to a high-throughput analysis of aminoglycoside
antibiotics and small molecules known to inhibit the Rev:RRE interaction.
The nine antagonists to be assayed were Neomycin B, Streptomycin, Pyronin
Y, Kanamycin, Tobramycin, Bekanamycin, Paromomycin, Amikacin and
Spectinomycin (Fig. 5.8). All compounds were purchased from Sigma-Aldrich as
lyophilised powders and used as provided without further separation or purification,
although solutions were filtered through a Millipore 0.22µ filter before use.
132
OHO
H2N
HO
NH2OO
NH2NH2
OHO
OHO
O
NH2OH
H2N OH
HO
OHO
H2N
HO
OH O
O
O
HO
NH2NH2
OH
NH2
HO
OHO
HO
HO
NH2OO
NH2NH2
OHO
OHO
O
NH2OH
H2N OH
HO
OHO
H2N
HO
NH2O
O
O
HO
NH2NH2
OH
NH2
HO
OH
H2N
HO
NH2O
O
O
HO
NH2NH2
OH
NH2
HO
OHO
H2N
HO
OH O
O
O
HO
NH2HN
OH
NH2
HO
HO
OH
OH
NH2
OHO
OHO
O
H2NNH
NH
NH2OH
O
HO
HNHO
HO HO
HO
O
O O
HNOH
HN
HO
O
O NN
Neomycin B Paromomycin
Streptomycin
Bekanamycin (Kanamycin B) Tobramycin Kanamycin (A)
AmikacinPyronin Y Spectinomycin
Figure 5.8: Small Molecule Antagonists of the Rev:RRE Interaction. The structures of the nine
small molecule antagonists assayed in this study are presented.
Paromomycin and Streptomycin contained a sulfate counter-ion in initial
stock solutions, and Neomycin B was supplied as a mixture of Neomycin B (70%)
and Neomycin C (30%). Pyronin Y was composed of Pyronin Y, Pyronin G, Pyronin
J and 45% methylene green (but no information was provided on the composition of
Pyronin elements of the mixture). Because they are mixtures, Neomycin B is referred
to as Neomycin and Pyronin Y as Pyronin for the rest of this chapter in relation to the
material used in the assays.
Solutions containing Rev or mixture of Rev and Neomycin at different
concentrations were tested over the surface to find values of Rev that elicited an
appreciable response on the surface but could be significantly inhibited by Neomycin.
An optimal concentration 15µM of Neomycin was found to reduce the plasmon
response of 8µM Rev to ~20% of the response of the Rev:RRE interaction alone.
Screening all antagonists at the same concentration as Neomycin (15µM), allows
direct comparison of inhibitor potency to Neomycin and also allows leeway in the
event a stronger inhibitor than Neomycin is found in this screen or future compound 133
screens.
Each antagonist was assayed in duplicate and was pre-equilibrated with Rev
(50mM Hepes (pH 7.4), 300mM NaCl, 3mM EDTA, 25ºC) at a final concentration of
8µM Rev and 15µM of the antagonist typically for a period of between 30min – 2h,
except for the second screening of Pyronin where Pyronin was tested 10min after
equilibrating as the apparent inhibition was noted to be similar to Neomycin. The
assays were conducted using two flow cells, the first a control channel with no RBE3
bound and the second an active channel containing the RBE3 RNA. The flow rate
used was 30µL/min to reduce any mass transfer limitations (Glaser,1993; Schuck and
Minton 1996; Schuck 1996) that may be present in the system and 40µL of the mixed
protein/antagonist solution was injected over the surface (INJECT command). Five
minutes after the time the injection was started, the flow rate was changed to 5µL/min
and 20µL of the regeneration solution (50mM Hepes (pH 7.4), 450mM NaCl, 0.1%
SDS) was injected, followed by 1min of the running buffer only at 5µL/min. The
surface was flushed by changing the flow rate to 50µL/min for 5min to remove any
remaining extra salt or SDS before changing to 30µL/min for 15min to allow the
surface to settle before the next injection.
-50
1950
3950
5950
7950
9950
11950
-10 10 30 50 70 90 110 130 150 170 190 210 230 250 270 290 310 330 350 370
RU
Res
pons
e
sTime Figure 5.9: Corrected Sensorgrams of Competitive Inhibition of Antagonists vs Rev. Shown is the
duplicated assay of the competitive binding of Rev (8µM) to immobilised RBE3 RNA in the presence
of small molecule antagonists (15µM each antagonist).
The data was analysed by subtracting the control channel from the
sensorgrams created on the active channel to remove bulk effects and correct for non-
specific binding to the SA chip surface (Fig 5.9) and the maximal response was 134
averaged from the two screens with the exception of Pyronin, with the difference in
response of the replication of the assay below 10% for all data except for Kanamycin
(11%) and Pyronin (30%).
The maximal responses were converted to percentage activity in the presence
of inhibitor by applying the formula:
% Activity = (RMaximal Inhibitor / RMaximal Rev) x 100
Where RMaximal Inhibitor is the maximum response of Rev in the presence of the
individual antagonists and RMaximal Rev is the maximum response of Rev in the absence
of the antagonists (Fig.5.10).
0
10
20
30
40
50
60
70
80
90
100
% A
ctiv
ity
Spectinomycin Amikacin
Paromomycin Bekanamycin
Streptomycin Neomycin
Pyronin Pyronin
Tobramycin Kanamycin Rev
Figure 5.10: SPR Analysis of Small Molecule Inhibition of the Rev:RRE interaction. Shown is the
relative response expressed as percentage of Rev activity on the RBE3 surface in the presence of small
molecule antagonist (average of two experiments). The Rev column is the activity of Rev in the
absence of antagonists. Pyronin is shown twice as its effect differed by more than 10% in the two
experiments. In one Pyronin was comparable to Neomycin and in the second was similar to
Bekanamycin.
The screen was also internally compared to Neomycin activity by applying the
formula:
% Inhibitioncf Neomycin = (RMaximal Rev - RMaximal Inhibitor)/ (RMaximal Rev - RMaximal Neomycin) x 100
135
Where RMaximal Neomycin is the maximum response of Rev in the presence of
Neomycin. (Fig. 5.11).
0
10
20
30
40
50
60
70
80
90
100
% In
hibi
tion c
fNeo
myc
in
Spectinomycin Amikacin Bekanamycin
Streptomycin Paromomycin TobramycinKanamycin Neomycin
Figure 5.11: Assay Data Expressed as Percentage Inhibition Compared to Neomycin. The
aminoglycosides ability to decrease Rev activity towards the RRE is expressed as a percentage
compared to Neomycin.
From both screens it was found that the order of potency was Neomycin >
Tobramycin > Paromomycin > Bekanamycin > Spectinomycin > Kanamycin >
Amikacin > Streptomycin (Fig. 5.10 and 5.11). Pyronin was the only compound to
show conflicting data. In the first assay Pyronin gave “apparent” strong inhibition of
activity, but in the second screen exhibited poorer inhibition to a level approximately
equal to Bekanamycin (~58% the activity of Neomycin). In figure 5.10 and 5.11 the
averaged data is presented except for Pyronin data that is presented individually in
figure 5.10 and is omitted from figure 5.11 due to the ambiguity of its inhibition.
The antagonists chosen to be tested in the high-throughput assay have all been
reported to inhibit the Rev:RRE interaction, hence inhibition values for these
compounds are known (Zapp et al., 1993; Hendrix et al., 1997) and agreement with
literature values may serve as a guide for the effectiveness of the assay for application
to unknown compounds. The assay successfully and accurately ascribed the order of
potency in the repeated screen in general agreement with other assay trends (Zapp et
al., 1993; Hendrix et al., 1997). In considering the structure-activity relationships
(SARs) of the screen it seems that shape as well as charge influences the recognition
of the RRE by the antagonists. The Neamine template is the common feature of the 136
137
most potent antagonists and seems critical in establishing the template on which
charged groups are presented to the RBE3 aptamer. However none of the inhibitors
were very potent. The important positions of the charges can be mapped by
comparing the differences in potency and the differences in substitution pattern (Fig
5.8). This was most apparent with the improved potency of the 4-6 disubstituted 2-
DOS (Neoribostamine-like) Tobramycin over Paromomycin. In some respects this is
surprising as Paromomycin is of the Neomycin family and different to Neomycin B
by only the 6-ethylamine (Neomycin) to 6-ethanolic (Paromomycin) functionality of
the substituted β-D-(+)-glucoring.
It is likely that the differences between Tobramycin and Paromomycin are not
a gain in potency of Tobramycin per se, but more a loss in potency of Paromomycin
compared to Neomycin, suggesting the 6-ethylamine functionality is very important
to binding to the RBE3 sequence, so much so that the Neomycin-core structure of
Paromamycin has a loss in potency to levels similar to Neoribostamine substitutions.
When comparing Tobramycin to the related 4,6–disubstituted DOS (Neoribostamine-
like) compounds Bekanamycin, Kanamycin and Amikacin, it is possible the 4’
functionality of the substituted β-D-(+)-glucoring may have greater affinity if it is not
a proton acceptor and the decreased activity of Kanamycin and Amikacin from
Bekanamycin is attributed to the 3-amine/3-hydroxyl substitution of the β-D-(+)-
glucoring. The decreased activity of Amikacin over Kanamycin can be attributed to
the DOS change of the 1-amine to a 1-amino-substituted functionality that
significantly adds size, shape and charge to the base Neamine substitution pattern.
The compounds Spectinomycin and Pyronin have more structural similarities
to each other than the aminoglycosides of the screen and both behaved poorly
compared to the aminoglycosides. Both are more rigid due to the planar 3,6-
dimethylamine-dibenzopyrilium structure of Pyronin and the oxopoly-cyclic structure
of Spectinomycin. However, finer comparisons with Pyronin are difficult as this
mixture exhibited strong inhibition in the first screen but in the second screen
inhibition was approximately equal to Bekanamycin, making quantitation of its
relative potency ambiguous. It is likely that, since Pyronin was a complex mixture,
that strong inhibition was noted in the first screen due to the longer duration the Rev
solution was pre-equilibrated with the Pyronin mixture. This may have caused
denaturation of the Rev protein during equilibration which led to a decrease in
138
activity for the RBE3 RNA surface and hence a decrease in plasmon response of Rev
to the RBE3 surface as opposed to competitive inhibition during the assay. In the
second screen, the order in which the inhibitors were screened was changed and
Pyronin was assayed first, dramatically decreasing the time the Rev protein was
exposed to the Pyronin mixture. Under this shorter time period the Pyronin mixture
showed far less apparent inhibition (~ 58% of Neomycin). The Pyronin solution was
also suspected to damage the RBE3 RNA surface, as the second competitive assay
screen had a lower response of Rev on the sensor chip surfaces. No further structure
activity relationships can accurately be drawn from Pyronin or Spectinomycin due to
their dissimilarity to the rest of the screen.
The least potent anatagonist was the substituted-biosamine Streptomycin that
possesses guanidinium-like imines similar to the RRE binding site of Rev but less
basic in nature. Since this compound is the least active in the screen, it is likely there
is a fundamental conflict in its shape and charge distribution for binding to the RRE.
Because of the differences in substitution to the other Neobiosamine aspects of the
aminoglycosides in the screen, it cannot be stated if poor inhibition by Streptomycin
is due to the differences of the core substituted-biosamine or the guanidinium-like
imines.
From the screen, the common functionality correlating with potency is the
Neamine-like aspects of the four most potent inhibitors and it is likely that the
Neamine base structure is a critical element to the inhibition of the Rev:RRE
interaction. This is in agreement with Zapp et al., (1993) who initially screened
aminoglycoside inhibitors and their derivatives, and Park et al., (1996) who
developed synthetic Neamine-based 4-5 disubstituted 2-DOS compounds that were
more potent than Neomycin (Fig 5.3). However this conclusion conflicts with that of
Hendrix and co-workers (1997), who claim the affinity of the aminoglycosides for the
RRE is similar for non-specific RNA and attribute binding to be a function of the
number of amines and overall charge. There are a few omissions from the Hendrix et
al., (1997) study that make their conclusions questionable. Firstly, though they go to
lengths to study Paromomycin, they do not include the Paromomycin data in their
pooled aminoglycoside SAR analysis (Figure 8 Hendrix et al., 1997) and from their
conclusions that charge alone was important, it would be predicted that Bekanamycin,
Tobramycin and Paromomycin would all have equal potency based on their similar
139
charges. The fact that their own experimental data does not agree with this suggests
their conclusion is incorrect.
It can be concluded from the studies here that the shape of the
aminoglycosides could be important to conform to the RBE3 structure and present
specific charge interactions similar to the guanidinium groups of Rev for RRE
recognition. A KD was not investigated for Neomycin or other small molecule
antagonists using the competitive assay, however the strength of the high-throughput
assay reported here is the speed with which it can be conducted on the same surface
of RNA, while relating inhibition to the control antagonist Neomycin. It would be
worth investigating more closely the SAR of a larger pool of aminoglycosides to
improve the structure-activity arguments made here. Ideally such a screen would
comprise more potent antibiotics including 4,5-disubstituted 2-DOS compounds that
were found to be so effective by Park et al., (1996). However, the inhibition of the
Rev:RRE by known aminogylcoside antibiotics is poor and other inhibitor classes and
strategies such as de novo design using cyclic peptides, peptide-nucleic acid
analogues and helix mimetics could prove more successful as an approach to
inhibition to develop a suitable therapeutic.
5.2.2 Assay of Helix Mimetics.
An attempt was made to measure the kinetics of the Rev:RBE3 association
directly using the same buffers and conditions as described above (a running buffer of
50mM Hepes (pH 7.4), 300mM NaCl, 3mM EDTA, 25ºC, and a wash solution of
50mM Hepes (pH 7.4), 450mM NaCl, 0.1% SDS), but an unacceptable level of non-
specific associations was apparent at lower protein concentrations on the SA surfaces.
The general practise when conducting a BIAcore-based assay is to subtract most non-
specific and bulk effects from a blank injection (buffer only) and subtracting the non-
specific events from a control channel that is similar to the active channel (BIAcore
1998a; BIAcore 1998b; BIAcore 1998c). However, non-specific interactions still
occurred when using different flow rates ranging from 5µL to 50µL and testing buffer
subtractions on control surfaces comprising the same surface as the active channels
but without captured RNA.
To reduce the non-specific interactions in this system (Fig. 5.12), a range of
strategies was devised and tested, (1) decreasing the amount of surface charges by
using low-density carboxymethyldextran chips and immobilising low amounts of
capture protein(s), (2) trying different capture proteins that are more chemically inert
in the system, (3) using agents in the buffers to mask either the surface, capture
protein or analyte charges and (4) removing bulk and remaining non-specific effects
by subtraction of controls.
Neutravidin and Streptavidin biotin-capture proteins were tested on Pioneer-
B1 sensor chip surfaces in order to derive a sensor chip platform with decreased non-
specific interactions. Neutravidin is a chemically altered form of Avidin with a pI
~6.3; the Pioneer-B1 chip has been developed with low carboxylation of the dextran
matrix and hence has a lower negative charge on the surface, expected to reduce non-
specific binding where analytes with a high degree of positive charges are used (such
as Rev and the Rev-based peptides 2 – 4). The chips were docked and surfaces
prepared as described in the Results section 5.2.1.
Figure 5.12: A stylisation of the Surface Plasmon Resonance Assay Showing Possible Non-specific Interactions. The figure represents biotin-labelled RBE3 RNA (ligand) captured onto a Neutravidin/Streptavidin protein (red) immobilised onto the sensor chip surface (gold) via a carboxymethyldextran layer (blue). The RNA is shown interacting with the helical Rev34-50 peptide in the centre of the figure (analyte) (figure generated using co-ordinates 1ETF from the pdb in Insight II). A change in the refractive index on analyte binding (∆θ) over time generates sensorgrams. Non-specific associations can form with RNA indicated at 1 to the RNA, 2 to the capture protein, or 3 to the carboxymethyldextran layer or gold film surface.
140
1 1 2 3
∆θ
The biotin-capture proteins, Streptavidin and Neutravidin were immobilised
141
onto Pioneer B1 chips using modified procedures based on BIAcore
recommendations that advise a 200 – 400µg/mL solution at pH 4.5 – 5 for
Streptavidin and 50µg/mL solution at pH 4.5 for Avidin. Neutravidin, as a chemically
altered form of Avidin, was assumed to have immobilisation conditions between
Avidin and Streptavidin but requiring a more acidic pH to avoid precipitation at pH
values close to the pI (~6.3). Streptavidin (Pierce batch 5063S) was reconstituted in
BIAcertified Acetate buffer (10mM pH 5.0, no salt, 4°C) to 300µg/mL based on the
mass provided by the supplier. Neutravidin (Pierce batch 0443W) was resuspended in
ddH2O to a stock concentration of 10mg/mL and was diluted in 10mM Acetate buffer
(pH 4.0, no salt, 4°C) to ~147.5µg/mL, confirmed spectrophotometrically (A1%280 =
1.66). Both proteins were incubated on ice for 30min after dilution and were
immobilised following a standard BIAcore protocol under continuous flow (5µL/min
HBS-EP buffer). The surface was activated using 25µL of EDC/NHS (final
concentrations 50mM and 200mM respectively) injected over the surface followed by
35µL of the capture protein before neutralising the surface with 35µL of
ethanolamine (1M, pH 8.5).
Stock biotin-RBE3 RNA (40µL aliquots, 129.8µg/mL, 40%v/v formamide)
was diluted to 200µL of 50mM Hepes pH 7.4, 300mM NaCl and annealed by heating
to 100ºC for 10min and slowly cooling to 25°C (~60min). The biotin-RBE3 RNA
solution was not further diluted as Pioneer-B1 chips are resistant to 40% formamide
(8%v/v in final conditions) and the RNA was captured onto the chip surfaces at a
flow rate of 5µL/min with injections of 200µL of the RNA solutions. The RNA-
captured surfaces were washed at 5µL/min with two washes of 1M NaCl (5µL) for
the Streptavidin-B1 surfaces and 2M MgCl2 (5µL) followed by 1M NaCl (5µL) for
the Neutravidin surfaces. The surface density of immobilised capture proteins was
different for Neutravidin and Streptavidin and the ability of the two capture proteins
to bind biotinylated RBE3 RNA in this system was also different. Neutravidin was
immobilised to a level of 5534RU on the Pioneer B1 surface and captured 890RU of
biotin-RBE3 RNA whereas Streptavidin was immobilised to a level of 1773RU on
the Pioneer B1 surface and captured 790RU of biotin-RBE3 RNA.
It was found that both the Neutravidin and Streptavidin Pioneer-B1 chips had
greatly reduced non-specific interactions and were far more sensitive to analytes than
142
SA chips. On the Streptavidin-B1 and Neutravidin-B1 chip, a control channel was
made following the same protocols as outlined above so that the surface was identical
to the active channel except for the omission of RBE3 RNA. The regeneration
solution of the high-throughput screening conditions (50mM Hepes (pH 7.4), 450mM
NaCl, 0.1% SDS) was found to damage the Neutravidin-B1 control surface and in
general regeneration buffer conditions were found to vary on all the different surfaces
(SA, Streptavidin-B1 and Neutravidin-B1), requiring optimisation on each chip and
surface. Ideal regeneration conditions were examined by experimenting with a range
of regeneration conditions and concentrations in the presence of a constant protein
(analyte) concentration on both the Neutravidin-B1 and Streptavidin-B1 chips using a
flow rate of 10µL/min and 1.14µM Rev protein (5µL injections) and washing the
surface with 5 – 10µL of various regeneration solutions. The buffered regeneration
conditions tested used a range of salts (150 – 750mM of NaCl, LiCl, MgCl2,
(NH4)2SO4 and Na2SO4) both with and without additives (SDS 0.005 – 0.1%, 5mM
DTT or 0.05% Triton X-100) in 50mM Hepes (pH 7.4), and unbuffered solutions of
the salts (0.5 - 2M) were also tested. It was found that the Neutravidin-B1 surfaces
were very sensitive to regeneration conditions with best regeneration conditions
found to be two washes of 2M MgCl2 and one wash of 1M NaCl (10µL injections)
and the Streptavidin-B1 surfaces regenerating from five washes of 1M NaCl (10µL
injections).
Because the Neutravidin-B1 surfaces could detect analytes at lower
concentrations, this sensor chip surface was investigated further for suitability to use
as an assay platform. It was also found that there was varied and unpredictable
behaviour of control channels (either the B1 surface with or without Neutravidin
immobilised) in the presence of Rev at different flow rates, complicating the
correction of non-specific interactions from an active RBE3 channel.
It was hypothesised that additives might be used to reduce non-specific
interactions without significant disruption of the Rev:RBE3 interaction. In order to
test this proposition, an experiment was devised where additives (Table 5.1) were
tested in both the running buffer and the solutions of Rev injected over the surface to
try to mask either the surface, capture protein or analyte charges that create non-
specific associations. The experiment used a 5µL injection of 1.14µM Rev in the
presence of additives (flow rate 5µL/min) onto all four channels of the the B1-Pioneer
chip where the channels were: surface only controls with no immobilised Neutravidin
(flow cell - FC 1 and 2), a Neutravidin control to measure the effectiveness against
Neutravidin non-specific associations comprised of a Neutravidin immobilised
surface but containing no RNA (FC 3) and the active RBE3 captured surface (FC 4).
The experiment was analysed for the change at equilibrium from the baseline (∆RU)
to measure the level of responses (corrected by subtraction of buffer-only injections
for each of the individual additives). The overall change was also measured by
conducting a control experiment under identical conditions without the additives
(RAll). The overall change was calculated by subtracting the experiments without
buffer additives from those with the identical experiment with the additives (overall
change = ∆RU – RAll), thereby gauging the effect of the additives (Fig 5.13 and Table
5.1).
baseline
Overall Change
RAll
∆RU
Figure 5.13: A stylisation of how Additives were Screened for Reducing Non-specific Binding.
The corrected curves (subtraction of a buffer only injection) are Rev in the absence of an additive on
the RBE3 surface (blue) and Rev in the presence of an additive on the RBE3 surface (red). The
subtraction of maximum response from baseline of Rev in the presence of an additive (∆RU) from the
maximal response of Rev in the absence of an additive (RAll) is reported as the overall change in
response.
In general a suitable additive is predicted to have overall changes that will be
negative on all channels, indicating the non-specific associations have been reduced,
lowering the response in the presence of the additives, but maintain a proportionally
strong response to the RBE3 surface and not significantly impede the RBE3:Rev
interaction on the surface. The best additives will therefore result in small changes on
the surface on the control channels (i.e. small ∆RU values for FC 1 – 3), but retain a
strong active channel response on FC 4. In order to gain an indication of the
selectivity of Rev for the RBE3 in the presence of an additive for the RBE3 over the
143
144
Neutravidin control channel, the data was further analysed by comparing the
difference between the response on the active channel (FC 4) and the Neutravidin
immobilised channel (FC 3) divided by the Neutravidin channel i.e. ([FC 4 – FC
3]/FC 3).
Compound tested Possible mode of action FC 1 – 4 ∆RU Overall change
3.5mM Spermidine Surface charge masking 18, 16, 25, 90 -38, -32, -33, -67 5mM (NH4)2SO4 Surface and Rev charge masking 40, 36, 65, 218 -16, -12, 7, 61
20mM Gu.HCl Surface charge masking 62, 55, 102, 204 6, 7, 44, 47 5mM ethylenediamine Surface charge masking 90, 81, 101, 197 34, 31, 43, 40
17mM Imidazole Surface charge masking 82, 72, 90, 166 26, 24, 32, 9 23mM Glycine Unknown 75, 66, 82, 156 19, 18, 24, -1
10mM Glutamate Unknown 60, 53, 88, 133 4, 5, 30, -24 10mM N-Acetyl-
Glucosamine Unknown 69, 62, 94, 170 13, 14, 36, 63
10µg/mL Calf Thymus
DNA
Rev charge masking 5, 6, 13, 69 -51, -42, -45, -88
8mM Hippuryl-Arg Surface charge masking 87, 79, 147, 214 56, 31, 89, 57 33.57mM Ammonium
Acetate Surface and Rev charge masking 68, 58, 121, 199 12, 10, 63, 42
4.4mM Lys-Pro Surface charge masking 83, 74, 137, 186 27, 26, 79, 29 1.75mM Spermidine,
5µg/mL + Calf Thymus
DNA
Surface and Rev charge masking 12, 11, 22, 63 -44, -37, -36, -94
Table 5.1: Effects of Buffer Additives on Non-specific Interactions. The table lists compounds
tested as buffer additives, concentration and possible mode of reducing non-specific interactions. The
flow cells were the standard B1 surface with no ligand (FC1, FC2), the Neutravidin surface (FC3) and
the RBE3 captured surface (FC4). The ∆RU column is the raw change from baseline to equilibrium of
Rev in the presence of the additives and the column “Overall change” is the difference to responses
without the additives. Omitted is Heparin (10µg/mL) as this was completely detrimental to Rev
causing precipitation before it could be applied to the sensor chip.
The best compound was calf thymus DNA with 4-fold greater selectivity and
small ∆RU values for FC 1 – 3 and negative values for the overall change. The next
best compound was spermidine with ~2.5-fold selectivity and relatively small ∆RU
values for FC 1 – 3 and negative overall changes, followed by the spermidine/calf
thymus DNA mixture and ammonium sulfate with a similar ~2-fold decrease and
145
favourable ∆RU values for FC 1 – 3, with lower overall changes for the
spermidine/calf thymus DNA mixture than ammonium sulfate. Upon this basis, calf
thymus DNA was selected as an additive for inclusion in all buffers and test solutions
to minimise non-specific associations as it reduced these interactions while allowing
the specific Rev:RBE3 response to be appreciably measured and the control surface
selected was the Neutravidin immobilised control being most like the active channels
except for the capture of RBE3 RNA.
An attempt was made to directly measure the interactions of Neomycin with
the RBE3 RNA as the Neutravidin-B1 surfaces were found to be sensitive enough to
measure small molecule interactions in contrast to the SA chips used in the high-
throughput screen that could not directly detect small molecule binding. Neomycin
was serially diluted in a lower salt running buffer comprising 25mM Hepes (pH 7.4),
150mM NaCl and 10µg/mL calf thymus DNA. Using this buffer, assays were
conducted in duplicate over a 21min cycle with varied flow rates. For association and
dissociation, a continuous flow of 10µL/min was used with a 30µL injection of
Neomycin solutions (0.01µM – 233µM). The surface was regenerated using a
combination of washing and changed flow, where after 100 seconds of free
dissociation the flow was changed to 50µL/min and then pulsed twice with 2M
MgCl2 (15µL) and once with 1M NaCl (15µL) and returning the flow rate to
10µL/min for 5min before the next injection as this milder form of washing was
sufficient to regenerate the surfaces in the presence of the aminoglycoside.
The Neomycin sensorgrams were corrected using a control channel
(Neutravidin only) (Fig 5.14) and the corrected data was found to have a significant
surface event in the association phase (1 and 2 of Fig. 5.14) that was not seen using
buffer alone (lower curves Fig 5.14). In the association phase there is an initial
binding event that is followed by a second, more pronounced surface event that could
be a conformational change of the RNA or multiple binding of the aminoglycoside or
even both (a conformational change brought about by multiple binding), as both a
conformational change upon ligand binding and multiple binding of Neomycin B has
been reported for the RRE RNA (Lacourciere et al., 2000; Cho and Rando, 1999).
It was found that no deterioration of the surface was detectable over the assay
period, but in some runs there was a very sharp spike in the association phase (Fig.
5.14). In such runs there was variation from the maximal response at equilibrium than
previous or subsequent runs at the same concentration. Such events have been termed
“memory effects” and have variously been hypothesised to occur at high analyte
concentrations from a carryover of analyte in the autosampler, needle, micro-fluidic
tubes/loops or integrated micro-fluidic cartridge (IFC), or from non-ideal regeneration
conditions (Stöcklein et al., 1998; BIAcore, 1998c) where high molarity regeneration
solution ions are incompletely washed from the surface before subsequent injections
of the analyte. This could cause a refractive index change since the higher salt
concentration on the surface cannot be subtracted from a buffer-only injection, but
such a hypothesis cannot easily be tested in the BIAcore system. The memory effects
observed here have no discernible pattern that would allow correction and could be
caused by either carryover or regeneration; in figure 5.14 the sensorgrams exhibiting
memory effects have an association phase “spike” at 1 of close to 300 RU that alters
the first association phase change well above replicated runs at the same
concentration and the equilibrium maximal response is observed to be lower by 12 –
16% at elevated concentrations, but as much as 32% at lower concentrations.
-50
0
50
100
150
200
250
300
350
-50 0 50 100 150 200 250 300 350 400
Time s
Res
pons
e
RU
2 3
1 4
Figure 5.14: Replicated Assay of Neomycin interaction with B1 Neutravidin:RBE3 surfaces. The
response of Neomycin (0.01µM – 233µM) interacting with RBE3 RNA is shown. The lowermost
sensorgram (red) is the response to buffer injection alone and was used to subtract bulk RI changes
from the other sensorgrams, as too was a blank channel (Neutravidin only). Shown is the change in the
association phase (1 and 2) before levelling of the response to equilibrium (3). It can be seen that some
of the lower and intermediate concentration data have a large spike in the first association phase (1)
and the maximum response at equilibrium was reduced (as shown at 4 with the lower values
misaligned for emphasis).
146
The data (not including those sensorgrams exhibiting memory-effects) would
not fit a Langmuir kinetic model (Lipshultz et al., 2000; BIAcore 1998c) but were
closer to a two-state (conformational change), although the data could not be fitted
adequately to a two-state reaction using the BIAcore BIAevaluation software. This is
likely a problem with the association phase that does not follow an expected shape for
the model, though there is a deficiency in the BIAcore 2000 machine as
conformational changes can only be fitted if the rate of change occurs over extended
time scales of several minutes (BIAcore, 1998d), whereas the observed changes are
completed within 50 seconds of injection.
A KD was calculated for Neomycin using the maximum response of all data
(including those exhibiting memory effects) from equilibrium (3 of Fig. 5.14) (Raw
data Table 5.2) using GraphPad Prism® (version 4.00 for Windows GraphPad
Software, San Diego California USA, www.graphpad.com). Initially the data was
fitted to one-site and two-site binding hyperbolic curves; both models describe the
binding of an analyte to a ligand following the law of mass action using the following
equations:
One site: Req = Rmax*[analyte]/(KD+[analyte])
Two site: Req = Rmax1*[analyte]/(KD1+[analyte]) + Rmax2*[analyte]/(KD2+[analyte])
Where the values Rmax are the maximal binding response and KD is
dissociation constant equivalent the concentration of analyte required reaching half of
Rmax (1 and 2 for individual sites). The two models were evaluated using Akaike’s
Information Criteria (AIC). AIC is similar to an F-test and balances the changing
goodness of fit as assessed by the Sum of Squares with the change in the number of
parameters to be fitted to the model, but AIC includes more complicated elements of
assessing the goodness of statistical data including entropy of information, maximum
likelihood theory and information theory (Motulsky and Christopoulos, 2003).
147
Conc (µM) Req Conc (µM) Req Conc (µM) Req0.1 4.00 5.83 40 100 187 0.1 8.00 10 55.50 117 193 1 18.80 10 52.60 117 171 1 23.00 58.3 167 175 229 1 19.40 87.4 138 175 229 1 22.90 87.4 165 233 269
5.83 27.00 100 183 233 271
Extending the
range of this fit to the upper limit of Req (205), the single site model is still favoured
and the fit is improved in terms of the r2 value (0.97), with an Rmax of 226 ± 16 and
higher KD of 29 ± 7µM. However the fit is strongly influenced by higher
concentration data points, raising the KD value (Fig. 5.16). The KD is also found to be
greater than the KD1 for the two site model of the entire data set (15 ± 11µM cf 29 ±
7µM), suggesting that if there are two sites where Neomycin binds, the second site
Req values may be part of the upper region of the fit around Req values of 205 and
skewing the data away from the lower range of the KD.
Table 5.2: Concentration and Req values for Neomycin Interactions
with Custom RBE3-Neutravin-B1 surfaces.
0 2.5×10-5 5.0×10-5 7.5×10-5 1.0×10-4 1.2×10-4 1.5×10-4 1.7×10-4 2.0×10-4 2.2×10-4 2.5×10-40
100
200
300
Conc
Req
Linear Region
1.0×10-7 1.0×10-6 1.0×10-50
10
20
30
40
50
60
Conc
Req
Figure 5.15: Uncorrected Non-linear Regression Analysis of Req vs Concentration for
Neomycin:RBE3 Binding (two site model). The top graph shows all the data fitted to the two site
model with a linear region indicated. The lower graph uses a log10 display of the concentration (X-axis)
and Req values below 131. The fit of the data below the predicted Rmax1 value (131) from the raw fit of
all the data assayed. KD = 1.8 ± 0.8µM, r2 = 0.85, n = 10.
148
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
50
100
150
200
250
Conc
Req
Figure 5.16: Non-linear Regression of Neomycin Req vs Concentration log10 (below Req = 205).
The fit of the data below the predicted Rmax1 value (131) from the raw fit of all the data assayed. KD =
29 ± 7µM, r2 = 0.97, n = 17.
A serious problem is found in the two site model as the data does not achieve
Rmax2 (Fig. 5.15) even at the high analyte concentrations used here, resulting in an
over prediction of Rmax2 (Rmax2 0.21e6 ± 25e6, a 77490% over predicted than the
largest Req value). The fitting of the second site to this high Rmax2 value causes a
mathematically related over prediction of KD2 (KD2 331mM ± 41M). Associated with
this high Rmax2 prediction, the higher concentration data (upper points) of the fit forms
a more linear relationship with a slope of 692800 and r2 0.91 (Fig. 5.15). Parameter
Uncorrected Single site
Uncorrected Two site
High Affinity Single site (131)
High Affinity Single site (205)
Rmax1 131 (74) 56 (7) 226 (16) KD1 15µM (11µM) 1.8µM (0.8µM) 29µM (7µM) Rmax2 352 (35) 0.21e+6 (25e+6) KD2 91µM (22µM) 331mM (41M)
Goodness r2 0.9672 0.9837 0.8534 0.9705 Abs SS 5860 2907 393.5 2690 Sy.x 17.56 13.08 7.013 13.39 AIC Probability 1.68% 98.32% 96.52% 83.59% Ratio 58.44 27.74 5.09 AICc 8.14 -6.65 -3.26
Table 5.3: Nonlinear Regression Analysis of Req vs Concentration for Neomycin:RBE3 Binding.
For the fit of the data the AIC = N·ln(SS/N)+2K and AICcorrected (AICc) = AIC·2K(K+1)/N-K-1, where
N is the number of data points and K is the number of parameters to be fitted. The Probability is AIC
weights, the Ratio is the evidence ratio based on absolute differences of AICc. Numbers in brackets are
the standard errors associated with the fit of individual parameters. The values r2 is the squared
multiple correlation coefficient, Abs SS is the absolute sum of squares and Sy.x is the standard
deviation of residuals.
149
150
A transformation of the data was conducted to form an upper limit of binding
by reducing the influence of the linear portion of the fit. This linear portion may be
influenced by non-specific binding to the fit due to many individual or combination
of factors in this assay including: multiple analyte binding to the specific RBE3 RNA
sites, heterogeneous analyte problems (since Neomycin used here is a mixture of
Neomycin B and Neomycin C), non-specific RNA association that is not corrected on
the control channel, or a heterogeneous ligand surface as the labelling technique is
random and the RBE3 sequence contains six U’s that are not part of the 3’ poly U tail
and could be biotinylated and provide non-specific binding affinity measured in the
BIAcore.
In order to reduce the effect of the linear region of the fit (Fig 5.15), a
correction factor was applied to all data following the equation:
Rnon-specific = 0.69 x C(µM)
Where Rnon-specific is the portion of Req that is due to non-specific associations,
0.69 is slope of the linear region expressed as 10-6M and C(µM) is the concentration
expressed in microM. The corrected values were then calculated by subtracting the
non-specific binding term from each point of Req:
Rcorrected = Req - Rnon-specific
This yields a value of Rcorrected, which is the Req value corrected for non-
specific associations. The results for the single site and two site fit for the corrected
data is presented in figure 5.17 and the table embedded therein. The AIC analysis
significantly favours the single site model and it can be seen that correcting for non-
specific associations has changed the upper curve shape forming a defined Rmax for
RBE3 interaction figure 5.17.
0 5.0×10-5 1.0×10-4 1.5×10-4 2.0×10-4 2.5×10-40
15
30
45
60
75
90
105
120
135
Conc
Rco
rrec
ted
Figure 5.17: Corrected Non-linear Regression Analysis (one site binding model) of Neomycin
interacting with RBE3 RNA (corrected data). Also shown in the figure is the regression analysis
using parameters as for the legend in Table 5.3. A 92% confidence interval of the corrected single site
model gives an Rmax of 118 ± 11 and the KD based on this range is between 13 ± 6µM.
Parameter
Corrected Single site
Corrected Two site
Rmax1 62 (38417) KD1 11.1µM (1mM) Rmax2 118 (6) 55.86 (0.4e+6) KD2 13µM
(3µM) 15µM (1.6mM)
Goodness r2 0.9235 0.9235 Abs SS 3007 3007 Sy.x 12.58 13.30 AIC Probability 96.43% 3.57% Ratio 27.00 AICc -6.59
The analysis of the corrected data (Fig 5.17) shows the two site and one site
model have similar values of the fit in terms of r2, sum of squares and standard
deviation of the residuals, with the one site model significantly favoured by the AIC
analysis. In agreement with the AIC analysis, the two site model predicts a lower
Rmax2 value than the Rmax1 value with a subsequent large error, and the values of KD1
and KD2 are similar, suggesting there is only one binding site for Neomycin and both
Rmax values are trying to be fit to the same maxima, therefore the one site model is
taken as correct.
It is worth noting that similar to fitting to the high affinity site in figure 5.15
the fitting to a single site below the Req value of 62 for the corrected data yielded a KD
of 1.2 ± 0.5µM, but the r2 value is similar to figure 5.15 (r2 0.85) and the plot does not
reach Rmax. Another aspect of the corrected fit in figure 5.17 is an outlying data point
(Req 127, Conc 58µM) and removal of this data point changes the fit to a two site
model with improved r2 value (0.96) and KD1 value ≤0.8µM and KD2 ≤54µM based
on the highest values of a 96% confidence interval of Rmax1 (37) and Rmax2 (125). The
removal of this point from the uncorrected data (Req 167, Conc 58µM) similarly
improves the fit (r2 0.99) and KD1 value ≤2.5µM and KD2 ≤432µM based on the
151
152
highest values of a 99% confidence interval of Rmax1 (56) and Rmax2 (674). Although
the removal of this data point improves the fit and final data analysis, the inclusion of
all points is practised here, and the results of the removal of this point and
independent investigation of the lower concentration data suggests the RBE3 RNA
aptamer may have two sites for binding Neomycin with a KD1 value ~1µM, however
this is not seen here without accepting too many problems with the data or removing
a data point. In this respect the one site model of the corrected data is taken as
appropriate based on the data with a KD for Neomycin of 13 ± 3µM. The maximum
error range is a 92% confidence interval of the Rmax value and is 13 ± 6µM.
The quoted KD values in the literature vary from as low as 210nM under
similar assay conditions used here (Hendrix et al., 1997) to 10 – 25µM for
competitive assays (Van Ryk and Venkatesan, 1999), with a commonly accepted
value range of 1 - 6µM (Zapp et al., 1993; Kirk et al., 2000; Chapman et al., 2002).
In this respect the KD found here is not unreasonable considering the variation of
literature values. The problems encountered with the fitting to the data is a reflection
of the imperfection of using data that contains errors due to non-specific binding and
a better analysis could be made using a non-specific RNA control channel and
assaying a broader range of concentrations below 100µM and especially below Req
values of 50, which should improve the fit for the lower value of the KD as suggested
from the “high affinity single site” analyses and the data analysed with the outlying
point removed.
In summary, Neutravidin (pI 6.3) was tested as a capture protein on B1-
Pioneer chips. Neutravidin is a deglycosylated form of Avidin and more neutral than
either Avidin (pI 10) or Streptavidin (pI ~5) and it was thought might exhibit less
non-specific interactions since RNA has a more acidic pI and most RNA binding
proteins such as Rev have a basic pI. In these experiments the use of Neutravidin was
found to elicit variability in behaviour with captured RBE3 RNA, and the
regeneration of these surfaces was difficult and inconsistent. Some of the problems
were more apparent when assaying Neomycin affinity for the RBE3 aptamer,
especially memory effects and non-specific binding. The memory effects occurred
when the surface was exposed to Neomycin over extended assay periods and was
characteristic with a sharp spike in the association phase followed by a different
response at equilibrium compared to repeated runs using the same concentration. The
153
non-specific interactions were minimised using 10µg/mL calf thymus DNA in the
running buffer and injected solutions of analytes, however there was evidence that
further non-specific associations were occurring on the Neutravidin-B1 surface as
assigning a KD for Neomycin required correcting for a linear region found in the plot
(Fig 5.15), which was attributed to non-specific binding and was more apparent at
elevated Neomycin concentrations. Despite these deficiencies in the system, the
corrected non-linear regression analysis found a KD for Neomycin of 13 ± 6µM.
The most stable surfaces were found be Streptavidin-B1 custom surfaces, and
these were used for the remainder of the study. The immobilisation level of
Streptavidin was 32% of Neutravidin, and was possibly due to the buffer conditions
used (pH 5), which is identical to the pI and so precipitation likely occurred reducing
the actual protein concentration used for immobilisation. However, the capture of
biotinylated RNA was much greater than for Neutravidin with only 11% less biotin-
RBE3 RNA captured on the Streptavidin-B1 surface despite the reduced
immobilisation of protein.
The Streptavidin-B1 surfaces were tested for ideal regeneration conditions,
and best running conditions with minimal non-specific interactions apparent using a
moderate flow rate (20µL/min) and calf thymus DNA at 10µg/mL concentration in
the Rev protein solution and running buffer similar to the Neutravidin-B1 surfaces.
The best regeneration conditions were found to be a series of five washes of 1M NaCl
with changed flow rates and resting of the surface for full regeneration and avoidance
of memory effects. The first wash was 5min after the end of the injection with a 20µL
injection (flow rate 20µL/min), followed by a change in the flow rate to 50µL/min for
1min and then a change in the rate back to 20µL/min before subsequent washes were
conducted. The surface was finally rested for 17min (constant flow rate of 20µL/min)
before a second assay point was taken, making the total run times of 50min for a
100µL injection.
In a non-competitive assay each peptide would need to be assessed
individually and the different affinities compared from the same conditions that
ideally would contain low concentrations of salts as higher salt conditions affect
Rev34-50 peptide helicity (Hendrix et al., 1997). A complication occurs under low salt
conditions since the full length Rev protein does not fold well or remain soluble at
ionic strengths below 300mM (as reported in Chapter 4). Since the flash folding
154
conditions of Chapter 4 allow for a lower protein concentration, it was thought the
Rev protein may retain activity in a lower salt buffer by flash folding. The Rev
protein was tested under increasingly stringent conditions between (150 – 500mM)
over appreciable assay time periods by dilution from 0.5mg/mL (39µM) Rev in
50mM Hepes (pH 7.4), 2M Gu.HCl, 500mM NaCl, 0.5mM EDTA at 4°C, to a
solution of 98.5nM Rev with the various salt buffers (50mM Hepes (pH 7.4) and
10µg/mL calf thymus DNA at 4°C) and then leaving this on ice for 30min before
further serial dilution with the various salt conditions in buffer (50mM Hepes (pH
7.4) and 10µg/mL calf thymus DNA at 4°C). At salt concentrations of 200mM and
150mM the response from repeated injections was not reproducible but from 300mM
solutions the response was found to be consistent. It was also found that flash folding
1:5000 from 0.5mg/mL (39µM) Rev in 50mM Hepes (pH 7.4), 2M Gu.HCl, 500mM
NaCl, 0.5mM EDTA at 4°C, to ~8nM, and then a further 1:10 dilution (~0.8nM)
concentration was acceptable in the BIAcore, generating sensorgrams detectable
following correction using an appropriate control with a similar dilution of the stock
buffer. There was no detectable signal when the protein was diluted directly from the
39µM stock to 0.8nM. There was little difference using the flash folded material or
Rev protein that had gone through serial dilution, with 5% difference from responses
to 30nM protein used from these different preparation pathways (Fig. 5.18a). The
response from flash folded material was suitable for up to four injections (200min
using 50min assay cycles) before the protein decreased response on the surface and
3h was the maximum time period any assay was conducted (meaning each
preparation was used for a maximum of three points) (Fig. 5.18b). Because the flash
folding conditions decreased the work-up time needed for successive dialysis this
method of protein preparation for the BIAcore assays was adopted for the
experiments described below.
The Rev34-50-based peptides 2 – 4 were supplied by Michael Kelso from the
laboratory of Professor Fairlie at the IMB (UQ). The three peptides were synthesised,
purified by RP-HPLC, characterised and provided in lyophilised form (2, 1.95mg; 3,
0.92mg; 4, 0.74mg). The peptides were reconstituted in 50mM Hepes (pH 7.4),
300mM NaCl, and separated into 100µL aliquots that were snap frozen in liquid
nitrogen and stored at -80°C until required.
20
40
60
80
100
120
140
160
180
-60 -52 -44 -36 -28 -20 -12 -4 4 12 20
Time s
Res
pons
e
RU
155
-25
75
175
275
375
475
575
-100 100 300 500 700 900 1100 1300 1500
Time s
Res
pons
e
RU
Figure 5.18: Ananlysis of Flash Folded Rev protein on Streptavidin-B1-RBE3 Chips. (a) Rev protein (30nM) from serial dilution and dialysis (green and blue upper sensorgrams) compared to flash folded material (red and blue lower sensorgram). The dialysed material had a maximum response of 165RU, whereas the flash folded protein had a minimum response of 156RU (5% variation). Both sensorgrams were corrected using dialysis buffer or the storage buffer following a similar dilution. (b) The flash folded Rev was assessed by repeated injection. The upper four most curves (uncorrected curves, slightly misaligned) highlight the reproducibility for 3hrs 20mins and beyond this time the protein deteriorated rapidly.
a
b
The assays were conducted at a flow rate of 20µL/min and the peptides or Rev
protein were diluted to running buffer conditions (50mM Hepes (pH 7.4), 300mM
NaCl and 10µg/mL calf thymus DNA) by serial dilution of the peptides or by flash
folding from 39µM directly to final conditions (maximum 1:5000 dilution) in the case
of the protein. Various concentrations of peptides (2, 10nM – 40µM; 3, 6nM – 93µM;
4, 2nM – 34µM) or protein (0.8nM – 197nM) were applied using the KINJECT
command (10min total contact time 2µL/min for peptides and 10µL/min for protein).
The data was corrected for non-specific binding and bulk effects by
subtracting a control channel (Streptavidin-B1 channel) and a blank injection (buffer
only) (Fig. 5.19). Protein concentrations were confirmed using the methods described
in Chapter 4 (Experimental 4.3.7) and peptide concentrations were based on
concentrations determined by Amino Acid Analysis conducted under contract by
Auspep (Melbourne).
Similar for the Neomycin-Neutravidin-B1 experimental data, the response at
equilibrium (Req) was plotted against concentration (Conc) and was fitted to the one
site and two site binding models using GraphPad Prism® (version 4.00 for Windows
GraphPad Software, San Diego California USA, www.graphpad.com), and the fits
were assessed using AIC.
-10
60
130
200
270
340
410
480
550
-10 60 130 200 270 340 410 480 550
RU
Res
pons
e
sTime
-100
100
300
500
700
900
-200 0 200 400 600 800 1000 1200
RU
Res
pons
e
sTime
-100
0
100
200
300
400
500
600
700
800
900
-200 0 200 400 600 800 1000 1200 1400
RU
Res
pons
e
sTime
-200
0
200
400
600
800
1000
1200
-200 0 200 400 600 800 1000
Time s
Res
pons
e
RU
Figure 5.19: Plots of Plasmon Response vs Time for Rev and Peptide interactions with RBE3
RNA. Corrected sensorgrams of Rev (top left), the native peptide 2 (top right), the succinylated
peptide 3 (bottom left) and the capped peptide 4 (bottom right).
It was found that data for all three peptides did not fit to a one site model
based on r2 values and the AIC analysis, but the Rev protein data was able to be fitted
to a one site model. The effect of correcting the data for a linear region, as found with
the Neomycin analysis, was also investigated by correcting data for the slope of upper
data points by ascribing a linear region in Microsoft® Excel (2000 Professional
Edition, www.microsoft.com) as follows: 2, 10x106 (r2 0.98, n = 6); 3, 4x106 (r2 0.96,
n = 10); 4, 10x106 (r2 0.90, n = 8) and Rev 1x109 (r2 0.87, n = 5). For the peptide 4 the
r2 value could be improved to 0.99 by sequential removal of points to n = 4 without
affecting the slope in Microsoft® Excel. The corrected Req values were then calculated
using the slope to transform the data as for the Neomycin investigation with the
following equations:
Rnon-specific = Slope x C(µM or nM) Rcorrected = Req - Rnon-specific
Where Slope is the slope of the linear portion of the plots x106 (except for the
Rev protein which is x109 – being nanomolar), Rnon-specific is the portion of Req that is
due to non-specific associations and C(µM or nM) is the concentration expressed in
microM or nanomolar – in the case of Rev. The corrected values were then calculated
by subtracting the non-specific binding term from each point of Req to give the value
Rcorrected.
156
Native Peptide (2).
The native peptide (2) was found to fit to all models except the single site
corrected fit in terms of the r2 values with best fits predicted from the AIC analysis to
be the two site model for both corrected and uncorrected data (Table 5.6). The
uncorrected data gives the best values for the fit, but the Rmax2 is not reached (1408)
(Fig 5.20) and the value is over-predicted by 160% of the highest Req value and this is
reflected in the large prediction for KD2. In contrast, though the corrected fit does not
reach Rmax2 (Fig 5.20) and is over predicted by less, being 107% of the highest
corrected Req value of 479. The corrected data KD1 value is similar to the uncorrected
KD1 value (15 ± 9nM cf 20 ± 7nM) and is lower due to a closer match and lower value
of Rmax2 rather than an improved Rmax1 value, as there are few data points below Rmax1
to improve the fit and strengthen the KD1 value. Examining the fit of the Neomycin
data, it was possible to divide the data based on the prediction of Rmax1 and
individually examine the hypothetical values of KD1 and KD2 – this was not possible
for the peptide 2 data because too few data points are within the region predicted to
be Rmax1, and therefore the high affinity KD values for peptide 2 are questionable in an
individual site analysis. In this respect the value of KD1 is weighted by assay points
above Req values greater than Rmax1 as there are few data points that are below Rmax1.
157
Conc (µM) Req Conc(µM)- Req
0.102 35.1 10 4300.402 82.6 12 4842.01 110 16.1 5652.01 193 20.1 6444.02 251 30.1 7585.02 309 40.2 8818.03 372
Linear Region 0 1.0×10-5 2.0×10-5 3.0×10-5 4.0×10-5 5.0×10-5
0
50
100
150
200
250
300
350
400
450
500
550
600
650
700
750
800
850
900
950
Conc
Req
0 5.0×10-6 1.0×10-5 1.5×10-5 2.0×10-5 2.5×10-5 3.0×10-5 3.5×10-5 4.0×10-5 4.5×10-50
100
200
300
400
500
Conc
Rco
rrec
ted
Figure 5.20: Non-linear Regression Analysis of Req vs Concentration for Peptide 2:RBE3 Binding (two site model). The upper graph is the uncorrected data and includes the embedded table of assay points (Req) and the concentration assayed for peptide 2. The right graph is the corrected non-linear regression analysis of Req vs concentration for Peptide 2:RBE3 Binding (two site model, all data).
Parameter
Uncorrected Single site
Uncorrected Two site
Corrected Single site
Corrected Two site
Rmax1 115 (9) 100 (15) KD1 20nM (7nM) 15nM (9nM) Rmax2 1194 (111) 1408 (58) 541 (42) 505 (23) KD2 17µM (3µM) 34µM (3µM) 5.6µM (1.4µM) 12µM (2µM)
Goodness R2 0.9703 0.9990 0.9334 0.9938 Abs SS 25129 852 17844 1661 Sy.x 47.80 9.727 40.28 13.58 AIC Probability <0.01% >99.99% <0.01% >99.99% Ratio N/A N/A AICc 34.10 20.96
Table 5.4: Nonlinear Regression Analysis of Req vs Concentration for the Native Peptide
2:RBE3 Binding. AIC table parameters here are the same as Table 5.3 and for subsequent tables. A
99% confidence interval of the Rmax value for the uncorrected data yields an Rmax1 of 114 ± 29, Rmax2
of 1408 ± 188; KD1 ≤42nMand KD2 of 34 ± 10µM.
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
250
500
750
1000
Conc
Req
Parameter
Uncorrected Single site
Rmax 1238 (60) KD 18µM (2µM) Goodness r2 0.9889 Abs SS 3915 Sy.x 23.65 AIC Probability 91.23% Ratio 10.40 AICc -4.68
Figure 5.21: Examination of the Low Affinity Site for Peptide 2 (Uncorrected data). The figure
comprises uncorrected data above Req = 251. The X-axis is a log10 display of the concentration to more
readily highlight the fit. Embedded is the table of the non-linear regression analysis for the single site
fit. A 99% confidence interval of the Rmax value for the uncorrected data yields an Rmax of 1238 ± 211
and KD of 18 ± 6µM.
The value of KD2 was examined by sequential removal of lower points until a
single site converged (Fig 5.21). For the uncorrected data this was four points with
the fit starting with Req = 251, and the results were similar for the single site fit (all
uncorrected data). The Rmax was not reached (Rmax of 1238, over predicted by 140%
of highest Req value) and the predicted KD was 18 ± 2µM. A 99% confidence interval
of the Rmax value (1238 ± 211) agrees with the Rmax2 value of the uncorrected fit of all
158
159
the data (1408) and so the single site analysis of the Rmax2 value suggests that though
the prediction of Rmax2 is high (1408, 160% above the highest Req value) it is not an
unreasonable prediction with the removal of data below Rmax1 that may influence the
value of Rmax2, but it should be noted that there are few points below Rmax1 anyway.
By way of comparison, the value of KD2 was also examined for the corrected data by
sequential removal of lower points until a single site converged (graph not shown).
The parameters were found to be almost identical to the single site analysis (all data,
Table 5.6) of the corrected model, but the Rmax (549 ± 19) was not reached and over
predicted by 115% of the highest corrected Req value (479).
In conclusion, both uncorrected and corrected data fit to a two site model. The
values of KD1 for the uncorrected and corrected data are similar (20 ± 7nM cf 15 ±
9nM) as there is little difference in using either uncorrected or corrected data to report
KD1 because correcting using the slope of the linear region has less effect on lower
Req values. In contrast there is a large difference in the KD2 values for the uncorrected
and corrected data (34 ± 3µM cf 12 ± 2µM) since the correction factor has a large
influence on lowering Rmax2.
The question of which fit to use, either uncorrected two site or corrected two
site, is a question of the level of interpretation that can be made based on the raw
data. On the one hand the Rmax2 value is not met in the uncorrected model but the
examination of the 99% confidence interval of the low affinity site suitably agrees
with the predicted Rmax2 value of 1408 ± 58. On the other hand, the corrected data
comes closer with the difference between Req value and the Rmax2 value and the value
itself is less at 12 ± 2µM than for the uncorrected data, however there are several
problems with accepting the corrected two site fit as appropriate in this case. In the
first instance there is no reason to assume the Rmax2 value for the uncorrected data is
not an unreasonable estimate without assaying peptide 2 at higher concentrations to
reach Rmax2, and it is noteworthy that even the corrected data does not reach Rmax2,
and so the corrected data cannot be stated as being any more or less accurate.
Secondly, the linear region is more subjective than for the Neomycin analysis where
the shape of the plot is distinctively different to a binding isotherm and Rmax2 was
over predicted by a much larger margin (77490% over predicted than the largest Req
value); for peptide 2 the shape is similar to a typical binding isotherm but merely has
not reached Rmax2, and though the r2 value of the subjective linear region suggests it is
real (0.98, n = 6), the Rmax2 value is not over predicted by such a large value as to be
unreasonable. It is possible at higher concentrations than assayed here that a linear
region is more apparent (as found for peptide 3 below), but it cannot be accurately
applied in the case of peptide 2. In this respect the values of the uncorrected two site
fit is used to report the affinity of peptide 2 for the RBE3 aptamer within the
concentration range investigated was KD1 = 20 ± 7nM and KD2 = 34 ± 3µM. The
largest error margin that can be reported is a 99% confidence interval of the Rmax
values, meaning only an upper limit for KD1 that can be ascribed due to few points
below Rmax1 with KD1 ≤42nM and KD2 = 34 ± 10µM.
Succinylated Peptide (3).
The succinylated peptide (3) data was especially difficult to fit due to a
pronounced linear region (Fig. 5.22) at higher concentrations, similar to the linear
region seen with Neomycin. This linear region was likewise attributed to non-specific
associations and a correction factor was applied based on the slope of the linear
region, and both one site and two site models are presented as uncorrected and
corrected non-linear regression analyses as for peptide 2 (Fig. 5.22 and Table 5.7).
Conc (µM) Req Conc (µM) Req
5.85x10-3 87.9 2.92 423 29.4 x10-3 119 5.85 471 58.5 x10-3 133 11.7 500
0.175 158 29.3 537 0.351 169 35.1 561 0.468 172 46.8 593 0.585 187 58.5 639 0.762 200 70.2 785 0.932 235 93.5 836 1.17 4104410 10
160
Linear Region
0 2.5×10-5 5.0×10-5 7.5×10-5 1.0×10-40
100
200
300
400
500
600
700
800
900
Conc
Req
0 2.5×10-5 5.0×10-5 7.5×10-50
100
200
300
400
500
600
Conc
Rco
rrec
ted
1.0×10-4
Figure 5.22: Non-linear Regression Analysis of Req vs Concentration for Peptide 3:RBE3 Binding (two site model, All data). The upper graph is the uncorrected data and includes the embedded table of assay points (Req) and the concentration assayed for peptide 3. The left graph is the corrected non-linear regression analysis of Req vs concentration for Peptide 3:RBE3 Binding (two site model, all data).
Table 5.5: Nonlinear Regression Analysis of Req vs Concentration for the Succinylated Peptide
3:RBE3 Binding (all data). The table parameters are as for table 5.3. A 90% confidence interval of
the corrected two site model yields Rmax1 = 91 ± 73, Rmax2 = 367 ± 74; KD1 ≤ 8nM, KD2 = 1 ± 0.7µM.
Parameter
Uncorrected Single site
Uncorrected Two site
Corrected Single site
Corrected Two site
Rmax1 453 (66) 91 (42) KD1 558nM (224nM) 0.5nM (4.5nM) Rmax2 658 (37) 6.4e+7 (2e+12) 452 (22) 367 (42) KD2 1.4µM (0.4µM) 16M (5.2e+5M) 554nM (132nM) 1µM (0.4µM)
Goodness r2 0.8670 0.9431 0.8476 0.9072 Abs SS 134263 57451 57357 34935 Sy.x 88.87 61.89 58.09 48.26 AIC Probability 1.04% 98.96% 23.10% 76.90% Ratio 95.26 3.33 AICc 9.11 2.41
It can be seen from Table 5.7 that the fit (Fig. 5.22) is imperfect and the r2
values are not ideal (0.85 – 0.94). The best r2 value is from the uncorrected analysis
of the two site model but the Rmax2 value (64x106 ± 2x1012) and KD2 value (16M ±
5.2x105M) are not reasonable with Rmax2 over estimated by 7,655,502% of the largest
Req value. From a log10 display of the X-axis (Conc) (Fig. 5.23) it can be seen the fit
is skewed by the middle (1 in Fig 5.23) and upper concentrations, with a sharper
increments than expected, as well as a small plateau followed by a linear increase
around Rmax2 (2 in Fig. 5.23), and these problems may be biasing the fit away from
the lower concentrations.
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
100200300400500600700800900
Conc
Req
2
1
Figure 5.23: Nonlinear Regression Analysis of Req vs Concentration (log10) for the Succinylated
Peptide 3:RBE3 Binding (all data, uncorrected). Shown in the fit are the points that skew the data
towards higher concentrations and away from the model. At 1 the typical rise for a two site model is
larger than expected and the mid point of the fit is linear. At 2 there is a second rise that does not
reach Rmax. 161
In terms of the shape of the fit, the data points that cause changes are: 410,
1.17µM; 639, 58.5µΜ; 785, 70.2µM; 836, 93.5µM (Fig. 5.23). The succinylated
peptide 3 is the only peptide assayed above 40µM and removal of these three end-
most points (making the analysis range to 47µM), and the further removal of the
1.17µM point significantly improves the fit as shown in figure 5.24 for the
uncorrected and corrected data.
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
50
100
150
200
250
300
350
400
450
500
550
600
650
Conc
Req
Parameter
Uncorrected Single site
Uncorrected Two site
Rmax1 107 (16) KD1 1.0nM (2nM) Rmax2 568 (30) 490 (18) KD2 1µM
(0.2µM) 2.4µM (0.4µM)
Goodness r2 0.9165 0.9886 Abs SS 40160 5467 Sy.x 55.58 22.29 AIC Probability <0.01% >99.99% Ratio N/A AICc 21.43
Parameter
Corrected Single site
Corrected Two site
Rmax1 96 (31) KD1 0.3nM (3.4nM) Rmax2 437 (24) 352 (32) KD2 0.6µM
(0.2µM) 1.2µM (0.4µM)
Goodness R2 0.8550 0.9442 Abs SS 41838 16106 Sy.x 54.67 36.64 AIC Probability 2.57% >97.43% Ratio 37.97 AICc 7.27
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
100
200
300
400
500
Conc
Rco
rrec
ted
Figure 5.24: Nonlinear Regression Analysis of Req vs Concentration for the Succinylated
Peptide 3:RBE3 Binding (selected data). The upper curve shows the uncorrected fit and analysis
and the lower fit is for the corrected data. The omitted points were Req = 410, 639, 785, 836.
162
163
Despite these improvements, the errors are still high and the removal of data is
especially considering incorrect in this instance as the assays are a first investigation
and so all data points must be taken as real and contribute to the fit. In this respect the
removal of the data points is useful to show where the fit may be wrong and help
identify possible problems to improve the assay, but is not used here as a practice. In
the case of the last three data points they are important as they more clearly defining
the linear portion of the fit.
Examining the uncorrected high affinity site below Req = 400 found a single
site with KD = 14 ± 7nM and Rmax of 191 ± 12 but the r2 value is unrealistic at 0.71
(data not shown). Similarly, examining the uncorrected low affinity data above 420
found a single site with KD = 1.2 ± 0.3µM and Rmax of 576 ± 15 with a poor r2 value
of 0.90 (data not shown), and because of the poor fit from independently examining
the individual KD regions, they cannot contribute to the analysis in a meaningful way.
It is concluded that the assay of the succinylated peptide was problematic
including an outlier and linear region, impeding the fit of the data. The data for the
uncorrected and corrected two site model was significantly favoured above a one site
model, however the uncorrected Rmax2, KD2 values and associated errors are
unrealistically large and the uncorrected fit cannot be taken as accurate. Correcting
for the linear region by transforming the data according to the slope of the linear
region allowed Rmax2 to be reached (367 ± 42) but the fit is non-ideal with the error
for KD1 being 9-fold greater than the reported value and the r2 value is 0.91 (KD1 = 0.5
± 4.5nM and KD2 = 1 ± 0.4µM). In lieu of these difficulties and poor r2 value, it is
more appropriate to report Rmax values and errors based on a 90% confidence interval
and KD values based on this Rmax range. Expressed in terms of a 90% confidence
interval, only an upper limit for the value of KD1 can be reported, from table 5.7 these
values are KD1 ≤ 8nM and KD2 1 ± 0.7µM.
Capped Peptide (4).
The fit to the data of the capped peptide 4 was found to follow a two site
model (Table 5.8) but was imperfect below Req values of 400 in both the uncorrected
and corrected fits (Fig. 5.25), and though the shape of the curve for the corrected data
is improved at higher concentrations, the fit to lower concentrations suffers.
Parameter
Uncorrected Single site
Uncorrected Two site
Corrected Single site
Corrected Two site
Rmax1 333 (23) 289 (26) KD1 41nM (10nM) 30nM (8nM) Rmax2 835 (41) 816 (38) 633 (20) 435 (25) KD2 625nM (153nM) 8.8µM (1.6µM) 257nM (41nM) 2.5µM (0.6µM)
Goodness R2 0.8828 0.9916 0.9301 0.9868 Abs SS 313812 22602 103169 19476 Sy.x 107.8 30.07 61.81 27.91 AIC Probability <0.01% >99.99% <0.01% >99.99% Ratio N/A N/A AICc 70.64 42.70
Table 5.6: Nonlinear Regression Analysis of Req vs Concentration for the Capped Peptide
4:RBE3 Binding (all data). The table parameters are the same as for table 5.3. A 99% confidence
interval of the Rmax values yields for the uncorrected data: Rmax1 = 333 ± 63, Rmax2 816 ± 105, KD1 41 ±
27nM, KD2 9 ± 5µM and for the corrected data: Rmax1 289 ± 73, Rmax2 436 ± 71, KD1 30 ± 22nM, KD2
2.5 ± 1.6µM.
0 5.0×10-6 1.0×10-5 1.5×10-5 2.0×10-5 2.5×10-5 3.0×10-5 3.5×10-5 4.0×10-50
100
200
300
400
500
600
700
800
900
1000
1100
Conc
Req
0 5.0×10-6 1.0×10-5 1.5×10-5 2.0×10-5 2.5×10-5 3.0×10-5 3.5×10-5 4.0×10-50
100
200
300
400
500
600
700
800
Conc
Rco
rrec
ted
Conc (µM) Req Conc (µM) Req2.09x10-3 22.3 0.523 401 4.18 x10-3 42.5 1.57 434 8.36 x10-3 63.9 2.09 458 10.5 x10-3 86.3 2.62 527 16.7 x10-3 111 3.35 586 21.0 x10-3 135 4.18 608 31.4 x10-3 157 6.28 648 52.3 x10-3 185 12.6 794 83.7 x10-3 208 16.7 843
0.126 232 20.9 818 0.167 257 24.1 960 0.209 283 30.3 982 0.251 302 33.5 1020 0.293 326 8.00 764 0.335 348
Linear Region
Figure 5.25: Non-linear Regression Analysis of Req vs Concentration for Peptide 4:RBE3 Binding (two site model). The upper figure is the uncorrected data and included is the embedded table of assay points (Req) and the concentration assayed for peptide 4, the Linear Region used for correcting the data is shown in the upper figure. The lower figure is the non-linear regression analysis of Req vs concentration for the corrected data.
164
Because of the poor fit at lower concentrations, a third site at the uppermost
region of the curve was suspected to cause a poorer fit to lower concentrations, and
so the data was separated into upper and lower regions based on the predicted Rmax1
value of 333 ± 23 for the uncorrected data and 289 ± 26 for the corrected data with a
one and two site analysis conducted on each (Fig 5.26). Parameter
Uncorrected Single site
Rmax 284 (7) KD 26nM (2nM) Goodness r2 0.9942 Abs SS 361.0 Sy.x 6.333 AIC Probability 95.84% Ratio 23.04 AICc -6.27 1.0×10-9 1.0×10-8 1.0×10-7 1.0×10-6
0
100
200
300
Conc
Req
Parameter
Corrected Single site
Rmax 281 (7) KD 25nM (2nM) Goodness R2 0.9942 Abs SS 252.8 Sy.x 6.261 AIC Probability 96.54% Ratio 27.86 AICc -6.65
1.0×10-9 1.0×10-8 1.0×10-7 1.0×10-60
100
200
300
Conc
Rco
rrec
ted
Figure 5.26: Comparison of High Affinity Sites for Peptide 4. The upper figure comprises data
below Req = 257 for the uncorrected data and the lower figure comprises data below Req = 255 for the
corrected data. Embedded are tables of the non-linear regression analysis for the single site fits.
It was found that when the data was below Req = 257 for the uncorrected data,
the fit converged with the single site prediction as shown in the upper graph of figure
5.26. Similarly for the corrected data, the fit converged below Req = 255 with a single
site prediction as shown in the lower graph of figure 5.26. It can be seen from the
tables embedded into figure 5.26 that both fits predicted the same KD value for a
single site model of the high affinity binding site with a KD of 26 ± 2nM for the
uncorrected data and 25 ± 2nM for the corrected model. The inclusion of another data
165
point for either fit led to a two site prediction and so this is the limit of the high
affinity binding site. The two values of the high affinity site KD (Fig. 5.26) are likely
so close because the correction factor does not have a large effect on lower
concentration data.
The Rmax2 and KD2 upper region of the two fits (uncorrected and corrected)
was examined by removing points sequentially from the lower concentrations until a
single site was apparent (Fig. 5.27). For the uncorrected data this encompassed the
highest 12 data points from Req = 458 with a KD of 2.8 ± 0.4µM. For the corrected
data, the single site was from the uppermost 13 points beginning with Req = 418 and a
KD range of 1.1 ± 0.2µM.
This compares favourably with the two site binding model for the corrected
data (all data) with a range of 2.5 ± 0.6µM and the 99% confidence interval of the
uncorrected data has a range of 9 ± 5µM and so the lower limit of this range is also
comparable to the upper limits of the single site analysis.
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
100200300400500600700800900
10001100
Conc
Req
Parameter
Uncorrected Single site
Rmax 1027 (34) KD 2.8µM
(0.4µM) Goodness R2 0.9323 Abs SS 25628 Sy.x 50.62 AIC Probability 52.28% Ratio 1.10 AICc -0.18
1.0×10-7 1.0×10-6 1.0×10-5 1.0×10-40
100
200
300
400
500
600
700
800
Conc
Rco
rrec
ted
Parameter
Corrected Single site
Rmax 715 (16) KD 1.1µM
(0.2µM) Goodness R2 0.9047 Abs SS 11261 Sy.x 32.00 AIC Probability 99.30% Ratio 141.65 AICc -0.91
Figure 5.27: Comparison of Low Affinity Sites for Peptide 4. The upper figure comprises data
above Req = 458 for the uncorrected data and the lower figure comprises data above Req = 418 for the
corrected data. Embedded are tables of the non-linear regression analysis for the single site fits. 166
167
Examining the uncorrected data, a third binding site is not found or
can be inferred from the data between the points used in the high and low affinity
binding sites as inclusion of these values with the high affinity site analysis did not
predict a significantly different Rmax2 value than seen with the entire data set and
likewise, inclusion of these values with the low affinity site analysis did not predict a
significantly different Rmax1 value than seen with the entire data set (Table 5.8).
Furthermore, examining a single site in the high and low affinity binding regions
found KD values that did not significantly differ from the strongly AIC predicted two
site analysis (Table 5.8). Therefore the data between the high and low affinity sites is
merely the point of inflection and there is only two binding sites on the RBE3 for the
peptide 4.
In summary, the binding of peptide 4 to RBE3 RNA followed a two site
model and from the fit using all the data, the predicted KD1 values are close to the KD
high affinity site values using the selected data (uncorrected 41 ± 10nM; corrected 30
± 8, compared to the high affinity analysis of 25/26 ± 2nM) and therefore the two site
fit is not significantly skewed by higher concentration data. The KD2 values are also a
reasonable range for the uncorrected and corrected data being 8.8 ± 1.6µM and 2.5 ±
0.6µM respectively. Though a little higher for the uncorrected data set compared to
the corrected, when a 99% confidence interval of Rmax2 is used to examine the
maximum possible range for KD2 being 9 ± 5µM and 2.5 ± 1.6µM for the uncorrected
and corrected data respectively, these differences are not unacceptable. Because the
fit using the uncorrected and corrected data is not greatly different, there is little
justification in applying the correction factor and the uncorrected data is taken as
accurate, being KD1 = 41 ± 10nM and KD2 = 8.8 ± 1.6µM and using a 99% confidence
interval of Rmax values the range is 41 ± 27nM and KD2 = 9 ± 5µM.
Rev Protein
The Rev protein was assayed at lower concentrations than the peptides 2 – 4
to avoid the problems seen with peptide 3 at elevated concentrations/responses. Fewer
assay points were taken of the Rev protein as each assay point was replicated with a
control blank injection (9 assay points and 9 blank injections) containing the same
quantities of the stock buffer conditions from which the protein was diluted in order
to avoid refractive index changes caused by a difference in buffer contents and
concentrations. The protein data was fit by non-linear regression and only converged
to the single site model, with a two site model yielding the same value as the single
site model (not shown). The correction factor for non-specific binding was applied to
the Rev protein data, and the uncorrected and corrected data are presented below (Fig
5.28). The uncorrected data did not reach Rmax and the value was over predicted by
153% of the highest Req value but there is no clearly defined linear region to justify
correcting the data and a linear fit of the five uppermost data points is poor (r2 0.87)
and to improve the assay, collection of higher concentration data points would be
necessary, however there is little reason to assume the predicted Rmax is not an
unreasonable value.
Conc (nM) Req Conc (nM) Req0.787 5 .46 78.7 325 7.87 7 8 78.7 312 39.4 206 118 4 19 39.4 201 197 5 01 78.7 378
168
0 5.0×10-8 1.0×10-7 1.5×10-7 2.0×10-70
100
200
300
400
500
600
Conc
Req
Parameter
Uncorrected Single site
Rmax 766 (74) KD 101nM (19nM) Goodness R2 0.9840 Abs SS 3324 Sy.x 21.79 AIC Probability >99.99% Ratio N/A AICc -19.20
0 5.0×10-8 1.0×10-7 1.5×10-7 2.0×10-70
50
100
150
200
250
300
350
Conc
Rco
rrec
ted
Parameter
Corrected Single site
Rmax 401 (44) KD 48nM (14nM) Goodness R2 0.9554 Abs SS 4081 Sy.x 24.15 AIC Probability >99.99% Ratio N/A AICc -19.20
Figure 5.28: Uncorrected and Corrected Non-linear Regression Analysis of Rev:RBE3 interaction. The
upper figure comprises the uncorrected data and embedded data points with regression analysis and the lower
figure is the corrected data and regression analysis. A 98% confidence interval of the uncorrected data is Rmax 766
± 222, KD 101 ± 58nM and a 95% confidence interval of the corrected data is Rmax 401 ± 105, KD 48 ± 34nM.
The fit of the corrected data was noted to improve with an outlying point
removed (78.7nM, 299) but removal of the same point in the uncorrected data does
not significantly change the fit, which along with the r2 value of the linear fit,
suggests that the correction factor is not representative. In this respect the uncorrected
data is taken as the best representation for the concentration range assayed with a KD
of 101 ± 19nM for the RBE3 aptamer and a maximum range based on the 98%
confidence interval of Rmax (544 – 989) is 44 – 159nM.
Since the controls used in this study cannot correct non-specific binding to
RNA, and since other SPR-based studies (Hendrix et al., 1997; Van Ryk and
Venkatesan, 1999) use non-specific RNA of similar size or mutated forms of the RRE
that are inactive as non-specific control channels, the effects of non-specific RNA
was investigated with the Rev protein in the conditions under which the assays were
conducted. The non-specific RNA (RNAneg) was commercially synthesised
biotinylated-RNA of 34-nucleotide length purchased from Prof Ross Smith (UQ).
RNAneg was captured parallel to RBE3 RNA on a custom B1-Streptavidin chip
control channel using identical methods for RNA capture as per the assays described
in this chapter for Rev and the peptides 2 – 4 and approximately equal concentration
of RNA were used (26µg).
The sensorgrams below show typical data for Rev (150nM) and the two RNA
species (Fig. 5.29). It can be seen that Rev has minimal interaction with the RNAneg
sequence (~25RU cf ~425RU for RBE3 RNA), and since the interaction with non-
specific RNA is so small (~6%), the overall effect on the KD values reported here is
likely to be minimal.
-50
0
50
100
150
200
250
300
350
400
450
-200 0 200 400 600 800 1000 1200
RU
Res
pons
e
sTime Figure 5.29: Rev interacting with RBE3 RNA [green] and RNAneg RNA [pink]. Clearly it can be
seen that Rev shows appreciable association and dissociation, while RNAneg RNA responds very
weakly to Rev.
169
170
The conclusion to be drawn from this experiment is though non-specific RNA
would be more suitable to subtract non-specific binding and is likely the best control
for this system, non-specific RNA interaction alone are unlikely to account for the
large and varied low affinity site observed here and further investigations would be
warranted to more clearly elucidate the second binding site to establish if it is a real
second site for peptides on the RBE3 aptamer or a non-specific association in the
system. It is also important for future assays to establish the concentration at which
the second site association is important and the exact nature of non-specific
associations in the assay system.
Summary
For all the assays conducted, the final values would have been improved if
more data points were taken using a refined concentration range based on the
difficulties encountered, however almost all peptides were completely depleted at the
end of the investigation. The Rev protein produced largest responses in the BIAcore
2000 machine, likely because of its greater size, allowing the Rev protein to be
assayed below concentrations where the low affinity state was significant for the
peptides and fit to a one site model. The most difficult peptide to assay was the
succinylated peptide 3, which had a broader concentration range and lower KD values,
however the fit of this data is also the most error-prone and the final results reported
here may contain a larger error than described.
The problems with the assay of 3 suggest there is something fundamentally
different with either the peptide as provided or the manner in which the peptide
behaves in the assay system. The difference to the peptides 2, 4 and Rev protein could
be related to the succinylated peptide itself, which may have greater non-specific
associations to the RNA, Streptavidin or surface carboxymethyldextran, but
structurally this is difficult to rationalise as the peptide sequences are the same for 2,
3 and 4, though the succinylated peptide 3 will have a different direction of its dipole,
and this may cause a difference in the way the peptide interacts within this assay
system. Future assays should be conducted uniformly below a 40µM threshold where
possible since the problems with 3 were exacerbated by assaying the peptide at
elevated concentrations compared to 2 or 4 and greater non-specific responses are
apparent at elevated concentrations.
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Comparing Neomycin, the three peptides and Rev protein affinity for the
RBE3 aptamer, the order of potency was found to be 3 > 2 > 4 > Rev > Neomycin
using the standard error values and using the maximum error ranges from the
confidence intervals of the fit the order of potency is 3 > 2 ≈ 4 ≥ Rev > Neomycin
(Table 5.9), and the maximum error range is likely a more accurate description of the
results as it encompasses a broader range of Rmax and Rmax2 which was problematic to
accurately ascribe.
Parameter Neomycin 2 Native
Peptide 3 Succinyl
Peptide 4 Capped Peptide
Rev Protein
KD or KD1 13 ± 3µM 20 ± 7nM ≤5nM10 ± 41 ٭nM 101 ± 19nM
KD2 N/A 34 ± 3µM 1 ± 0.4µM 8.8 ± 1.6µM N/A
Range KD/KD1
KD2(µM)/CI
7-19µM
-- (92%)
≤42nM
24-44 (99%)
≤8nM٭
0.3-1.7 (90%)
14-68nM
4-14 (99%)
43-159nM
-- (98%)
Literature KD 1-6µM; 0.2-
25µM
Peptides reported range varies from 2.3 – 40nM,
but reported as high as 121 – 490nM using SPR 0.4 – 10nM
Table 5.7: Non-linear Regression Analysis of SPR Data. Assays were conducted in 50mM Hepes
buffer (pH 7.4), 300 mM NaCl, 10µg/mL calf thymus DNA for all samples except Neomycin (25mM
Hepes (pH 7.4), 150mM NaCl and 10µg/mL calf thymus) at 25°C. A two site binding model was used
for all assays with all peptides fitting to a two site model, and Neomycin and Rev fitting to a single site
model. The Range is based on the confidence interval range from the individual analysis (indicated as
CI in the table). The succinylated peptide٭ is more accurately ascribed as the maximum range (≤8nM)
as the confidence interval was greater than 10%. From the literature cited earlier in this chapter, the
standard range is 1-6µM for Neomycin, but reported as broad as 10 – 25µM and as low as 0.2µM. Also
reported is the range for peptides corresponding to those used here and Rev, this literature is cited
below.
Other groups reporting kinetic data for either Rev or the peptide(s) used here
show no definitive value or general agreement for KD values. From gel mobility shift
assays, Harada et al., (1997) report KD values of 6 – 40nM for succinylated peptides
equivalent to 3, Kjems et al., (1992) (who provided the constructs used in this study)
report 10 – 20nM for both peptides and Rev protein (similar affinity). Kjems et al.,
(1992) do recognise that most gel shift assays report between 1 – 3nM for the Rev
protein. For the succinylated peptide 3, fluorescence anisotropy experiments (Kirk et
al., 2000) yielded an IC50 of 12.5nM and KD of 2.3nM, whereas Kumangai et al.,
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(2001) reported 44 – 86nM for this peptide. For SPR assays, West and Ramsdale
(1997) report 2.5 – 121nM values for the succinylated peptide 3, while Hendrix et al.,
(1997) report values of 215 – 490nM and Van Ryk and Venkatesan, (1999) claim
0.04nM for Rev, and both Hendrix et al., (1997) and Van Ryk and Venkatesan,
(1999) found an upper limit to the concentration they could assay of both Rev34-50
peptides and Rev respectively, with Hendrix et al., (1997) suggesting the problem is a
surface transport limitation under assay conditions similar to those used here. The
paper of Chapman et al., (2002) is perhaps the most revealing as they report a KD of
5nM for their modifications of a filter binding assay but could not assign a KD when
trying to validate the assay using SPR, because multiple binding was found above a
certain threshold - similar to what was found in this study for which the correction
factor was applied. In this respect the data reported here has encountered similar
problems that is found in the literature and though the KD values are higher than the
generally accepted range, they suitably agree with the literature based on the broad
ranges reported. It is worth noting that if a single site analysis only is conducted for
the uncorrected Neomycin and peptide 4 data below Rmax1 values, the KD values are
lower and closer to literature values. For other peptide analytes there were few data
points below Rmax1 to examine, but this trend could be the same. In this regard, the
differences present in the literature may be complicated by a two-state binding mode
that alters the apparent dissociation constant if the data is fitted to a single state, or if
data beyond the high-affinity state are included (Chapman et al., 2002). Therefore the
assessment of which binding model (single or two-state) and the concentration of
Rev34-50 peptides or the Rev protein assayed should be carefully considered when
studying the Rev:RBE3 interaction.
It is possible that the RRE recognition sequence (Rev34-50) that constitutes the
peptides may bind non-specifically to the RBE3 aptamer, as the literature reports that
as many as three aggregates of Rev multimerise on stem loop II of the RRE, and eight
over the entire RRE structure (Cook et al., 1991; Daly et al., 1989). It is also possible
that the three observed in gel mobility shift assays and two-state binding observed
here may be attributed to multiple binding at higher analyte concentrations as found
by Van Ryk and Venkatesan, (1999) and Chapman et al., (2002). For Neomycin
multiple binding may not be seen because of the inherent weak association the
aminoglycoside makes with the RBE3 aptamer, and so all other binding may be
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removed with correction of the data. Alternately the single site observed for
Neomycin may be a reflection of the difficulty assaying a small analyte and lower
affinity sites may not be detectable in this system.
The low affinity state (KD2) for the peptides exhibited a range of values from 1
– 34µM (0.3 – 44µM based on the total range of the confidence intervals). For the
peptides 2 and 4 the values were similar to Neomycin interactions for the RBE3
aptamer, whereas 3 exhibited the greatest affinity (~1µM). It is difficult to ascribe this
variation solely to a preference for a second site or non-specific binding of one
analyte over another to the RBE3 and this two site binding possibly includes non-
specific multiple binding to RBE3 RNA and residual non-specific binding to the
carboxymethyldextran-Streptavidin surface at high concentrations of analyte. A third
possibility included in the low affinity state may be a heterogeneous ligand surface
due to the random labelling technique, where one or more biotinylated Us are
incorporated towards the middle of the RBE3 sequence rather than the 3’ ends and
hence have lower affinity for analytes. It should also be stated that the KD2 values
could not be ascribed with confidence since a range of analyte concentrations was
made, but not all the analytes could be assayed at an elevated concentration, as the
peptides were a limiting resource. In this respect none of the analytes were assayed at
concentrations that would saturate binding and reach their individual Rmax2 or KD2
values and therefore the values of Rmax2 and KD2 may also contain an error, and single
site non-linear regression analysis of the peptides 2 and 4 suggests the range may
actually be narrow (0.9 – 6.7µM).
Within the BIAcore machine the nature of the low affinity site (whether it is a
problem attributable to the RNA ligand or the surface) is difficult to elucidate. Further
experiments that might prove useful would include an RNAneg control and achieving
Rmax2 to properly ascribe this value. However, the RNAneg control experiment
suggested the non-specific contribution of RNA is minor to Rev and this could also
be the case with the peptides, and there appears to be cases of continued non-specific
interactions in the literature despite using an RNA control (Hendrix et al., 1997; Van
Ryk and Venkatesan, 1999). It would also be useful to include unlabelled RBE3 RNA
at various concentrations with analyte injections in order to compete with low
affinity/non-specific binding, or to inject different concentrations of unlabelled RBE3
RNA over the surface during ligand dissociation in order to out compete the non-
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specific interactions or titrate them away from the surface and gain a greater
understanding of what is occurring in the assay system and this might prove a useful
investigation in the future. If the problems of the low affinity state can be attributed to
the random labelling of the RNA creating a heterogeneous ligand surface, then ligand
surface generation using the methods of Van Ryk and Venkatesan, (1999) would be
worth investigating; where a synthetic biotinylated poly A stretch is captured onto the
surface and in vitro transcribed RNA is base-paired to the poly A stretch (with a
complimentary poly T/U stretch) to generate a controlled, homogeneous surface.
5.2.3 Does Helix Induction Increase Affinity for the RBE3 RNA
Aptamer?
Because the RRE-binding domain of HIV-1 Rev interacts in a highly specific
manner with the RRE and adopts a fully helical conformation in the complex, it was
used to test in a significant biological context the proposition that helix induction can
increase ligand affinity. Compound 1 (Fig 5.6) is believed to nucleate helicity by
presenting its carbonyl oxygen atoms in the correct orientation to hydrogen bond to
three amide hydrogen atoms in a peptide sequence attached to its N-terminus.
Circular dichroism (CD) spectra (kindly supplied by Dr Michael Kelso, IMB)
for compounds 1 - 4 were collected under low salt conditions. The cyclic template 1
did not show any significant absorption alone between 200-250nm, but does exhibit a
characteristic CD maximum at 270 nm as reported (Austin et al., 1997). It was found
that the native peptide 2 has a dominant minimum at 208nm, indicative of random
coil structure. The N-terminal succinylated analogue 3 was observed to have a
prominent second minimum at 222nm, which was even more prominent for 4. Using
the mean residue elipticity at 222nm, the percentage helicity was calculated as 54%
(4), 27% (3) and 15% (2) respectively, indicating that the N-terminal template 1 is an
effective N-terminal nucleator that enhances helicity of the Rev34-50 peptide sequence,
overcoming the problem of loss of helical conformation when shortening a minimal
polypeptide-binding unit away from the helix-stabilised environment provided by the
protein.
Though 3 is an order of magnitude lower, the assay of the compounds 2, 4 and
the full-length Rev protein show the order of affinity for bound RBE3 RNA to be 2 ≈
4 ≥ Rev when considering the absolute range using the confidence interval of the
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analysis, as assayed in 50mM Hepes buffer (pH 7.4), 300 mM NaCl, 10µg/mL calf
thymus DNA at 25°C. The BIAcore assays conducted here cannot confirm that fixing
of helicity in a shorter peptide based on Rev34-50 increases affinity for the RBE3 RNA
target. However, the structure (including helicity) of peptides and proteins is affected
by the environment that surrounds it, including the buffer additives, pH, temperature,
salt ions and concentration of these ions as well as the concentration of the
peptide/protein itself (Ru et al., 2000; Cameron et al., 1997; Collins 1995; Wiggins
1995; Goto and Aimoto 1991; Washabaugh and Collins 1986; Tiffany 1975). In the
case of the buffer conditions used in this study, the increased temperature (25ºC)
generally favours a less helical content by lowering the energy for motion, and
Hendrix et al., (1997) report a helical content reduced from 64% (5ºC) to 29% (30ºC)
for a peptide equivalent to 3, and their assays were conducted at lower temperatures
to avoid this loss of helicity. The buffers used here also had a higher salt
concentration of chaotropic halide ions (such as sodium), which typically disfavour
helicity, but have also been reported to favour helicity through formation of salt
bridges and desolvation of the peptide/protein (Bradley et al., 1990). It is likely the
peptides 2 and 4 had the same helical content under the assay conditions due to the
temperature the assays were ran and chaotropic nature of the buffer, and hence their
similar KD values are not surprising.
A question that needs to be addressed in future work is the relevance of
inducing helicity in an intracellular environment comprised largely of potassium and
phosphate ions (~140mM and 100mM respectively) in contrast to interstitial and
plasma ion concentrations that are largely comprised of sodium and chloride ions
(~142–147mM and 105–114mM respectively) (Marieb 1989). The nucleolus itself
appears to be isotonic to the rest of the cell (~150mM), and suffers under hypotonic
solutions (Zatsepina et al., 1997), and so a therapeutic with a target inside this
compartment would likely encounter similar conditions to the cell itself. In this
respect it would be worthwhile to assay the peptides in lower salt conditions than
used here and also examine the temperature and circular dichroic properties of the
peptides under typical intracellular ionic conditions.
The finding that helicity did not increase affinity for the target in this assay
system does not show that helix-nucleating caps cannot be used to facilitate the
design of new inhibitors of helix-mediated biological processes. In the first instance,
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the CD data convincingly demonstrate that helix nucleation has occurred, and in this
study no detrimental effects (loss of activity) of the capped peptide 4 were observed
when compared to the native peptide 2 and Rev protein. Furthermore, since the
peptides used here were of an appreciable size for native helicity to occur the
comparison itself may have been flawed and ultimately future work should also
examine reducing the size of the peptides to a more minimal binding unit that would
be more significant as a potential therapeutic.
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Chapter 6: General Discussion and Conclusions
178
This thesis comprised a path of research to overcome initial obstacles
necessary to develop suitable inhibitors against both Rev:RRE and HIV-1 PR
function with the long term goal of examining combination therapy with inhibitors to
these targets. The specific aims were therefore to facilitate future drug design and
development and comprised:
The acquisition and analysis of high-resolution crystal structures of HIV-1 PR
protein crystal complexes to a pre-organised β-strand substrate and its hydrolysed
products, determined using cryo-crystallographic techniques (Chapter 2).
To design, and synthesize novel HIV-1 PR inhibitors using a combinatorial
approach from cyclic peptide templates (Chapter 3).
To develop a reliable RNA-Rev binding assay and, to use an established
fluorometric HIV-1 PR assay to test prospective Rev and HIV-1 PR antagonists in
vitro (Chapters 3, 4 and 5).
The work in Chapter 2 addresses the first aim of the thesis: the acquisition,
analysis and comparison of high-resolution crystal structures for complexes of HIV-1
PR with substrate, inhibitor, and products bound within the active site. It was
hypothesised that structures for HIV-1 PR bound to a pre-organised β-strand substrate
and its hydrolysed products would provide new information about substrate
recognition and the mechanism of catalysis.
It was found that for the bicyclic substrate used, the N-terminal cycle half
locks the substrate into the active site and does not move throughout catalysis
allowing efficient electron transfer during acid-base catalysis and proton exchange
with the active site residues. In contrast the C-terminal cycle moves during catalysis
by a maximum of 1.0Å and is pivoted towards the active site.
The inhibitor was found to adopt a different binding mode due to the charge,
hybridisation state and lengthened isostere backbone, displacing the inhibitor more
towards S3 than either the substrate or product N-terminal cycles. The inhibitor was
found to pucker about the scissile bond compared to the substrates and products, with
the hydroxyl of the isostere 1.0Å away from the position of the substrate carbonyl and
1.6Å away from the analogous hydroxyl of the product. It was also found that when
binding to a ligand, HIV-1 PR alters interactions and structure about the flap region
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and through the conserved water to satisfy all H-bonding interactions, with a more
optimal interaction of the water molecule with the substrates compared to the
inhibitor. Despite these differences between the ligands and HIV-1 PR, the enzyme
and respective ligands maintain almost identical H-bonding to HIV-1 PR with a
maximum difference of 0.3Å seen near the flexible cyclic linker regions at the ends of
the cycles. In contrast to common beliefs, no evidence was found to support the
hypothesis that the flaps close more tightly on an inhibitor compared to a substrate
and suggesting that substrate binding is the rate-limiting step of catalysis.
A definitive mechanism of catalysis could not be unambiguously elucidated
from this structural data alone. However several important findings were made with
implications for catalysis and these were incorporated into current proposals. The
initial step in catalysis was found to occur with D125 acting as the Lewis acid and
interacting with the substrate carbonyl. The catalytic water molecule was modelled
using the product hydroxyl atom, assuming that like the substrates, this water
molecule moves little during catalysis. In this model, the catalytic water molecule is
bonded to D25 or additionally bonded to G27 but not D125 and is not activated from
between the catalytic residues. The water molecule or hydroxyl ion must attack the Re
face of the substrate at 107˚ as predicted for ideal SN2 nucleophilic attack (Burgi et
al., 1973).
Within the literature, problems understanding how a substrate is recognised
and processed has impeded inhibitor design and elucidation of the catalytic
mechanisms of proteolytic enzymes (Laco et al., 1997; Rose et al., 1996). For drug
design it is useful to understand the finer details of how the substrate and enzyme
change through the catalytic profile from the E-S to the E-P state in order to design
inhibitors that are more substrate-like or conform to the possible changes the enzyme
may undergo through catalysis that may influence inhibitor recognition. From the
literature, the changes that occur to the substrate and enzyme through the catalytic
profile are not well known or understood, with few incidents of HIV-1 PR E-S and E-
P complexes reported to shed light on these changes (Prabu-Jeyabalan et al., 2000;
Laco et al., 1997; Rose et al., 1996). This study is the first report of the complete
catalytic profile of HIV-1 PR using the same ligand sequence from E-S through the
E-I (an analogue of the transition state) to the E-P state and may assist future drug
design.
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The second aim, addressed in Chapter 3, was to design and synthesise novel
HIV-1 PR inhibitors. A significant problem in protease inhibitor design is optimising
one inhibitor region independently from another in the presence of cooperative effects
(Hofmann et al., 1988; Epps et al., 1990; Ridky et al., 1996; Majer et al., 1997).
Cooperativity occurs when designed changes to the inhibitor initiate changes in the
substrate binding cleft of the enzyme such that adjacent pockets or peripheral loops of
the enzyme change shape resulting in reduced potency (Epps, et al., 1990; Abbenante
et al., 1995; March et al., 1996; West and Fairlie, 1995; Todd and Freire, 1999).
Since the N-terminal cycle was found in Chapter 2 to be anchored through catalysis,
it was hypothesised that the binding mode of HIV-1 PR from S1 - S3 could be closely
maintained, dampening the effects of cooperativity and allowing independent probing
of the substrate binding pockets at S1’ and S2’.
The structure of a potent HIV-1 PR – N-terminal cyclic peptide inhibitor
(Tyndall et al., 2000) was superimposed onto the E-S, E-I and E-P complex revealing
the suitability of the structure to serve as design template having an N-terminal
domain from S1 – S3 occupying similar atom positions as those of the study in
Chapter 2. This template structure was used to define the limits of the possible
substitutions and a combinatorial chemical approach was taken, where the P1’
fuctionality was maintained with an isoamyl group while the S2’ pocket was probed
using a series of aromatic sulfonamides and ureas. Similarly, a benzylsulfonamide
was maintained at P2’ while a series of primary amines was used to probe the S1’
pocket in an attempt to; use bulky groups such as aromatics to fill the pocket, short
aliphatics to accommodate shrinkage of the S1’ pocket through mutations and flexible
aliphatics to adapt to possible shape changes in the S1’ pocket. These compounds
were added to a larger pool of compounds synthesised by Dr Robert Reid (IMB) to
form a library that was tested for HIV-1 PR antagonism in a fluorometric HIV-1 PR
assay (Bergman et al., 1995; Abbenante et al., 1995).
The investigation found that the macrocyclic template was a good tool to
overcome cooperativity, while maintaining a low inhibitor size. For substitution at
P2’, the N-alkyl sulfonamides were the more potent inhibitors, especially a 2-
substituted sulfonyl napthylene functionality which best filled the S2’ pocket and
directed the aromatic group along the line of the S2’ pocket and out towards the
solvent interface. In contrast the N-alkyl ureas were less potent as they likely directed
their substituents along the H-bonding channel that the peptide backbone normally
occupies, rather than entering the substrate binding pocket S2’. The best compounds
from the S1’ probe were small and bulky about the isostere linkage or had extended
hydrophobic bulk projecting further into the pocket. Extended aliphatic chains at P1'
were designed to accommodate changes at S1’ and had significantly lower potency.
Attempts to use a P1’ substitution, to bridge from S1' into the S3’ subsite and utilise a
conserved Arginine, resulted in the poorest inhibitor in the series.
The work in Chapter 3 may provide a springboard to further compounds that
are even more substrate-like and better avoid or minimise emerging viral resistance to
inhibitors. Several features of the current molecules are attractive for minimising
resistance including: (1) a small (n=4) cycle, and (2) substitutions at the P2’ position
with the 2-sulfonyl napthylene and other 2-sulfonyl constituents, such as quinolines
and coumarins that may direct their functionalities along the trough of the P2’ pocket,
and may have improved bioavailability properties (Fig 6.1).
It may be worthwhile to form a parallel library and modify the template at P3,
as the amine modification used in this study improved potency and may similarly
improve bioavailability (Fig 6.1). The direction that future work could take is more
towards the chemical modification of the cyclic inhibitors to address bioavailability
issues, followed by in vitro resistance studies – a track being pursued within the
Fairlie laboratory (Tyndell et al., 2000; Glenn et al., 2002). These efforts will
hopefully yield a successful inhibitor to advance to clinical trials in the not-too-distant
future.
Figure 6.1: The Possible Future Work of N-terminal Cyclic Inhibitors. An n=4 N-terminal cycle is
shown substituted at P2’ (yellow) with a 2-substituted sulfonyl napthylene and an isoamyl at P1’
(green), with adjacent groups that may substitute in these regions to form a small library, including a 2-
substituted sulfonyl quinoline (P2’). From the study a P3 amine (blue) was found to improve potency
and would be a worthwhile modification to add to the template for a parallel library.
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182
The third aim of the thesis was to develop an RNA-Rev binding assay to test
prospective Rev antagonists in vitro. Work towards this objective is summarised in
Chapters 4 and 5. Chapter 4 entailed the expression, labelling (RNA) and purification
of recombinant Rev protein and RNA, while Chapter 5 explores the use of this
material in two BIAcore assays.
The first BIAcore assay was a competitive assay for HIV-1 RBE3 (ligand)
immobilised onto streptavidin (SA) chips binding to competing analytes (Rev and
small molecule antagonists). By establishing parameters of the assay using
Neomycin, a screen of known small molecule antagonists were tested using constant
concentrations of antagonists and Rev. It was found that the order of inhibition was
Neomycin > Tobramycin > Paromomycin > Bekanamycin > Spectinomycin >
Kanamycin > Amikacin > Streptomycin and the key factor to potency in the screen
was the core Neamine functionality, however, all the compounds tested were poor
inhibitors of the Rev:RRE interaction.
The second assay used custom B1-Neutravidin-RBE3 and B1-Streptavidin-
RBE3 surfaces to directly assess the affinity of Neomycin, Rev and three peptides
based on the RRE binding domain of Rev (Rev34-50). The three peptides comprised
the native sequence, a succinylated form, and the native sequence incorporating a
helix-stabilising end group or cap. The proposition examined was that helix induction
can increase the affinity of shortened peptides for their biologically important targets.
It was found that non-specific associations occurred on the RNA surfaces, requiring
the use of calf thymus DNA as a masking agent and the application of a correction
factor for Neomycin and the succinylated peptide where a linear region of the binding
isotherms was observed. When rigorously analysed it was found for Neomycin and
Rev binding to only a single site on the RBE3 aptamer, but all peptides fit to a two
site model in the assay system.
Neomycin was found to bind with a KD of 7 – 19µM and from the high
affinity data of the two-state model, the succinylated peptide had the highest affinity
for the RBE3 aptamer (≤8nM). Based on maximum confidence intervals, the helix-
inducing cap and native peptide were found to be similar in potency, but the Rev
protein was found to be the weakest in the series. Upon this basis, the assay data
could not support the hypothesis that helix stabilisation can increase affinity for a
biologically important therapeutic target. However, the assay conditions themselves
are somewhat flawed as the temperature (25°C) and salt concentration (300mM
NaCl) are sufficiently high to affect the peptide secondary structure.
To date, our arsenal against RNA therapeutic targets is limited, yet RNA has
potential for therapeutic intervention of pathogens, disease and inflammatory states,
e.g. the control of AU-rich sequences of proto-oncogene and cytokine transcripts are
potential targets for helix mimetic therapeutics (Shaw and Kamen 1986; Sachs 1993).
In this regard it would be worthwhile to develop new helix inducing caps and study
how far we can truncate a capped peptide and maintain helicity and efficacy against
the RNA target and test the potential of capped peptides to directly serve as a
therapeutic.
Evolutionary theories accept RNA as playing a crucial role in the origins of all
nucleic acid containing life and cell dependent elements such as viruses and
transposons by serving as a hybrid of protein-like functionality and nucleic acid
coding (Schroeder et al., 2000; Wank et al., 1999). It is believed that some inhibitors
and antibiotics were intimately associated with RNA and proteins in this RNA-
protein prebiotic world (Schroeder et al., 2000; Wank et al., 1999). Included in this
class of inhibitors are cyclic peptides such as cyclosporins, viomycin and related
capreomycins, and aminoglycoside hybrids like vancomycin. It may be possible to
create new cyclic peptides similar to those used in Chapter 2 and 3, but instead of
constraining them in an extended conformation, they would be designed to mimic the
turn of an α-helix since, for all intents and purposes, a cyclic peptide can be viewed
as a loop or turn of an α-helix as shown in Figure 6.2.
Figure 6.2: How a Cyclic peptide can Serve as a Helix Mimetic. A helix of Rev34-50 is shown with two possible means of designing cyclic peptides to mimic regions. The lower region (orange) is between successive helix turns and the upper region (orange) is shown within a helix turn (image created from Battiste et al., 1995).
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184
Using the NMR models of the Rev:RRE complex (Battiste et al., 1995; Ye et
al.,1996; Bartel et al., 1991) and what we know are the key residues of the RRE
recognition sequence of Rev, it should be possible to "tune" a macrocyclic template to
be specific and potent, and thereby develop a whole new approach to RNA drug
design and inhibition. A library of such inhibitors could be synthesised quickly,
screened as described in Chapter 5 and rapidly cycled through rounds of drug design
and development to produce antagonists to Rev:RRE function.
The SPR work in Chapter 5 is novel for several reasons. In the first instance,
this is the first time a high-throughput BIAcore assay has been developed for RNA.
Typically fluoroescence anisotropy is the more common method for high-throughput
RNA assays, for screening libraries of compounds (Xavier et al., 2000) though radio-
ligand binding assays have been employed successfully for a very large library
(Chapman et al., 2002) and fourier transform ion cyclotron resonance mass
spectrometry is emerging as a tool (Hofstadler et al., 1999; Griffey et al., 1999;
Swayze et al., 2002).
A question worth asking is can SPR compete with more common or newer
emerging methods as a means of screening libraries of compounds that interact with
RNA? This study has shown that a robust high-throughput screen can be conducted
using an archetype internal standard (such as Neomycin for the Rev:RRE association)
and this is perhaps the greatest strength that SPR has for RNA applications. Once
established, screening by SPR can be conducted quickly and efficiently, unlike more
elegant and labour intensive studies such as kinetic and equilibrium studies using
SPR. However, SPR is labour intensive and technical. A great deal of time is needed
to learn all the aspects of SPR interactions and investigate the limitations of the
system before the potential of the system can be realised. In this study alone, more
than 250 SPR experiments were conducted to derive the assays reported. In this
respect SPR high-throughput screening assays will likely be conducted by researchers
who are willing to invest the time, energy and money to make the assay system work.
I do not envisage SPR eclipsing other more traditional biophysical characterisation
methods or newer emerging screening applications, but rather SPR will prove to be
another tool in our arsenal to more quickly arrive at our goals.
Throughout this study, non-specific and surface events were challenging to
overcome and control, requiring the development of strategies and techniques such as
185
lower density ligand and matrix surfaces, the use of buffer additives, the subtraction
of controls and buffer bulk effects as well as correction of data for non-specific
associations. Though SA and B1-Pioneer chips were used, another surface that could
prove very useful to overcome surface problems is BIAcore lipid chips that allow
capture proteins to be immobilised by impregnating them into a micelle and forming
a bilayer on the sensorchip surface. Using such a surface, the charged RNA would not
be predicted to associate with the lipophilic surface and the only significant problem
would arise if the analyte is found to be excessively hydrophobic (this might only
require stabilising the protein with mild surfactants). Therefore, future work could
extend this study using lipid chips and would form a good rounding of the work and
the basis of a publication for a techniques paper on SPR methods for handling RNA.
The advent of HIV-1 PR and reverse transcription inhibitors has had an
immense therapeutic impact, reducing the viral load and hence the onset of AIDS-
defining illnesses and increasing life expectancy (Egger et al., 2002). The a priori
concept in this study is that combination therapy with Rev:RRE antagonists and HIV-
1 PR inhibitors may similarly form a successful basis for combination therapy.
Being a Retrovirus, HIV-1 has evolved a clever means of replication that
maximises the efficiency of virion production while minimising the size of the
genome. By encoding multiple proteins within the same coding regions and dividing
the replication cycle into early and late phases (mediated by the concentration of
Rev), and subsequently releasing late proteins by activation of an internally coded
protease, HIV-1 maintains efficient virion production from a minimal genome size.
The Rev:RRE and HIV-1 PR steps of HIV-1 replication represent closely associated
and interdependent stages leading towards virion production and maturation since the
vRNA that Rev exports from the nucleus encodes HIV-1 PR and it’s substrates. This
interdependence makes the inhibition of Rev:RRE and HIV-1 PR function attractive
to combination therapy and continuation of the work conducted in this thesis can lead
to suitable testing of the potential synergistic inhibition of HIV-1 replication using
Rev:RRE and HIV-1 PR inhibitor and perhaps a novel therapeutic approach to HIV-1
infection.
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