JAIDS Journal of Acquired Immune Deficiency Syndromes:
The A62V and S68G Mutations in HIV-1 Reverse Transcriptase Partially Restore the Replication Defect Associated With the K65R Mutation
Svarovskaia, Evguenia S PhD*; Feng, Joy Y PhD†; Margot, Nicolas A MA†; Myrick, Florence BS*; Goodman, Derrick BS*; Ly, John K MS†; White, Kirsten L PhD†; Kutty, Nilima BS†; Wang, Ruth BS†; Borroto-Esoda, Katyna MS*; Miller, Michael D PhD†
From the *Gilead Sciences, Inc, Durham, NC; and †Gilead Sciences, Inc, Foster City, CA.
Received for publication October 11, 2007; accepted March 11, 2008.
Supported by Gilead Sciences, Inc.
Previously presented in part at the 14th Conference on Retroviruses and Opportunistic Infections, February 25-28, 2007, Los Angeles, CA.
All authors are employees of Gilead Sciences, Inc.
E.S.S. and J.Y.F. contributed equally to the work.
Correspondence to: Dr. Evguenia Svarovskaia, PhD, Gilead Sciences, Inc, 4 University Place, 4611 University Drive, Durham, NC, 27707 (e-mail: email@example.com).
Background: The K65R mutation in human immunodeficiency virus type 1 reverse transcriptase can be selected by abacavir, didanosine, tenofovir, and stavudine in vivo resulting in reduced susceptibility to these drugs and decreased viral replication capacity. In clinical isolates, K65R is frequently accompanied by the A62V and S68G reverse transcriptase mutations.
Methods: The role of A62V and S68G in combination with K65R was investigated using phenotypic, viral growth competition, pre-steady-state kinetic, and excision analyses.
Results: Addition of A62V and S68G to K65R caused no significant change in human immunodeficiency virus type 1 resistance to abacavir, didanosine, tenofovir, or stavudine but partially restored the replication defect of virus containing K65R. The triple mutant K65R+A62V+S68G still showed some replication defect compared with wild-type virus. Pre-steady-state kinetic analysis demonstrated that K65R resulted in a decreased rate of incorporation (kpol) for all natural dNTPs, which were partially restored to wild-type levels by addition of A62V and S68G. When added to K65R and S68G, the A62V mutation seemed to restore adenosine triphosphate-mediated excision of tenofovir to wild-type levels.
Conclusions: A62V and S68G serve as partial compensatory mutations for the K65R mutation in reverse transcriptase by improving the viral replication capacity, which is likely due to increased incorporation efficiency of the natural substrates.
Human immunodeficiency virus type 1 (HIV-1) infection is currently effectively managed with antiretroviral therapies including highly active 3-drug combinations. The emergence of drug resistance mutations is a major limitation to long-term treatment efficacy. Drug resistance mutations may be coselected with other mutations that have little or no effect on susceptibility of the virus to the drug but result in compensation of the decreased replication capacity associated with the drug resistance mutations. These compensatory mutations have been best characterized in the context of HIV-1 protease inhibitor-associated resistance mutations.1 Often, these compensatory mutations are naturally occurring polymorphisms in antiretroviral treatment-naive patients but are enriched among treatment-experienced patients with drug resistance mutations.
The K65R mutation in HIV-1 reverse transcriptase (RT) can be selected during therapy containing didanosine (ddI), abacavir (ABC), tenofovir disoproxil fumarate [the prodrug of tenofovir (TFV)], or stavudine (d4T), resulting in reduced susceptibility to these drugs.2-10 Virus containing the K65R mutation also has strongly impaired replication capacity.11-13 The S68G and A62V RT mutations have been shown to be coselected with K65R in clinical studies,6,8 in cell culture drug selection experiments,14 and through an analysis of the Monogram Biosciences HIV-1 RT genetic database.15 The frequency of A62V+K65R+S68G triple mutants constituted about 12% of K65R cases in study 903.8 A similar frequency (43/354, 12%) of the triple mutant was found among patient viral sequences containing K65R in the Stanford database. Higher frequencies of the K65R with the individual mutations A62V and S68G are observed. The S68G mutation is a polymorphic change in RT, which is present in a minority of treatment-naive and treatment-experienced individuals, but it is strongly enriched among isolates expressing K65R (~37%) (http://hivdb.stanford.edu).16 A possible compensatory role of the S68G mutation in the presence of K65R to improve viral fitness has been suggested.6,8,14 In contrast, the A62V mutation is rarely observed in treatment-naive patients and is usually reported as part of a multinucleoside resistance (MNR) complex.17 The Q151M MNR complex containing A62V, V75I, F77L, F116Y, and Q151M has been shown to reduce susceptibility to all currently approved nucleoside reverse transcriptase inhibitors (NRTIs) except TFV disoproxil fumarate.17 Another MNR complex contains an insertion at position 69 of RT in combination with M41L, A62V, K70R, L210W, T215Y/F, and K219E/Q and increases resistance to all approved NRTIs.17 The role of A62V in MNR is not well defined; however, it has been suggested that the A62V mutation contributes to increased excision of NRTIs when present in MNR complex containing insertions at position 69.18 Another recent study proposed that the presence of the A62V mutation favors positioning of RT in the pretranslocational state, which also would be consistent with favoring excision of chain terminators.19 In the Stanford database, A62V is present in ~15% of isolates with K65R but in less than 3% of those without K65R (http://hivdb.stanford.edu).16
In this study, we performed drug susceptibility assays and RT-catalyzed incorporation and excision assays to evaluate the role of the S68G and A62V mutations in combination with K65R. We also measured relative viral fitness of K65R alone and in combination with A62V and S68G to determine the effect of these mutations on the replication capacity of K65R mutant virus. The results demonstrate that A62V and S68G act as partial compensatory mutations for the K65R mutation in RT by improving the viral replication capacity, which is likely due to the increased incorporation efficiency of natural dNTPs.
TFV and emtricitabine (FTC) were synthesized by Gilead Sciences. Lamivudine (3TC) was obtained from Moravek Biochemicals (Brea, CA), and ABC was obtained from GlaxoSmithKline (Research Triangle Park, NC). Zidovudine (AZT) and ddI were purchased from Sigma-Aldrich (St Louis, MO), and d4T was supplied by Bristol-Myers Squibb (New York, NY). All 4 natural dNTPs were from GE Healthcare BioSciences (Piscataway, NJ). AZT triphosphate was purchased from TriLink Bio-Technologies (San Diego, CA). TFV diphosphate (TFV-DP) and FTC triphosphate were synthesized by ChemCyte, Inc (San Diego, CA). Carbovir triphosphate, the active metabolite of ABC, was synthesized at Sierra Bioresearch (Tucson, AZ). Oligonucleotides used in pre-steady-state analysis D19-mer and DNA36-mers were synthesized and PAGE purified by Integrated DNA Technologies, Inc (Coralville, IA) (Table 1). The DNA primer was 5′-32P labeled and annealed to DNA templates as previously described.20
Recombinant HIV-1 RT Enzyme Construction and Purification
Wild-type HXB2D RT enzyme corresponding to the p51 and p66 subunits was cloned into pET14b vector lacking the thrombin cleavage site but containing an N-terminal 6-His tag linker as previously described.21 K65R, A62V+K65R, K65R+S68G, and A62V+K65R+S68G mutations were introduced using the QuickChange XLII Site-Directed Mutagenesis Kit (Stratagene, La Jolla, CA). The resulting pETp66 plasmids encoded an HIV-1 RT open reading frame containing either 440 amino acids (p51 subunit) or 560 amino acids (p66 subunit) with an N-terminal His tag. Protein expression of each subunit was induced in Escherichia coli BL21 (DE3) cells (Stratagene), purified, and associated to yield p51/p66 heterodimers as previously described.21 The active site concentrations were 52%, 42%, 56%, and 48% for wild-type, K65R, A62V+K65R, and A62V+K65R+S68G RTs, respectively, for 2′-deoxyadenosine 5′-triphosphate (dATP) incorporation into a DNA/DNA 19/36-mer as determined by pre-steady-state burst experiments as previously described.22
Phenotypic Analyses and Site-Directed Recombinant Viruses
The susceptibilities of the HIV-1 viruses to TFV, FTC, AZT, 3TC, d4T, ABC, and ddI were determined in MT-2 cells as described previously.14,23,24 Briefly, 1.2 × 106 MT-2 cells were infected with either wild-type HIV-1 (HXB2D) or mutant virus and incubated for 5 days starting with an initial concentration of 1.7 × 104 cells/well. The antiviral effects of the compounds tested were measured by determining the HIV-1 cytopathic effect by using the vital dye XTT (Sigma-Aldrich). Effective concentrations which inhibited 50% of viral replication (EC50) were determined using SigmaPlot (Systat Software, San Jose, CA). Site-directed recombinant viruses were generated by cotransfection in the Sup-T1 cell line of the HIV-1 proviral molecular clone pHXB2Δ2-261RT (plasmid DNA provided by C. Boucher, Utrecht Medical Center, Utrecht, The Netherlands) with a polymerase chain reaction (PCR) product corresponding to the first 300 amino acids of HIV-1 RT on which the mutation(s) of interest had been inserted by site-directed mutagenesis. Replication-competent viruses generated by homologous recombination were harvested after 10-20 days.25,26 The nucleotide sequence corresponding to the first 300 amino acids of RT for the obtained viruses was confirmed by genotypic analysis using an ABI 3100 genetic analyzer (Applied Biosystems, Inc, Foster City, CA).
Growth Competition Assay
To generate mutant viruses for growth competition assays, a fragment of the pETp66 plasmid containing wild-type or mutant RT gene was amplified using primers containing Xba I and Xma I restriction sites and then cloned into pxx-LAI, an HIV-1LAI backbone.27 The resulting recombinant proviral clones were transfected into 293T cells using TransIT-LT1 Transfection Reagent (Mirus Bio Corporation, Madison, WI). Virus was collected 48 hours posttransfection, filtered, aliquoted, and frozen at −80°C. Titers of viral stocks were determined using the Magi assay.28 Viral stocks were diluted and mixed at 1:1 ratio to a total of approximately 1.5 × 102 infectious units and used to infect 1.5 × 105 MT-2 cells in 1 mL for 2 hours with resultant multiplicity of infection (MOI) of 0.01. To remove unbound virus, cells were pelleted and washed twice with 3 mL of phosphate-buffered saline, resuspended in 5 mL of medium, and cultured at 37°C. On days 4, 7, 11, and 15, cells were pelleted and the supernatant was removed and replaced with 1.5 × 105 MT-2 cells in a total volume of 10 mL. Viral RNA was extracted from 140 μL of the supernatant at each time point using QIAamp Viral RNA Mini Kit (Qiagen, Valencia, CA). Viral RNA samples were subjected to DNase digestion for 60 minutes using TURBO DNAfree kit (Ambion, Austin, TX). The percentage of mutant viral RNA present at each time point was determined using MultiCode RTx PCR (EraGen, Madison, WI).
MultiCode RTx PCR Assay
To differentiate between 2 competing viruses, silent changes were introduced into the xx-LAI plasmid by site-directed mutagenesis to generate a wild-type virus with marker mutations (F-xx-LAI).27 These silent mutations change codons of amino acids 6 and 7 of RT gene from GAA and ATC of xx-LAI to GAG and ACG of F-xx-LAI. For each growth competition experiment, viruses with 2 different backbones were combined. The percentage of mutant virus for each time point during the competition experiment was determined using MutiCode RTx allele-specific PCR that targeted marker mutations as previously described but with modifications.29,30 Briefly, the PCR conditions involved 20-μL reaction mixtures in 1× ISOlution buffer (EraGen), 5 mM dithiothreitol, Titanium Taq DNA polymerase (Clontech, Palo Alto, CA), and SuperScript III RT (Invitrogen, Carlsbad, CA) at the manufacturer's recommended concentration. The MultiCode RTx RT-PCR assays were carried out using the Roche LightCycler 480 (Roche, Indianapolis, IN) with the following cycling conditions: 5 minutes at 54°C of reverse transcription step, 2 minutes of denaturing at 95°C, and 1 cycle of 5 seconds at 95°C, 5 seconds at 47°C, and 20 seconds at 72°C, followed by 45 cycles of 5 seconds at 95°C, 5 seconds at 57°C, and 20 seconds at 72°C with optical read. A thermal melt with 2.5 optical readings per degree Celsius from 60 to 95°C was performed directly after the last 72°C step of thermal cycling. Allele-specific primers were as follows: the F-xx-LAI (GG)-specific primer was FAM- isoC- TGCTGACATTAGTCCTATTGAGACG; the xx-LAI (AT)-specific primer was HEX-isoC-ACGAGACCATTAGTCCTATTGAAACT; and the reverse primer was TGTCAATGGCCATTGTTTAACTTTTGG. PCR primers were used at the following final concentrations: 2 forward allele-specific primers at 200 nM and reverse primer at 400 nM. The percentage of a mutant was determined based on standard curves generated using SigmaPlot (Systat Software).
For selected samples, the percentage of mutant viruses in the population was also determined at day 10 of the growth competition experiment using clonal analysis. Viral RNA samples were prepared as described above and RT-PCR amplified using the following primers: 3HIV3665; 5′-TATCTGGTTGTGCTTGAATGATTCCTAATGCAT-3′, 5HIV2053, 5′-GGTACAGTATTAGTAGGACCTACAC-3′; and 3HIV2896, 5′-CCCACTAACTTCTGTATGTCATTGACAGTCCAGC-3′. Resultant PCR products were cloned into TopoTA as per manufacturer's protocol (Invitrogen). DNA from 25 to 30 bacterial colonies per sample was sequenced. The percentage of mutant virus in the population was determined as number of sequences containing the K65R mutation divided by the total number of colonies analyzed.
Relative Fitness Calculations
Relative fitness value was calculated as previously described (1+s) = exp[1/t × ln (Mt/Wt × Wt0/Mt0)], where t is time in days and Mt, Mt0, Wt, and Wt0 are fractions of mutant virus at initial and time of measurement and fractions of wild-type virus at initial and time of measurement, respectively.31
Pre-Steady-State Kinetic Analyses
Transient kinetic experiments were performed by the rapid quench method as described previously using a KinTek Instrument Model RQF-3 rapid-quench-flow apparatus.22 All the concentrations described below are final concentrations unless noted otherwise. Briefly, the reactions were carried out by mixing a solution containing the preincubated complex of HIV-1 RT (100 nM) and 5′-32P-labeled D19/D36 primer/template duplex (300 nM) with a solution containing 10 mM MgCl2 and various concentrations of the dNTP. The templates and a primer that were used in this study are listed in Table 1. When single-turnover conditions were used, the reactions were carried out by mixing a solution containing the preincubated complex of HIV-1 RT (200-250 nM) and 5′-32P-labeled D30/D45 duplex (50 nM) with a solution of 10 mM MgCl2 and various concentrations of the dNTP. The reactions were quenched with 0.3 M EDTA at time intervals ranging from 3 milliseconds to 2 minutes. The products from each quench reaction were resolved by electrophoresis (16% acrylamide, 8 M urea) followed by Phosphor Imaging (Bio-Rad Personal Molecular Imager FX). Data were fitted by nonlinear regression (KaleidaGraph 3.51). Under burst conditions, the product formation occurred in a fast exponential phase, followed by a slower linear phase. Data were fitted into a burst equation: [product] = A[1 − exp(−kobsdt) + ksst], where A represents the amplitude of the burst that correlates with the concentration of enzyme in active form, kobsd is the observed first-order rate constant for dNTP incorporation, and kss is the steady-state rate constant. Data from single-turnover experiments were fit to a single exponential equation: [product] = A[1 − exp(−kobsdt). The dissociation constant (Kd) for dNTP to the RT•DNA complex is calculated by fitting the data to the following hyperbolic equation: kobsd = kpol[dNTP]/(Kd + [dNTP]), where kpol is the maximum rate of dNTP incorporation and [dNTP] is the corresponding concentration of dNTP.
ATP-Mediated Excision Assay
TFV chain-terminated DNA primers were generated using a 31-nucleotide DNA primer L31 (5′-CTA CTA GTT TTC TCC ATC TAG ACG ATA CCA G-3′),32 which was 5′ end labeled with [γ-33P] adenosine triphosphate (ATP) (GE Healthcare BioSciences), using T4 polynucleotide kinase (New England Biolabs, Ipswich, MA), annealed to a 1.5-fold excess 50-nucleotide DNA template WL50 (5′-GAG TGC TGA GGT CTT CAT TCT GGT ATC GTC TAG ATG GAG AAA ACT AGT AG-3′),32 and chain terminated with TFV-DP using wild-type RT enzyme at 37°C for 30 minutes as previously described.21,26 The TFV-terminated primers were gel purified and annealed to 1.5-fold excess of WL50 as described.21 Excision was initiated with 3.2 mM ATP (Invitrogen) that was pretreated with Tth pyrophosphatase (Roche) in the presence of 3 μM of the next complementary nucleotide dATP (the estimated physiological concentration)33 that was also pretreated with Tth pyrophosphatase. After 120 minutes of the excision reaction, the RT enzyme was heat inactivated and the unblocked primers were extended with 500 μM dNTPs using Klenow polymerase (New England BioLabs, Beverly, MA) as previously described.26 Reactions were stopped by addition of an equal volume of formamide-loading buffer. Reaction products were separated through 7 M urea containing 16% polyacrylamide sequencing gels. Bands were quantified with a Storm860 PhosphorImager (GE Healthcare BioSciences) and ImageQuant TL software (Molecular Dynamics, Sunnyvale, CA). Degradation of the blocked primer band did not occur when each RT enzyme was incubated with the primer template at 37°C in absence of ATP. The percentages of full-length, rescued primers were determined from 3 or more independent experiments.
Effect of the A62V and S68G Mutations on Susceptibilities of K65R HIV-1 Virus to NRTIs
In vitro drug susceptibilities of wild-type virus and K65R mutant, alone or in combination with A62V and/or S68G, were determined to address the effect of these mutations on resistance to NRTIs. Results of in vitro drug susceptibility assays for TFV, AZT, FTC, 3TC, ddI, d4T, and ABC are shown in Table 2. The virus harboring only the K65R mutation had a 2.9-fold increase in the EC50 of TFV compared with that of the wild-type control (P < 0.01). This level of resistance to TFV was not significantly affected by addition of A62V or/and S68G RT mutations to K65R mutant virus with changes in EC50 of 1.1-, 1.1-, and 1.2-fold above the EC50 of K65R alone for A62V+K65R, K65R+S68G, and A62V+K65R+S68G mutants, respectively (data not shown). Similarly, no significant changes in resistance levels were observed for FTC, 3TC, ddI, d4T, and ABC with addition of A62V or/and S68G mutations to K65R virus. The K65R mutant virus exhibited full susceptibility to AZT with EC50 levels 0.6-fold lower than the EC50 for the wild-type control. Susceptibility to AZT was maintained with the addition of the S68G mutation to K65R virus. However, addition of the A62V mutation seemed to reduce the susceptibility levels to AZT. For the triple mutant A62V+K65R+S68G, the AZT susceptibility was 2.6-fold above the wild-type control.
The A62V and S68G Mutations Partially Restore the Replication Defect of the K65R RT Mutant HIV-1
Because the A62V and S68G mutations did not significantly change the resistance levels of K65R mutant HIV-1 to most NRTIs, we hypothesized that these mutations were coselected with K65R to restore the replication fitness of the K65R virus. To test our hypothesis, a growth competition assay was developed that allowed determination of the replication fitness for each mutant in combination with wild-type or K65R viruses. We created a pair of HIV-1 LAI-based constructs that contained silent mutations at the third positions of codons encoding amino acids 6 and 7 of RT. These silent changes served as markers to monitor the replication of each virus and were quantified by real-time allele-specific PCR during the course of the growth competition experiments. The relative replication capacity of each mutant was measured in competition with wild-type virus. Control experiments demonstrated that construction of the wild-type virus with the silent mutations did not affect the viral fitness (Table 3). Virus stocks of the K65R, A62V+K65R, K65R+S68G, and A62V+K65R+S68G mutants were coinfected with wild-type virus at a 1:1 ratio. As expected, the proportion of K65R compared with wild-type virus decreased from 50% to 23% by day 11, indicating reduced replication capacity of the K65R virus in the absence of drug (Fig. 1A). The replication defects of each mutant were further quantified by calculating relative fitness (1+s) values (Table 3).31 The order of viral fitness was as follows: K65R < A62V+K65R+S68G < wild type, with significant difference at each comparison (P < 0.05). The double mutants A62V+K65R and K65R+S68G seemed more fit than K65R alone but less fit than triple mutant; however, differences in relative fitness were not statistically significant.
To verify that quantification of mutant virus growth was accurate using detection of the silent marker mutations by real-time PCR, we performed clonal analysis of 3 viral samples at day 11 from growth competitions between K65R and wild-type viruses. Sequencing of a total of 129 clones indicated that identical number of clones contained K65R (21 clones) and marker mutations, indicating presence of K65R on between 10% and 24% of the clones as calculated using binominal confidence interval calculator with alpha of 0.05. Overall detection of K65R mutant virus by the real-time PCR-based assay was in close agreement with the results of clonal analysis, indicating that the rate of recombination under the conditions used did not significantly affect quantification of mutant viruses during growth competition experiments.
To confirm that the double and triple mutants have replication advantages over K65R alone, we performed direct growth competition experiments between K65R and the double or triple mutants. In each case, the proportion of K65R virus was reproducibly diminished in each experiment, indicating the stronger replication capacity of the double mutant (data not shown) and triple mutant over K65R alone (Fig. 1B). Even though addition of the A62V and S68G mutations to the K65R virus resulted in increased viral replication capacity, the A62V+K65R+S68G triple mutant still remained replication defective when competed with wild-type virus (Fig. 1C, Table 3).
To further evaluate the replication defect of the triple mutant, we performed competition experiments in the presence of several concentrations of TFV (Fig. 1D). The results indicated that the A62V+K65R+S68G triple mutant had a replication advantage over wild-type virus only in the presence of TFV at concentration ≥7 μM, presumably due to drug inhibition of wild-type virus but preserved growth of the resistant mutant. However, in the absence of TFV or using low concentrations of TFV, the triple mutant continued to display a replication defect as compared with wild-type virus.
S68G and A62V Improved the Incorporation Efficiency of dATP and dGTP for K65R HIV-1 RT
The single-nucleotide incorporation of natural dNTPs was studied using pre-steady-state kinetic analyses. The results are summarized in Table 4. Compared with wild-type RT, the presence of the K65R mutation decreased the incorporation efficiency of the natural 2′-deoxynucleoside 5′-triphosphate (dNTP), dATP, 2′-deoxyguanosine 5′-triphosphate (dGTP), thymidine 5′-triphosphate (TTP), and 2′-deoxycytidine 5′-triphosphate (dCTP) by 2.0-, 4.1-, 2.1-, and 1.6-fold, respectively, mainly due to the decreases in kpol. These findings are consistent with results published by other groups.34,35 The addition of S68G and A62V+S68G on a background of K65R showed increasing improvements on the kpol values for all 4 dNTPs. Compared with K65R, the triple mutation led to a 3.2-, 2.3-, 4.1-, and 1.6-fold increase in the rate of incorporation for dATP, dGTP, TTP, and dCTP, respectively. These were accompanied by a 3.6-fold increase in the Kd value for TTP and relatively minor changes in the Kd values for dATP, dGTP, and dCTP. Overall, the A62V+S68G mutations completely restored the incorporation efficiency (kpol/Kd) of dATP for K65R RT to the wild-type level and partially restored it for dGTP (Fig. 2). The incorporation efficiency of TTP and dCTP was not significantly different between K65R and the double or triple mutants because the effect of the increased kpol was diminished by an increased Kd value.
Of note, many kinetic measurements using DNA or RNA polymerase, the absolute values for kinetic constants, are sequence dependent; however, the relative ratio of wild type to mutant enzyme, which are the key factors to determine the effect of a mutation, remains relatively constant.11,35,36
S68G and A62V Showed No Impact on the Resistance Factors for ddNTP Analogues by K65R
The single-nucleotide incorporation of 4 clinically relevant nucleotide analogues was also studied using pre-steady-state kinetic analysis. The results are summarized in Table 4 and Figure 3. In addition to the 3 kinetic parameters kpol, Kd, and kpol/Kd used for evaluation of the natural dNTPs, 2 more parameters were calculated. The selectivity factor, defined as the ratio of kpol/Kd of a natural dNTP to its analogue, demonstrates the degree of preference an enzyme has toward the natural dNTP as compared with its analogue. The resistance factor is defined by the ratio of selectivity factors of wild-type and mutant RT. This parameter indicates how the RT mutation affects the enzyme's ability to discriminate the NRTI analogue from natural dNTPs.20,35,37,38 Similar to earlier reports,11,34-36 the K65R mutant RT had resistance factors of 11-, 3.3-, 3.6-, and 5.1-fold, respectively, to TFV-DP, carbovir triphosphate, AZT triphosphate, and FTC triphosphate (the active metabolites of TFV, ABC, AZT, and FTC, respectively) (Fig. 4). The resistance was mainly caused by a decrease in the kpol for these analogues. When A62V+S68G was introduced into a background of K65R, 2.2- to 5.6-fold increases in kpol values were observed for the NRTIs, accompanied by moderate increases in Kd values. This resulted in an increase in the incorporation efficiencies for TFV-DP and carbovir triphosphate that were partially offset by the increase in incorporation efficiencies of their corresponding natural substrates resulting in only slight reductions in the resistance factors, which were not significantly different than K65R alone (Fig. 3).
ATP-Mediated Excision of TFV
The ATP-mediated excision of TFV was measured. Wild-type RT showed moderate levels of TFV excision (Fig. 4). In contrast, K65R and K65R+S68G RT showed significantly reduced excision when compared to wild-type enzyme, agreeing with previously published results for K65R.21,39 The A62V+K65R+S68G RT, on the other hand, seemed to restore excision of TFV to wild-type levels. To address a possibility that effects of mutations on excision properties of RTs are specific to a given template-primer sequence, we performed excision assays using an alternative primer template (data not shown), resulting in excision levels in the following order WT > A62V+K65R+S68G > K65R.
HIV-1 virus containing the K65R RT mutation can be selected by ABC, ddI, TFV, and d4T and is associated with reduced susceptibility to these NRTIs and decreased replication capacity. The K65R mutation is frequently accompanied by the A62V and S68G RT mutations in HIV-1 sequence databases and in patients failing therapy. In this study, we performed direct viral growth competition experiments to compare the replication fitness of viruses with K65R, with and without A62V and S68G. Our results demonstrated that the reduced replication fitness of K65R was partially restored by addition of the A62V or S68G mutations. We explored the mechanism underlying the compensatory effect of the A62V and S68G mutations on K65R RT using pre-steady-state kinetic analysis. These results demonstrated that K65R resulted in a decreased rate of incorporation (kpol) for all natural dNTPs. The incorporation efficiency (kpol/Kd) of dATP was completely restored to a wild-type level for the triple mutant A62V+K65R+S68G; the incorporation efficiency of dGTP was partially restored. The compensatory effect of the A62V and S68G mutations observed in the cell culture competition experiments is consistent with the finding of improved incorporation efficiency of dATP and dGTP by the triple mutant. An earlier pre-steady-state kinetic report on A62V for the incorporation of dGTP and a dioxolane analogue-b-D-dioxolane guanosine 5′-triphosphate (DXG-TP) also showed increases in both the kpol and Kd values and an increased incorporation efficiency for dGTP.40
The K65R and associated A62V and S68G RT mutations are all located in the flexible β3-β4 fingers loop domain located near the polymerase active site.41 During dNTP incorporation, part of the fingers subdomain rotates toward the palm subdomain and inward toward the primer/template and the polymerase active site.22,41-43 The K65R change in this domain affects multiple enzymatic properties of RT. The K65R mutation alters the incorporation rate of dNTPs and NRTIs, processivity, and excision of NRTIs and also results in an overall decrease in replication capacity.13,21,34,39 It has been hypothesized using molecular models that K65R mediates these effects by repositioning the bound substrate or potentially by generating additional contacts that may decrease the mobility of this fingers loop domain.39 The A62V and S68G changes may partially alleviate the K65R-mediated reduction in the flexibility of the fingers loop and active site substrate positioning. This hypothesis is in close agreement with the observed effects of K65R, A62V, and S68G on the binding affinity of dNTP and ddNTP for the RT-DNA complex.
The A62V and S68G mutations did not significantly alter the single-nucleotide incorporation assay resistance factors for K65R HIV-1 RT incorporation of NRTIs. This result is in agreement with cell culture-based phenotypic analysis of NRTI susceptibilities. The only discrepancy is in the case of AZT. However, it is also not clear whether these small changes in susceptibility to AZT for the triple mutant A62V+K65R+S68G would have any clinical relevance when compared to the high-level resistance achieved by the thymidine analog mutations selected by AZT (>100-fold).
Increased NRTI excision by RT containing thymidine analogue mutations is a mechanism of resistance that explains the decreased HIV-1 susceptibility of the corresponding virus to most NRTIs.44,45 For K65R, however, there is decreased NRTI excision compared with wild-type RT leading to greater stability of the chain terminator and reduced resistance.21,35,39 In the current work, the A62V mutation seemed to restore wild-type levels of TFV excision, similarly to the restored dNTP incorporation. The excision data can be explained mechanistically based on a recent report that showed that the A62V mutation favors positioning of RT in the pretranslocational state that favors excision.19 The observed increases in TFV excision associated with the triple mutant relative to K65R may also serve to counteract the slight increases in TFV incorporation efficiency observed in pre-steady-state kinetics, which were not reflected in a change in phenotypic susceptibility to TFV.
In conclusion, the studies described here have characterized the role of the A62V and S68G HIV-1 RT mutations in combination with K65R. Collectively, our analyses suggest that the A62V and S68G mutations play a compensatory role and partially rescue the replication defect of the K65R mutant but do not have a significant effect on resistance levels to NRTIs. The mechanism for the increase in replication capacity of the A62V+K65R+S68G triple mutant as compared with K65R alone lies in the enhanced incorporation efficiency of the natural substrates dATP and dGTP. The triple mutant, however, still remains replication impaired relative to wild-type HIV-1.
We especially thank Michael J. Moser (Eragen Biosciences, Madison, WI) for valuable advice for design of MultiCode RTx primers for growth competition assay. We thank Jeanette Harris, Joshua Waters, Damian McColl, Yuao Zhu, Rebecca Ledford, and Swami Swaminathan for helpful discussions throughout this project.
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© 2008 Lippincott Williams & Wilkins, Inc.
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