Regulation of in vitro human T cell development through interleukin-7 deprivation and anti-CD3 stimulation
© Patel et al.; licensee BioMed Central Ltd. 2012
Received: 15 March 2012
Accepted: 20 June 2012
Published: 16 August 2012
The role of IL-7 and pre-TCR signaling during T cell development has been well characterized in murine but not in human system. We and others have reported that human BM hematopoietic progenitor cells (HPCs) display poor proliferation, inefficient double negative (DN) to double positive (DP) transition and no functional maturation in the in vitro OP9-Delta-like 1 (DL1) culture system.
In this study, we investigated the importance of optimal IL-7 and pre-TCR signaling during adult human T cell development. Using a modified OP9-DL1 culture ectopically expressing IL-7 and Fms-like tyrosine kinase 3 ligand (Flt3L), we demonstrated enhanced T cell precursor expansion. IL-7 removal at various time points during T cell development promoted a slight increase of DP cells; however, these cells did not differentiate further and underwent cell death. As pre-TCR signaling rescues DN cells from programmed cell death, we treated the culture with anti-CD3 antibody. Upon pre-TCR stimulation, the IL-7 deprived T precursors differentiated into CD3+TCRαβ+DP cells and further matured into functional CD4 T cells, albeit displayed a skewed TCR Vβ repertoire.
Our study establishes for the first time a critical control for differentiation and maturation of adult human T cells from HPCs by concomitant regulation of IL-7 and pre-TCR signaling.
KeywordsT cell development Interleukin-7 T cell receptor Vbeta repertoire
Generation of mature human T cells from adult bone marrow (BM) CD34+HPCs in vitro may overcome two major limitations in T cell therapy, namely HLA disparity and immune tolerance. Patients undergoing chemotherapy or with HIV infection suffer from prolonged lymphodepletion leading to opportunistic infections and mortality . Hematopoietic stem cell transplant (HSCT) has been used to reconstitute the immune system in such patients . However, T cells take the longest time to recover after HSCT . Thus ex vivo differentiation of T cells using an in vitro OP9 stromal cell line expressing Notch ligand, Delta like-1 (DL1), has been of tremendous interest [3–5]. Recent reports showed that the OP9DL1 stromal cell culture system established by Zuniga-Pflucker can support terminal maturation of cord blood (CB) and post natal thymus derived CD34+HPCs [6, 7]. In case of immune rejection of CB HPCs due to HLA disparity or limited availability, BM CD34+HPCs may serve as a convenient source as they can be obtained from patient’s own BM [8, 9]. We and others have demonstrated that adult BM-derived CD34+HPCs, from both normal adults and patients undergoing chemotherapy, yields a low number of T cell precursors in vitro[10–12]. T cell development of adult human BM derived CD34+HPCs in the OP9 DL1 culture system is less well studied due to low cellular yields when compared to the CB counterparts. In addition, terminal T cell differentiation from adult human BM derived CD34+HPCs in vitro has not yet been demonstrated [10, 13].
We have previously reported that lentivector-modified OP9 cell lines expressing various cytokines and growth factors supported enhanced HPC and dendritic cell precursor expansion and differentiation . To overcome the limited proliferation of BM HPCs in vitro, we modified a previously defined LmDL1 cell line (Lentivector-modified OP9 expressing DL1) , to ectopically express T cell developmental factors IL-7 and Flt3L, and established LmDL1-FL7 cell line. We found that LmDL1-FL7 provided a proliferative advantage to adult BM CD34+HPCs over LmDL1 cell line supplemented with soluble recombinant hIL-7 and hFlt3L.
During T cell development, the CD34+CD8-CD4- double negative (DN) thymocytes differentiate through CD3-CD8+ immature single positive stage (ISP) in mice and CD3-CD4+ ISP in humans, followed by CD3loCD4+CD8+ double positive (DP), CD3+TCRαβ+DP and then CD3+TCRαβ+CD4+ or CD8+ mature single positive T cells [15, 16]. We observed that the transition of CD3lo DP to CD3+TCRαβ+ DP stage, an intermediate stage that precedes the terminal maturation to CD8 or CD4 T cell lineage, is inefficient during adult BM T cell development in vitro. IL-7 plays an inhibitory role during DN to DP transition in mice [18–22] and signaling via CD3/Pre-TCR complex plays a permissive role in transition from CD3lo DP to CD3+ TCRαβ+ DP [23–25]. Thus, we hypothesized that the inefficient pre-TCR signaling is either due to continued presence of IL-7 or due to inefficient stimulation via CD3 receptor. Here we report that intermittent IL-7 withdrawal alone did not result in efficient differentiation to CD3+TCRαβ+ DP stage. Importantly, taking a combination approach of IL-7 withdrawal and activating pre-TCR signaling using anti-CD3/CD28 antibodies, we demonstrate for the first time in vitro differentiation of adult BM HPCs to CD3+TCRαβ+ DP stage and subsequent functional maturation of CD4 T cells. Our findings provide a better understanding of the factors involved in proliferation and differentiation of BM derived HPCs to mature T cells in vitro.
OP9-DL1 cells ectopically expressing Flt3L and IL-7 support enhanced T cell precursor expansion
To examine the differentiation and expansion potential of adult human BM CD34+ HPCs co-cultured with LmDL1 exogenously supplemented with recombinant human Flt3L (5 ng/mL) and IL-7 (5 ng/mL), or co-cultured with LmDL1-FL7, we determined the proliferation rate of the incubated cells by counting total number of suspension cells at various time points in three independent experiments. The result showed that CD34+ HPCs cells, when co-cultured with LmDL1-FL7 for 35 days, expanded up to five fold more than those co-cultured with LmDL1 supplemented with recombinant Flt3L and IL-7 (Figure 1F, representative of three donors). Thus, LmDL1-FL7 was superior to LmDL1 in supporting T cell precursor proliferation.
Adult BM CD34+ HPCs co-cultured on LmDL1-FL7 or LmDL1 supplemented with IL-7 and Flt3L follow similar T cell differentiation kinetics but do not undergo functional T cell maturation
IL-7 deprivation alone does not induce efficient DN to DP transition
IL-7 withdrawal does not increase T cell receptor excision circle (TREC) in the developing T cell precursors
Adult human HPCs can differentiate to DP T cells and adopt a CD4 T cell lineage in vitro upon IL-7 deprivation followed by anti-CD3 stimulation
Vβ repertoire analysis of the in vitro generated CD4 T cells
In vitro adult human BM HPC-derived functional T cells have great potential for therapeutic applications, as this approach provides donor HLA-matched T cells that may be genetically engineered to fight infections, cancer or to treat immunodeficiencies. Murine HPCs, human CB and post-natal thymic HPCs undergo full maturation in the OP9-DL1 culture system [34, 35]. However, adult human BM HPCs undergo limited proliferation and are arrested at CD3loCD4+CD8+ DP stage of T cell development [10, 13]. In this report, we demonstrated that cell-based IL-7 and Flt3L provided a proliferative advantage to adult BM HPCs over exogenously supplemented recombinant IL-7 and Flt3L. We then took a combination approach of IL-7 withdrawal and activating pre-TCR signaling using anti-CD3/CD28 antibodies, which successfully overcame the arrest in CD3loCD4+CD8+ DP stage and induced differentiation to CD3+TCRαβ+CD4+CD8+ DP stage, and subsequent maturation to CD4 T cells. Our findings provide a better understanding of the factors involved in proliferation and differentiation of adult BM-derived HPCs in vitro.
LmDL1-FL7 was superior in supporting T cell precursor proliferation when compared to LmDL1 supplemented with exogenous Flt3L and IL-7. The mechanism that enhanced precursor proliferation on LmDL1-FL7 remains to be elucidated. It is possible that concentration or cell-based modifications, or both, could contribute to the enhanced proliferation. As all three cell lines, LmDL1, LmDL1-FL and LmDL1-FL7 expressed high levels of mDL1 (Figure 1), differential DL1 expression level does not seem to play a role. Flt3L is expressed as a soluble as well as membrane bound form , and glycosylated form of IL-7 has been reported . Besides soluble factors, cell-cell interactions play a critical role in T cell development. Our results appear to point to the importance of cell-based modification of cytokines, as use of glycosylated IL-7 for clinical trials is being considered . Previous studies demonstrated that a high dose of IL-7 has a modest effect on increasing the absolute cell number during T cell development [18, 21]. These studies support our view that exogenously added cytokine dose has limited effects on T cell development.
While the T cell development potential such as occurrence of CD8 ISP and DP cells were comparable for both culture systems, some differences exist, such as CD3 expression and development of TCRγδ cells. Cell-free or cell-cell signaling of the cytokines may account for the differences in proliferation and differentiation of the two systems. Nevertheless, differentiation to DP stage was inefficient and neither system supported terminal T cell maturation. Under both culture conditions, precursor proliferation rate declined beyond 30–35 days suggesting a discontinued dependence for IL-7 and Notch signals, consistent with previous reports [39, 40], as such, this culture system alone does not support continued differentiation of adult human T cell precursors to CD3 and TCRαβ-bearing DP cells.
Signaling through IL-7/IL-7R supports survival and proliferation through DN3 stage in murine T cell development and the same is true for human T cell development [20, 41]. In transgenic mice, expression of IL-7 under the control of lck promoter at low levels enhances proliferation of developing αβ T cells, but at high levels, it reduces proliferation and displays a marked block in DP transition . Recent studies further support that IL-7R signals impair differentiation of CD8 ISP to DP cells in Zap70−/− and IL-7Rα transgenic mice , and IL-7R signals inhibit the expression of HMG domain transcription factors TCF-1, LEF-1 and RORγt, factors important for pre-T to DP transition . In addition, IL-7 suppresses anti-CD3 antibody induced differentiation to DP stage in fetal thymus organ culture of Rag1-deficient mice . Thus, we hypothesized that IL-7 withdrawal prior to ISP might be necessary for efficient DP transition. While IL-7 has been reported to display an inhibitory role in DP transition in murine T cell development, our results showed that the intermittent removal of IL-7 in the in vitro co-culture only had a minimal effect on human T cell DP transition.
The mechanism by which IL-7 inhibits T cell development is unclear. We observed abundant transcripts of TCF and LEF in T cell precursors at various time points. Thus, it seems unlikely that IL-7 withdrawal promotes T cell development by de-repressing transcription of the above factors. Our data suggest that IL-7 does not inhibit TREC formation, neither does it directly inhibit pre-TCR signaling. Interestingly, an increase in CD4 surface expression post IL-7 withdrawal may play a role in how these cells respond to anti-CD3 stimulation. As in human T cell development, CD4ISP precede DP stage, it is possible that increased CD4 may account for increased responsiveness to anti-CD3 stimulation. Alternative possibilities are, IL-7 mediates its effect through STAT-5 on transcription of genes necessary for pre-TCR expression and function, directly inhibits pre-TCR activation, or interferes with the TGFβ signaling pathway . Detailed evaluation of these possibilities requires further investigation.
IL-7 mediates survival and proliferation of DN thymocytes . In addition, IL-7 is required for TCRγδ gene rearrangement and also induces TCRβ chain rearrangement [44–46]. In order to progress to the next DP maturation stage, DN/ISP thymocytes must seize the rearrangement and express a functional TCRγδ or TCRαβ . It is known that signaling via a functional TCR mediates allelic exclusion, survival and progression to SP stage . Interestingly, in mice IL-7 signaling is inhibited at DP stage by down-regulating the IL-7Rα. In humans, IL-7R is expressed but its binding partner γC is down-regulated and STAT-5 responsiveness is lost . Hence it is tempting to speculate that IL-7 signaling down-regulation might be an additional way of terminating rearrangement and preventing survival of T cells with non-functional TCRs. As both IL-7 and TCR signaling deliver survival signals, the down-regulation of IL-7 signaling ensures shutdown of an alternative survival pathway and selects for cells that respond to TCR signals. Clearly, the change in IL-7/IL-7R signaling is physiologically important and the reason for such regulation might reside on the intracellular signaling of IL-7/IL-7R on T cell activation and fate decision: proliferation, death or differentiation.
During T cell development, the appearance of ISP is dominated by CD4 ISP in human and CD8 ISP in mouse; pre-TCR signals drive proliferation, TCRα rearrangement, followed by the appearance of CD8 ISP in mouse and CD4 ISP in human. Interestingly, we observed CD8 ISP derived from human CD34 HPCs in vitro; we found a lack of proliferative burst and minimal rearrangement in the TCRα locus. Thus, the CD8 ISP may not be true ISP generated by pre-TCR signals, rather a result of cytokine-mediated CD8 expression .
IL-7 withdrawal is necessary but not sufficient for further differentiation to DP stage, and anti-CD3 stimulation plays a key role in inducing CD3+TCRαβ+DP transition and subsequent maturation to CD4 T cells. Our findings further advance the experimental system required for in vitro modeling of adult human T cell differentiation, and will help develop novel approaches toward generating functional T cells from adult HPCs.
Human CD34+ cells and cell lines
The adult BM CD34+ HPCs from normal donors were purchased from AllCell Inc. (San Mateo, CA, USA). Control PBMCs were obtained from Civitan Regional Blood Center Inc. (Gainesville, FL) reviewed and approved by University of Florida Health Science Center Institutional Review Board (#507-1997, UF IRB-01). All studies involving human subjects are conducted in accordance with the guidelines of the World Medical Association's Declaration of Helsinki (most recent revision). The mouse fetal stromal cells (OP9) were purchased from the American Type Culture Collection (ATCC, Manassas, VA) and maintained as previously described . The engineered LmDL1 and LmDL1-FL7 cell lines were generated by transducing cells with lentivectors encoding DL1, Flt3L and IL-7, respectively. The stromal cells were maintained in α-MEM (Invitrogen/Gibco BRL, Grand Island, NY) supplemented with 15-20% fetal bovine serum (FBS, Invitrogen/Gibco BRL) and 1% Penicillin-Streptomycin (Mediatech Inc., Manassas, VA). IL-7 cytokine secretion was measured by using human IL-7 ELISA kit (Ray Biotech, Inc) and soluble Flt3L in culture was measured using human Flt3L ELISA kit (Assay Biotechnology Company, Inc, Sunnyvale, CA). Cell free supernatants were harvested from LmDL1 and LmDLFL7 cells cultured for 48 hrs (80-90% confluent), in a 12 well plate containing 1 ml of media. The samples were read on model 680 microplate reader (Bio-Rad). The surface expression of mouse DL1 and Flt3L was analyzed by flow cytometry with Alexa Fluor 647-conjugated anti-DL1 Ab (Biolegend) and purified anti-Flt3L Ab (Abcam Inc. Cambridge, MA) conjugated with Zenon-Alexa 488 according to manufacturer’s instructions (Invitrogen).
LmDL1 stromal cell and CD34+ HPC co-culture
The CD34+ HPCs were seeded into 24-well-plate at 1x105 cells/well containing a confluent monolayer of LmDL1 or LmDL1-FL7 cells. The cocultures were maintained in complete medium from day 1, consisting of α-MEM with 20% FBS and 1% Penicillin-Streptomycin, supplemented with 5 ng/ml IL-7 (PeproTech, Inc. Rocky Hill, NJ) and 5 ng/ml Flt3L (PeproTech, Inc.) as indicated. The cocultures were replenished with new media every 2–3 days. The cells in suspension were transferred to a new confluent stromal monolayer once the monolayer began to differentiate or when developing cells reached 80-90% confluent. The cells were transferred by vigorous pipetting, followed by filtering through a 70 μm filter (BD/Falcon, BD Biosciences, Sparks, MD) and centrifugation at 250 RCF, at room temperature for 10 min. The cell pellet was transferred to a fresh confluent monolayer. The cells were harvested at the indicated time points during the T cell development for analysis.
Monoclonal antibodies and flow cytometry
The surface expression of mouse DL1 and Flt3L was analyzed by flow cytometry with Alexa Fluor 647-conjugated anti-DL1 antibody (Biolegend) and purified anti-Flt3L antibody (Abcam Inc. Cambridge, MA) conjugated with Zenon-Alexa 488 according to manufacturer’s instructions (Invitrogen). The antibodies used for surface staining of T cell development included CD4 (clone RPA-T4 Pacific blue), CD8 (clone RPA-T8 PE), CD3 (clone UCHT1, Pacific Blue, clone SK7, PE-Cy7), TCRαβ (clone T10B9.1A-31, FITC), CD1a (clone HI149, APC), CD7 (clone M-T701, FITC, PE), and intracellular staining for Ki67 (clone B56, FITC), and isotype IgG1κ, which were from BD Biosciences (San Jose, CA). anti-CD127 (clone 40131-FITC) was from R&D systems (Minneapolis, MN). Vβ repertoire analysis was performed using IOTest® Beta Mark TCR Vβ Repertoire Kit according to manufacturer’s instructions (Beckman Coulter, Fullerton, CA). For flow cytometric staining, cells were first washed with PBS plus 2% FBS and blocked with mouse and human serum at 4°C for 30 min. Cells were incubated with antibodies per manufacturer’s instructions. For each fluorochrome-labeled Ab used, appropriate isotype control was included. After antibody staining, the cells were washed twice and fixed with 2% para-formaldehyde. Intracellular staining was performed using BD cytofix/cytoperm kit, according to the manufacturer’s protocol. Data was acquired using BD FACS Diva software (version 5.0.1) on a BD FACSAria or a BD LSR and analyzed using the Flowjo software (version 188.8.131.52, Tree Star, Inc. Pasadena, TX).
T cell stimulation and effector function analysis
To stimulate naïve T cells, a protocol for long term stimulation was followed using anti-CD3/CD28 beads (Dynal/Invitrogen, San Diego, CA) per manufacturer’s instructions. The cells and the beads were mixed and plated into a 96 well plate at 37°C for 2–3 days in X-Vivo 20 (Gibco) media, on day 3, 12.5 U of IL-2, 5 ng/ml of IL-7 and 20 ng/ml of IL-15 were added and the cells were cultured for additional 11–12 days. The in vitro expanded CD4 T cells were stimulated with PMA and Ionomycin (Sigma-Aldrich, St. Louis, MO), and analyzed for the release of IFN-γ, IL-4 and IL-17. Briefly the cells were incubated with 25 ng/ml PMA and 1 μg/ml ionomycin for one hour followed by the addition of 6 μg/ml monensin (Sigma-Aldrich) to inhibit Golgi-mediated cytokine secretion. After 4–5 hours of incubation, the cells were harvested and surface stained for CD4 (clone RPA-T4, Pacific blue), CD8 (clone SK1, APC-Cy7), CD3 (clone SK7 PE-Cy7), CD25 (clone M-A251, PE) and intracellular stained for IFN-γ (clone 25723.11, FITC), IL-4 (clone MP425D2, APC), FOXP3 (clone PCH101, Alexa 647); the above antibodies were from BD Biosciences, and IL-17 (clone 64CAP17, PE) antibody was from e-Biosciences. The data were collected by flow cytometry using BD FACSAria and analyzed using Flowjo.
RNA was harvested from cells using TRI-Reagent (Sigma-Aldrich) and 1 ug RNA was reverse transcribed into cDNA by using Two-step AMV RT-PCR kit (Gene Choice, MD). The following primers were used for the PCR: mGAPDH, F (Forward) 5’-TCA CCA CCA TGG AGA AGG C-3’ and R (Reverse) 5’-GCT AAG CAG TTG GTG GTG CA-3’; mDL1, F 5’-GCT CTT CCC CTT GTT CTA ACG-3’ and R 5’-CAC ATT GTC CTC GCA GTA CC-3’; Flt3L, F 5’-AAG GAT CCG CAG GAT GAG GCC TTG-3’ and R 5’-CGG CGA CAG GAG GCA TGA G-3’; IL-7, F 5’-TTC TCG AGT TAT CAG TGT TCT TTA GTG-3’ and R 5’-AAG CGG CCG CCA CCA TGT TCC ATG TTT CT-3’; huGAPDH, F 5’-CCG ATG GCA AAT TCG ATG GC-3’ and R 5’-GAT GAC CCT TTT GGC TCC CC-3’; hLEF-1, F 5'-CGA CGC CAA AGG AAC ACT GAC ATC-3' and R 5'-GCA CGC AGA TAT GGG GGG AGA AA-3'; hTCF-1, F 5'-CGG GAC AGA GGA CCA TTA CAA CTA GAT CAA GGA G-3', and R 5'-CCA CCT GCC TCG GCC TGC CAA AGT-3'; Rag-1, F 5'-CAG CGT TTT GCT GAG CTC CT-3' and R 5'-GGC TTT CCA GAG AGT CCT C-3'; Rag-2 F 5'-GCA ACA TGG GAA ATG GAA CTG-3' and R 5'-GGT GTC AAA TTC ATC ATC ACC ATC-3'. After 30 cycles of amplification (95°C for 30 seconds, 55°C for 30 seconds, and 72°C for 60 seconds), PCR products were separated on a 2% agarose gel.
T cell receptor excision circle (TREC) analysis
The TREC and RAG2 sequences were amplified by nested PCRs using two outer primers in the first round PCR, followed by adding two inner primers in the second round PCR using genomic DNA of PBMCs as templates, and cloned into pSTblue and verified by DNA sequencing. The two 5’ primers for TREC amplification were: outer 5'-AAT CTA GAG CAT GTT GCT TGA ACT CCT C-3', and inner 5'-AAT CTA GAG TAG CAT AAT TTC CTG GTT GAC-3'; the two 3’ primers for TREC amplification were: outer 5'-AAT CTA GAC CAA GGT GAA TCC TCT GAT C-3', and inner 5'-AAT CTA GAG TCC CAC ACT CCG TGC TG-3'. The two 5’ primers for RAG2 amplification were: outer 5'-AAG GAT CCA GCT GTG AAT TGC ACA GTC-3', and inner 5'-AAG GAT CCG CAA TCC TGA CTC AAA CTA AC-3'; the two 3’ primers for RAG2 amplification were: outer 5'-AAG GAT CCA GTT GAA TAG AAT GGT ACC-3’ and inner 5'-AAG GAT CCG TAA TCC AGT AGC CTG TCT C-3'. Cell lysates were prepared by proteinase K digestion (100 μg/mL) at 56°C for 1 hr, followed by heat inactivation at 95°C for 10 minutes. In brief, 1.5 μL of cell lysates equivalent to 100 ng DNA or 15,000 cells, were used as template for PCR amplification. The following primers were used for the PCR reaction for TREC: F 5’-CAG AGG GGT GCC TCT GTC A-3’ and R 5’-CTG TGA AAC ACT CCC CAG C-3’, and RAG2: F 5’-TCT TGG CAT ACC AGG AGA CA-3’ and R- 5’-AGT GGA ATC CCC TGG ATC TT-3’. PCR conditions were 95°C for 10 minutes, followed by 35 cycles of 95°C for 30 seconds, 55°C (RAG2) 59°C (TREC) for 60 seconds, 72°C for 60 seconds, with a final extension at 72°C for 10 minutes. PCR products were analyzed on 1% agarose gel.
The statistical analysis was performed using Student’s t-test and GRAPHPAD PRISM 5 software.
We thank S. Williams, W. Chou, Dr. Shuhong Han and Yuling Yeh for technical assistance, Dr. Yung Chang and Dr. Lizi Wu for critical comments and reading the manuscript, and N. Benson for help with flow cytometry analysis. ESP was supported by NIH T32 graduate training grant. The study was supported by amfAR grant 107768, Yongling Foundation, and flow cytometry fund of UF Shands Cancer Center.
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