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Clinical Science

Highly active antiretroviral therapy corrects hematopoiesis in HIV-1 infected patients: interest for peripheral blood stem cell-based gene therapy

Baillou, Claude; Simon, Anne; Leclercq, Virginie; Azar, Nabih; Rosenzwajg, Michele; Herson, Serge; Klatzmann, David; Lemoine, François M

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Author Information

From the Biologie et Thérapeutique des Pathologies Immunitaires, UMR CNRS 7087 C.E.R.V.I. and Internal Medicine Department, UPMC, Hospital Pitié Salpêtrière, Paris, France.

Correspondence to F. M. Lemoine, Biologie et Thérapeutique des Pathologies Immunitaires, UMR CNRS 7087-C.E.R.V.I.-UPMC-Groupe hospitalier Pitié Salpêtrière, 83, Boulevard de l'Hôpital, 75651 Paris Cedex 13, France.

Received: 9 August 2002; revised: 25 October 2002; accepted: 5 November 2002.

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Objectives: To study, in asymptomatic HIV-1-infected (HIV+) patients, whether peripheral blood hematopoietic progenitor/stem cells (PBPC) mobilized by granulocyte colony stimulating factor (G-CSF), can be used as a source of cells for retroviral gene therapy.

Design: PBPC from two groups of HIV+ patients (treated or untreated by highly active antiretroviral therapy) and from seronegative donors were mobilized with G-CSF.

Methods: PBPC collected by leukapheresis were enriched for CD34 cells, immunophenotypically and functionally characterized, cultured and infected with retroviral vectors. HIV proviral integration was studied on fresh and cultured cells.

Results: G-CSF moderately and transiently increased the viral load in untreated patients only, and induced in both groups of HIV+ patients mobilization of percentages and numbers of CD34 cells comparable to those of seronegative volunteers. The most immature CD34 cell subset, the clonogenic progenitor and long-term culture initiating cells were significantly decreased in leukapheresis products and CD34-enriched fractions from untreated HIV+ patients but not in those from treated HIV+ patients. Cell cycle activation and growth factor responses of CD34 cells from both groups of HIV+ patients were not different from those of the control group. Culture and retroviral infection of CD34 cells from HIV+ patients did not enhance HIV replication, and yielded transduction levels similar to those obtained using CD34 cells from seronegative donors.

Conclusions: G-CSF-mobilized PBPC can be safely used for HIV retroviral gene therapy in asymptomatic treated patients while highly active antiretroviral therapy would control the G-CSF-induced increase in viral load and correct the defective hematopoiesis observed in untreated patients, without inhibiting the retroviral transduction of PBPC.

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Highly active antiretroviral therapy (HAART) for HIV infection is widely beneficial to HIV-positive (HIV+) patients whatever the clinical status of the disease [1]. However, HAART is a cumbersome treatment for the patients and is also responsible for numerous side-effects [2,3]. Over 30% of patients may escape the long-term efficacy of HAART because of lack of compliance and the development of viral drug resistance [2,4]. Thus, it may be important to envisage alternative therapeutic approaches, such as gene therapy, that may offer to patients the possibility of stopping or reducing HAART and that will provide a modality for long-term control of the disease [5–7]. Because HIV gene therapy will require durable and stable expression of the therapeutic gene, this strategy will be based mainly on the use of retroviral vectors that can integrate the gene of interest into the genome of host cells. Hematopoietic stem cells (HSC), because of their high proliferative properties and ability to reconstitute both the hematopoietic and immune systems, represent ideal target cells for HIV gene therapy in as much as their retroviral transduction using oncoviral or lentiviral vectors has now been achieved [8–13]. Furthermore, HSC-based gene therapy using retroviral vectors has recently been carried out for inherited immunodeficiencies with indubitable proof of efficacy [14]. For HIV gene therapy, one can envision using either, in a limited number of cases, HSC obtained from allogeneic seronegative donors or, in most cases, autologous HSC. In the latter case, it would be important to demonstrate that HSC from HIV+ patients harvested either from bone marrow or, more easily, from peripheral blood after granulocyte colony stimulating factor (G-CSF) mobilization are not functionally defective or infected by HIV. Indeed, conflicting results have been published concerning the infection and the diminution of the most primitive hematopoietic compartment by HIV [15–18].

For these reasons, it appears essential to answer the following questions with a view to develop a gene therapy strategy based on the use of autologous HSC collected from the peripheral blood of HIV+ patients after G-CSF mobilization: (i) is G-CSF administration to HIV+ patients a reasonable and innocuous procedure to mobilize peripheral blood hematopoietic progenitor/stem cells (PBPC)? (ii) can we safely harvest sufficient numbers of PBPC from HIV+ patients? (iii) are the PBPC in leukapheresis products (LP) and CD34 cell-enriched fractions from HIV+ patients immunophenotypically and functionally different from those of normal donors? (iv) can we detect HIV proviral integration in PBPC? (v) is it possible to manipulate PBPC in vitro without favouring HIV replication? (vi) can we transduce with retrovirus PBPC of HIV+ patients at levels comparable to those possible with PBPB of seronegative donors, whatever the therapeutic status (treated or not by HAART) of the patients? Thus, we designed a pre-clinical trial consisting of mobilizing PBPC with G-CSF in two groups of asymptomatic HIV+ patients untreated (n = 6) or treated (n = 10) by HAART. Leukapheresis was performed and PBPC enriched by positive CD34 selection using a clinical device. The PBPC present in LP and CD34 cell-enriched fractions were studied by flow-cytometry and by short- and long-term culture assays. Furthermore, growth factor responses and cell cycle activation of CD34 cells as well as their susceptibility to transduction by retrovirus were evaluated. Analysis of HIV proviral integration was also performed by semi-quantitative PCR on fresh cell suspensions and cultured cells. Results were compared with those from a control group of seronegative donors.

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Materials and methods

Patients and controls

Fifteen male HIV+ patients were recruited in the Internal Medicine department between March 1997 and June 2001. Written informed consent was obtained from all study participants as approved by the local institutional board and the French National Agency for AIDS Research. Study participants had confirmed HIV type 1 infection, were asymptomatic and either untreated (n = 6) or treated with HAART (n = 10), one patient belonged to both groups (No. 2 and No. 7). No significant weight differences were observed between the two groups of patients. Patients with anaemia (haemoglobin < 100g/l) or with thrombocytopaenia (< 1 × 1011/l), patients receiving antibiotics, prophylaxis of opportunistic infections or aspirin, patients with hepatitis B or C or cytomegalovirus viraemia were excluded from the study. Clinical status of the patients is presented in Table 1. Six healthy seronegative volunteers were studied as controls.

Table 1
Table 1
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Mobilization regimen and leukapheresis procedure

Patients received 10 μg/kg per day of G-CSF (Neupogen, Roche, Meylan, France) subcutaneously for 5 days according to procedures used in both autologous and allogeneic grafts. Only mild side-effects such as bone pain and myalgia were noted and easily controlled by standard doses of analgesic drugs. Mononuclear cells were collected from 8–10 l blood using the Fenwal CS-3000 Plus Blood Cell Separator (Baxter, Maurepas, France). Leukapheresis duration varied from 2 to 3 h and the mean leukapheresis volume was 54 ± 0.7 ml. Harvested cells were then transported to the laboratory and used for in vitro studies.

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CD34 cell purification

CD34 cells were enriched from LP using the immunomagnetic Isolex 300i system (Baxter) according to the manufacturer's instructions and as described previously [19]. Then, CD34 cell enriched fractions were washed and counted (cell counts were adjusted for dead cells as revealed by Trypan blue staining). The purity of CD34 cells was determined by flow cytometry. All of the following studies, except retroviral transduction, were carried out with fresh CD34 cells.

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Immunophenotyping of CD34 subsets

CD34 cell subsets were studied by two- or three-colour flow cytometry after staining of either 1 × 106 cells harvested from LP or 1 × 105 cells harvested from the CD34-enriched fraction or from cultured CD34 cells (see below) using appropriate directly-conjugated monoclonal antibodies (mAb). The following murine anti-human mAb directly conjugated to fluorescein isothiocyanate (FITC), phycoerythrin (PE) or phycoerythrin-cyanine-5 (PECy5) were used: CD4–FITC, CD10–FITC, CD7–PE, CD19–PE, CD45–PE and CD34–PECy5 (Beckman-Coulter, Villepinte, France), CD38–PE and HLA-DR-PE from (Becton Dickinson, Le Pont de Claix, France) and CD90-FITC (Thy-1) from Pharmingen, San Diego, California, USA). Negative controls (Beckman-Coulter) were appropriate irrelevant isotype-matched mAb. After staining, cells were fixed in phosphate-buffered saline containing 4% paraformaldehyde, maintained at +4°C for 24 h and then analysed using an EPICS Elite flow-cytometer (Beckman-Coulter); the proportion of each subsets was determined by using EPICS Elite software.

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Liquid cultures and cell cycle analysis of CD34 cells

Enriched CD34 cells (1 × 106 cells/2 ml) were cultured for 1–3 days at 37°C in X-vivo 10 medium (Biowhittaker, Emerainville, France) containing 10% foetal calf serum and the following recombinant human growth factors: interleukin-3, (100 U/ml; Genzyme, Cambridge, Massachusetts, USA), interleukin-6 (100 U/ml; a gift from L. Aarden, Amsterdam, the Netherlands), stem cell factor (300 ng/ml; a gift from Amgen, Thousands Oaks, California, USA), Insulin-like growth factor I (50 ng/ml; Valbiotech, Paris, France), basic fibroblast growth factor (50 ng/ml; Pepro Tech Inc, London, UK) and Flt3-ligand (300 ng/ml; a gift from Immunex, Seattle, Washington, USA). We have previously shown that these culture conditions can strongly favour the cycling of CD34 cells and their retroviral infection [10,20].

Cell cycle analysis was performed on CD34 cells just after purification and after 1–3 days of liquid culture using Coulter Reagents Kit (Beckman-Coulter) that contains both reagents for cell lysing and permeabilizing (DNA-Prep LPR) and reagents for DNA staining with propidium iodide (DNA-Prep Stain) as described elsewhere [20].

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Short-term culture assay

For colony forming cell (CFC) assays 2.5 × 104 cells from the bulk LP, 5 × 102 cells from the CD34-enriched fraction or from CD34-cultured cells were plated in methylcellulose duplicate cultures as described [21]. Burst forming unit erythroid (BFU-E) cells, colony forming unit granulocyte-macrophages (CFU-GM) and colony forming unit-macrophages (CFU-M) were counted under an inverted microscope between culture days 12 and 14.

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Long-term culture initiating cell (LTC-IC) assay

For LTC-IC assays, 2 × 104 cells from bulk LP or 240 cells from the CD34-enriched fraction were seeded in 96-well tissue culture plates (Costar supplied by D. Dutscher, Brumath, France) containing LTC medium (StemCell Technologies, Vancouver, British Columbia, Canada) and a pre-established feeder layer of 40 Gy-irradiated MS-5 cells [22]. After 5 weeks of co-culture, non-adherent and adherent cells obtained after trypsinization in each well were pooled and assayed for their CFC content as described above. The colonies obtained in these secondary methylcellulose cultures are the product of LTC-IC [23].

The frequency of LTC-IC was assessed by limiting dilution analysis using a described method [24]. Briefly, bulk LP or CD34-enriched fraction were seeded on MS5 cells as above at cell concentrations varying from 2.5 × 102 to 5 × 104 and from 2.5 to 960 in 100 μl of LTC medium, respectively. Thirty wells were prepared for each dilution. Wells were maintained at 37°C in a 5% CO2 atmosphere and fed weekly by changing half of the medium. After 5 weeks of culture, the supernatant from individual wells was harvested and adherent cells were overlaid by 100 μl of methylcellulose culture medium in order to determine the CFC content [21]. The frequency of LTC-IC was then calculated by Poisson statistic analysis as described [25].

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Retroviral transduction of CD34 cells
Packaging cell lines and retroviral supernatant

H293-GALV/Thy-1 1704 cells (1704 cells), a stable clonal human packaging cell line derived from human embryonic kidney 293 cells, produce GALV pseudotyped viral particles (5 × 105–106 particles/ml) containing the human Thy-1 (CD90) reporter gene (unpublished data). H293-GALV 1166 cells (1166 cells) used for mock-infection are similar to 1704 cells except that these cells produce GALV pseudotyped viral particles devoid of retroviral vectors. Viral supernatants were collected as described elsewhere [10].

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Transduction protocol

Because in preliminary experiments we did not observe differences in the cycling activation of fresh or frozen CD34 cells, the transduction protocol was performed using aliquots of frozen CD34 cells. After thawing, CD34 cells were cultured for 48 h, as described above, in order to activate them into cycling. Then, CD34 cells (1 × 105 cells/ml) were infected or mock-infected for 48 h with supernatant harvested from either 1704 cells or 1166 cells using a previously published protocol [10].

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Transduction efficiency analysis

Twenty-four h after infection or mock-infection, cells were counted and stained either with CD34–PECy5 and CD90–PE (Pharmingen)-conjugated mAb or with irrelevant isotype-matched mAb as negative controls. The percentage of CD34 cells expressing the Thy-1 reporter gene was determined by flow-cytometric analysis.

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HIV-1 viral load and detection of HIV-1 provirus integration

For plasma viral load, plasma was separated from whole blood within 6 h of collection and stored at −80°C. Plasma was collected 2 weeks before mobilization, at the onset of G-CSF treatment, and at days 5, 13, 20, 30, 60 and 90 following G-CSF administration. HIV-1 RNA was measured with a quantitative RNA reverse transcription (RT)–PCR) according to the manufacturer's instructions (Amplicor HIV-1 Monitor assay, Roche Diagnostics, Branchburg, New Jersey, USA). Each sample was tested in duplicate, the lower limit of detection was 200 RNA copies/ml.

Detection of HIV-1 provirus integration was performed on fresh and cultured cell suspensions and on colonies from methylcellulose cultures. In the latter case, colonies were plucked with a micropipet and pooled by groups of 10 by category, i.e., BFU-E, CFU-GM or CFU-M. Cells were first washed, lysed in lysis buffer, digested with 100 μg/ml proteinase K (Boehringer, Mannheim, Germany) at 56°C for 1 h and then nested PCR and semi-quantitative nested PCR were performed using P3, P4, P5 and P6 Pol primers as described [26,27]. Relative virus amounts in different samples were estimated by endpoint dilution of lysates of 1 × 106/ml HIV-1-infected cells in lysates of 1 × 106/ml HIV-1 negative A301 T-cell line. Serial dilutions of HIV-1 infected 8E5/LAV cells (1 virus copy/cell) in parental A301 cell lysates showed that the assay could detect 1 HIV DNA copy per 3 × 104 unfixed cells and 10–100 HIV DNA copies per 3 × 104 4% paraformaldehyde-fixed cells. PCR was also performed with β-actin primers or β-globin primers as amplification (5 min at 92°C, followed by 35 cycles of 1 min at 92°C, 1 min at 58°C and 1 min at 72°C) and DNA content control [27]. Amplified products (20 μl) and molecular weight markers were loaded onto 2% agarose and stained with ethidium bromide for UV visualization of the 233 base pair (bp; P5/P6) pol, 358 bp β-actin and 268 bp β-globin PCR products.

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Statistical analysis

When data were compared between groups of HIV+ patients (treated versus non-treated) and the control group, statistical analyses were performed using the unpaired Student's t test. Comparisons of data collected from the same patients were analysed by the paired Student's t test. Statistical significance was taken at the 5% level (P < 0.05).

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Effects of G-CSF administration on granulocytes and CD4 T-cell counts

To evaluate the stimulating effect of G-CSF, granulocyte and CD4 T-cell counts were studied at days 0, 5, 13, 20, 30 and 60 following G-CSF administration (day 0 corresponds to the beginning of treatment). Results indicate that 5 days of G-CSF treatment significantly increased by 9 ± 1-fold (P < 0.0001, paired t test) and by 2 ± 0.2-fold (P < 0.001, paired t test) the granulocyte and CD4 T-cell counts, respectively. These cell counts returned to baseline levels as early as day 13. The effects of G-CSF on both granulocytes and CD4 T-cell counts were not significantly different between the two groups of patients and were superimposable on those obtained with seronegative donors [28,29]. These data indicate that G-CSF can efficiently mobilize granulocytes and, to a lesser extent, CD4 T cells in asymptomatic HIV+ patients.

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Effect of G-CSF administration on HIV-1 viral load

To study any effect of G-CSF on HIV replication, plasma viral load was measured by quantitative RT–PCR at different time points before and after its administration. Surprisingly, the viral load significantly increased 1.8–5-fold (P ≤ 0.02, paired t test) on day 5 post G-CSF in the six non-treated patients (Fig. 1). In all cases, viral load returned to baseline level as early as day 13 and remained stable until at least day 90. In the 10 treated patients, no effect of G-CSF was observed, the viral load remaining under the threshold of detection (< 200 copies/ml). These findings indicate that G-CSF in the absence of antiretroviral therapy transiently increases the viral load, such a side-effect being controlled by HAART.

Fig. 1
Fig. 1
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Percentage and total number of CD34 cells in leukapheresis products and CD34 cell-enriched fractions from HIV+ patients

The percentage of CD34 cells in the different LP and in the CD34 cell-enriched fractions obtained from non-treated and treated patients after 5 days of G-CSF treatment was compared to a control group of normal donors receiving the same mobilization regimen. Table 2 shows that the mean percentages of CD34 cells in LP and CD34-enriched fractions from the non-treated group, the treated group and the control group were 1.6 ± 0.15%, 1.8 ± 0.4%, 1.5 ± 0.35% and 83 ± 3%, 85 ± 3%, 90 ± 2%, respectively. Statistical analysis revealed no significant differences (P > 0.05; unpaired t test) between the groups of patients. A similar observation was made when total numbers of CD34 cells were compared. These data indicate that the percentage and absolute number of CD34 cells in LP as well as after immunomagnetic selection were comparable in both groups of patients and not different from that of normal volunteers. Ours findings indicate that G-CSF mobilizes CD34 cells equally well in asymptomatic HIV+ treated or non-treated patients as in seronegative volunteers.

Table 2
Table 2
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Immunophenotypic analysis of CD34 cell subsets in leukapheresis products and CD34-enriched fractions from HIV+ patients

Cells from LP or from CD34 cell-enriched fractions were stained with various mAb and analysed by flow cytometry to determine the percentages of different CD34 cell subsets such as CD34+/CD38 cells, CD34+/CD90+ cells or CD34+/HLA-DRlow cells that contain the most primitive progenitors and CD34+/CD10+ cells, CD34+/CD7+ cells, CD34+/CD19+ cells that contain more specifically lymphoid-committed progenitors [21,30,31]. Fig. 2 shows that CD34+/CD38− cells, CD34/CD90 cells and CD34/HLA-DRlow cells within the CD34 cell-enriched fractions were significantly decreased (P < 0.05, unpaired t test) in non-treated relative to treated patients or to control subjects. In contrast, no significant differences (P > 0.05, unpaired t test) were observed for lymphoid committed CD34 cells or for those that expressed CD4. Interestingly, the distribution of the various CD34 cell subsets in treated patients did not differ from those in the control group. Similar results were noted when CD34 cell subsets from LP were analysed (data not shown). As CD34 cell percentages are similar in LP and CD34-enriched fractions for both HIV+ groups, these findings suggest a qualitative imbalance of the most primitive CD34 cell compartment in asymptomatic HIV+ non-treated patients that is not found in treated patients.

Fig. 2
Fig. 2
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Functional analysis of hematopoiesis in leukapheresis products and CD34-enriched fractions from HIV+ patients

The clonogenic progenitor (CFC) and LTC-IC content of LP and CD34-enriched fractions obtained from non-treated and treated patients were assessed by short- and long-term culture assays, respectively. The data in Table 3 indicate that CFC frequency in both LP and CD34 fractions from non-treated patients was reduced (P < 0.05, unpaired t test) relative to that in treated patients. Thus, cloning efficiencies (mean ± SEM) of CFC in CD34 fractions from non-treated and treated patients were 16.8 ± 0.7 % and 35.7 ± 2.3 %, respectively. When compared to the control group, it was found that the cloning efficiency of CFC from non-treated patients was reduced, whereas CFC cloning efficiency from treated patients was not different from that of normal donors. More precise analysis of CFC growth in the CD34-enriched fractions indicate that in non-treated patients BFU-E, CFU-GM and CFU-M were reduced in the same proportion (P < 0.05, unpaired t test) compared to treated patients or normal donors. Of note, no differences regarding the morphology and the size of CFC were observed between the two groups of patients and the control group. LTC-IC, which represent more primitive progenitors, were studied in both LP and CD34 fractions by long-term culture assay and their frequency was determined by limiting dilution analysis. The results (Table 3) showed that the LTC-IC content of both LP and CD34 fractions from non-treated patients was reduced (P < 0.05, unpaired t test) relative to treated patients or to the control group. By contrast, the LTC-IC frequency of treated patients was not statistically different from that of the control group. Thus, the mean LTC-IC frequency in the CD34 fractions from non-treated patients, treated patients and normal donors were 1/40, 1/23 and 1/24, respectively. Taking into account these data, calculation of the total number of CFC and LTC-IC present in the CD34 fractions from non-treated patients compared with those from treated patients or normal donors was significantly (P < 0.05, unpaired t test) reduced by approximately three- and twofold, respectively.

Table 3
Table 3
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Overall, these data show a significant decrease of the CFC and LTC-IC content in LP from non-treated patients but not from treated patients. Although our study was really comparative for one patient only (No.2 and No.7), the results strongly suggest that defective hematopoiesis observed in non-treated patients can be corrected by HAART.

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Cell cycling analysis and growth factor responses of CD34 cells

One of the requirements for retroviral gene transfer is that target cells are dividing [32]. For this reason we studied the cell cycling activation and growth of CD34 cells in response to a combination of growth factors previously used to efficiently transduce CD34 cells, CFC and LTC-IC from normal donors [9,10,20]. We found that only 3.3 ± 1% and 3.5 ± 0.5% of CD34 cells from non-treated and treated patients, respectively, were in S+G2/M phase before stimulation. After 48 h culture, CD34 cells were highly cycling, 43 ± 3% and 47 ± 2% of them being in S+G2/M phase. As compared with the control group, the cell cycling activation of CD34 cells from non-treated or treated patients did not differ. Under these culture conditions, cell proliferation of CD34 cells from either non-treated or treated patients were superimposable and total numbers of cells were expanded by around fourfold at day 3. Overall, these data indicate that CD34 cells from asymptomatic HIV+ patients exhibit similar growth factor responses to those of CD34 cells from normal donors.

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Detection of HIV-1 provirus integration in LP, CD34-enriched fractions and progenitor cells

HIV-1 provirus was studied by PCR in cells from LP, CD34-enriched fractions and CD34 cell cultures. While HIV-1 provirus was positive in 13 of 16 and 9 of 16 LP and CD34 cell-enriched fractions, respectively (Table 4), the fraction of positive cells decreased in CD34-cultured cells. Indeed, only one out of 16 cultures was weakly positive after 3 days of culture. Interestingly, HIV-1 DNA detection did not appear to be less frequent in treated patients than in untreated patients. HIV-1 provirus was also studied in methylcellulose colonies derived from short- and long-term cultures. For each patient, between 10 and 15 pools of 10 CFC or 10 LTC-IC-derived colonies from LP, CD34 cell-enriched fractions or cultured CD34 cells were studied by PCR. Results from cultured CD34 cells showed that in almost 90% of patients none of the 100–150 colonies tested per patient were HIV-1 positive (Table 4). However, positive detection in CFC and LTC-IC pooled colonies (at least one positive colony in the pool) was observed in 12.5% and 7.5 % of patients, respectively. In fact, frequency of positivity corresponds to 1–2 pool(s) out of 10–15 pools tested. To determine more precisely the frequency of infected progenitors in positively detected patients, 10–20 individual colonies of each category were analysed for HIV-1 provirus, visualization of β-actin signal allowing assessment of the presence of genomic DNA. Interestingly, none of the individual colonies were positive except for patient No.8 (see below), strongly suggesting that detection of HIV-1 provirus in pooled colonies might be due to the presence of few contaminating long-term surviving HIV-positive cells as we used a very sensitive PCR technique. For patient No.8, where eight out of 10 pools of 10 colonies were negative, we found that approximately 13% of the individual colonies tested within the two remaining positive pools of colonies were positive for HIV-1 provirus. This led for this patient, who was the oldest HIV+ patient in the study, to a frequency of positive detection of approximately 2.7%. Taking into account that HIV-1 provirus in hematopoietic progenitors was positively detected for one out of 16 patients, this would indicate that HIV-1 infection would occur in less than 0.2% of hematopoietic progenitors, if any. Furthermore, our findings clearly show that our culture conditions do not promote HIV-1 replication.

Table 4
Table 4
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Retroviral transduction of CD34 cells from HIV+ non-treated and treated patients

We investigated whether CD34 cells from either treated or non-treated patients could be infected and transduced using defective retroviral vectors carrying the human thy-1 reporter gene. Fig. 3 shows a representative FACS analysis of transduced CD34 cells from one HIV+ patients and one control patient. Overall, retroviral transduction efficiency of CD34 cells from three non-treated patients, seven treated patients and three control donors were 41.5 ± 5.3%, 41.2 ± 2.7% and 39.7 ± 5.4%, respectively. These results indicate that retroviral transduction of CD34 cells from asymptomatic HIV+ patients can be achieved at levels that are as good as those obtained with CD34 cells from seronegative donors, even when patients have been treated with HAART. Furthemore, high expression of the thy-1 reporter gene, as shown in the Fig. 3, allows further enrichment of transduced cells by immunomagnetic beads (unpublished data).

Fig. 3
Fig. 3
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Despite the large progress of HAART on HIV clinical course, HSC-based gene therapy for HIV infection still remains an important alternative or complementary therapeutic approach. Autologous G-CSF-mobilized PBPC collected by leukapheresis and then enriched for CD34 cells appear to be the most convenient source of HSC in HIV+ patients. Whether HIV gene therapy should be envisaged in either HIV+ untreated or treated patients is also debatable. For these reasons we studied G-CSF mobilization of PBPC from asymptomatic HIV+ patients treated or not by HAART. First, we found in both groups of patients that G-CSF administration during 5 days led to an important increase in granulocyte counts at levels comparable to those obtained in seronegative volunteers. We also observed a significant increase in the CD4+ T-cell counts. Furthermore we detected a significant increase in the viral load in non-treated patients only. Mobilization of CD34 cells from HIV+ patients using similar doses of G-CSF has already been reported by others and our data concerning the effects of G-CSF on granulocyte and CD4 T-cell counts are in line with most of these reports [33–36]. However, increase of the viral load induced by G-CSF was not generally mentioned [33,35,36] except by another group [37]. Thus, we and others have observed that G-CSF treatment transiently increases the viral load. Such apparent conflicting observations might be due to the sensitivity of the viral load detection method or to the clinical and therapeutic status of the patients studied. More interestingly, our data point out that in HAART-treated patients, G-CSF did not lead to detectable increased viral load even by using a very sensitive RT–PCR (≤ 200 RNA copies/ml). The mechanisms by which G-CSF might increase the viral load remain to be determined. Recently it has been shown that an essential step contributing to mobilization of CD34 cells by G-CSF is related to the cleavage of stromal cell vascular cell adhesion molecule-1 by neutrophil proteases released from neutrophils accumulating in the bone marrow after G-CSF treatment [38]. Then, one may suggest that G-CSF would also mobilize T cells by a similar mechanism resulting in augmented numbers of infected CD4 T cells in the blood and consequently in increased viral load. As such a side-effect of G-CSF on viral load appeared to be controlled by HAART, we first conclude that G-CSF can be safely used in treated patients.

Concerning mobilization of CD34 cells, we found that G-CSF was as efficient for HIV+ patients, whatever their therapeutic status, as for seronegative donors. It is admitted that a minimum of 2 × 106 CD34 cells/kg is necessary to reach the minimal number of PBPC required for prompt and sustained engraftment in autologous and allogeneic transplant settings [39]. In this report, the total amount of CD34 cells obtained in both groups of HIV+ patients was sufficient and not different from normal donors as reported by others [33,34,36,40]. However, because LP were contaminated by HIV-infected cells (see Table 4), it appeared mandatory to further enrich CD34 cells. Consequently, recovery of CD34 cells was dramatically reduced after the purification step, but was almost sufficient as more than 1 × 106 CD34 cells/kg were obtained (see Table 2). Hopefully, the required number of CD34 cells would theoretically be reached by performing a second round of leukapheresis.

Although quantitative analysis of mobilized CD34 cells showed no differences between the two groups of patients, qualitative studies of LP and CD34-enriched fractions showed significant defects of PBPC from non-treated patients. Few reports have studied the immunophenotype and function of G-CSF-mobilized CD34 cells from HIV+ patients. Surprisingly, no significant defects of PBPC, compared to seronegative donors, have been described [33,34,36,40]. In the present report, by carrying out a complete study of PBPC, i.e., analysis of CD34 cell subsets and of CFC and LTC-IC frequency in LP, we indirectly detected impairment of bone marrow hematopoiesis that seems to occur early in the clinical course of HIV infection. In a series of HIV+-treated and non-treated patients at a more advanced clinical stage, such loss of primitive hematopoietic progenitors in the bone marrow has already been reported [41]. Furthermore, using the SCID/Hu Thy/Liv murine model, Jenkins et al. [17] have clearly demonstrated that HIV can interrupt thymopoiesis and multilineage hematopoiesis. Thus, there are several pieces of evidence showing that HIV can induce an imbalance of primitive hematopoiesis even if there is no detectable anemia or thrombocytopenia.

In this report, we have also studied by PCR HIV-1 proviral integration in purified CD34 cells. HIV-1 provirus detection in these cells was not surprising as CD34 cells were not highly purified (purity ranged from 64 to 96%) and consequently contaminated by HIV-infected cells that progressively disappeared after a few days of liquid culture (see Table 4). Of particular interest was the analysis of the progeny of hematopoietic progenitors obtained after short- and long-term culture assays. Indeed, detection of HIV-1 provirus was found in few cases when pooled CFC and LTC-IC-derived colonies were analysed, and in one case only (patient No.8) when individual CFC and LTC-IC-derived colonies were studied. In the latter case, the frequency of HIV-positive colonies was estimated at approximately 2.7%. Thus, our data, in line that of with others [33,35,40], suggest that PBPC are not or are extremely rarely directly infected by HIV-1. In fact, our results reinforce the hypothesis that suppression of hematopoiesis during HIV infection relies on indirect mechanisms such as infection of regulatory cells, including T cells, macrophages and stromal cells or the inhibitory effects of HIV-1 structural and regulatory proteins [16]. Indeed, when we studied PBPC from asymptomatic treated patients, defective hematopoiesis was not found. Although our study was really comparative for one patient only, who studied before and after HAART (No.2 and No.7), our data strongly suggest that control of HIV-1 replication by HAART results in an improvement, even to a return at normal levels, of hematopoiesis of HIV+ patients. Other reports have also shown that HAART improves recovery of hematopoietic activity in the bone marrow [42] and blood [18,43] of HIV+ patients. Whether HAART acts directly in vivo on hematopoeitic progenitors is not known. However, it has been shown that protease inhibitors stimulate the formation of CFC derived from bone marrow CD34 cells in normal donors and HIV+ patients [44]. Nevertheless, despite efficient antiretroviral therapy, defective hematopoiesis seems to persist in HIV+ patients at a more advanced clinical stage [41] as well as in simian/human immunodeficiency virus-infected macaques [45]. Furthermore, replication-competent non-latent HIV-1 can be detected in circulating monocytes from patients on prolonged HAART [46,47]. The latter observation, according to our results, seems more in favor of a de novo cell infection by a reservoir of resting infected CD4 T cells rather than an active production of monocytes by latently infected CD34 cells. Nevertheless, one can conclude that HAART can correct HIV-induced suppression of G-CSF-mobilized PBPC at least in asymptomatic HIV+ patients. In view of developing an HIV gene therapy strategy based on retroviral transduction of PBPC, ex vivo manipulation of CD34 cells is necessary. In spite of the defect of hematopoiesis depicted in PBPC from HIV+-untreated patients we found, surprisingly, that CD34 cells from both groups of HIV+ patients could be activated into cycle and then retrovirally transduced at similar levels than CD34 cells from seronegative donors. Moreover, no increase of HIV-1 replication was detected in either CD34 cells or progenitor cells during their ex vivo manipulation (i.e., growth factor stimulation in liquid culture for 2–3 days). These encouraging data, in line with the findings of others [34,35,48], demonstrate the feasibility of retroviral gene transfer into CD34 cells and PBPC from HIV+ patients. In addition, the absence of decreased transduction efficiency in CD34 cells from HAART-treated patients indicates that antiretroviral drugs, whatever their combination (i.e., presence or not of protease inhibitors), do not block in vitro the infection of CD34 cells with defective recombinant retroviral vectors.

Overall, this pre-clinical trial shows that PBPC can be safely mobilized by G-CSF in asymptomatic treated patients, as HAART controls the G-CSF-induced increase in viral load and corrects the defective hematopoiesis observed in untreated patients. Because HAART does not preclude transduction of CD34 cells by retroviral vectors, PBPC-based HIV gene therapy can be envisaged in such patients.

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The authors thank J.C. Gluckman for critical reading of this manuscript, V. Bon Durand and C. Desmyter for expert technical assistance, M. Bonmarchand and A. Choquel for monitoring HIV patients, L. Aarden for his generous gift for interleukin-6, and AMGEN and Immunex for the gift for stem cell factor and Flt3-ligand, respectively.

Sponsorship: Supported by a clinical grant from the French National Agency for AIDS Research.

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HAART; hematopoiesis; HIV-1; G-CSF; CD34; retroviral gene therapy

© 2003 Lippincott Williams & Wilkins, Inc.


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