Early and highly active antiretroviral therapy (HAART) has been widely used over recent years in patients with primary HIV infection (PHI) in an attempt to control viraemia rapidly, subsequently improve the long-term outcome of the disease and eventually eradicate HIV from the infected organism. However, it has now been demonstrated that a reservoir of latently infected resting CD4 T cells is established early after contamination, both in the lymph nodes and peripheral blood, and can persist for months in patients with undetectable plasma viraemia under HAART [1–5]. Therefore, it might be of interest to evaluate viral replication in peripheral blood mononuclear cells (PBMC) and lymphoid tissue. The quantification of cell-associated proviral HIV-1 DNA might provide an opportunity to assess the capacity of HIV replication of latently infected cells.
In 1996, the Agence Nationale de Recherche sur le SIDA (ANRS) 053 trial, was initiated to assess the efficacy of HAART given early and for at least 18 months, to patients with symptomatic PHI . The present study was aimed at quantifying HIV-1 RNA and DNA both in PBMC and lymph node mononuclear cells (LNMC) in those of the patients enrolled in the ANRS 053 trial whose plasma and PBMC HIV-1-RNA loads were undetectable after 18 months of HAART. More specifically, we wanted to ascertain the decrease of cell-associated HIV-1 DNA and to determine the potential usefulness of cell-associated HIV-DNA monitoring in the follow-up of HAART-treated HIV-infected patients.
Patients and methods
From March 1996 to September 1997, 64 patients were enrolled in the ANRS 053 trial. The methods and results of this trial have been described previously . In brief, patients were enrolled if they presented with symptomatic PHI within the 4 weeks before inclusion and had evidence of recent seroconversion at the time of inclusion.
HAART consisted of the combination of zidovudine 500 mg/day, lamivudine 300 mg/day, and ritonavir 1200 mg/day after a titration period of 7–14 days, all given twice a day. Patients who were intolerant to ritonavir were offered to switch to indinavir 2400 mg/day, and remained in the study. The ad hoc Ethics Committee approved the therapeutic trial and patients gave their written informed consent before enrolment.
In patients with both plasma HIV-1-RNA levels less than 50 copies/ml and PBMC HIV-1-RNA levels less than 50 copies/106 cells at month 18, HIV-1 DNA was quantified in PBMC sampled at baseline and then every 6 months. These patients were also offered to undergo excision lymph node biopsy between months 18 and 21 for HIV-1-RNA and -DNA quantification in LNMC.
HIV-1 quantification techniques
In plasma, HIV-1 RNA was measured using the Amplicor HIV-1 Monitor 1.5 Test (Roche Diagnostic Systems, Neuilly sur Seine, France) according to the manufacturer's instructions, and with a limit of quantification of 50 copies/ml.
PBMC were isolated by performing a Ficoll gradient (MSL; Eurobio, Les Ulis, France) and were stored at −80°C. For HIV-1-RNA quantification, samples were incubated for 1 h at 37°C with DNase I- RNase-free and extraction was performed following the manufacturer's instructions. For HIV-1-DNA quantification, extraction was achieved with the Whole Blood Specimen Preparation Kit (Roche Diagnostic Systems). After extraction using an input of 1–3 million cells, HIV-1 RNA and DNA were quantified using the Amplicor HIV-1 Monitor 1.5 Test. For DNA, an internal quantitative standard (courtesy of S. Kwok, Roche Molecular Systems, Alameda, CA, USA) was used along with the commercial kit.
Lymph nodes were minced with a scalpel and the cells teased out in RPMI-1640 (Life Technologies, Cergy-Pontoise, France). After the Ficoll gradient and storage at −80°C of the 1–3 million LNMC pellets, HIV-1 RNA and DNA were quantified using the same method as for PBMC. Lower limits of quantification were 50 copies/106 cells and 10 copies/106 cells for HIV-1 RNA and DNA, respectively, in PBMC as well as in LNMC. The samples that showed a positive quantification signal below the limit of quantification were considered to contain detectable RNA or DNA.
Analysis of variance for repeated measurements was performed to compare viral loads within the same compartment at different times. Pearson correlation coefficients were calculated between plasma HIV-1 RNA and cellular HIV-1-RNA and -DNA titres at baseline and at month 18. P values less than 0.05 were regarded as statistically significant.
Eighteen months after enrolment, 45 of the 64 patients enrolled were still receiving HAART, and 27 (25 men and two women; median age 33 years, range 21–53) had plasma HIV-1-RNA levels less than 50 copies/ml for a median time of 12 months (range 6–15); 25 patients also had PBMC HIV-1-RNA levels less than 50 copies/106 cells. Fourteen patients accepted excision lymph node biopsy, including two patients with hardly detectable PBMC HIV-1 RNA at month 18 (50 and 58 copies/106 cells); biopsy was nevertheless performed on the basis of the previous result (< 50 copies/106 cells at month 12). Lymph node samples were suitable for HIV-1 quantification in 11 patients out of 14.
The median time between the first PHI symptom and the start of treatment was 24 days (range 11–55). At baseline, median CD4 and CD8 lymphocytes counts were 607/μl (range 207–1542) and 1213/μl (range 560–4884), respectively.
The results of the quantification of HIV-1 RNA and DNA in plasma, PBMC and LNMC are displayed in Table 1.
The median decrease in plasma HIV-1 RNA was −3.6 log/ml between baseline and month 18 (P < 0.005). The number of patients with undetectable PBMC HIV-1 RNA increased from four out of 21 (19%) at baseline to 23 out of 25 (92%) at month 18. The decrease in PBMC HIV-1-RNA levels was sharp during the first 6 months, and plateaued later on until month 18. HIV-1 RNA showed a median decrease of 1.1 log in PBMC over the 18 month follow-up (P < 0.05). Of the 11 patients tested for HIV-1 RNA in LNMC, four were below the limit of quantification (36.4%).
HIV-1 DNA was detected in PBMC at baseline in all but two patients (92.3%). At month 18, one patient still had undetectable PBMC HIV-1 DNA (after detectable loads at months 6 and 12), and there was no available sample for the other patient. HIV-1 DNA showed a median decrease of 1 log over the 18 month follow-up (P < 0.001) and was still detectable in 92.3% of patients at month 18. In LNMC, all available samples had detectable HIV-1-DNA loads and titres were above the lower limit of quantification. HIV-1-DNA titres in LNMC were two- to sixfold higher than in PBMC in four patients, identical in one and twofold lower in two.
In PBMC, HIV-1-DNA titres correlated significantly with HIV-1 RNA, both at baseline (r = 0.52, P < 0.02) and at month 18 (r = 0.46, P = 0.02). No other correlation was observed at any timepoint, either between PBMC HIV-1 DNA and plasma HIV-1 RNA, PBMC HIV-1 RNA and plasma HIV-1 RNA, LNMC HIV-1 RNA and PBMC HIV-1 RNA or between LNMC HIV-1 DNA and PBMC HIV-1 DNA.
We studied cell-associated HIV-1 RNA and HIV-1 DNA in an attempt to search for viral parameters that could be used routinely and could provide valuable information in patients with persistently undetectable plasma viral loads after prolonged HAART started at the time of PHI. Heavier methods that allow us to distinguish spliced from unspliced RNA and integrated from unintegrated DNA are not in favour for routine use. Moreover, different studies [5,7,8] did not conclude that these parameters were obviously more relevant in these patients.
In our population of newly infected patients with controlled viral replication, the decreases in viral RNA and DNA in PBMC were slower than those in plasma RNA, which were previously reported by Perrin et al. . After a progressive decrease over time in PBMC, HIV-1 DNA was found to be below the limit of quantification in only four out of 26 (15.4%) samples at month 18, whereas all LNMC samples tested scored positive for HIV-1 DNA. As observed in studies using a combination of zidovudine and didanosine in recently infected patients [4,9], DNA loads could be undetectable in PBMC but not in LNMC. In chronic HIV-1-infected patients [7,8,10–12], a similar slow and weak decrease was observed in PBMC DNA loads, without their reaching undetectable levels in either PBMC or LNMC when tested.
Even using cell-sorted CD4 lymphocytes, other authors [7,13,14] have also shown that HIV-1 proviral DNA decreased by more than 1 log after 1 year of treatment without reaching undetectable levels. Moreover, recent studies [4,5,7,9–11,15–17] have shown that replication-competent virus can still be demonstrated in residual latently infected resting CD4 T cells, despite a lack of detectable plasma viral activity. This is of serious concern because the reservoir serves as a potential source of reactivation of viral replication and remains a major obstacle for the eradication of HIV-1 in infected individuals receiving HAART [1–3]. Earliest and reinforced therapy may be tried out to control or decrease the size of this residual pool below a threshold at which the immune system alone could control it .
The utility of proviral HIV-1-DNA monitoring was not clearly demonstrated in this 18 month follow-up of HAART-treated primary-infected patients. However, this finding could be reconsidered when using other therapeutic strategies, such as structured treatment interruptions, reinforced treatment or additive immunotherapy, which could help to improve immune responses and flush out the latently infected cells [19–21]. Finally, the diffusion of drugs in the lymph nodes should be taken into account and studied to ensure the local efficacy of antiretroviral treatment.
The authors would like to thank the patients who participated in the study, Pascal Bonot and Didier Bachellerie for excellent technical assistance, Claudine Pardon (GlaxoWellcome, France) and all the clinical investigators: Hôpitaux de Bordeaux: J.M. Ragnaud, P. Morlat, D. Malvy, D. Lacoste, H.Dutronc, J.L. Pellegrin; Hôpital de Toulouse: P. Massip, B. Marchou, A. Bicart-See, M. Obadia, L. Cuzin, J. Puel; Hôpital de Nantes: F. Raffi, E. Billaud, V. Reliquet, S. Billaudel; Hôpital Cochin, Paris: D. Séréni, C. Lascoux, A. Krivine; Hôpital de Tours: P. Choutet, F. Bastides, J.M. Besnier, F. Barin; Hôpital de Besançon: H. Gil, C. Drobatcheff, Y. Bourezane, B. Hoen, A. Bassignot; Hôpital Bichat Claude Bernard, Paris: C. Leport, J.L. Vildé, F. Vachon, E. Bouvet, M.H. Prevot, A. Villemant, N. Meslem, A.G. Saimot, F. Brun-Vézinet; Hôpital de Grenoble: M. Micoud, O. Bouchard, P. Leclercq, B. Chanzy; Hôpital de Rennes: C. Arvieux, M.L. Lombart, F. Cartier, C. Michelet, A. Ruffault; Hôpital Saint Louis, Paris: D. Ponscarme, J.M. Molina, F. Timsit, P. Lesprit, J. Modaï, P. Morel, F. Ferchal; Hôpital de Versailles: A. Greder, M. Harzic; Hôpital de Nancy: T. Doco-Lecompte, Ph. Canton, A. Le Faou; Hôpital de Cavaillon: J.P. Igual, M. Carrière, F. Duluc; Hôpital de Mantes la Jolie: V. Leclerc; Hôpital de la Pitié Salpétrière, Paris: F. Bricaire, M.A. Valentin, J.M. Huraux; Hôpital d'Amiens: S. Redeker, A. Smaïl, G. Duverlie; Hôpital de Mâcon: J.P. Kisterman, F. Peronnet; Hôpital de Villejuif: C.Minozzi, D. Vittecoq, E. Dussaix; Hôpital de Dijon: M. Buisson, P. Chavanet, P. Pothier; Hôpital d'Angers: J.M. Chennebault, C. Payan; Hôpital de Nice: C. Sohn, P. Cassuto, O. Bernard; Hôpital de Saint-Antoine, Paris: C. Gentil, J. Frottier, L. Morand-Joubert; Hôpital de Tourcoing: M. Valette, Y. Mouton, P. Wattre; Hôpital de Montpellier: V. Baillat, J. Reynes, M. Segondy. The first two authors contributed equally to this study.
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