HM Medical Clinic

 

Treatment-interruption-v7-submitted

Evolution of Drug-resistant Viral Populations during Interruption of
Antiretroviral Therapy
Running title: Evolution and fitness of drug-resistant viruses Dongning Wang1, Charles B. Hicks2, Neela Goswami2, Emi Tafoya3, Ruy M. Ribeiro3, Fangping Cai1, Alan S. Perelson3 and Feng Gao1* 1Duke Human Vaccine Institute and 2Department of Medicine, Duke University Medical Center, Durham, NC 27710, USA 3Theoretical Biology and Biophysics, Los Alamos National Laboratory, Los Alamos, NM 87545, USA. *Corresponding author 3072B MSRB II, DUMC 103020 Duke Human Vaccine Institute Duke University Medical Center Durham, NC 27710, USA Tel: 1-919-668-6433 Fax: 1-919-681-8992 E-mail: [email protected] Abstract
Analysis of a large number of HIV-1 genomes at multiple time points after antiretroviral treatment (ART) interruption allows determination of the evolution of drug-resistant viruses and viral fitness in vivo in the absence of drug selection pressure. Using a parallel allele-specific sequencing (PASS) assay, potential primary drug-resistant mutations were studied according to drugs used in five individual patients by analyzing over 18,000 viral genomes. A three-phase evolution of drug-resistant viruses was observed after termination of ART. In the first phase, viruses carrying various combinations of multiple drug-resistant (MDR) mutations predominated with each mutation persisting in relatively stable proportions while the overall number of resistant viruses gradually increased. In the second phase, viruses with linked MDR mutations rapidly became undetectable and single-drug-resistant (SDR) viruses emerged as minority populations while wild-type viruses quickly predominated. In the third phase, low- frequency SDR viruses remained detectable as long as 59 months after treatment interruption. Mathematical modeling showed that the loss in relative fitness increased with the number of mutations in each viral genome, and that viruses with MDR mutations had lower fitness than viruses with SDR mutations. No single viral genome had seven or more drug-resistant mutations, suggesting that such severely mutated viruses were too unfit to be detected or that the resistance gain offered by the seventh mutation did not outweigh its contribution to the overall fitness loss of the virus. These data improve understanding of evolution and fitness of drug-resistant viruses in vivo and may suggest strategies for improved management of ART-experienced patients. Despite significant advances in antiretroviral therapy (ART), some HIV- infected patients still fail treatment due to drug resistance, poor adherence, drug toxicity, and suboptimal drug metabolism (4, 35). Among these causes, emergence of drug-resistant mutations plays a central role in ART failure (14, 35). In addition, the presence of pre-existing drug-resistant viruses correlates with poor responses to ART (18, 28, 36-39). Multiple drug-resistant (MDR) mutations have been shown to exist in various complex linkage patterns in viral genomes (3, 26, 34) and these MDR viruses can play an important role in ART failure. Discontinuation of ART in patients who have developed drug resistance leads to disappearance of drug-resistant mutations in the viral population within weeks and replacement with wild-type (WT) viruses (7-9, 20, 40). However, when a more sensitive allele-specific PCR (ASPCR) method was used, minority drug-resistant viruses (0.01% - 20%) could be detected in most patients for months or even years following ART discontinuation (12, 33). The evolution of both majority and minority linked MDR virus populations after treatment interruption, however, has only been characterized previously in a limited number of viral genomes in HIV-1-infected persons. Fitness of drug-resistant viruses has been extensively studied in vivo and in vitro (6, 10, 11, 13, 15, 23-25, 27). An improved understanding of viral fitness among various drug-resistant viruses may have implications for development of better treatment strategies and may be a factor in antiretroviral drug discovery. However, only a limited set of mutations and small number of viral genomes have been studied previously in HIV-1-infected individuals. A more thorough evaluation of drug-resistant mutations in the absence of drug selection pressure by analyzing large viral populations in patients who discontinued failing ART provides an opportunity to investigate evolution, fitness and reservoirs of the drug-resistant viruses, especially those with linked MDR mutations. We recently developed a highly sensitive parallel allele-specific sequencing (PASS) assay that can detect minority drug-resistance populations present at frequencies as low as 0.01% to 0.1% of the viral population by simultaneously analyzing thousands of viral genomes in a single assay (3). In addition, this new technology allows identification of linkages among MDR mutations on individual viral genomes. In this study, we characterize the evolution of MDR mutations and their linkage relationships among a large viral population from longitudinal samples collected from individual patients following treatment interruption. These data are then used to model the evolution and fitness of various drug-resistant viruses, and to estimate the time to emergence of the WT virus following treatment interruption. Materials and Methods
Patient plasma samples. The study population consisted of HIV-1-
infected persons followed at the Duke University Medical Center between 2001 and 2003. Residual plasma remaining after clinical viral load testing were stored at -80o C. Patients selected for this study were identified from the HIV Patients Sample Repository database after reviewing characteristics of their treatment history. To be included in the study, patients had to have been followed in the Duke HIV/AIDS Clinic for at least one year, during which time they underwent ART interruption and thus had both on-treatment and off-treatment samples available at different time points to be studied. Based on patient responses to previous ART regimens and/or results from population-based genotypic drug- resistance testing, samples were selected from patients experiencing treatment failure with viral load rebound. Patients were excluded if fewer than two blood samples after treatment interruption were available for analysis. Written informed consent was obtained from all the individuals whose blood samples were collected. The study was approved by the Duke University Institutional Review Viral RNA extraction and cDNA synthesis. One milliliter of each sample
was then concentrated by ultra-centrifugation at 32,000 rpm for 3 hours at 4o C. The virus pellet was then resuspended with 400 µl of PBS and the viral RNA (vRNA) was extracted using PureLink viral RNA/DNA mini kit (Invitrogen, Carlsbad, CA). The vRNA was eluted into 17 µl of the elution buffer and used for cDNA synthesis using Superscript III (Invitrogen, Carlsbad, CA) and primer RTuni1 5'-CCAATCCCCCCTTTTCTTTTAAAATTGTG-3' in a 40 µl reaction. An appropriate amount of cDNA was used for the PASS assay to obtain an optimal number of viral genomes in each assay. Detection of drug-resistant mutations by PASS. The PASS assay was
performed as previously described (3). Briefly, 20 µl of 6% acrylamide gel mix, containing 1 µM acrydite-modified reverse primer, 5'Acr- AATCCCTGCATAAATCTGACTTGCCCAAT-3', cDNA template (5 ul to 18.5 ul), 0.3% diallyltartramide, 5% rhinohide, 0.1% APS, 0.1% TEMED and 0.2% BSA, was used to cast a gel on a bind-saline (Amersham Biosciences, Piscataway, NJ) treated glass slide. The in-gel PCR amplification was then performed in a PTC-200 Thermal Cycler with a mix of 1 µM forward primer, 5'- TTAGCTTCCCTCAGATCACTCTTTGGCA-3', 0.1% Tween-20, 0.2% BSA, 1x PCR buffer, 250uM dNTP mix, 3.3 units of Jumpstart Taq DNA polymerase (Sigma, St. Louis, MO), and H2O (up to 200ul) under a sealed SecurSeal chamber (Grace Bio-Labs, Inc., Bend, OR). The PCR condition was: 94 o C for After PCR amplification, single-base extension (SBE) was performed with mutant and wild-type (WT) bases distinctively labeled with Cy3 and Cy5, respectively, using the primers that annealed just upstream of the mutation sites of interest. To detect multiple drug-resistant mutations on the same viral genome for linkage analysis, the amplified viral genomes (polonies) in each gel were then sequentially interrogated by 6-12 SBE reactions for targeted drug-resistant mutations. Mutation sites were selected for analysis based on the treatment history of the patients and genotypic test results when they were available. Between 6 and 12 drug-resistant mutation sites were assayed for the amplified viral genomes in each sample by repeating SBEs. After each SBE, the gel was scanned with a GenePix 4000B Microarray Scanner (Molecular Devices, Sunnyvale, CA) to acquire images. PASS data analysis. The two channel images (Cy5 for WT and Cy3 for
mutant) acquired from each PASS assay were first cropped with Picture Window Pro3.5 to remove the edge area containing no signal. The cropped images were then analyzed with the Progenesis PG200 (Nonlinear Dynamics, Durham, NC) software. After background subtraction, normalization and spot filter setting, only the unambiguous spots at either channel were included for further analysis. The normalized pixel count data at multiple mutation sites for each spot were exported into an Excel file with a unique number. By comparing each spot's normalized values at both channels, the position was classified as WT or mutant. Finally, the linkage pattern of all mutations on each viral genome was determined by compiling mutation information at all analyzed sites with the Linksys program Frequency of mutants in the population. At each sampling time a
number of sequences, N, are analyzed. If as in Fig.1, we find 9 out of 9 sequences all have the M184V mutation, the frequency of M184V mutants in the sample is 100%, but other sequences not containing M184V may be present in the population and just not sampled. For example, if the frequency of non-M184V sequences is 1/(N+1), i.e. 10% in this case, then the probability of observing 9 out of 9 sequences with M184V, computed from the binomial distribution, Binom(9,9,p=0.9) is 0.39. One can also use the binomial distribution to determine with what frequency non-M184V sequences would need to be present in order to obtain with 95% probability no non-M184V mutants in a sample of size N. As discussed in more detail in reference (21), this is the solution of Binom(0,N,p) = 0.95. For N=9, if non-M184V sequences were present at p=0.5% or less one would expect 9 out of 9 sequences to be M184V with >95% Viral fitness comparison. A conventional method (11, 15, 41) of
analyzing the difference in growth rates (or fitness) between mutant and WT was used to evaluate the relative fitness between strains. In this method, one assumes the wild type (W) and mutant (M) virus grow according to Here the number of cells infected by WT and mutant virus was assumed to be proportional to the amount of wild type and mutant virus, respectively (i.e., the virus and infected cells were assumed to be in quasi-steady state (31)). Cells infected with WT virus were assumed to grow at rate r, and to be cleared at rate δ. Cells infected with mutant virus were assumed to grow at rate r' and to be lost at rate δ'. The solution for these equations, which gives the evolution of the viral levels for each strain, between times t1 and t2 is The ratio of mutant to WT virus at the two times was used to calculate d, the difference in the net growth rates, where d=(r'-δ')- (r-δ). This difference, d, has been called the log-relative fitness (41), and is computed here by the formula where f(t)= M(t)/W(t) and ln denotes the natural logarithm. For simplicity and to be consistent with much of the virology literature, d will simply be called the relative fitness rather than the more precise log relative fitness. The relative fitness can also be related to the selection coefficient, s, by the formula d=ln (1+s) (41). Note that when s<<1, d is approximately equal to s and this approximation has been used to calculate s (11). Equation (1) is written such that we are assessing the fitness of the mutant relative to the WT. Since our measurements were made after interruption of therapy, we expect the drug- resistant mutants to be less fit than WT and d will be negative, and its value can be viewed as the fitness loss of the mutant relative to the WT. For each patient, the relative fitness was calculated over the time frame where both WT and mutant virus were observed. For this time frame, we write ln(f(t2)/f(t1))=d Δt which is the equation of a straight line with Δt=t2-t1 as the independent variable. Using linear regression one can then estimate d (41). Note that if the fitness of the mutant or WT changes over time, Eq. (2) can be interpreted as the mean relative fitness of the mutant averaged over the time interval, i.e., We analyzed the data by first grouping all sequences that had the same number of mutations to calculate the relative fitness of a "strain" with i mutations, i =1, 2, 3… versus the WT. Since we had the most data for strains with one or two mutations in PID811, we also computed the relative fitness of these strains individually for this patient. We calculated confidence intervals for the relative fitness estimates using a bootstrap method. In each case, we generated 1000 new datasets, with the same number of total genomes as the data and with a distribution of WT and mutants as given by the data. That is, we used a multinomial distribution with the probabilities given by the data to calculate for each bootstrap replicate the new number of WT, 1-point, 2-point, etc. mutant genomes. In those few cases where the number of genomes for a given strain in the bootstrap dataset was chosen as zero, we imputed 1 genome (note that Eq. 2 cannot be applied when one of the frequencies is zero). Implicitly this is the same as assuming that the lowest frequency of detection is 1/N (where N is the total number of genomes analyzed at a given time point) and that the next genome sampled would be of that strain (1). We then used Eq. 2, as before, to calculate the relative fitness in these bootstrap datasets. We picked the 2.5-percentile and the 97.5-percentile of these estimates as the limits of the 95% confidence interval. Estimation of the time for WT virus appearance after treatment
interruption. The time at which the wild type strain emerged was estimated by a
method introduced by Asquith and Mclean (1) in the context of determining the time that CTL escape mutants arise. The fraction of the wild type strain, p(t), is where g = M(0)/W(0), and d is the difference of viral growth rates, as before. Equation 3 can be transformed to a linear equation as where G=ln (g). To estimate the parameters d and G, we used the data after treatment interruption in Supplementary Table 1. From this data we computed the fraction of WT virus, p(t), at various times and then used linear regression (Eq. 4) to estimate d and G. We then define the time when the WT strain emerges as the time its frequency reaches 1% of the population. We can determine this time by choosing p(t)=0.01 and calculating t in Eq. (4), using the best-fit parameters d and G from the linear regression described above. Clinical characteristics of the study subjects
Five patients who interrupted antiretroviral therapy, either per physician recommendation or by self-choice, were selected for analysis (Table 1). All were followed regularly in the HIV/AIDS clinic and had periodic viral load testing during the duration of their treatment interruption. In all cases, plasma viral loads quickly increased following treatment interruption. Multiple off-therapy samples and one on-therapy sample (0-61 weeks before treatment interruption) were available for assessment in each individual. All patients had received therapy with three or more antiretroviral agents over a period of 37 to >260 weeks prior to treatment interruption. Drug-resistant mutations were identified in a clinical population- based HIV genotyping resistance test in two patients (PID811 and PID908) either while still on a failing ART regimen or just after treatment interruption commenced, confirming the presence of drug-resistant mutations in both individuals. Clinical data including prior ART regimens, duration of ART, dates of samples collection, viral load test results, and the results from clinical HIV resistance testing are summarized in Table 1. Dynamic population changes of individual drug-resistant mutations
Viral RNA was extracted from longitudinal samples in each patient and drug-resistant mutations were assayed by PASS. A total of 18,451 viral genomes (an average of 802 per time point) were analyzed. In general, samples collected during non-suppressive ART that only partially suppressed viral replication had a low viral load and only few or no viral genomes were detected in four patients (PID811, PID908, PID268 and PID295). In each of two cases (PID895 and PID295) with low viral loads (1,111 copies/mL and 213 copies/mL, respectively), only one viral genome was detected. These samples were not included in the Six to twelve mutation sites in each patient were selected for analysis based on potential resistance patterns that may have been selected by the particular agents being used during the time of treatment failure. When available, clinical genotypic resistance test results were also used to identify mutations of interest. Four to eight drug-resistant mutations were detected in four of the five patients; no drug-resistant mutations were detected in the fifth patient PID295 (Table 1). Drug-resistant mutations were first analyzed individually for each patient. Patient PID811 was treated with zidovudine (AZT), lamivudine (3TC) and abacavir (ABC). The patient had also received nevirapine (NVP) in an earlier regimen. The viral load was 2,104 copies/mL in an on-therapy sample collected 10 weeks before treatment interruption, and it continuously increased to 178,017 copies/mL at week 32 after ART interruption. Eight primary and one secondary drug-resistant mutations were detected by a clinical genotype resistance test in a sample obtained three weeks prior to treatment interruption (Table 1). The PASS assay was completed on one on-therapy sample and five off-therapy samples collected within 32 weeks after treatment interruption. On average, 304 viral genomes (range 9 to 822) were analyzed at each time point. Based on the treatment history and available genotypic resistance tests, eleven primary drug- resistant mutations to reverse transcriptase inhibitors (RTIs) and one to protease inhibitor (PI) were assessed. Of these, eight (M41L, D67N, K65R, K70R, K103N, M184V, L210W and K219Q/E) were detected and four (L74V, Y115F, T215Y/F and I84V) were not (Table1). K103N (RTI mutation) and the I84V (PI mutation) were included for analysis based on the genotypic test result and the previous treatment history, respectively. While the patient was still on therapy, the viral load was low and only nine viral genomes were analyzed. At this time, all viruses had the M184V mutation (Fig. 1). The proportion of observed viruses with the M184V mutation was relatively stable (89% -100%) for the first nine weeks following treatment interruption, and then dramatically declined over the next 23 weeks during which time WT viruses predominated (Fig. 1 and Fig. 2A). After week 24, the M184V mutation became undetectable. Similar results were observed for the other seven identified mutations, although they were present in varying proportions in the viral population. The exception to this pattern was the K103N mutation, which initially decreased to 5% at week 24 but then became a higher proportion (26%) at week 32 (Fig. 2A). The K013N substitution can be caused by either AAC or AAT allele. Since the AAC allele is about 100 fold higher than the AAT allele in patients who have developed the K103N mutation (32), we only analyzed the AAC allele. Thus, the total percentage of the K103N mutation might be higher in this patient. The K103N mutation has been detected in patients months after treatment termination and the percentage in the viral population generally decreased over the time (17, 33). The cause of the sudden increase of the K103N virus at one time point in PID811 was unclear. Fewer time point samples from three other patients were available for analysis. However, evolution of drug-resistant mutations in these patients was very similar to what was observed in PID811. Two samples in the early weeks after treatment interruption were analyzed in PID908 (Table 1 and Fig. 2C). Three mutations in the reverse transcriptase gene (M184V, D67N and K70R) and four in the protease gene (M46I, G48V, V82A, I84V) were detected. M184V was present in all detected viral genomes while other mutations accounted for various fractions of the viral population when the patient was on therapy. Similar to the pattern seen in PID811, the proportions of the various mutations either persisted at the same level (M184V) or modestly decreased (others) at week 6 but then all declined substantially by week 11 when WT virus began to emerge (Fig. 2C). In PID268, the first sample was collected on the day the patient discontinued ART. Fifteen viral genomes were analyzed at this time point and all had the RTI-resistance mutation M184V and the PI-resistance mutation V82A (Fig. 2E). Interestingly, neither the M184V mutation nor the V82A mutation was detected in other three samples obtained later during the course of treatment interruption (34-52 weeks). Instead, three RTI-resistance mutations (D67N, K70R and K219QE) were detected at week 32 and, as noted in PID811, they decreased to very low levels by week 52. Samples were only available 61 or 39 weeks before treatment interruption in PID895 and PID295, respectively. Both samples had low viral loads (1,111 copies/mL in PID895 and 213 copies/mL in PID295), suggesting that viral replication was well controlled when ART was stopped. Four drug-resistant mutations were detected in PID895 and all were present at very low frequencies (<0.6%) even at nine weeks after treatment interruption. The sum of all detected resistant viruses accounted for less than 1% of the total viral population at any time point during treatment interruption (Fig. 2G). No mutations were detected at any time points (between week 13 and week 58) after treatment interruption in PID295 (Table 1). Of note, three RTI-resistance mutations (D67N, K70R, M184V) and one PI-resistance mutation (V82A) were consistently detected in other patients (Table 1 and Fig. 2) but not in this patient. The infrequent detection of minority resistant mutations in these two patients suggests that selection of drug-resistant mutations was limited at the time treatment was discontinued. Linkage analysis of MDR mutations.
The ability of the PASS assay to detect different mutations present in each viral genome allows determination of the linkage patterns of MDR mutations among a large population of individual viral genomes. Viral genomes with linked MDR mutations were identified in three individuals (PID811, PID908 and PID268), all of whom had higher levels of viral replication at or shortly before treatment interruption. In all three patients, nearly all detected on-therapy viruses carried two or more drug-resistant mutations (Tables 2-4). Linkage analysis of eight drug-resistant mutations in PID811 revealed 35 different linkage patterns (Table 2). During the first 15 weeks following treatment interruption, viruses with as many as 6 linked MDR mutations were identified but at very low frequencies, while the majority of detected viruses had 2, 3 or 4 linked MDR mutations. Nearly all these MDR viruses became undetectable by weeks 24 and 32. In contrast, single drug-resistant (SDR) viruses were rare in the first 9 weeks after treatment interruption (only M184V mutants were consistently detected). After week 15, nearly all drug-resistant viruses had only a single mutation, almost always the K103N or M41L mutation. Similar results were observed in PID908. A total of 28 linkage patterns were identified, and the MDR viruses predominated in the viral population from on-therapy to week 6 after interruption (Table 3). These viruses then rapidly decreased in frequency and were replaced by WT viruses and minority SDR viral populations (<2%) by week 11. Interestingly, viruses with only the M184V mutation accounted for 39% of the viral population at week 6 and increased further to 57% of the viral population by week 11. In the third patient PID268, 15 viral genomes were analyzed at the time of treatment discontinuation, and all had both M184V and V82A mutations (Table 4). However, neither M184V nor V82A were detected in subsequent samples (weeks 34, 41 and 52), perhaps because no samples were available during the early time period following ART interruption. Instead, low frequencies of linked MDR mutations (D67N/K219QE and K70R/K219QE) were detected at week 34, but all viruses had only SDR mutations which continuously decreased over the Most linked MDR mutation patterns were present as minority populations (<2%), but a few linkage patterns were more common. For example, four linkage patterns (M184V/K103N/K70R/K219QE, M184V/K103N/M41L, M184V/K103N and M184V/K219QE) in PID811 accounted for more than 25% of the population at week 3 and week 9 (Table 2). Similarly, three linkage patterns (M184V/V82A/G48V/I84V, M184V/V82A/G48V and M184V/V82A) accounted for much higher proportions of viruses than others in PID908 (Table 3). These results indicate that some drug resistant viruses with particular linked MDR mutation patterns might have a significant fitness advantage. In all three patients, viruses with 7 or more mutations were not found at any time points, suggesting such viruses are too unfit to survive, i.e., that the resistance gain offered by the seventh mutation did not outweigh its contribution to the overall fitness loss of the virus, or do not exist in sufficient quantity for detection. In the three patients who carried linked MDR viruses, those with combinations of four or five linked mutations were noted to gradually decrease over time after therapy interruption, but viruses with one or two mutations transiently increased during the first 6-10 weeks and then decreased thereafter (Fig. 3). Viruses with five or six mutations occurred too infrequently for conclusions to be drawn. However, some linkage patterns, for example, M184V/K103N/K70R/K219QE and M184V/K103N/M41L in PID811 continued to increase in the viral population during the first 9 weeks after ART interruption and persisted at relatively high frequency (19% and 4%, respectively) at week 15 (Fig. 3D and Table 2). Interestingly, predominant viruses with linked MDR mutations shared the same mutations (M184V, K103N and M41L in patient PID811; M184V and V82A in both patients PID908 and PID268 who were treated with RTIs and PIs). The mutations present in the dominant viruses with linked MDR mutations were generally those present at higher frequencies as SDR mutations (Tables 2-3). These results suggest that some drug-resistant viruses with different combinations of MDR mutations existed in higher frequencies, probably because they could be easily selected under these particular ART regimens and had greater fitness. Three-phase evolution of drug-resistant mutations during treatment
Three patients with higher viral loads (PID811, PID908 and PID268) had samples available at the time of or shortly before treatment interruption. When the combined data from these three patients were analyzed, a three-phase evolution of drug-resistant mutations was observed following treatment interruption. In the first phase (the first 9 weeks after treatment interruption), the proportions of drug-resistant viruses present were stable or slightly decreased (Fig. 2A and 2C; Tables 2 and 3) while the absolute number of viruses with resistant mutations increased (Fig. 2B and 2D). The predominant viruses have MDR mutations. Viruses with only SDR mutations were uncommon. In this phase, WT viruses were very rare or undetectable. In the second phase (9-32 weeks after treatment interruption), the proportion and number of drug-resistant viruses rapidly decreased and they were quickly replaced by WT viruses (Fig. 2A-2D; Tables 2 and 3). In this phase, viruses with MDR drug-resistant mutations decreased markedly while viruses with minority SDR mutations became the major drug-resistant population. In the third phase (32-34 weeks after treatment interruption), drug-resistant viruses became very rare in the viral populations (<1%) and only viruses with SDR mutations were detectable (Fig. 2A and 2E). Fitness of drug-resistant viruses
To determine the fitness of various groups of drug-resistant viruses, we first classified them into seven groups based on the number (0 through 6) of mutations each virus carried (Supplementary Table 1). Each mutation was counted as one, so viruses in each group might contain different combinations of mutations. For each patient, t1 was chosen as the first time for which data was available for both WT and mutant strains, and t2 chosen as a subsequent time. Then, we estimated the relative fitness (d) by fitting Equation 2 to the data. Viral genomes with 5-6 mutations were not included for analysis since there were too few of them for comparison. The estimated relative fitness (d) of viral strains with different numbers of mutations was computed, together with the respective 95% confidence intervals (Figure 4). In patients PID811 and PID908, from whom viruses with MDR mutations were detected at multiple time points, we can compare the cost of increased number of mutations in relation to the WT virus (Figure 4). The loss in relative fitness increased linearly in PID811 (p=0.00016), whereas in PID908 the three point mutant did not lose as much fitness as predicted by a linear loss (i.e., a positive quadratic term for fitness loss was significant, p=0.02) (Figure 4). Various combinations of drug-resistant mutations at different time points were observed in PID811 (Table 6). This allowed for calculation of the relative fitness by taking into account the different combinations of mutations. Equation 1 was applied to determine the relative fitness based on which drug-resistant mutations were present in the viral genome. Among 41 strains of drug-resistant viruses detected in PID811, the majority of them carried MDR mutations (2-6) while only six had SDR mutations (Table 2). Over time, the frequency of the fittest strains increased (Table 6). For instance, from day 63 to 105, the M184V virus was present with low fitness (d=-0.104) relative to the WT; however, SDR mutants with M41L or K103N emerged between days 105 and 168, and they were more fit (d=-0.016 or -0.012) than the M184V mutant, which was lost in this time window. This also was seen for the double mutant population, where from days 105 to168, all double mutants except K103N/M41L were lost, and the K103N/M41L double mutant was at least six times more fit than any other double mutants in the previous time period. Estimation of timing for WT virus emergence during treatment interruption
WT viruses gradually increased and became dominant after treatment interruption in patients PID811, PID908 and PID268 (Tables 2-4). It would be informative to estimate the time at which the WT viruses emerged in each patient. Using the values of d and G in Eq. 4, the time for WT viruses to emerge after treatment was estimated. Here we define the time of emergence as the time the WT virus reaches 1% of the population, when it was undetectable before. The time for WT virus to reach 1% of the population was 56 days in PID811 (based on data collected until week 24), 50 days in PID908, and 44 days in PID268. On average, WT viruses emerged 50 days after treatment interruption. However, these times depend on our definition of "emergence". If instead of a frequency of 1%, we define it as the time it takes the WT to reach 0.1%, then the estimated times are 23 and 35 for PID811 and PID908, respectively. For PID268, we cannot calculate the time it takes WT to reach 0.1%, because the estimated proportion of WT at time 0 is already 0.3%. Note that for PID268, we have no data between weeks 0 and 34, i.e. day 238, which leads to high uncertainty in frequency estimates early after therapy interruption. Discussion
Interrupting antiretroviral therapy in HIV-infected persons whose treatment regimens are not achieving viral suppression provides an opportunity to study the evolution of drug-resistant mutations and viral fitness in vivo during the period after drug selection pressure is removed. In this study, WT, SDR and linked MDR mutations were analyzed from over 18,000 viral genomes at multiple time points in five treatment-interrupted patients using the PASS assay. These data demonstrated a three-phase evolution of drug-resistant mutations and complicated patterns of linked MDR mutations. Mathematical modeling showed that the relative loss of viral fitness increased with the number of mutations in the viral genome, and that WT viruses reached an estimated frequency of 1% after 1-2 months post treatment interruption. Understanding of the evolution of viral populations following treatment interruption has been hindered by inability to characterize a large number of viruses and the limited availability of samples from appropriate time points. Analysis of thousands of viral genomes by PASS from three patients at multiple time points (up to 59 weeks) following treatment discontinuation demonstrated a three-phase evolution pattern of drug-resistant mutations. In the first phase, HIV viral load quickly rebounded shortly after ART interruption. These early rebound viruses were almost exclusively those with linked MDR mutations, although the proportion of drug-resistant viruses within the population remained stable or decreased slightly. WT viruses were generally undetectable at this time, although our model predicted they started to accumulate. In the second phase, the proportions and the numbers of viruses with MDR mutations rapidly decreased while viruses with SDR mutations gradually increased. In this phase, WT viruses quickly increased and became predominant. In the third phase, the vast majority of viruses are wild type and the only detected drug-resistant viruses were those with SDR mutations present at very low frequencies (<1%). This phase was observed to last more than a year. Our data indicated that the relatively stable proportion of drug-resistant mutations observed in the first phase was maintained by a continuous increase of drug-resistant viruses in blood following ART interruption, while the rapid disappearance of these drug-resistant viruses was caused by the increase of more fit WT viruses and possible decreased release of drug-resistant viruses into the blood in the second phase. Similar evolution patterns of drug-resistant viruses have been reported previously (7, 8). In these studies, the drug susceptibility of the viral populations was determined by a phenotypic resistance assay after ART discontinuation. Viruses in these patients were fully resistant to ART over the first 15 weeks following ART discontinuation, but they were quickly replaced by drug susceptible viruses over a very short period and remained drug- susceptible thereafter. However, drug-resistant mutations were not determined in Although viruses with linked MDR mutations are less fit than the WT virus, they continuously replicate early on when drug-selection pressure has been removed since initially they are the only detectable viruses. This results in the observed early increase of the number of drug-resistant viruses and the persistence of the proportion of drug-resistant viruses in the first stage. However, due to the impaired fitness of these viruses in the absence of drug selection pressure, they are subsequently outcompeted by WT viruses, which become the dominant population beginning about 10 weeks after treatment interruption. The drug selection pressure declines relatively quickly after ART interruption since the half-lives of antiretroviral drugs are only a few days even for long lasting NNRTIs (2, 5, 19, 29). Therefore, the presence of mostly drug-resistant viruses during the first nine weeks after treatment interruption cannot be due to the continued suppression of WT viruses by residual antiretroviral effect in these patients, but rather may reflect the fitness costs of resistance, as calculated here, and the low frequency of WT when treatment is interrupted. An area of considerable potential importance is the effect of linked MDR mutations present in a single viral genome as opposed to various drug-resistant mutations being present as single mutations in different virions within a patient. This phenomenon has not been well studied to date due to the difficulty of assessing a large number of viral genomes within any one HIV-1 infected individual (26, 34). Using the recently developed novel PASS assay (3), we were able to perform linkage analysis of drug-resistant mutations from thousands of viral genomes in each patient. Many different genetic variants with various combinations of linked MDR mutations were identified in patients in whom MDR mutations were detected. Viruses with 5-6 mutations were detected but they were present at much lower frequencies, suggesting that they had impaired fitness. In contrast, viruses with 2-4 mutations were the majority viruses at the time of ART failure and during the early stage of treatment interruption. Population genetic analysis also indicated that a few virus populations with specific combinations of some drug-resistant mutations were more predominant than others. The precise reason why some mutations predominate with certain combinations is not clear, but likely is the result of pathways of improved viral fitness and the interplay between mutations best suited for resistance to particular drugs that are being administered or have been previously used. Although hundreds of viral genomes at each time point were analyzed by PASS during treatment failure and treatment interruption, no viral genomes were found to have 7 or more drug-resistant mutations. There may be two possibilities for this. Firstly, the resistance gain offered by the 7 or more mutations did not outweigh its contribution to the overall fitness loss of the virus. The fitness cost for such viruses was too high for them to outcompete viruses with fewer mutations to be detected when the drug selection pressure was not present. Secondly, they may not be developed because viruses didn't need to have as many mutations in their genomes to be fully resistant to the combinations of Analysis of a large number of virus genomes after removal of drug selection allows us to evaluate virus fitness in vivo. Our mathematical model showed a positive correlation between the number of mutations in the viral genome and the loss of viral fitness. This is in agreement with our observation that viruses with 5-6 drug-resistant mutations were rare and viruses with more than 6 mutations were not detected in any patients. Viruses with more drug- resistant mutations can theoretically be more advantageous for resistance to combinations of antiretroviral drugs, but the severe fitness cost render such viruses incapable to replicate and/or compete with other more fit viruses with fewer mutations. When viruses with multiple drug-resistant mutations were analyzed, we found that viral fitness was lower in viruses with MDR mutations compared to viruses that only carried SDR mutations. Our model also showed that the timing of the emergence WT viruses varies somewhat from patient to patient but the average time to reach 1% WT across the three most closely studied patients (PID811, PID908 and PID268) was 50 days. Our study has some limitations, reflecting the source of the patient population as a clinical cohort. Treatment interruption is no longer considered clinically acceptable so the total number of such patients that can be studied is limited. Patients were treated with different drug combinations and had various previous treatment histories. The samples were collected at different time points for each subject. The analysis was performed for select drug-resistant mutations and the effect of potential compensatory mutations (16, 30, 39) was not assessed. A well-controlled cohort in which a larger number of patients are treated with the same drugs and samples are collected at similar schedules would be ideal, but such a population is no longer feasible. In summary, a comprehensive study of the dynamics of drug-resistant virus populations following treatment interruption and determination of the reservoirs for the linked MDR viruses represent important opportunities for improved understanding of ART resistance and its management. This work was supported by grants from the National Institutes of Health (GM065057, AI64518, AI067854, AI28433, RR06555, P20-RR18754, and K24- AI01608), the National Science Foundation (Grant No. NSF PHY05-51164) and portion of the work were done under the auspices of the U. S. Department of Energy under contract DE-AC52-06NA25396. References:
Asquith, B., and A. R. McLean. 2007. In vivo CD8+ T cell control of
immunodeficiency virus infection in humans and macaques. Proc Natl Acad Sci U S A 104:6365-70.
Boffito, M., L. Else, D. Back, J. Taylor, S. Khoo, M. Sousa, A. Pozniak,
B. Gazzard, and G. Moyle. 2008. Pharmacokinetics of
atazanavir/ritonavir once daily and lopinavir/ritonavir twice and once daily over 72 h following drug cessation. Antivir Ther 13:901-7.
Cai, F., H. Chen, C. B. Hicks, J. A. Bartlett, J. Zhu, and F. Gao. 2007.
Detection of minor drug-resistant populations by parallel allele-specific sequencing. Nat. Methods 4:123-125.
Clavel, F., and A. J. Hance. 2004. HIV drug resistance. N. Engl. J. Med.
Cressey, T. R., G. Jourdain, M. J. Lallemant, S. Kunkeaw, J. B.
Jackson, P. Musoke, E. Capparelli, and M. Mirochnick. 2005.
Persistence of nevirapine exposure during the postpartum period after intrapartum single-dose nevirapine in addition to zidovudine prophylaxis for the prevention of mother-to-child transmission of HIV-1. J Acquir Immune Defic Syndr 38:283-8.
Croteau, G., L. Doyon, D. Thibeault, G. McKercher, L. Pilote, and D.
Lamarre. 1997. Impaired fitness of human immunodeficiency virus type 1
variants with high-level resistance to protease inhibitors. J Virol 71:1089-
Deeks, S. G., R. Hoh, T. B. Neilands, T. Liegler, F. Aweeka, C. J.
Petropoulos, R. M. Grant, and J. N. Martin. 2005. Interruption of
treatment with individual therapeutic drug classes in adults with multidrug- resistant HIV-1 infection. J Infect Dis 192:1537-44.
Deeks, S. G., T. Wrin, T. Liegler, R. Hoh, M. Hayden, J. D. Barbour, N.
S. Hellmann, C. J. Petropoulos, J. M. McCune, M. K. Hellerstein, and
R. M. Grant. 2001. Virologic and immunologic consequences of
discontinuing combination antiretroviral-drug therapy in HIV-infected patients with detectable viremia. N. Engl. J. Med. 344:472-80.
Devereux, H. L., M. Youle, M. A. Johnson, and C. Loveday. 1999.
Rapid decline in detectability of HIV-1 drug resistance mutations after stopping therapy. AIDS 13:F123-7.
Goudsmit, J., A. de Ronde, E. de Rooij, and R. de Boer. 1997. Broad
spectrum of in vivo fitness of human immunodeficiency virus type 1 subpopulations differing at reverse transcriptase codons 41 and 215. J Virol 71:4479-84.
Goudsmit, J., A. De Ronde, D. D. Ho, and A. S. Perelson. 1996. Human
immunodeficiency virus fitness in vivo: calculations based on a single zidovudine resistance mutation at codon 215 of reverse transcriptase. J Virol 70:5662-4.
Hance, A. J., V. Lemiale, J. Izopet, D. Lecossier, V. Joly, P. Massip, F.
Mammano, D. Descamps, F. Brun-Vezinet, and F. Clavel. 2001.
Changes in human immunodeficiency virus type 1 populations after treatment interruption in patients failing antiretroviral therapy. J. Virol. 75:6410-7.
Harrigan, P. R., S. Bloor, and B. A. Larder. 1998. Relative replicative
fitness of zidovudine-resistant human immunodeficiency virus type 1 isolates in vitro. J Virol 72:3773-8.
Hirsch, M. S., F. Brun-Vezinet, B. Clotet, B. Conway, D. R. Kuritzkes,
R. T. D'Aquila, L. M. Demeter, S. M. Hammer, V. A. Johnson, C.
Loveday, J. W. Mellors, D. M. Jacobsen, and D. D. Richman. 2003.
Antiretroviral drug resistance testing in adults infected with human immunodeficiency virus type 1: 2003 recommendations of an International AIDS Society-USA Panel. Clin. Infect. Dis. 37:113-28.
Holland, J. J., J. C. de la Torre, D. K. Clarke, and E. Duarte. 1991.
Quantitation of relative fitness and great adaptability of clonal populations of RNA viruses. J Virol 65:2960-7.
Huigen, M. C., P. M. van Ham, L. de Graaf, R. M. Kagan, C. A.
Boucher, and M. Nijhuis. 2008. Identification of a novel resistance
(E40F) and compensatory (K43E) substitution in HIV-1 reverse transcriptase. Retrovirology 5:20.
Iarikov, D. E., M. Irizarry-Acosta, C. Martorell, C. A. Rauch, R. P.
Hoffman, and D. J. Skiest. Use of HIV resistance testing after prolonged
treatment interruption. J Acquir Immune Defic Syndr 53:333-7.
Johnson, J. A., J. F. Li, X. Wei, J. Lipscomb, D. Irlbeck, C. Craig, A.
Smith, D. E. Bennett, M. Monsour, P. Sandstrom, E. R. Lanier, and W.
Heneine. 2008. Minority HIV-1 drug resistance mutations are present in
antiretroviral treatment-naive populations and associate with reduced treatment efficacy. PLoS Med 5:e158.
Kikaire, B., S. Khoo, A. S. Walker, F. Ssali, P. Munderi, L. Namale, A.
Reid, D. M. Gibb, P. Mugyenyi, and H. Grosskurth. 2007. Nevirapine
clearance from plasma in African adults stopping therapy: a pharmacokinetic substudy. AIDS 21:733-7.
Lawrence, J., D. L. Mayers, K. H. Hullsiek, G. Collins, D. I. Abrams, R.
B. Reisler, L. R. Crane, B. S. Schmetter, T. J. Dionne, J. M. Saldanha,
M. C. Jones, and J. D. Baxter. 2003. Structured treatment interruption in
patients with multidrug-resistant human immunodeficiency virus. N Engl J Med 349:837-46.
Lee, H. Y., E. E. Giorgi, B. F. Keele, B. Gaschen, G. S. Athreya, J. F.
Salazar-Gonzalez, K. T. Pham, P. A. Goepfert, J. M. Kilby, M. S. Saag,
E. L. Delwart, M. P. Busch, B. H. Hahn, G. M. Shaw, B. T. Korber, T.
Bhattacharya, and A. S. Perelson. 2009. Modeling sequence evolution
in acute HIV-1 infection. J Theor Biol 261:341-60.
Liu, J., M. D. Miller, R. M. Danovich, N. Vandergrift, F. Cai, C. B. Hicks,
D. J. Hazuda, and F. Gao. 2011. Analysis of low frequency mutations
associated with drug-resistance to raltegravir before antiretroviral treatment. Antimicrob Agents Chemother. Mammano, F., C. Petit, and F. Clavel. 1998. Resistance-associated loss
of viral fitness in human immunodeficiency virus type 1: phenotypic analysis of protease and gag coevolution in protease inhibitor-treated patients. J Virol 72:7632-7.
Mammano, F., V. Trouplin, V. Zennou, and F. Clavel. 2000. Retracing
the evolutionary pathways of human immunodeficiency virus type 1 resistance to protease inhibitors: virus fitness in the absence and in the presence of drug. J Virol 74:8524-31.
Markowitz, M. 2000. Resistance, fitness, adherence, and potency:
mapping the paths to virologic failure. JAMA 283:250-1.
Martinez-Picado, J., M. P. DePasquale, N. Kartsonis, G. J. Hanna, J.
Wong, D. Finzi, E. Rosenberg, H. F. Gunthard, L. Sutton, A. Savara, C.
J. Petropoulos, N. Hellmann, B. D. Walker, D. D. Richman, R.
Siliciano, and R. T. D'Aquila. 2000. Antiretroviral resistance during
successful therapy of HIV type 1 infection. Proc. Natl. Acad. Sci. USA Martinez-Picado, J., A. V. Savara, L. Sutton, and R. T. D'Aquila. 1999.
Replicative fitness of protease inhibitor-resistant mutants of human immunodeficiency virus type 1. J Virol 73:3744-52.
Metzner, K. J., S. G. Giulieri, S. A. Knoepfel, P. Rauch, P. Burgisser,
S. Yerly, H. F. Gunthard, and M. Cavassini. 2009. Minority quasispecies
of drug-resistant HIV-1 that lead to early therapy failure in treatment-naive and -adherent patients. Clin Infect Dis 48:239-47.
Moore, K. H., J. E. Barrett, S. Shaw, G. E. Pakes, R. Churchus, A.
Kapoor, J. Lloyd, M. G. Barry, and D. Back. 1999. The
pharmacokinetics of lamivudine phosphorylation in peripheral blood mononuclear cells from patients infected with HIV-1. AIDS 13:2239-50.
Nijhuis, M., R. Schuurman, D. de Jong, J. Erickson, E. Gustchina, J.
Albert, P. Schipper, S. Gulnik, and C. A. Boucher. 1999. Increased
fitness of drug resistant HIV-1 protease as a result of acquisition of compensatory mutations during suboptimal therapy. AIDS 13:2349-59.
Nowak, M. A., and R. M. May. 2000. Virus Dynamics: Mathematical
Principles of Immunology and Virology. Oxford University Press, Oxford, Palmer, S., V. Boltz, F. Maldarelli, M. Kearney, E. K. Halvas, D. Rock,
J. Falloon, R. T. Davey, Jr., R. L. Dewar, J. A. Metcalf, J. W. Mellors,
and J. M. Coffin. 2006. Selection and persistence of non-nucleoside
reverse transcriptase inhibitor-resistant HIV-1 in patients starting and stopping non-nucleoside therapy. AIDS 20:701-10.
Palmer, S., V. Boltz, N. Martinson, F. Maldarelli, G. Gray, J. McIntyre,
J. Mellors, L. Morris, and J. Coffin. 2006. Persistence of nevirapine-
resistant HIV-1 in women after single-dose nevirapine therapy for prevention of maternal-to-fetal HIV-1 transmission. Proc. Natl. Acad. Sci. USA 103:7094-9.
Palmer, S., M. Kearney, F. Maldarelli, E. K. Halvas, C. J. Bixby, H.
Bazmi, D. Rock, J. Falloon, R. T. Davey, Jr., R. L. Dewar, J. A. Metcalf,
S. Hammer, J. W. Mellors, and J. M. Coffin. 2005. Multiple, linked
human immunodeficiency virus type 1 drug resistance mutations in treatment-experienced patients are missed by standard genotype analysis. J. Clin. Microbiol. 43:406-13.
Richman, D. D. 2006. Antiviral drug resistance. Antiviral. Res. 71:117-21.
Richman, D. D., J. M. Grimes, and S. W. Lagakos. 1990. Effect of stage
of disease and drug dose on zidovudine susceptibilities of isolates of human immunodeficiency virus. J Acquir Immune Defic Syndr 3:743-6.
Shafer, R. W. 2009. Low-abundance drug-resistant HIV-1 variants: finding
significance in an era of abundant diagnostic and therapeutic options. J Infect Dis 199:610-2.
Simen, B. B., J. F. Simons, K. H. Hullsiek, R. M. Novak, R. D.
Macarthur, J. D. Baxter, C. Huang, C. Lubeski, G. S. Turenchalk, M. S.
Braverman, B. Desany, J. M. Rothberg, M. Egholm, and M. J. Kozal.
2009. Low-abundance drug-resistant viral variants in chronically HIV- infected, antiretroviral treatment-naive patients significantly impact treatment outcomes. J Infect Dis 199:693-701.
Svarovskaia, E. S., J. Y. Feng, N. A. Margot, F. Myrick, D. Goodman,
J. K. Ly, K. L. White, N. Kutty, R. Wang, K. Borroto-Esoda, and M. D.
Miller. 2008. The A62V and S68G mutations in HIV-1 reverse
transcriptase partially restore the replication defect associated with the K65R mutation. J Acquir Immune Defic Syndr 48:428-36.
Verhofstede, C., F. V. Wanzeele, B. Van Der Gucht, N. De Cabooter,
and J. Plum. 1999. Interruption of reverse transcriptase inhibitors or a
switch from reverse transcriptase to protease inhibitors resulted in a fast reappearance of virus strains with a reverse transcriptase inhibitor- sensitive genotype. AIDS 13:2541-6.
Wu, H., Y. Huang, C. Dykes, D. Liu, J. Ma, A. S. Perelson, and L. M.
Demeter. 2006. Modeling and estimation of replication fitness of human
immunodeficiency virus type 1 in vitro experiments by using a growth competition assay. J Virol 80:2380-9.
Figure legend:
Fig. 1. Detection of the M184V mutation after treatment interruption. The M184V
mutation was determined by PASS in plasma samples collected on therapy (week -10) and off therapy (week 3 to week 32) in patient PID811. Green and red dots indicate mutant and WT bases detected in amplified individual viral genomes, respectively. Viral loads (VL), numbers of viral genome detected, numbers of mutants in the viral population, and percentages of mutant viruses in the sequenced genomes are shown above and underneath of the PASS images. Due to limited sampling of the viral population the percentages of mutants observed may differ from the percentage of mutants in the population (see Fig. 2. Evolution of drug-resistant mutations following treatment interruption.
Detected individual drug-resistant mutations were plotted as percentages (A, C, E and G) and absolute numbers (B, D, F and H) in the viral population at each time point in each individual. Drug-resistant mutations are indicated by unique symbols as shown at the far right for each patient. The RTI mutations are shown as solid lines and the PI mutations are shown as dotted lines. The WT virus and viral load data are indicated as red triangles and black crosses, respectively. WT viruses were not included in patient PID895 in order to show the differences among minority drug-resistant virus populations (G and H). The G48V mutation was not analyzed at week 11 and is not shown in the plots (C and D) Fig. 3. Dynamic changes of linked MDR viruses following treatment interruption.
The viral population changes over time were compared in PID811 (A), PID908 (B) and PID268 (C). The numbers of linked drug-resistant mutations in viruses are indicated by different symbols and colors as indicated in each graph. The population changes of select predominant viruses with linked MDR mutations were compared in PID811 (D). Fig. 4. Relationship between relative fitness loss of drug-resistant mutants
compared to the WT and the numbers of drug-resistant mutations. The vertical bars indicate the 95% confidence interval range. Note that for PID268, the error bars are so small that they cannot be easily visualized. The dashed lines represent the best fit for the trend of fitness loss with higher number of mutations.

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