In most patients infected with HIV-1, highly active antiretroviral treatment (HAART) decreases plasma viremia to < 200, or even < 50 copies/ml. This reduction translates into clinical benefits [1,2]. Plasma viremia levels result from a dynamic process with continuous cycles of viral replication, and rapid turnover of viruses and virus-producing cells . More refined analysis of viral decay in HAART-treated patients identified a second-phase decay, which allowed computation of the time required for virus eradication. This time was estimated at 2–3 years, with the provision that viral reservoirs with longer half-lives might increase this lapse of time . These data are challenged by the finding, when sensitive assays are used, that plasma viremia is still detectable for years at low levels in a large percentage of adherent patients on combined therapy [5–7]. A reservoir of long-lived infected resting CD4 T lymphocytes was also identified through the use of refined co-culture methods [8–10]. In patients with plasma viremia levels below 200 or 50 copies/ml, estimates of the average half-life of the pool of infected resting CD4 T cells range from 34 months in those starting HAART during the chronic phase of HIV-1 infection  to 6 months in those treated at the time of acute HIV-1 infection . These data and others [7,13–14] suggest that initiation of treatment at the time of acute infection and when CD4 T cells are high in chronically infected patients leads to a more efficient control of HIV-1 replication.
In this study, we used newly developed highly sensitive virological assays for the quantification of plasma viremia and cell-associated viral RNA and DNA to evaluate the clearance of infected cells and to identify factors associated with the level of residual viral load in a selected group of adherent patients treated at the time of acute HIV-1 infection.
Materials and methods
Selection of patients
Patients belonging to previously described cohorts [15,16] were selected on the basis of the following criteria: initiation of combined antiretroviral therapy during acute HIV-1 infection, treatment for at least 24 months, sustained plasma HIV-1 RNA < 200 copies/ml and availability of at least eight samples of frozen peripheral blood mononuclear cells (PBMC) including baseline sample
Plasma viremia quantification
Plasma HIV-1 RNA was quantified using first the standard Amplicor HIV-1 Monitor assay version 1.5 (Roche Diagnostic System, Basel, Switzerland). The samples with RNA < 200 copies/ml were retested using a modified version of Amplicor HIV-1 Monitor assay [7,17]. Briefly, 1 ml plasma was centrifuged at 50 000 ×g for 80 min at 4°C. After removal of plasma, the next steps were performed according to the manufacturer's instructions except that the concentration of the internal quantitative standard was reduced 15-fold and the volume of specimen diluent was reduced from 400 to 55 μl. The substrate incubation time was extended to 15 min. The limit of detection for RNA was 3 copies/ml and the mean coefficient of variation was 40% (range 23–52) for 5–50 copies/ml.
Collection of peripheral blood mononuclear cells and lymph node mononuclear cells
PBMC were isolated from blood treated with ethylenediamine tetraacetic acid by gradient centrifugation (Ficoll Hypaque, Pharmacia, Dubendorf, Switzerland) and samples of 3 × 106 cells were stored in liquid nitrogen. Inguinal biopsy was performed under local anesthesia. Lymph node tissue was minced with a scalpel in tissue culture medium and the cells were teased out using small tweezers. Lymph node mononuclear cells (LNMC) were isolated by gradient centrifugation (Ficoll Hypaque) and samples of 3 × 106 cells were stored in liquid nitrogen.
CD3, CD4 and CD8 lymphocyte cell counts were determined in PBMC and LNMC by flow cytometry (Coulter EPICS IV, Basel, Switzerland) using fluorescein-conjugated DAKO-T3 and DAKO-T8, and R-Phycoerythrin DAKO-CD4 (Dako, Glostrup, Denmark).
Cell-associated HIV-1 RNA and DNA quantification
Cell-associated HIV-1 RNA and DNA were quantified on the same cell portion using the reagents of the Amplicor HIV-1 Monitor assay (Roche) as previously described . Briefly, 3 × 106 cells (PBMC or LNMC) were lysed in lysis buffer containing RNA internal quantification standard at 50% of the concentration recommended. For measurement of cell-associated HIV-1 RNA, the nucleic acid preparation was incubated with 20 U DNAase I RNAase-free (Boehringer, Manheim, Germany) for 1 h at 37°C, then for 5 min at 95°C and diluted in 80 μl specimen diluent. Then, 50 μl RNA preparation was amplified according to the manufacturer's instructions. For measurement of cell-associated HIV-1 DNA, the nucleic acid preparation was incubated for 1 h at 37°C with 10 μg RNAase A DNAase-free (Sigma, Buchs, Switzerland) and diluted in 80 μl specimen diluent. Then, 50 μl DNA preparation was added to mastermix buffer containing 25 copies DNA internal quantification standard (provided by Roche Diagnostic Research and Development, Alameda, California) and amplified. The limit of detection was approximately 3 copies/106 cells for RNA and 5 copies/106 cells for DNA. The mean coefficient of variation for 10–1000 copies/106 cells was 12% (range 3–26) for cell-associated RNA and 18% (range 2–29) for cell-associated DNA. Results are expressed in log10 copies/106CD4 T cells.
Associations between variables were assessed using Pearson correlation coefficient and comparisons between parameters were assessed using paired Wilcoxon-test. Association of variables with the mean levels of plasma viremia, cell-associated RNA and DNA from 6 to 33 months after treatment initiation was performed using univariate and multivariate linear regression models. Samples with undetectable plasma viremia, cell-associated RNA or DNA were assigned a value of 0.1 (−1 log10) copies/ml. Linear regression models were used to estimate the mean decay in log10 RNA or DNA copies/106CD4 T cells over follow-up time from initiation of therapy. Half-life of cell-associated DNA and RNA was calculated using the formula: −log10(2)/slope.
Patients with acute HIV-1 infection
Table 1 reports the characteristics of the 15 patients included in the study. All study patients had symptomatic acute HIV-1 infection and initiated antiretroviral therapy, on average, 20 days (range 7–40) after onset of symptoms. The most common symptoms were fever (73%), asthenia (53%), cutaneous rash (40%) and adenopathy (40%). The initial treatment regimen was changed during the follow-up in three patients; zidovudine was replaced by stavudine in two patients because of anemia and indinavir was replaced by other protease inhibitors owing to nephrolithiasis in two patients. All patients included in the study signed the informed consent form approved by the local ethics committee.
Evolution of plasma viremia, and cell-associated RNA and DNA in blood
Figure 1 shows mean plasma viremia and cell-associated RNA and DNA levels as a function of time. Plasma HIV-1 RNA decreased in all patients to < 200 copies/ml by 6 months after initiation of treatment. Using an assay with a detection limit for HIV-1 RNA of 3 copies/ml and assigning a value of 0.1 (−1 log10) copies/ml to undetectable samples, the mean plasma viremia was 0.72 log10 copies/ml at 6 months of therapy and decreased to −0.09 log10 copies/ml at 24 months. Plasma viremia remained persistently below this detection threshold in nine (60%) patients for a period of 12 to 27 months; in five patients it fluctuated between 3 and 50 copies/ml and in one patient it fluctuated between 50 and 200 copies/ml.
The mean cell-associated RNA in PBMC was 3.35 log10 copies/106 CD4 T cells at baseline and decreased to 0.95 log10 copies/106 CD4 T cells at 24 months. PBMC-associated RNA remained persistently below the detection threshold of 3 copies/106 cells in three patients (patients 7, 11, 15) for a period of 23 to 33 months. Two other patients (patients 4, 12) had undetectable PBMC-associated RNA in more than 50% of the samples collected after 6 months of treatment. PBMC-associated RNA was detectable in all samples analyzed in the remaining eight patients.
The mean PBMC-associated DNA was 3.50 log10 copies/106CD4 T cells at baseline and decreased to 1.27 log10 copies/106 CD4 T cells at 24 months. PBMC-associated DNA was detectable (> 5 copies/106 cells) at all time-points analyzed in 11 (73%) patients. One patient had persistent undetectable PBMC-associated DNA between 6 and 24 months; however his baseline PBMC-associated DNA was low (1.11 log10 copies/106 CD4 T cells).
At baseline, the level of plasma viremia was highly correlated with the level of both PBMC-associated RNA (r = 0.77, P = 0.001) and PBMC-associated DNA (r = 0.85, P < 0.001). A strong correlation was also found between PBMC-associated RNA and DNA (r = 0.84, P < 0.001).
To account for multiple measurements in the same patient, the mean values for virological parameters were calculated in samples collected after 6 months of treatment for each patient. The mean plasma viremia was 0.05 log10 copies/ml (range, −1.00 to 2.13), the mean PBMC-associated RNA was 1.09 log10 copies/106 CD4 T cells (range, −1.00 to 2.45) and the mean PBMC-associated DNA was 1.84 log10 copies/106 CD4 T cells (range, −1.00 to 3.73). The mean level of PBMC-associated DNA under therapy was strongly correlated with the mean level of PBMC-associated RNA (r = 0.76, P = 0.001) and the mean level of plasma viremia (r = 0.67, P = 0.006). The correlation between the mean levels of plasma viremia and PBMC-associated RNA was weaker (r = 0.54, P = 0.04).
Cell-associated RNA and DNA in lymph nodes
As lymphoid tissues are the main site of HIV-1 replication, we also determined cell-associated HIV-1 RNA and DNA levels in LNMC from inguinal lymph node biopsies collected after a mean of 14 months (range, 8–19) of therapy. LNMC were available for 14 patients (insufficient recovery for one patient). Table 2 reports virological parameters in blood and lymph node samples collected at the same time point. Nine patients had plasma viremia below the threshold of 3 copies/ml and four of them had PBMC-associated RNA below the threshold of 3 copies/106 cells. The mean levels of cell-associated RNA were similar in blood and lymph (0.97 and 1.05 log10 copies/106 CD4 T cells, respectively) and were significantly correlated (r = 0.61, P = 0.02). Five patients (patients 4, 7, 11, 12, 15) had undetectable LNMC-associated RNA. In these five patients, PBMC-associated RNA was undetectable in 50–100% of the samples collected after 6 months of treatment.
Cell-associated HIV-1 DNA was detectable in all patients, except in PBMC of patient 11 and in LNMC of patient 7. Mean cell-associated DNA levels in blood and in lymph nodes were similar (1.67 and 1.64 log10 copies/106 CD4 T cells, respectively;P = 0.95, paired t test) and were significantly correlated (r = 0.66, P = 0.01). In each compartment, mean cell-associated DNA levels were significantly higher than cell-associated RNA levels (PBMC: 1.66 and 0.97 log10 copies/106 CD4 T cells, respectively;P = 0.015 paired t test; LNMC: 1.64 and 0.91 log10 copies/106 CD4 T cells;P = 0.013, paired t test). A strong correlation was observed between cell-associated RNA and DNA in both blood (r = 0.77, P = 0.001) and lymph nodes (r = 0.80, P = 0.001).
Decays of cell-associated DNA and RNA
To address the issue of clearance of HIV-1-infected cells, the levels of cell-associated DNA and RNA were evaluated in each patient 9 to 12 times (median 10) over the course of the 24–33 months of treatment. A two-phase decay rate in cell-associated DNA was observed after treatment initiation, with an inflexion point at 3 months (Fig. 1). The first phase was characterized by a rapid decay with a mean slope of −0.36 log10 copies/106 CD4 T cells per month [95% confidence interval (CI) −0.46 to −0.26] corresponding to a half-life of 0.84 months (95% CI 0.65–1.16) (Table 3). The second mean decay rate was slower, with a slope of −0.045 log10 copies/106 CD4 T cells per month, corresponding to a half-life of 6.6 months. The mean slope of the second-phase decay was statistically different from zero (P < 0.001). The lower and upper 95% CI for the slope (−0.069 and −0.022, respectively) correspond to half-lives of 4.4 and 13.8 months, respectively. In most patients, the second-phase DNA decay was remarkably linear, but slopes varied markedly among individuals (Fig. 2).
The two-phase linear regression model used in this analysis fitted well the individual data for the 15 patients (mean adjusted r2 0.73; range 0.35–0.94). To provide a more accurate estimate of mean decay, individual slopes were weighted proportionally to the inverse of the variance of each slope estimate. Similar results were obtained, with a first-phase cell-associated DNA decay of −0.30 log10 copies/106 CD4 T cells per month (95% CI −0.37 to −0.22) and a second-phase decay of −0.046 log10 copies/106 CD4 T cells per month (95% CI −0.069 to −0.023).
The second-phase decay rate of cell-associated DNA was highly correlated with the mean level of cell-associated RNA over the 6 to 33 months after treatment initiation (r = 0.77, P = 0.001). In addition, the five patients (patients 4, 7, 11, 12, 15) with undetectable LNMC-associated RNA and more than 50% of samples with undetectable cell-associated RNA in blood presented a more rapid second-phase decay of cell-associated DNA. The median slope was −0.060 log10copies/106 CD4 T cells per month (range −0.13 to −0.040) for these five patients compared with −0.027 log10 DNA copies/106 CD4 T cells per month (range −0.061 to 0.026) for the other patients (P = 0.05). No significant association was found between the second-phase decay of cell-associated DNA and the mean level of plasma viremia among the 15 patients (P = 0.45). Finally, the second-phase DNA decay was correlated with the level of LNMC-associated RNA measured, on average, 14 months after treatment initiation (r = 0.62, P = 0.02).
There was also a two-phase decay rate of cell-associated RNA (Fig. 1). The first phase occurred during the initial month of treatment and was characterized by a sharp decrease with a mean slope of −2.02 log10 copies/106 CD4 T cells per month (95% CI −2.70 to −1.34), corresponding to a half-life of 0.15 month (Table 3). In contrast, the second-phase decay was extremely slow, with a mean slope of −0.022 log10 copies/106 CD4 T cells per month (95% CI −0.037 to −0.007), corresponding to a half-life of 13.7 months.
Predictors of residual plasma viremia, cell-associated RNA and DNA
The association between baseline virological parameters and the mean levels of plasma viremia, cell-associated RNA and DNA from 6 to 33 months of treatment were assessed using linear regression models (Table 4). Baseline PBMC-associated DNA level was the best predictor of the mean levels of plasma viremia and PBMC-associated RNA and DNA, whereas baseline plasma viremia was not significantly associated with long-term outcome. These results suggest that cell-associated virological markers may be better predictors of long-term virological outcome than plasma viremia in treated patients. We also found that LNMC-associated RNA and DNA were highly associated with the mean level of PBMC-associated RNA and to a lower extent with the mean level of PBMC-associated DNA.
In this investigation, the kinetics of cell-associated HIV-1 DNA and RNA was assessed in patients treated at the time of acute HIV-1 infection. Although 9 of 15 patients achieved a persistent plasma viremia < 3 copies/ml after 1 year of treatment, all but three had evidence of continuous or intermittent HIV-1 replication based on the detection of cell-associated unspliced viral mRNA and/or genomic RNA in PBMC. Following a rapid decay during the first month of treatment, the second-phase decay rate of cell-associated RNA was extremely slow, with a mean half-life of 13.7 months. The kinetics of cell-associated DNA also presented a two-phase decay characterized by a rapid decay rate in the first 3 months followed by a slower decay rate, corresponding to a mean half-life of 6.6 months. This second-phase DNA decay was characterized by large interindividual differences and was strongly correlated with the mean level of cell-associated RNA in PBMC collected from 6 to 33 months after treatment initiation.
Several of the findings deserve comment. The marked difference in the first decay rates between cell-associated RNA and DNA (−2.02and −0.36 log10 copies/106 CD4 T cells per month, respectively) is likely to result from the respective contribution of various types of HIV-1-infected cells. Indeed, the rapid removal of acutely infected cells containing high numbers of cell-associated RNA copies and but few copies of cell-associated DNA results in a steep slope of cell-associated RNA, whereas cells containing few copies of cell-associated RNA, such as infected monocytes, infected resting CD4 T cells and cells containing defective viral DNA, contribute to the slower decay of cell-associated DNA.
Low levels of cell-associated RNA in LNMC indicate that, among patients on HAART with undetectable plasma viremia, viral replication is inhibited at a similar extent in blood and lymph nodes after an average treatment period of 1 year. Finally, the finding that levels of cell-associated RNA and DNA in lymph nodes were strongly correlated with the mean level of cell-associated RNA in blood has direct consequences for the monitoring of aviremic patients. In such patients, an evaluation of treatment efficacy can be derived from the iterative measurements of PBMC-associated RNA and DNA. These measurements rely on simple assays performed on small amount of biological material. In addition, a baseline assessment of cell-associated DNA in blood was the best predictor of viral activity markers (plasma viremia, cell-associated DNA and RNA) after 6 to 33 months of antiretroviral treatment. No significant association was found between baseline plasma viremia and any of the three markers. If this is confirmed in patients starting therapy during the chronic phase of HIV-1 infection, initial cell-associated DNA determinations could provide useful information to guide antiretroviral treatment.
The clearance of HIV-1-infected cells in patients on HAART has been previously evaluated by longitudinal measurements of the reservoir of infected resting CD4 T cells [11,12]. The methodology used in these investigations is based on sophisticated co-culture procedures requiring the collection of large volumes of blood. The cell-associated DNA assay, while much easier and simpler to implement, differentiated neither pre-integration DNA complex from integrated proviral DNA, nor replication-competent from defective viral DNA. As unintegrated DNA has a short half-life , its interference with the assessment of the second phase decay rate should be minimal in a population with very low plasma viremia. Cell-associated DNA was measured in lymphomononuclear cells without differentiating between their lineage and activation state, in contrast to previous investigations focusing on the pool of long-lived infected resting CD4 T [11,12]. While we have reported results in relation to the concentration of CD4 T cells, the main target of HIV-1 , we cannot exclude that a proportion of viral DNA was derived from other cell types. Despite differences in methodology, we found an average DNA half-life similar to that reported previously in patients in whom HAART was initiated during primary HIV-1 infection [12,20]. The mean 6.6 months half-life of infected cells measured in this study corresponds to that reported for uninfected CD4 T cells . As our measure of DNA half-life included both cells with short half-life (acutely infected cells, infected monocytes) and cells with long half-life (infected resting CD4 T cells, cells containing defective proviral DNA), the relatively short mean half-life of cell-associated DNA reported here does not exclude the persistence of de novo cell infections.
Hence, our data provide two lines of evidence supporting the persistence of de novo cell infection in patients with primary HIV-1 infection taking HAART and with very low or undetectable plasma viremia. First, the very large interindividual variation in the second phase of cell-associated DNA decay rate suggests that cell re-infection persists over time in most patients despite initiation of HAART at the time of acute HIV-1 infection. Second, the high correlation between the second-phase DNA decay rate and the mean level of cell-associated RNA 6 to 33 months after treatment initiation identifies persistent low level of viral replication as the likely culprit responsible for the slow clearance of cell-associated DNA. This argument is further supported by the faster DNA decay rate observed in the five patients who had undetectable cell-associated RNA in lymph nodes and in a majority of PBMC samples. Other investigations have also documented low levels of viral replication in patients on HAART [6,7,22–25]. Our results extend previous observations showing that decay of long-lived infected resting CD4 T cells is correlated with ‘bumps’ of plasma viremia . As most of our patients had undetectable plasma viremia, the data presented here provide, through the measurement of cell-associated RNA, a marker of higher sensitivity.
New treatments provide opportunities to achieve even greater inhibition of viral replication, with most patients achieving undetectable plasma viremia [26,27]. In these circumstances, treatment efficacy can be evaluated further by the assessment in blood of cell-associated HIV-1 RNA and DNA.
We thank K. Zollinger for excellent technical help and Dr S. Kwok (Roche Molecular Systems, Alameda, California, USA) for providing internal quantification standards for cell-associated DNA assay.
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Keywords:© 2000 Lippincott Williams & Wilkins, Inc.
primary HIV infection; antiretroviral therapy; viral decay; reservoir; lymph nodes