Turriziani, Ombretta PhD*; Bucci, Mauro BSc*; Stano, Armando BSc*; Scagnolari, Carolina PhD*; Bellomi, Francesca PhD*; Fimiani, Caterina MD†; Mezzaroma, Ivano MD†; D'Ettorre, Gabriella MD‡; Brogi, Andrea MD‡; Vullo, Vincenzo MD‡; Antonelli, Guido PhD*
Advances in HIV treatment have reduced morbidity and mortality rates in HIV-infected individuals.1 Although highly active antiretroviral therapy (HAART) can keep HIV-1 RNA plasma levels below the detection limit, it fails to eradicate the virus completely, even after years of uninterrupted therapy.2,3
Viral persistence is considered to be the result of 2 main mechanisms: the establishment of a latently infected CD4 cell compartment and residual low-level viral replication.4-6 Recent studies have demonstrated that any viral variant, including drug-resistant strains that circulate for certain times during antiretroviral treatment, may be archived, remaining conserved in the latent viral reservoir.7-14 The presence of drug-resistant variants in the latent cell compartment may be important when antiretroviral therapy is changed or interrupted. A better understanding of the composition of the latent virus reservoir in terms of wild type and drug resistance might help to improve the strategy for successful treatment of highly exposed patients.
In this study, we compared the mutational resistance patterns of the cell-free HIV and cell-associated archived provirus in the blood. The analysis was undertaken in 95 patients who, having undergone long-term treatment, were failing to respond to antiretroviral therapy. The main objective was to see whether many years of selection for a drug-resistant virus led to a genotype of HIV-1 in peripheral blood mononuclear cell (PBMC) DNA coinciding with that in plasma RNA or whether discordance exists between these 2 compartments.
MATERIAL AND METHODS
The samples used in this study were derived from routine genotypic resistance testing at the HIV drug resistance monitoring service of the University “La Sapienza,” Rome, Italy, where a genotyping service has been established since 2004 and where, in addition to plasma, PBMC DNA has been stored for all samples. Plasma RNA and PBMC DNA were collected from 95 HAART-experienced patients. All were infected with the HIV-1 subtype B strain and had been receiving antiretroviral treatment for a long time (mean = 7 years, range: 3-12 years). In particular, the therapeutic histories of 39 patients included 3 classes of drugs (nucleoside reverse transcriptase inhibitors [NRTIs], nonnucleoside reverse transcriptase inhibitors [NNRTIs], and protease inhibitors [PIs]); 31 patients had received NRTIs and PIs, 19 were PI naive but NRTI and NNRTI experienced, and 6 had been treated with NRTIs only.
At the time of the analysis, the median CD4 count was 287 cells/μL (range: 2-1507 cells/μL) and the median viral load was 4.4 log copies/mL (range: 1.69-5.69 log copies/mL).
The 5 samples analyzed in this study were also available after the interruption of treatment. Specifically, plasma and PBMC samples were harvested before (T0) and 3 months after (T1) interruption of therapy.
Isolation of Peripheral Blood Mononuclear Cells
PBMCs were isolated from venous blood samples and separated on Lympholyte-H (Cedarlane Laboratories Limited, Burlington, Ontario, Canada) gradients. The isolated cells were pelleted by centrifugation, washed twice with phosphate-buffered saline, and then counted in Turk staining; 5 × 106 cells were pelleted and kept as dry pellets at −80°C.
RNA was extracted from 140 μL of plasma using a QIAmp Viral RNA kit (Qiagen, Milan, Italy) in accordance with the manufacturer's instructions. The HIV-1 reverse transcriptase (RT) and protease genes were amplified and sequenced using a TruGene HIV-1 Genotyping kit (Bayer Health Care LLC, Tarrytown, NY) and an OpenGene automated DNA sequencing system (Bayer Diagnostics, Milan, Italy). The results were interpreted using GuideLine Rules 9.0 (Bayer Health Care LLC). Briefly, the entire protease gene (codon 1-99) and codon 37 through 248 of the HIV-1 RT gene were sequenced. Viral RNA was converted to 1.3 kb of complementary DNA (cDNA) and amplified by polymerase chain reaction (PCR) in a single-tube RT-PCR reaction. The reaction products were then added, without purification, to a set of sequencing reaction tubes. Sequencing was undertaken using CLIP (TruGene HIV-1, Bayer Diagnostics), a DNA sequencing technique for direct sequencing of small quantities of amplified template.
DNA was extracted from 5 × 106 PBMCs using the Dneasy Tissue kit (Qiagen). To check the DNA extraction, the β-actin gene was amplified as described previously.15 The RT-PCR step provided in the TruGene HIV-1 was modified for viral DNA. Specifically, 2 μL of the extract, containing approximately 0.3 μg of DNA, was added to the PCR reaction. Samples were denatured for 5 minutes at 94°C, followed by 20 cycles at 94°C for 30 seconds, 57°C for 30 seconds, and 68°C for 90 seconds and then 17 cycles at 94°C for 30 seconds, 60°C for 30 seconds, and 70°C for 2 minutes, with a final incubation at 70°C for 7 minutes. The PCR products were sequenced by TruGene HIV-1 in accordance with the manufacturer's instructions.
The McNemar-Bowker test was used to measure the agreement between the numbers of mutations in PBMC DNA and in plasma RNA. We calculated the proportion of discordant samples (DSs) and the corresponding 95% confidence intervals.
The analysis of the mutation patterns associated with drug resistance detected in PBMC HIV DNA and plasma HIV RNA revealed that 8 of the 95 patients' samples analyzed showed virologic failure (as indicated by the rebound of viral load), without demonstrating any resistant mutations in both blood compartments. Careful anamnesis by the clinicians revealed that these patients were not adherent to antiretroviral therapy.
Eighty-five (89%) of the 95 patients' samples showed resistance mutations in HIV RNA and HIV DNA. Of these 85 samples, however, only 23 (27%) showed the same number and type of mutations in both compartments. The mutations at the codon recognized as important in the genotypic resistance pattern differed between HIV RNA and HIV DNA in 62 patients (73%; 95% confidence interval: 62% to 84%), with 2 patients (2%) having evidence of genotypic resistance in HIV RNA only. On the basis of these results, the patients' samples that demonstrated a replicating virus different from that archived in the PBMCs were arbitrarily defined as DSs.
Genotypic analysis revealed that in the 33 patients in the DS group (51%), the different numbers or types of DNA mutations led to different interpretations of drug susceptibility assay results. Furthermore, in 51% of DSs, there were greater numbers of resistance mutations in plasma HIV RNA compared with HIV DNA, whereas in 37%, a greater number of mutations were recorded in PBMCs. In the latter group of patients, some RT mutations (M41L, E44D, K65R, K70R, L74V, K101E, K103N, V106M, Y181I, Y188L, M184V, V179D, G190A, L210W, and T215Y) and some PI mutations (L10F, K20M, M36I, M46I, I54V, L63P, G73S, A71V, and V82A) were present only in PBMCs and not in plasma.
In 7 patients (11%), the numbers but not the types of mutations in HIV DNA and HIV RNA were the same (Fig. 1).
Analysis of Discordant Samples and Mutations Within Classes of Inhibitors
Analysis of the numbers of mutations detected in the 2 compartments revealed that more than 47% of samples had at least 5 mutations associated with NRTI or PI resistance. As shown in Figure 2, 32 samples had ≥5 NRTI mutations in PBMC DNA and 37 samples had ≥5 NRTI mutations in plasma HIV RNA. Only 8 plasma samples did not contain NRTI mutations. Similar results were obtained for the numbers of PI mutations.
With regard to the number of mutations responsible for NNRTI resistance, no mutations were detected in 28 PBMC DNA samples or in 22 plasma samples. As indicated in Figure 2, 5 mutations associated with NNRTI resistance were detected in plasma virus in only 1 patient, and, again, in only 1 patient, 5 NNRTI mutations were found in PBMC DNA. More usually, 2 mutations associated with NNRTI resistance were seen.
It is important to note that for NNRTIs, there is a trend toward a significant difference in the distribution of DNA and RNA mutations (McNemar-Bowker test, P = 0.056), whereas there were no differences in the case of NRTIs and PIs.
Analysis of the Frequency of Specific Mutations
Analysis of the frequency of mutations associated with NRTI resistance showed that the resistance mutation most frequently detected in our samples was M184V (76% of samples), followed by T215Y/F/S (70%), M41L (66%), D67N (51%) L210W (47%), and V118I (41%) (Fig. 3A). The high incidence of M184V reflects the wide use of lamivudine (3TC). In fact, 94% of the samples analyzed in this study came from patients who were 3TC experienced, and in 61% of these patients, 3TC was also part of their current therapeutic regimen. The selection of thymidine analogue mutations such as T215Y/F/S, M41L, D67N, and L210W was also attributable to 98% of patients being treated with thymidine analogues. Specifically, 28 patients (44%) were currently treated with a therapeutic regimen containing a thymidine analogue.
To evaluate the distribution of each mutation in the 2 compartments, we verified that the mutation was harbored in both compartments or in only 1 of the 2 analyzed.
The results, shown in Figure 3B, indicated that in most of the DS samples harboring a virus with NRTI mutations, these occurred in HIV RNA and PBMC DNA. A modest number of samples (range: 1-11 samples) showed mutations only in plasma RNA, with a few patients harboring the NRTI resistance mutations only in PBMC DNA (range: 1-9 samples). Interestingly, the K70R mutation was more often present in the sequence derived from PBMC DNA than in that derived from plasma RNA. The therapeutic histories of patients harboring mutations only in PBMC DNA showed compounds that could be responsible for the selection of the archived mutation. Intriguingly, the only 2 PBMC DNA samples containing a K65R mutation came from patients who had never been treated with drugs like didanosine, abacavir, or tenofovir, which are known to be the main cause of the appearance of this mutation.
As far as mutations associated with NNRTI resistance are concerned, higher incidences were observed for K103N (34% of sample) and G190A (33% of sample) (see Fig. 3C). Fourteen of the 22 samples harboring the virus with K103N and 15 of the 21 samples harboring G190A showed these mutations in both compartments. Further, some NNRTI mutations, such as V179D and V108I, were observed in the plasma virus or the cellular-associated virus but never in both compartments (see Fig. 3D). Intriguingly, 2 samples containing NNRTI mutations were from patients who were NNRTI naive. Specifically, in 1 case, K101E and G190A were found in plasma RNA, and in the other case, the plasma virus contained V106M and V179D and the PBMC DNA showed a G190A mutation.
Analysis of PI mutations showed that L63P and L10F occurred most frequently in our population (see Fig. 3E). Specifically, L63P was observed in 76% of samples and L10F was seen in 61%. Analysis of the distribution of PI mutations showed that samples harboring a virus with primary mutations, such as M46I, V82A, I84V, and L90M, demonstrated these mutations in both compartments; these mutations occurred in PBMC or plasma in only a few samples (see Fig. 3F). It is worth noting that the presence of PI primary mutations was detected in association with current therapeutic regimens containing PIs. Indeed, only 5 patients displayed primary PI mutations, even if PIs were not part of their current therapy.
Altogether these results clearly demonstrate that in patients receiving extensive antiretroviral treatment and with therapeutic failure, drug-resistant variants can easily be found archived in PBMCs and that the PBMC compartment does not necessarily reflect the plasma compartment. This could imply indirectly that PBMCs may constitute a reservoir for drug-resistant variants and might replenish plasma with drug-resistant HIV variants in certain situations.
Within the framework of this study, we had the opportunity to address and gain new insights into this issue. Indeed, various PBMC and plasma samples were available from 5 patients included in the analysis whose therapy was interrupted for 3 months. Hence, it was possible to analyze drug resistance mutations in both compartments both before and after the interruption in treatment. The results, shown in Figure 4, demonstrated that the interruption of treatment resulted in the resistant virus being replaced in the plasma by a wild-type virus, as shown by a decrease in the number of RNA mutations in those patients with a dominant wild-type population in PBMCs. No specific pattern of mutations was seen to disappear after therapy interruption. Indeed, all RT and PI mutations detected in plasma RNA disappeared after 3 months' interruption of therapy. Specifically, RT mutations detected in plasma virus were M41L, D67N, M184V, L210W, and T215Y/F/S (3 subjects); T69D (2 subjects); V118I (1 subject); K219Q/E (1 subject); K103N (1 subject); V108I (2 subjects); Y181I (2 subjects); and G190A (1 subject). The PI mutations were L10F (2 subjects); K20M, M36L, I54V, L63P, A71V, G73S, I84V, and L90M (all in 1 subject); and M46I (2 subjects).
When drug-resistant strains were archived in the cellular reservoir, the interruption of therapy did not reduce the number of RNA mutations. In particular, in the case of samples harboring resistant variants in PBMCs, 1 patient showed mutations with resistance to all NRTIs and NNRTIs before therapy was interrupted, together with mutations responsible for resistance to 4 different PIs. These mutations were detected in plasma RNA and PBMC DNA. After the interruption in treatment, the only mutation that had disappeared from plasma RNA was L74V, the only mutation without a counterpart in PBMC DNA, whereas the other mutations were still present in both compartments. In another patient, before the interruption in treatment, the virus from plasma RNA showed a pattern of mutations that are responsible for resistance to all NRTI and NNRTI analogues, whereas the virus from PBMCs showed only those mutations responsible for resistance to NRTIs. After the interruption in treatment, the plasma virus was still resistant to NRTIs but had recovered its sensitivity to NNRTIs, whereas the PBMC DNA mutation pattern remained unchanged.
Although advances in HIV treatment have reduced the morbidity and mortality rates among HIV-infected individuals, all current therapeutic regimens have failed to eliminate the latent reservoir, and it is known that eradication of the virus is not possible with current antiretroviral drugs.1,2 Antiretroviral therapy must be administered for a patient's entire life, and this is complicated by the side effects of the drugs and the emergence of drug resistance.16-18 HIV-infected individuals in whom therapeutic regimens have repeatedly failed often harbor a virus with multidrug-resistant mutations. Currently, plasma is the only compartment used routinely for drug resistance testing, because the analysis of viral RNA is supposed to reflect the virus that is currently replicating. Several studies in patients on successful HAART but with a history of pre-HAART drug resistance have shown that any viral variant replicated for a time during the infection enters the reservoir and remains archived, however.7-14 Further, a recent study by Chew and coworkers19 indicates that PBMC analysis is a reliable predictor of drug resistance. The persistence of drug-resistant mutants in the reservoir may be of particular importance when the circulating virus does not reflect the archived population and when patients have more than 1 salvage therapy.
In this retrospective study, we compared the mutational resistance patterns of the cell-free HIV and cell-associated archived provirus in the blood of 95 HIV-infected patients in whom antiretroviral regimens had failed. We observed that in approximately 73% of these individuals, the HIV-1 drug resistance mutations detected in plasma were different from those in PBMCs, which suggested that analysis of plasma virus would not be sufficient to understand the drug-resistant status of the patient completely in some cases. As expected, and in accordance with the findings reported by other authors,8,10,11,20 our results showed that there were a greater number of mutations in plasma than in PBMCs in most patients. Our data also showed that in 37% of samples, PBMCs contained the greatest numbers of mutations. These data indicate that although the plasma virus represents the material of choice for the detection of drug resistance, as reported by other authors,11,21 the PBMC HIV sequence can, at least in some cases, contain the greatest variety of mutations. In agreement with other studies,8,11,12,20 the additional mutations found in the cellular compartment probably arose from previous therapy; these patients had, in fact, been treated previously with drugs able to select the mutations found in the PBMCs.
It is important to underline that this analysis is based on the use of a routine assay that does not allow the detection of minor viral variants. In this regard, a previous study8 showed that when mutation analysis was performed by cloning PBMC HIV DNA and HIV RNA from plasma, additional RT and protease mutations were found in PBMCs. Hence, we cannot rule out the possibility that the PBMCs studied here contain a mixture of viral variants more heterogeneous than that detected using our method.
In the population studied, many NRTI and PI mutations were found in plasma and PBMC DNA. These data probably reflect the therapeutic histories of these patients. In fact, it is important to stress that most of our patients had received extensive treatment (mean = 7 years); as expected, all their antiretroviral drug histories showed the use of nucleoside analogues and 77% had received at least 1 PI. Furthermore, most of them received HAART at a time when resistance testing was not being undertaken, which could allow replication of the resistant variant. This hypothesis is in agreement with the data published by Verhofstede and coworkers,14 who suggested that the probability of finding a resistant variant within the reservoirs depended, at least in part, on the period over which this variant was able to replicate. Thus, a delay in changing a failed therapeutic regimen might favor drug-resistant mutants entering the reservoirs.
The presence in most of the patients studied of a low number of RT mutations that conferred resistance to NNRTIs was probably attributable to the fact that these drugs were not only less well represented than NRTIs and PIs in the therapeutic regimens but were used for shorter times than other drugs. Most samples contained 1 or 2 NNRTI-resistant mutations, and these were detected in plasma and PBMCs. NNRTIs are known to be drugs with low genetic barriers, and a single mutation in the NNRTI-binding pocket may lead to broad cross-resistance to currently licensed agents in this drug class. Second-generation NNRTIs, such as TMC125, show promise of being active against viruses with 1 or 2 mutations;22 hence, such therapies could be effective in patients harboring these viral variants.
The extensive previous treatment of our study population can easily explain the presence of the resistance mutations in plasma and PBMC DNA or, in some cases, only in PBMCs. Another possible explanation, however, could be the differing abilities of the variants to replicate. It is known that some resistant mutations affect viral fitness;23 thus, the possibility cannot be excluded that a drug-resistant variant might also display a different ability to persist in the reservoirs. In this regard, it is worth noting that the K70R mutation was the only mutation found more frequently in PBMCs than in plasma. This mutation was one of the earliest substitutions observed with the emergence of genotypic resistance to zidovudine, and it was preferentially selected during zidovudine monotherapy.24,25 Those patients harboring K70R only in a proviral sequence had undergone prolonged treatment with zidovudine as monotherapy or in combination (>4 years), and their therapeutic regimen did not include thymidine analogues at the time of the resistance testing. This observation supports the hypothesis from Ruff and coworkers9 that the clones containing this mutation persist once archived, despite continued drug-selective pressure for further RT mutations.
Furthermore, the persistence of this viral mutant in the reservoirs for so long a time corroborates the hypothesis, previously suggested by other authors, 26-28 regarding the persistence of wild-type HIV-1; this, as is known, has the potential to re-emerge from the latent reservoir even after up to 10 years of antiretroviral therapy. These authors suggested, and our data support, the possibility that memory T cells or their progeny can survive for years, and probably decades, allowing the persistence of an archival wild-type or resistant mutant.9,26-28
Some mutations, such as V179D and V108I, have never been observed in both compartments. This is difficult to explain, except for the possibility that the viral strains displaying these mutations are not easily archived or require a longer time to be archived in PBMCs. Because these mutations were seen in the PBMCs of just 1 patient, however, it is difficult to hypothesize from so few samples.
Our data also demonstrate that in patients who have received extensive treatment, the dominant population harbored in PBMCs is represented by a drug-resistant variant and the resistance mutation detected in PBMC DNA differed from that detected in plasma RNA. These data do not seem to be consistent with the findings published by other authors, who reported a substantial correlation between HIV drug-associated resistance mutations in plasma and PBMCs.12,20 A possible explanation for this discrepancy lies in the different numbers of patients examined, together with the difference in the definition of DSs. Indeed, in our study, we arbitrarily defined the discordance, because just 1 mutation was considered sufficient to define a DS. Because the mutations detected are drug resistant, we consider that the presence or absence of one of these mutations might lead to a change in drug susceptibility or a different evolution of the drug resistance pattern. In some patients, for example, mutations such as M184V or K103N, selected by a previous regimen, were detected only in PBMCs; because these mutations are known to be responsible for resistance to cytidine analogues and to NNRTIs, respectively,29-32 their presence in the reservoirs might further accelerate the emergence of a drug-resistant mutant. These data, together with previously reported findings,9 suggest that this phenomenon could be responsible for the short-term responses to reinitiation of therapy documented after an interruption of treatment.33-36 Although relating to only a small number of patients, our data demonstrate that the intracellular viral population seems to affect the replacement of the mutant by a wild-type virus. In fact, when the intracellular dominant viral population is represented by a drug-resistant variant, a treatment interruption of 3 months does not seem to be long enough to enable a wild-type virus to re-emerge. These data, which need to be confirmed in a large cohort of patients, support the findings of Lambotte and coworkers,13 who suggested that the lymphocytes' HIV reservoir contributes to the plasma viral load rebound that occurs when therapy is stopped, but do not seem to be in agreement with other findings.10,20 Indeed, the authors of these reports demonstrated that the discontinuation of therapy resulted in the rapid disappearance of mutations from plasma but not from DNA. Our results can be explained by taking into account the facts that our patients, who showed no reductions in plasma RNA mutations, had longer therapeutic histories (mean = 6.5 years) than those reported in the other reports and had more than 1 previously failed therapeutic regimen. Thus, we suppose that resistant viral variants represent the dominant population in these individuals and that because wild-type viruses are present in only small amounts, it takes longer to replace the resistant viral population. We consider, however, that the crucial point arising from all these studies is the persistence of the mutations in the cellular reservoir, and this could affect any subsequent salvage regimen.
This study had certain limitations. It was performed using a commercial kit based on direct sequencing and, as stated previously, this did not allow the detection of minor viral variants. Furthermore, being a retrospective study, it was performed on total PBMCs. We consider that further information on the compartmentalization of resistant viral strains could be derived from an analysis of the resistance patterns in different blood cells. It is known that although CD4+ T cells seem to be the main HIV-1 pool,37 other cells, such as monocytes, contribute to viral persistence during HAART. Hence, a comparison of these 2 cellular compartments could provide further information relating not only to the persistence of the resistance mutants but to the dynamics of HIV infection.
In conclusion, our study reinforces the data obtained by other authors and strongly indicates that in patients who have received extensive treatment, PBMC DNA represents a reliable guide to drug resistance. Any therapeutic decisions regarding changes in regimens, treatment interruption, or “recycling drugs” should take into account the fact that drug-resistant forms of the virus in a given patient could be archived in the reservoirs and might re-emerge when conditions become more favorable.
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