JAIDS Journal of Acquired Immune Deficiency Syndromes:
Low-Level K65R Mutation in HIV-1 Reverse Transcriptase of Treatment-Experienced Patients Exposed to Abacavir or Didanosine
Svarovskaia, Evguenia S PhD*; Margot, Nicolas A MA†; Bae, Andrew S BA*; Waters, Joshua M BS*; Goodman, Derrick BS*; Zhong, Lijie PhD†; Borroto-Esoda, Katyna MS*; Miller, Michael D PhD†
From *Gilead Sciences, Inc., Durham, NC; and †Gilead Sciences, Inc., Foster City, CA.
Received for publication January 17, 2007; accepted June 14, 2007.
Supported by Gilead Sciences, Inc.
Previously presented in part at the XV International HIV Drug Resistance Workshop, Sitges, Spain, June 13-17, 2006.
All authors are employees of Gilead Sciences, Inc.
Correspondence to: Michael Miller, PhD, Gilead Sciences, 333 Lakeside Drive, Foster City, CA 94404 (e-mail: email@example.com).
Background: Prior abacavir (ABC) or didanosine (ddI) therapy can result in the L74V/I or K65R mutation in HIV-1 reverse transcriptase. Preexisting K65R may have an impact on the treatment response to tenofovir disoproxil fumarate (TDF).
Methods: An allele-specific polymerase chain reaction (AS-PCR) assay was developed to detect K65R with a lower limit of quantitation of 0.5%.
Results: Among baseline plasma samples from 63 treatment-naive patients, no K65R was detected by AS-PCR. Among baseline samples from 154 treatment-experienced patients, 8 had K65R and 44 had L74V/I by population sequencing. Low-level K65R was detected in an additional 11 patients by AS-PCR, 3 of whom subsequently developed full K65R. Baseline K65R correlated with absence of thymidine analog mutations (TAMs; P = 0.003) and use of ABC or ddI (P = 0.004). Patients with full or low-level K65R at baseline or with L74V/I showed a diminished TDF response. Multivariate analyses confirmed that multiple TAMs, K65R, and L74V/I were independent predictors of diminished TDF response.
Conclusions: Prior therapy with ABC or ddI can result in a population genotype that shows K65R or L74V/I but does not reveal low-level K65R present in some patients. Subsequent treatment intensification with TDF resulted in a poor virologic response and may result in expansion of the preexisting K65R mutant.
Overlapping drug resistance profiles can result in cross-resistance among members of the same class of antiviral drugs and subsequent poor treatment responses in patients with cross-resistance. Use of HIV-1 genotyping and phenotyping assays is part of clinical practice to assess the presence of resistance and potential cross-resistance to other drugs. Current genotyping assays are based on population sequencing of the HIV-1 quasispecies within an infected individual and have limits of detection for minority variants of approximately 20% to 30%.1 However, minority variants at lower frequencies than detectable by standard sequencing may have an impact on treatment response, as has been shown for resistance within the nonnucleoside reverse transcriptase (NNRTI) inhibitor class.2-4
The K65R mutation in HIV-1 reverse transcriptase (RT) is associated with didanosine (ddI), abacavir (ABC), stavudine (d4T) and tenofovir disoproxil fumarate (TDF) therapy. TDF seems to select exclusively for a K65R mutation; however, some reports have also shown the development of a K70E mutation that may represent a temporally intermediate resistance pattern for TDF.5-7 Both ddI and ABC can select for a K65R or L74V mutation in RT, as evidenced by monotherapy studies in which resistance was characterized.8-10 For ABC, an M184V mutation typically occurred first (independent of lamivudine [3TC] use), followed by K65R or L74V, and possibly Y115F. At the dose of 300 mg administered twice daily, ABC selected more frequently for L74V than for K65R and both patterns did not generally occur in the same patients. Similar observations have been made for combination therapy with ABC, 3TC, and efavirenz (EFV).11 In the case of ddI, the pattern of resistance was similar to that of ABC; a K65R or L74V mutation may develop.10 In this monotherapy study, clonal analysis was performed from 3 patients who had evidence of both patterns. In all clones analyzed, the mutation patterns appeared on independent genomes. These findings may be related to the biochemical observations showing that an RT mutant containing both mutations is severely impaired in its enzymatic function.12
Patients treated with ddI or ABC who had developed a detectable K65R mutation would not be considered candidates for subsequent TDF therapy. Patients with no detectable resistance or an L74V mutation would be considered candidates because the L74V RT mutant of HIV-1 remains fully susceptible to tenofovir in vitro.13 However, clinical data have suggested a reduced TDF response in patients with an L74V mutation.14,15 We have recently developed an allele-specific (AS) real-time polymerase chain reaction (PCR) assay for the detection of low-level K65R.16 The current study assesses the presence of low-level K65R among patients pretreated with ddI or ABC and its potential effect on subsequent treatment with TDF.
Clinical Study Description
Studies 902 and 907 were randomized, placebo-controlled, double-blind phase 2/3 studies of the safety and efficacy of TDF intensification therapy. All patients had >400 copies/mL of HIV-1 RNA and >8 weeks of prior stable background antiretroviral therapy to which TDF or placebo was added. For studies 902 and 907, the maximum baseline viral loads were 100,000 and 10,000 copies/mL, respectively. The primary endpoint for efficacy was the time-weighted average change in HIV RNA at week 24 (DAVG24) for both studies. The clinical and virologic results from studies 902 and 907 have been previously published.15,17-20 Population genotypic analyses were performed on plasma samples (at baseline and end of study) from patients with a viral load >400 copies/mL (Virco Laboratories, Mechelen, Belgium). During the course of these studies, 14 patients had developed a detectable K65R mutation through up to 96 weeks of TDF therapy.
Single Genome Sequencing
HIV-1-positive patient samples were diluted, if needed, to viral loads less than 10,000 copies/mL. The HIV-1 viral RNA was then isolated, and an RT reaction using custom primers was performed to produce a complementary DNA (cDNA) product. A cDNA dilution that gives approximately a 30% yield of PCR product is required; thus, varying dilutions of cDNA were amplified through 2 rounds of PCR using custom primers in sets of 11 and then qualified by agarose gel electrophoresis to find the correct dilution. According to Poisson distribution, a cDNA dilution yielding 3 of 10 positive PCR products (∼30%) contains 1 copy of cDNA per positive result about 80% of the time.21 After establishing the correct cDNA dilution, 95 reactions were amplified through 2 rounds of PCR to attain 20 to 30 (∼30%) clones. These clones were then purified, cycle sequenced using Applied Biosystems (Foster City, CA) Big Dye Terminator chemistry, sequenced using Applied Biosystems genetic analyzer technology, and analyzed as previously described.22
Allele-Specific Polymerase Chain Reaction
Viral RNA was extracted from 140 to 750 μL of patient plasma using the QIAamp viral RNA kit (Qiagen,Valencia, CA) and eluted in 50 μL of water. To generate RT-PCR product, 20 μL of viral RNA was first reverse transcribed at 42°C for 60 minutes using HIV-1 specific primer 5′-TATCTGGTTGTGCTTGAATGATTCCTAATGCAT and SuperScript II RT (Invitrogen, Carlsbad, CA) in a total volume of 50 μL. Amplification of 20 μL HIV-1 DNA was carried out using rTth DNA polymerase XL (Applied Biosystems) in a total volume of 100 μL with the following PCR conditions: 1 cycle of 95°C for 3 minutes; 35 cycles of 95°C for 15 seconds, 66°C for 40 seconds, and 72°C for 2 minutes; and 1 cycle at 72°C for 10 minutes. The forward and reverse primers for the amplification were as follows: 5′-GAAATGATGACAGCA TGTCAGGGAGT and 5′-TATCTGGTTGTGCTTGAATGATTCCTAATGCAT. The obtained RT-PCR products were diluted and tested for the presence of K65R using AS-PCR, as previously described.16 Briefly, PCR primers were used as follows: curative primer, 5′-GAAAATCCAT ACAATACTCCAGTATTTGCCATAAAGA; K65K primer, 5′-ACAGGTAGTATTTGCCATAAAGAA; K65R primer, 5′-GACATGAGTATTTGCCATAAAGAG; and reverse primer 5′-CCTGCIGGA TGTGGTATTCCTA, where “I” is a deoxyinosine. For each assay, PCR primers were used at the following concentrations: forward curative primer at 5 nM, AS primers at 100 nM, and reverse primer at 400 nM. The PCR conditions involved 25-μL reaction mixtures in 1 × ISOlution buffer (EraGen Biosciences, Madison, WI) and Titanium Taq DNA polymerase (Clontech, Palo Alto, CA) at the manufacturer's recommended concentration. PCR cycling steps were carried out with the rapid ramping rate of 20°C per second unless otherwise specified on the Roche LightCycler 1.2 (Roche, Indianapolis, IN) and were as follows: 2 minutes at 95°C; 3 cycles of 5 seconds at 95°C, 5 seconds at 61°C, and a ramp of 1°C per second to 10 seconds at 72°C; 1 cycle of 5 seconds at 95°C, 5 seconds at 45°C, and a ramp at 1°C per second to 20 seconds at 72°C; 80 cycles of 5 seconds at 95°C, 5 seconds at 55°C, and 20 seconds at 72°C (fluorescence read); and melt at 60°C to 95°C with a ramp at 0.4°C per second ramp (fluorescence step read). Fluorescence data were collected in 2 channels corresponding to the labels on AS primers for K65R and K65K. Fluorescence data were exported and analyzed with MultiCode-RTx analysis software (EraGen Biosciences). Delta threshold cycle (Ct) values were calculated as Ct of K65R amplification minus Ct of K65K amplification. Standard curve RNA samples generated from viral mixtures containing 0% to 50% of K65R site-directed HIV-1 RT mutant virus were processed in parallel with patient plasma samples. Delta Ct values of standard curve samples were used to quantify the presence of the K65R allele in each plasma sample (SigmaPlot/SigmaStat 9.01; SYSTAT Software, San Jose, CA).
All HIV-1 RNA statistical analyses were performed using SAS (version 8.1; SAS Institute, Cary, NC). P values <0.05 were considered significant. Multivariate linear regression analyses were performed to evaluate the impact of different mutations along with other baseline parameters on HIV-1 RNA response. A stepwise method was applied with a significance level of 0.15 for entry and staying in the model. P values were not adjusted for multiple comparisons.
A total of 437 patients were initially evaluated for the development of resistance and had baseline genotypic data from studies 902 and 907 combined. Each patient was exposed to TDF for a period of 24 to 96 weeks. During this follow-up period, 14 patients developed the K65R mutation by population sequencing. Within this group, there was a striking negative association with the presence of baseline thymidine analog mutations (TAMs [M41L, D67N, K70R, L210W, T215Y/F, and K219Q/E/N]), with none of the 311 patients with baseline TAMs developing K65R (Table 1). Among patients without baseline TAMs, patients with baseline L74V showed a positive correlation with the development of K65R (4 of 10 patients; P = 0.014), although there were few patients within this group.
Single-Genome Sequencing Analysis
We further investigated the 4 patients with baseline L74V who subsequently developed K65R by single-genome sequencing (SGS) analysis to determine whether there was any detectable K65R at baseline and whether it was expressed on the same genome as L74V. Baseline plasma samples were available for 3 of these 4 patients. SGS was performed for these 3 patients at baseline, with 91, 77, and 63 individual genomes analyzed for patients 4055, 2076, and 1278, respectively (Table 2). Of the 91 baseline HIV-1 RT genomes sequenced from patient 4055, all expressed L74V/I with no detectable K65R at baseline. By week 8, K65R was observed in 3.8% of RT genomes and then in 72% of RT genomes by week 16. Interestingly, a substantial fraction of these K65R genomes also included an L74I mutation that was only detected as a minor population at baseline (2%); all clones containing L74V were wild type at K65. For patients 2076 and 1278, however, low-level K65R was detected at baseline in 2.6% and 6.3% of the RT genomes analyzed, respectively. By week 4 of TDF intensification, these percentages had increased to 46% and 56%, respectively, with further increases at week 8. There was a concomitant decrease in the fraction with L74V in these patients, and none of the RT genomes analyzed expressed the K65R and L74V mutations together. At baseline, patient 1278 was taking ddI, EFV, saquinavir, and ritonavir, whereas patient 2076 was taking ABC, 3TC, and amprenavir. Their HIV RNA responses at week 24 were −0.13 and −0.22 log10, respectively.
Allele-Specific Real-Time PCR
Given the observations by SGS of low-level K65R associated with 2 patients treated with ddI or ABC, we developed a higher throughput assay for the detection of low-level K65R using AS primers and real-time PCR.16 Starting from RT-PCR products of plasma HIV-1, this AS-PCR assay demonstrates a lower limit of detection for K65R mutant genomes of 0.5% for subtype B HIV-1. Plasma samples from 63 treatment-naive HIV-1-infected patients showed no detectable K65R minority species in our assay with its cutoff of 0.5%. Plasma samples from 154 treatment-experienced patients from study 907 were also analyzed. Within this group, we analyzed all patients from study 907 with L74V/I by population sequencing (n = 44) and a random sample of patients without L74V/I who were taking ddI or ABC (n = 51) or not taking ddI or ABC (n = 51), including 10 of the 14 patients who had developed K65R (the other 4 patients had no remaining baseline samples). As positive controls, we also analyzed patients with K65R by population sequencing, all 8 of whom were taking ABC (n = 4), ddI (n = 3), or ABC and ddI (n = 1) on study entry.
As expected, all patients with K65R by population sequencing showed >50% K65R by AS-PCR (7 of 8 patients >90% K65R; Fig. 1). In addition, we detected low-level K65R by AS-PCR in 11 (7.5%) of 146 analyzed patients, including the 2 patients who showed low-level K65R by SGS. Among the 11 patients with low-level K65R, the percentage of minority species K65R ranged from 0.7% to 25% of viral sequences as quantified by AS-PCR. The patient with 25% K65R by AS-PCR did not show detectable K65R by population sequencing, suggesting that the limit of K65R detection by population sequencing is >25%. The remaining patients had K65R percentages of <11%, with 8 of 11 patients having <3%. Ten of these 11 patients with low-level K65R were taking ddI (n = 5), ABC (n = 3), or ddI and ABC (n = 2) on study entry. Overall, the presence of baseline K65R, at the population sequencing level or as a minority variant, correlated positively with the use of ABC or ddI (P = 0.004) and negatively with the presence of baseline TAMs (P = 0.003) (Table 3). Among patients with baseline L74V/I by population sequencing, there was a trend toward the detection of K65R by AS-PCR, with 6 of 44 samples analyzed showing low-level K65R (13.6% vs. 4.9% without L74V/I; P = 0.088).
Development of K65R
Among the 10 analyzed patients who had developed K65R by population sequencing, 3 had detectable K65R in their baseline sequence by AS-PCR. As described above, SGS analyses of 2 of these patients also showed low-level K65R at baseline and then subsequent expansion of K65R on addition of TDF treatment. The third patient with low-level K65R had the Q151M complex of multinucleoside resistance mutations at baseline (A62V, V75I, F116Y, and Q151M) and then developed full K65R by week 12. The remaining 7 patients who had developed K65R did not have detectable K65R at baseline by AS-PCR. Development of K65R in these patients likely reflects de novo K65R selection because of incomplete viral load suppression and TDF therapy in combination with other NRTIs.
Treatment Response to Tenofovir Disoproxil Fumarate
HIV RNA responses to intensification treatment with TDF are shown in Figure 2. Among TDF-treated patients with baseline genotypic data in these studies, the mean viral load reduction was −0.62 log10 copies/mL (DAVG24). Patients with baseline TAMs, L74V, or K65R by population sequencing showed diminished responses relative to the overall treatment response. The effects of baseline TAMs and K65R on the treatment response to TDF have been previously described in a subset of the patients analyzed in the current study.15 Patients with low-level K65R that was only detected by AS-PCR also showed a strongly reduced mean response to TDF treatment (−0.11 log10 copies/mL, n = 11). Although most patients with baseline L74V also had multiple TAMs, a few patients with baseline L74V in the absence of TAMs also showed a reduced treatment response to TDF of −0.31 log10 copies/mL.
We performed a multivariate statistical analysis using baseline genotypic, HIV disease, and demographic data as parameters for HIV RNA response at week 24. As shown in Table 4, treatment with TDF, baseline HIV-1 RNA levels, and baseline CD4 cell counts were all significant predictors of HIV RNA response. The 4 TAMs (M41L, D67N, L210W, and T215Y) predicted a reduced HIV RNA response in agreement with previous studies.15 The K65R mutation at baseline by population sequencing showed the strongest negative effect on treatment response, with a parameter estimate of +0.61 log10 copies/mL nearly ablating all the TDF treatment effect of −0.66 log10 copies/mL. Independent of K65R and TAMs, the L74V mutation also showed a negative effect on HIV RNA response, with a parameter estimate of +0.29 log10 copies/mL. Finally, as previously described, the M184V mutation showed a modest positive effect on treatment response of −0.13 log10 copies/mL.
New technologies are permitting the characterization of minority variants of HIV-1 within an infected individual.23-25 Some of these techniques, such as SGS or clonal analysis, are highly labor-intensive but reveal complete sequence information for an individual viral genome.21 Others, such as AS-PCR or ligation-amplification PCR,26 provide for higher throughput analysis of viral subpopulations but do not provide complete sequence information and must be developed individually for each mutation being assessed. We have developed an AS-PCR technique specifically for the K65R mutant of HIV-1 RT and assessed plasma samples from treatment-naive and treatment-experienced HIV-1-infected patients for the presence of low-level K65R. In no case have we observed >0.5% K65R among plasma samples from 63 treatment-naive patients. The cutoff of 0.5% was set based on the background signal observed with RT-PCR products of a clonal virus population containing only wild-type HIV-1, as previously described.16 A specific cutoff must be established for each mutant in AS-PCR, and this can vary, in our experience, from 0.05% to 0.5% for a given HIV-1 RT mutant. Similar mutant-specific cutoffs have been described by others with analogous AS-PCR systems.25,27,28 From these studies, it seems that interrogation of K65R is associated with a fairly high degree of nonspecific signal presumably attributable to the specific surrounding nucleotide sequence at this position. Thus, interpretation of a true positive signal must be carefully distinguished from background in these systems, and appropriate cutoffs must be determined. Of note, we observed high background with some subtype B HIV-1 patient sequences corresponding to AAA AAG at codons 64 and 65. Because this is the predominant sequence for subtype C viruses, our AS-PCR assay is not currently amenable to subtype C analysis. Subtype specificity has also been reported by others for PCR-based assays.23
Once detected, the clinical relevance of minority HIV-1 variants needs to be established. In the case of minority variants of nonnucleoside reverse transcriptase inhibitor (NNRTI)-resistant viruses, at least 3 studies have demonstrated that the presence of minority populations of resistant virus has a negative effect on subsequent treatment with an NNRTI-based regimen. In Adult Clinical Trials Group (ACTG) 398, initial efficacy observations showed that NNRTI-experienced patients who had no detectable NNRTI-associated resistance mutations by population sequencing demonstrated an inferior treatment response to a new EFV and protease inhibitor (PI)-based regimen as compared with those naive to NNRTIs.29 Similar observations have been made in multiple other clinical studies with NNRTIs. In ACTG 398, a subset of the NNRTI-experienced patients was analyzed by SGS, and 6 of 10 subjects showed evidence of NNRTI-resistant minority variants (3% to 30% of genomes).4 In a more recent study, AS-PCR was used to detect K103N, Y181C, or M184V among treatment-naive patients who failed a regimen of ABC, 3TC, and EFV.27 Among 70 virologic failures analyzed, 7 patients had 1 or more of these mutations only detectable by AS-PCR at baseline. A strong correlation was shown between the presence of these mutations as minority species at baseline and subsequent virologic failure (P = 0.005). Of note, 2 patients only had the M184V mutation detected at baseline, suggesting that even a minor population of this mutant may jeopardize the treatment response to a regimen of ABC, 3TC, and EFV. Finally, multiple studies of single-dose nevirapine administered intrapartum have demonstrated the development of nevirapine resistance in a substantial fraction of mothers and subsequent impaired treatment responses to nevirapine-based regimens.2,30-33
In the current study, low-level minority variants of K65R were observed in 11 (7.5%) of 146 treatment-experienced patients analyzed. There was a significantly inferior treatment response to TDF among these patients (−0.11 log10 copies/mL) as compared with the overall patient population. Among these patients, 3 went on to develop a full K65R mutation on addition of TDF therapy. Among the other patients not developing a full K65R mutation, there was limited follow-up and/or addition of other antiretrovirals (eg, lopinavir) that precluded further longitudinal analysis of resistance development. Overall, combining patients with K65R by population sequencing and those with K65R by AS-PCR, there was a positive correlation of baseline K65R with concurrent ABC or ddI use (P = 0.004) and a negative correlation with TAMs (P = 0.003). These results are consistent with the established resistance profiles of ABC and ddI and with multiple publications showing negative interactions between K65R and TAMs.34-37 Of note, we did observe 1 patient who had K65R and multiple TAMs on the same genome (M41L, D67N, L210W, and T215Y). Thus, this negative correlation is not absolute, as has been shown by others.38
The vast majority of patients with detectable K65R in our study (18 of 19 patients) by population sequencing or AS-PCR had prior exposure to ddI or ABC. Recent observations have demonstrated that d4T can also select for K65R in vitro and in vivo, however.39,40 The single patient with K65R in our study who was not taking ddI or ABC was taking d4T and 3TC as NRTIs. Although d4T typically selects for TAM-associated mutations, selection of K65R may be favored under certain NRTI combinations (eg, d4T, 3TC, nevirapine) and may have clinical consequences for continued or new use of NRTIs such as TDF.
Interestingly, 6 of the 11 patients with low-level K65R had L74V/I at baseline, and the presence of L74V in the absence of TAMs was significantly associated with the development of K65R. Thus, the relation between K65R and L74V/I is of interest. Both ABC and ddI can select for either mutation, and, in general, these mutations are seen on different viral genomes. Within an individual, however, both mutants may exist as competing mutant populations and L74V is generally the dominant population. The results of our SGS analysis of 3 patients are in agreement with the earlier clonal analyses and demonstrated the typical genomic exclusivity of K65R and L74V. In patient 4055, however, we did observe K65R and L74I on the same viral genome, whereas L74V was always observed with wild-type K65. A recent publication has also demonstrated the occurrence of K65R and L74V/I on the same genome.38 These observations of K65R and L74V/I together on the same genome seem to be rare, however, and may reflect the presence of additional mutations potentially compensating for the demonstrated reduced replication capacity of this double mutant.12
Previous multivariate analyses of treatment response to TDF have indicated that the L74V mutation was associated with a reduced treatment response to TDF.14,15 These results were initially puzzling, because L74V itself has no demonstrable phenotypic effect on TDF susceptibility in vitro.13 In the current analyses with a larger data set of TDF-treated patients, we confirm that L74V is a predictor of reduced treatment response. In most patients, L74V is associated with multiple TAMs, and those patients have a strong reduction in treatment response that seems to be primarily attributable to the effects of the TAMs. Among those patients without TAMs, however, there is still an apparent decrease in TDF treatment response (−0.31 log10 copies/mL). Our current data suggest that the presence of low-level K65R among some of these patients may be partially responsible for the reduced treatment response to TDF in these patients. Minority variants of other NRTI mutations (eg, M184V, TAMs) may also have an impact on subsequent therapies. Of note, the treatment responses observed in this study are limited by the intensification design of the clinical study. Minority variants of K65R and other NRTI mutations need to be further assessed for clinical relevance under conditions of optimized new therapies and in less advanced patient populations to determine their clinical relevance in these settings. Overall, however, our studies describe the potential for further expansion of minority K65R variants that were preexisting because of prior ABC or ddI therapy and their negative impact on treatment responses to TDF.
The authors thank the study personnel, investigators, and patients from studies 902 and 907. They also thank Hans Reiser for review of this manuscript and Margaret Benton for administrative assistance and document preparation.
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This article has been cited 12 time(s).
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allele-specific polymerase chain reaction; K65R; nucleoside reverse transcriptase inhibitor resistance; quasispecies; tenofovir; tenofovir disoproxil fumarate
© 2007 Lippincott Williams & Wilkins, Inc.
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