|Home | About | Journals | Submit | Contact Us | Français|
Immunodeficiency after peripheral blood progenitor cell (PBPC) transplantation may be influenced by graft composition, underlying disease, and/or pre-treatment. These factors are difficult to study independently in humans. Ex vivo culture and genetic manipulation of PBPC grafts may also affect immune reconstitution, with relevance to gene therapy applications. We directly compared the effects of three clinically relevant autologous graft compositions on immune reconstitution after myeloblative total body irradiation in rhesus macaques, the first time these studies have been performed in a large animal model with direct clinical relevance. Animals received CD34+ cell dose-matched grafts of either peripheral blood mononuclear cells, purified CD34+ PBPCs, or purified CD34+ PBPCs expanded in vitro and retrovirally transduced. We evaluated the reconstitution of T, B, natural killer, dendritic cells, and monocytes in blood and lymph nodes for up to 1 year post-transplantation. Animals receiving selected-transduced CD34+ cells had the fastest recovery of T-cell numbers, along with the highest T-cell-receptor gene rearrangement excision circles levels, the fewest proliferating Ki-67+ T-cells in the blood, and the best-preserved thymic architecture. Selected-transduced CD34+ cells may therefore repopulate the thymus more efficiently and promote a higher output of naïve T-cells. These results have implications for the design of gene therapy trials, as well as for the use of expanded PBPCs for improved T-cell immune reconstitution after transplantation.
Early after transplantation, all arms of the immune system are impaired, which can result in numerous infections and impaired antitumor activity in the allogenic setting . These complications primarily arise from an abnormal T-cell compartment, which correlates with thymic dysfunction [2, 3]. The T-cell compartment can be reconstituted through either a thymic-dependent or a thymic-independent pathway [4–6]. The thymic-dependent pathway results in de novo generation of naïve T-cells with a more diverse T-cell receptor (TCR) specificity repertoire, whereas the thymic independent pathway relies on peripheral expansion of transplanted mature T-cells  with a consequently more limited TCR repertoire.
Analysis of immune recovery in humans undergoing transplantation is complicated by many factors that affect their immune system besides the actual transplantation, particularly their age, underlying disease, and prior chemotherapy or radio-therapy exposure. Different types of progenitor cell grafts have been used for both autologous and allogenic transplantation, but their impact on immune reconstitution has not been fully characterized. Assessment of the impact of selected CD34+ cells versus unselected peripheral blood progenitor cells (PBPCs) on kinetics and quality of immune reconstitution post-transplantation has been difficult in human clinical trials due to patient heterogeneity, particularly regarding prior chemo-therapy. Ex vivo culture of CD34+ cells has been performed in a number of clinical trials to deplete tumor cells, expand progenitor cells, or transduce target cells with gene transfer vectors, but there is little information regarding the impact of these manipulations on any immune parameters besides basic hematologic engraftment [7–9].
Here, we used the rhesus macaque model to study immune reconstitution after myeloblative total body irradiation (TBI) followed by different clinically relevant graft regimens. In this study, the animals received autologous CD34+ cell dose-matched grafts of either peripheral blood mononuclear cells, purified CD34+ PBPCs, or purified CD34+ PBPCs that had been expanded in vitro and retrovirally transduced. We performed a detailed phenotypic analysis of cell recovery post-transplantation, monitored the levels of T-cell-receptor gene rearrangement excision circles (TRECs), and evaluated thymic architecture. This provides the most extensive and detailed data set available on immune reconstitution following autologous transplantation in primates, human or nonhuman, for the first time also including information on the impact of ex vivo culture and/or transduction on the behavior of the graft. Our results indicate significant differences in the kinetics and characteristics of immune recovery following these different types of grafts and suggest that ex vivo culture of purified CD34+ cells prior to transplantation may be beneficial for T-cell immune reconstitution, as well as allowing genetic modification to correct disorders of immunity and hematopoietic function.
All animals used in this study were colony-bred rhesus macaques (Macaca mulatta) of Indian origin maintained and used in accordance with guidelines of the National Institutes of Health Guide for the Care and Use of Laboratory Animals (Department of Health and Human Services [DHHS] publication no. NIH85−23). The protocol was approved by the Animal Care and Use Committee of National Heart, Lung, and Blood Institute, NIH/DHHS. Healthy animals of either sex at ages 3−5 years were selected (Table 1). These animals were free of known infectious or immunologic diseases.
Hematopoietic progenitor cells were mobilized with granulocyte-colony stimulating factor (G-CSF) (10 μg/kg; Amgen, Thousand Oaks, CA) in combination with stem cell factor (SCF) (200 μg/kg; Amgen) administered by subcutaneous injection for 4 days. Mobilized PBMCs were collected by leukapheresis and isolated using density gradient centrifugation. CD34+ cell enrichment was performed using the 12.8 immunoglobulin-M (IgM) anti-CD34 biotinylated antibody and MACS streptavidin microbeads (Miltenyi Biotec, Auburn, CA, http://www.miltenyibiotec.com). The purity of the MACS-sorted CD34+ cells was analyzed by flow cytometry. The cells were frozen in 90% fetal bovine serum (FBS) mixed with 10% dimethyl sulfoxide. The viability of the cells after thawing was >95% as measured by trypan blue staining.
The Moloney murine leukemia virus-derived retroviral vector G1Na containing the neo gene was used for transduction . The retroviral supernatant was produced and harvested as described . The biologic titer was between 2 × 105 and 5 × 105 biologically active vector particles per ml. For transduction, retroviral supernatant was harvested from subconfluent producer cells cultured for 12−18 hours in Dulbecco's modified Eagle's medium (DMEM) (Mediatech, Herndon, VA) supplemented with 10% FBS (Atlanta Biologicals, Norcross, GA, http://www.atlantabio.com), 4 mM l-glutamine, penicillin (50 mg/ml), and streptomycin (50 mg/ml) at 37°C in 5% CO2. Fresh vector supernatant was passed through a 0.22-μm filter (Milli-pore, Bedford, MA, http://www.millipore.com) to remove cellular debris before use . CD34+-enriched cells were cultured at a starting concentration of 2 × 105 cells per ml in the supernatant, supplemented with 100 ng/ml SCF, 100 ng/ml fms-like tyrosine kinase 3 ligand (Flt-3L), and 100 ng/ml megakarocyte growth and development factor (MGDF) (Amgen), in flasks coated with the CH-296 fragment of fibronectin (Retronectin; Takara, Shiga, Japan, http://www.takara.co.jp). Every 24 hours, nonadherent cells were harvested, spun down, resuspended in fresh vector supernatant and cytokines, and added back to the same fibronectin-coated flask. After 96 hours, cultured cells were harvested, first by removal of all nonadherent cells followed by application of 0.25% trypsin (Gibco, Grand Island, NY, http://www.invitrogen.com) to remove all remaining adherent cells from the flask. Cells were combined, washed, counted, and cryopreserved as described above.
The animals received 500 cGy TBI daily for 2 days, delivered at a rate of 8.8 cGy/min via a cobalt-60 teletherapy irradiator (Eldorado 8). The next day, cryopreserved autologous PBMCs, CD34+-selected PBPCs (purity 88%−93%), or CD34+ PBPCs that had been ex vivo-expanded and retrovirally transduced (purity 89%−93%) were reinfused. The CD34+ cell numbers infused into each animal was matched between the three groups (Table 1). For the animals receiving cultured and transduced cells, the starting CD34+ cell numbers placed into culture was matched to the other groups, not the cell numbers harvested at the end of 96 hours. The graft was thawed and re-infused via a central venous catheter. A day later, the animals were started on G-CSF at 5 μg/kg intravenously daily until the total white blood cell count reached 6,000 cells per μl. Hematopoietic recovery was monitored by daily complete blood counts.
At the end of culture and transduction, CD34-enriched cells were plated at concentrations of 500, 103, and 104 cells per ml and analyzed by colony-forming unit (CFU) assays using MethoCult M4230 methylcellulose medium (Stem Cell Technologies, Vancouver, BC, Canada, http://www.stemcell.com) supplemented with 5 U/ml erythropoietin (Amgen), 10 ng/ml GM-CSF (Sandoz, East Hanover, NJ), 10 ng/ml recombinant human interleukin (rhuIL)-3, (Sandoz), and 100 ng/ml rhuSCF (Amgen) at 37°C in 5% CO2. Colonies of more than 50 cells were counted on days 10−14, and 15 to 20 individual CFUs were plucked from the plates for polymerase chain reaction (PCR) analysis.
DNA was extracted from blood samples using phenol-chloroform . The primers and conditions used for neo and β-actin PCR have been described . In vitro transduction efficiency was estimated by nested PCR on DNA isolated from individual clonogenic progenitors . Engraftment with transduced cells was assessed by quantitative PCR (qPCR) on peripheral blood granulocyte and mononuclear cell DNA using an ABI Prism 7700 sequence detector (Applied BioSystems, Foster City, CA, http://www.appliedbiosystems.com) .
Blood was drawn and a lymph node was removed for baseline analysis prior to mobilization. All animals were followed for 1 year after transplantation (Fig. 1). Blood was drawn every month up to 9 months and thereafter at 12 months post-transplantation. Inguinal or axillary lymph nodes were removed at 1, 3, 6, and 12 months post-transplantation. Thirteen to 18 months after transplantation, the animals were euthanized and underwent necropsy to collect all lymphoid tissues.
PBMCs were isolated from citrated venous blood by density gradient sedimentation using Ficoll-Hypaque (Pharmacia, Uppsala, Sweden). Mononuclear cells were isolated from lymph node samples by gentle mechanical disruption of tissues in RPMI (HyClone, Logan, UT, http://www.hyclone.com) supplemented with 10% heat-inactivated FBS, 2 mM l-glutamine, penicillin, and streptomycin. The cells were passed through a 100 μm mesh filter to remove any remaining tissue fragments.
Cells (0.25−1.0 × 106) were resuspended in wash buffer (phosphate-buffered saline-0.5% bovine serum albumin) and incubated for 15−25 min at 4°C with appropriately titrated directly conjugated monoclonal antibodies (supplemental online Table 1). The antibodies that were not commercially conjugated were conjugated in the laboratory using standard protocols developed (http://drmr.com/abcon/index.html). Stained cells were washed and resuspended in 1% paraformaldehyde. For intracellular analysis of Ki-67 expression, cells were first stained for the cell surface markers CD4, CD8β, and CD95, before incubation in 2× fixation/permeabilization solution (BD Biosciences, San Jose, CA, http://www.bdbiosciences.com). Cells were then stained with Ki-67 Abs. Four-parameter flow cytometric analysis was performed on a FACSCalibur instrument (BD Biosciences), and data were analyzed using Flow Jo software (Tree-star Inc., San Carlos, CA).
CD4+ and CD8+ T-cells were stained as described above and immediately sorted on a modified FACSVantage SE/DiVa (Becton Dickinson) by gating on either CD4+ or CD8+ cells within the CD3+ cell population. This isolation procedure resulted in >98% pure populations of T-cell subsets. Cell pellets were frozen, and TREC levels were measured by qPCR directly on cell lysates as described . Real-time PCR was performed on the ABI Prism 7700 sequence detector (Applied BioSystems).
At necropsy, tissues from the following sites were harvested: tonsils, thymus, lymph nodes (submandibular, axillary, mediastinal, mesenteric, inguinal, iliac), spleen, Peyer's patches, and bone marrow. A portion of each tissue was snap frozen in OCT and kept at −80°C; the remaining tissue was fixed in 10% formalin and paraffin-embedded. Immunohistochemical staining with anti-CD3 (LabVision), anti-CD20, anti-high molecular weight keratin, anti-Ki-67 (all from DakoCytomation, Glostrup, Denmark, http://www.dakocytomation.com) were performed using Envision Plus detection system (DakoCytomation) with 3′-diaminobenzidine tetra hydrochloride as chromogen on a DakoAutomated immunostainer.
Statistical analysis was conducted with the program S-plus (StatSci Division, MathSoft Inc.). Simple two-group comparisons were conducted using t tests, using logarithmic or square root transformations as necessary. To examine differences in responses over time, linear models were fit to all measurements less than 160 days after transplantation that had different mean responses for the groups, and the correlations of measurements from the same animal were modeled using animal-specific random effects. These models were used to test for group differences in T-cell populations and for the percentage of T-cells that were Ki-67+.
Healthy rhesus macaques underwent mobilization of PBPCs with G-CSF and SCF followed by leukapheresis. They were conditioned with myeloablative TBI and transplanted with PBMCs, purified CD34+ PBPCs, or purified CD34+ PBPCs that had been retrovirally transduced with a vector containing the bacterial neomycin resistance gene (neo) and cultured for 4 days in the presence of Flt-3L, MDGF, SCF, and Retronectin (Fig. 1; Table 1). The number of CD34+ cells within each graft was adjusted to be the same for the three groups. For the animals receiving selected-transduced cells, the number of CD34+ cells placed into culture was adjusted. The total expansion during culture was 1.3−1.5-fold. The purity of the grafts for the CD34-selected and CD34-selected/transduced groups was 88% or higher, with less than 3% residual T-cells. The culture conditions used did not include any cytokines capable of supporting viability or proliferation of T-cells, and during similar cultures of rhesus CD34+ cells, we have found a 6.5 ± 2.6-fold decrease (n = 6) in T-cells by day 4 of culture.
All animals engrafted (leukocytes >1,000/μl, neutrophils >500/μl) between day 8 and day 14 post-transplantation. Up to 10% of circulating granulocytes and mononuclear cells contained the vector early after transplantation in the selected-transduced group. This level stabilized thereafter to a level of 0.1%−2%, comparable to levels found previously [14, 15]. One animal in this group required euthanasia 6 months after transplantation due to radiation-induced lung fibrosis.
The absolute numbers of total CD3+ T-cells in the animals combined from all groups prior to transplantation were 1,622 ± 234 CD3+ T-cells per μl of blood (mean ± SEM). One month after transplantation, all animals showed dramatic depletion of T-cells (Fig. 2A). Animals that received selected-transduced PBPCs had 424 ± 82 CD3+ T-cells per μl 1 month after transplantation, whereas animals receiving selected PBPCs or unselected leukocytes had 936 ± 533 T-cells per μl and 970 ± 337 T-cells per μl, respectively. Even though the selected-transduced group showed the lowest number of CD3+ T-cells 1 month after transplantation, they had the fastest recovery of CD3+ T-cells compared with the other two groups (Fig. 2A). Numbers of CD3+ T-cells on the square root scale from month 1 to month 5 suggest differences between the selected-transduced group and the other groups (one-way ANOVA p value = .09).
In all groups, there was a rapid expansion of CD8+ T-cells, whereas the recovery of CD4+ T-cells showed slower kinetics. This led to an inversion of the CD4+/CD8+ ratio found during the first 3 months after transplantation, as previously reported in humans [16, 17]. Detailed phenotyping of CD8+ and CD4+ T-cell subsets was performed. Optimal delineation of CD4+ naive and memory T-cells was achieved by identifying naive cells as a uniform CD95lowCD28high, β7 integrinint population as previously described  (Fig. 3A). Effector memory CD4+ T-cells were identified as CD95high β7 integrinint and CD28−, and the rest of the CD4+ T-cells were defined as central memory T-cells (Fig. 3A). Naïve CD8+ T-cells showed a homogeneous phenotype of CD95lowCD28intCD11alow, and the remaining CD8+ T-cells were defined as memory phenotype  (Fig. 3B).
As all animals studied were older juveniles at the time of study entry, naïve T-cells were the most prevalent T-cell phenotype both in the CD4+ and CD8+ T-cell populations prior to transplantation. Naïve, memory, and effector CD4+ T-cell counts were 722 ± 154 cells per μl, 260 ± 34 cells per μl, and 17 ± 4 cells per μl, respectively (mean ± SEM), whereas there were 251 ± 29 CD8+ naïve T-cells per μl and 110 ± 15 memory CD8+ T-cells per μl. After transplantation, there was a dramatic change in the ratio of naïve to memory T-cell pheno-type in all animals. One month after transplantation, memory T-cells were predominant in both the CD4+ and CD8+ T-cell populations (Fig. 2B–2F). The numbers of T-cells with a naïve phenotype increased gradually over time in all groups. The fastest recovery for both naïve CD4+ and CD8+ T-cells was observed in the animals that received a selected-transduced graft compared with the two other groups (p = .0364 for CD4 and p = .0356 for CD8) (Fig. 2B, 2D). There were no apparent differences in T-cell recovery between the sexes included in the respective groups. However, due to the limited numbers of subjects in each group, a solid evaluation of this is not possible.
To estimate thymic output, we measured TRECs within sorted CD4+ and CD8+ T-cell populations from all animals. Irrespective of the graft type, 1 month after transplantation, there were very low numbers of TRECs, with a median reduction of 22-fold compared with pretransplant levels in all animals combined (Fig. 4A, 4B). By 3 months post-transplant, TREC levels had increased by a median of 18-fold compared with the first month. Although at the majority of early time points post-transplantation (<7 months), the selected-transduced animals had the highest numbers of TRECs in CD4+ T-cells (Fig. 4A), only the 4-month time point reached a statistically significant difference (p = .0055). At 5−7 months, TREC levels in all groups were shown to exceed the TREC levels found at baseline, suggesting preferential colonization by recent thymic emigrants for naïve T-cell production.
To address this further, we examined the numbers of dividing peripheral blood T-cells by staining for the nuclear cell cycle-associated antigen Ki-67. The numbers of Ki-67+ CD4+ and Ki-67+ CD8+ T-cells were less than 10% prior to transplantation in all groups (Fig. 4C, 4D). At 1 month after transplantation, the mean Ki-67 expression levels in CD4+ and CD8+ T-cells were 37.8% and 33.9%, respectively, in the group that had received selected PBPC, 17.1% and 23.2% in the group that received an unselected graft, and 9.1% and 8% in the selected-transduced group. After the peak of Ki-67+ T-cells observed at 1−2 months in the selected and unselected groups, the numbers of Ki-67+ T-cells declined and reached low levels similar to those found in the selected-transduced group. No significant differences were found for numbers of Ki-67+ CD4+ T-cells between the groups (Fig. 4C) (p = .1099 using a random effects model). However, there were significantly fewer Ki-67+ CD8+ T-cells within the first 4 months of transplantation in the selected-transduced group compared with the two other groups combined (Fig. 4D) (p = .0356 using a random effects model). As expected, the vast majority of Ki-67+ T-cells had a memory phenotype. The lowest frequency of Ki-67+ T-cells together with the highest TREC levels observed in the selected-transduced group suggest that this group had greater thymic output of de novo-generated T-cells and thus less peripheral T-cell expansion.
Histological analysis was performed on tissues obtained at necropsy 13−18 months post-transplantation to evaluate the integrity of the lymphoid organs without having to disrupt their architecture. There were no discernible differences in the architecture or cellularity of lymph nodes, tonsils, spleen, bone marrow, or Peyer's patches between the groups. Lymph nodes showed follicular and paracortical hyperplasia in all animals, and the white pulp of the spleen was hyperplastic with prominent follicles.
Notable differences were observed in the thymus between the three groups (Fig. 5). In the selected-transduced group, all animals showed preserved lobular architecture and well-defined cortical and medullary areas (Fig. 5A, 5B). The thymus was composed of CD3+ T-cells, a high proportion of which were proliferating in the cortex as determined by Ki-67 staining, whereas no or few proliferating cells were noted in the medulla (data not shown). In the other two groups, the majority of the animals instead showed various degrees of thymic atrophy, that is, fat replacement, decreased thickness of the cortex, and cystic changes of the thymic epithelium (Fig. 5C–5F). These findings were particularly prominent in two out of three animals in the unselected group. In the latter group, the number of CD3+ T-cells was reduced, and so was the proliferative rate. The degree of thymic atrophy could not be assessed in one animal in the selected group that died of sepsis, with pleuritis and pericarditis that also incorporated the thymus 11 months after transplantation.
B-cells, defined in our study as CD20+ CD3− CD14− cells (supplemental online Fig. 1), recovered quickly within a 2-month time period after transplantation using any of the graft regimens (supplemental online Fig. 1). For all groups, we observed supranormal absolute B-cell numbers throughout the year post-transplantation, as described earlier in clinical studies [19, 20]. There were no statistical significant differences in the reconstitution pattern between the groups after transplantation.
We did not detect differences in NK-cell reconstitution after transplantation between the groups. Here, we defined NK-cells as either CD3− CD14− CD56+ or CD3− CD14− CD16+ (supplemental online Fig. 1). In our study, overall, the CD56+ NK-cell population was smaller than the CD16+ NK-cell population (p = .0017 using a t test on the baseline values, pooling all the data across groups; supplemental online Fig. 1).
We assessed the recovery of DCs and monocytes after transplantation. The two subsets of DCs were identified as CD11c+ CD123− myeloid DCs (MDCs) and CD11c− CD123+ plasma-cytoid DCs (PDCs), which are both HLA-DR+ and CD3−, CD14−, CD20− (Fig. 6G) Here, we found that CD11c+ MDCs were the most predominant subset in blood, which has been described previously  (Fig. 6A− 6F). The absolute numbers of circulating MDCs and PDCs in all the animals combined prior to transplantation were 70 ± 20 cells per μl of blood and 8 ± 2.4 cells per μl of blood, respectively. These levels were not found to be changed at 1 month after irradiation and transplantation in any of the groups of transplanted animals (Fig. 6). However, 2−4 months post-transplantation, an increase in the numbers of both MDCs and PDCs was observed. The animals that received selected-transduced PBPCs showed the highest peak value of both subsets of DCs. However, this was not found to be statistically different from the other groups.
The ratio of MDCs and PDCs was reversed in lymph nodes compared with blood. Irrespective of time point pre- or post-transplant, PDCs were the predominant DC subset in lymph nodes. At 1 month post-transplantation, there was an increase in proportions of both DC subsets in the lymph nodes compared with the levels found prior to transplantation in all groups (Fig. 6). The frequencies of DCs were found to return to values observed pre-transplantation 3−6 months post-transplantation (Fig. 6). CD14+ monocytes, both in blood and in lymph nodes, showed a pattern similar to that of the DC subsets (Fig. 6). There was an increase above baseline in the absolute numbers of circulating monocytes within the first 5 months post-transplantation (Fig. 6) and an increase in the percentage of monocytes residing in the lymph node within the first 3 months. This early recruitment of antigen-presenting cells (APCs) after transplantation may be important in driving T-cell development. The highest levels of DCs in blood and lymphoid tissue, as well as monocytes in lymph nodes, were found in animals that had received the selected-transduced grafts. This may account in part for the superior T-cell recovery found in this group.
In the present study, we compared the kinetics and characteristics of immune reconstitution after autologous transplantation of either PBMCs, CD34+ selected PBPCs, or CD34+ selected, cultured, and retrovirally transduced PBPCs in rhesus macaques. Each graft type was reinfused after myeloblative TBI. In all three groups, initial hematologic reconstitution with recovery of neutrophils, red blood cells, and platelets was comparable and rapid. All animals had markedly decreased thymic output of naïve T-cells for 1−2 months after transplantation, as previously described . The most striking and surprising finding of our study was multiple lines of evidence indicating that animals transplanted with cultured and retrovirally transduced PBPCs exhibited enhanced T-cell immune reconstitution compared with recipients of other graft types. This group had the fastest recovery of total CD3+ T-cells and both CD4+ and CD8+ naïve T-cells, accompanied by higher TREC numbers, less peripheral expansion of memory T-cells, and better preserved thymic architecture.
The thymic-dependent pathway relies on an adequate supply of progenitors able to home to the thymus and able to effectively proliferate and differentiate. The differences observed in this study cannot be explained by the CD34+ cell dose, since the starting number of CD34+ cells administered was equivalent in all groups and the ex vivo expansion of CD34+ cells was only 1.5-fold at most (Table 1). Besides, multiple primate and murine studies have reported a loss in the number of true repopulating stem cells during ex vivo culture under similar cytokine conditions [23–25]. In addition, patients that received unmarked expanded cells have been shown to suffer from engraftment failure . Thus, the higher TREC levels and better preserved thymic architecture in animals transplanted with the cultured-transduced CD34+ cells may result from the specific characteristics of the graft. In vitro treatment of CD34+ PBPCs with a cytokine cocktail similar to that applied in this study promoted the development of lymphoid progenitor cells that repopulate the thymus [26, 27]. The culture conditions we used during transduction of the CD34+ PBPCs included fibronectin support and media supplemented with SCF, Flt3 ligand, and MDGF. Fibronectin support prevents apoptosis without inducing active cell cycling [28–31] and helps maintain hematopoietic stem cell activity and function . SCF has also been shown to promote survival without resulting in proliferation or differentiation . Flt3 ligand has little effect in vitro on early progenitors, whereas in vivo administration leads to major alterations of different hematopoietic organs [34, 35]. MDGF is a specific growth factor for the platelet lineage . It enhances the survival of hematopoietic stem cells and augments their proliferation [37, 38].
We found that the culture condition used for transduction does not induce proliferation or maturation of contaminating T-cells within the graft. In addition, there were low levels of contaminating T-cells in our grafts prior to in vitro transduction, and the overall expansion of the CD34+ cells was very low (Table 1; Fig. 3A). Therefore, the animals in the selected-transduced group received fewer mature T-cells than the animals in the other two groups, and thus it is unlikely that re-infused cultured T-cells contributed to their faster recovery. The most likely pathway for the generation of T-cells is the thymic-dependent pathway in this group, which was supported by less peripheral expansion of T-cells, higher TREC levels, and preserved thymic architecture [39, 40]. The observed differences between the groups could be due to thymic production of naïve T-cells induced by feedback signals from the T-cell depleted periphery, therefore inducing higher thymic activity [41, 42]. The highest numbers of DCs and monocytes were also observed in the selected-transduced group, which may also have an impact on T-cell homeostasis, as APCs enhance induction of T-cell activation and proliferation in the periphery. The interaction between DCs and naïve T-cells is essential for the induction of primary T-cell responses against antigens and therefore crucial for the defense against numerous infections that may follow the immune suppression after transplantation. In this limited population of animals, differences observed in immune reconstitution did not translate into a different frequency of infections. In our clean facility with careful veterinary care and infection prophylaxis, we rarely encounter serious or fatal infections in our animals. Therefore, we do not have any evidence whether the higher output of naïve T-cells found in the selected-transduced group led to better response to pathogenic insult. Further studies addressing this are ongoing. As three out of four animals that had preserved thymic tissue at the end of the study were females, there is a possibility that sex can have an impact. However, there was no apparent trend that the female subjects in the respective group had better recovery of T-cells than the males. The neomycin resistance phosphotransferase is one of the most common marker genes used in gene transfer experimentation. Previous studies have suggested that neo gene expression could have deleterious effects on cell proliferation [43–45], but more definitive transplantation studies did not show any detrimental effects on engraftment or proliferation . We preferred to use a vector containing neo over, for example, enhanced green fluorescence protein as the latter might be more immunogenic [47, 48]. As the levels of cells in vivo showing retroviral insertion after transplantation was low, it may be unlikely that the retroviral transduction itself accounts for the improved immune reconstitution. In addition to the animals receiving selected-transduced cells in the current study, we have observed well-preserved thymic architecture in two animals (not included in this study) that received CD34+ cells transduced with a different retroviral vector (RD114 pseudotyped and containing the LacZ gene). Therefore, the in vitro culture procedure of CD34+ cells per se, rather than the retroviral vector used, may most profoundly promote lymphoid progenitor cell development and subsequent thymic repopulation. These results are of potential relevance to the rapid immune reconstitution observed in previously reported X-SCID gene therapy trials [49–51]. On the other hand, the early and enhanced immune recovery due to ex vivo culture and manipulation may increase the risk of a subsequent mutational event, which may precede the lymphoproliferation observed in a limited number of in children treated with CD34+ gene therapy for X-SCID. Further studies are necessary to define and optimize culture conditions contributing to improved immune reconstitution.
In this study, we found that nonhuman primates receiving purified and in vitro-transduced CD34+ cells after myeloblative TBI repopulated thymus efficiently, which promoted a higher output of naïve T-cells. Our detailed phenotypic results may serve as a platform for future studies regarding further interventions to impact on the immune reconstitution process, as well as a valuable reference for gene therapy, cell therapy and transplantation trials.
K.L. and R.S. contributed equally to the manuscript. R.S. was supported by a postdoctoral scholarship grant from the Deutsche Krebshilfe. We thank Amgen for supplying G-CSF and SCF, Kirin for MGDF, Immunex for Flt3-L, and Takara Shuzo for Retronectin. We also thank Louis Picker (Oregon National Primate Research Center, Oregon Health & Science University, Beaverton, Oregon) for advice on the staining panel used for phenotyping, Mario Roederer and Joanne Yu (ImmunoTechnology Section, Vaccine Research Center, NIH, Bethesda, Maryland) for supplying some of the conjugated, validated reagents used in this study, Stephanie Sellers and Andre LaRochelle for assistance with phenotyping, and Thomas Fountaine III for assistance with immunohistochemical stains.
The authors indicate no potential conflicts of interest.