Human skin cells support thymus-independent T cell development (original) (raw)
Design of the skin cell construct. Thymic epithelial cells are thought to provide the crucial signals for positive selection that allow double-positive T cell precursors to mature into single-positive T cells (17). It was therefore important to ensure that the construct contained large numbers of healthy epithelial keratinocytes. Initial experiments in which fragments of human skin were allowed to grow into artificial matrices produced constructs with mostly fibroblasts; additionally, these cultures were contaminated with T cells that were present in the skin samples.
We found the most successful approach was to expand the fibroblasts and keratinocytes separately and then combine them together onto artificial 3-dimensional Cellfoam cell growth matrices (Cytomatrix). Cellfoam is 3-dimensional tantalum-coated carbon matrix originally designed as an artificial bone matrix and used in previous xenogenic in vitro thymus explant cultures (Figure 1A) (8). Matrices seeded with keratinocytes and fibroblasts were cultured together using an intermediate cell culture medium until the cells reached confluence on the matrices (5–6 days), as assessed by immunofluorescence microscopy (IF). Human bone marrow–derived AC133+ hematopoietic precursors were then introduced into the colonized matrices. AC133+ input cell populations were at least 95% AC133+ and invariably negative for CD3+ T cells by flow cytometry analysis. These complete constructs were maintained in culture for 3–4 weeks in the presence of the prolymphopoietic cytokines IL-7 and IL-15, as well as Flt-3 ligand.
Structure of 3-dimensional skin cell cultures. (A) Scanning electron micrograph of the Cellfoam 3-dimensional matrix. Image courtesy of Cytomatrix LLC. (B and C) IF demonstrating the morphologies of (B) fibroblasts stained with vimentin antibody and (C) keratinocytes stained with antibodies to cytokeratins when these cells were grown alone on matrices. (D) When grown together, keratinocytes (orange) and fibroblasts (green) occupied distinct sites on the matrices. (E) DCs, identified by intense staining with HLA-DR antibodies, were observed only if bone marrow progenitor cells were added to the matrices. (F) DCs were often found adherent to the surface of vimentin+ fibroblasts. Magnification: ×2 (E and F), ×5 (A), ×10 (B–D).
IF of the constructs showed that fibroblasts grew as adherent cells on the matrices and also formed thin cellular projections that spanned adjacent matrix components, whereas keratinocytes grew as flattened, nonstratified, plate-like cells (Figure 1, B and C). When cultured together, keratinocytes and fibroblasts occupied nonoverlapping sites on the matrices (Figure 1D). Viable populations of both keratinocytes and fibroblasts were present on the matrices after 3 weeks of culture in Iscove’s medium, a medium well suited to T cell development but not normally supportive of fibroblast and keratinocyte growth (Figure 1D).
A viable population of epithelial keratinocytes was found to be required for successful T cell development. IF was used to estimate the proportion of fibroblasts and keratinocytes occupying sites on the matrix. In a particular focal plane, the proportion of adherent cells staining for both vimentin and epithelial keratins (Figure 1D, orange) was estimated and compared to the proportion of cells staining only for vimentin (Figure 1D, green). Optimal T cell production was observed when keratinocytes composed at least 40% of the adherent cell types on the matrices. Multiple experiments (8 experiments, using 2 different skin donors and 4 different HPC donors) confirmed that T cells were never produced from HPCs in matrices colonized only with fibroblasts (data not shown). In contrast, matrices colonized with keratinocytes alone produced low numbers of T cells (mean, 9% of T cells produced from matrices containing both keratinocytes and fibroblasts and using identical skin and HPCs; n = 3; data not shown). However, keratinocytes grown in the absence of feeder fibroblasts were smaller and fewer in number by the end of the culture period. Thus, the lack of efficient T cell differentiation may be a result of low keratinocyte viability as opposed to an intrinsic inability of keratinocytes to support T cell development.
DCs, distinguished by their high expression of MHC class II and low expression of CD14, were demonstrable by IF only if HPCs were added to the construct, indicating that these cells are of bone marrow origin (Figure 1, E and F).
Characterization of T cells produced in the skin cell cultures. Cells harvested from the constructs described above were analyzed by flow cytometry. Matrices colonized with both keratinocytes and fibroblasts robustly supported the development of cells that ultimately expressed T lymphocyte cell surface markers, including the CD3/T cell receptor complex. Optimal T cell production was observed when matrices contained at least 40–50% keratinocytes, as described above. From 1 × 104 precursor cells, individual 9-mm × 9-mm × 1.5-mm constructs produced 7.8 × 105 ± 2.4 × 105 total cells (mean ± SD; n = 6), 5–33% of which were CD3+ T cells. Individual constructs yielded between 4 × 104–1.7 × 105 T cells (mean, 7.9 × 104 ± 2.8 × 104; n = 6 representative experiments, each containing 12–40 individual matrices). In addition to T cells, we observed production of CD14loHLA-DRhi DCs (approximately 20–40%), CD14+ myeloid cells, and a variable number of CD56+ cells (data not shown).
We next studied the kinetics of T cell maturation. During development in the coculture system, progenitor cells initially expressed CD34 but lost expression of this marker as increasing levels of CD3 were acquired (Figure 2A). This loss of CD34 and gain of CD3 is also seen during normal T cell development in the thymus (21). Progenitor cells in our system were initially negative for CD3, CD4, and CD8. Transient production of double-positive CD4+CD8+ T cells expressing CD3 was noted between days 7 and 12 of culture; after day 14 of culture, double-positive cells were not detectable. By day 21, single-positive CD4+ and CD8+ T cells were present (Figure 2A). Normally developing thymocytes also progress from double-negative CD4–CD8– cells to double-positive CD4+CD8+ cells and subsequently to mature single-positive CD4+ or CD8+ cells (21). Double-positive CD4+CD8+ T cells, demonstrably present in our skin cell construct, are a cell population normally found exclusively in the thymus (21). By varying the culture conditions, we were able to influence the final production of CD4+ versus CD8+ cells. Irradiation of the construct prior to precursor addition, performed initially to prevent stromal overgrowth, produced larger numbers of single-positive CD4+ T cells (Figure 2B; duplicate conditions shown in Figure 2A). Pretreatment of the construct with IFN-γ prior to the addition of HPCs, performed initially to increase MHC class II expression by epithelial cells, led to increased production of single-positive CD8+ cells (Figure 2C). A more equal distribution of CD4+ and CD8+ cells was produced in the absence of these treatments (Figure 2D). Production of double-positive intermediate CD4+CD8+ cells was similar under all 3 culture conditions (data not shown).
HPCs differentiate into T cells in skin cell cultures. (A) Maturation of surface markers during T cell development in skin cell cultures. Matrices were irradiated prior to the addition of HPCs and produced primarily CD4+ T cells. (B–D) Alterations in culture conditions influenced the production of CD4+ versus CD8+ cells. (B) Irradiation of the construct prior to the addition of HPCs supported differentiation of proportionately more CD4+ cells. (C) Treatment of the skin cell construct with IFN-γ before the addition of HPCs increased the percentage of CD8+ cells produced. (D) In the absence of these treatments, a more equal distribution of CD4+ and CD8+ cells was observed. Cells were harvested from the matrices at 28 days. Similar results were seen in duplicate experiments. The percentage of cells in each quadrant is shown. (E) Expression of αβ TCR versus γδ TCR by CD3+ cells produced in skin cell cultures. (F) Qualitative RT-PCR for expression of TREC (T) and GAPDH (G) of input AC133+ HPCs and output T cells produced in skin cell cultures.
Skin and gut have been reported to support the development of T cells with the γδ TCR, whereas αβ T cells are primarily produced by the thymus (22). We therefore examined newly generated T cells for expression of the αβ versus γδ TCR. Flow cytometry analysis showed that consistently more than 95% of the CD3+ cells produced in our construct were αβ TCR T cells (Figure 2E).
Newly produced T cells contain T cell receptor excision circles. The T cells observed in our cocultures could have arisen from either de novo T cell differentiation or simply from expansion of very low numbers of mature T cells contaminating the skin cell or HPC populations. To test whether the T cells produced in our system are in fact naive, newly produced T cells, we tested output cells from our construct for the presence of T cell receptor excision circles (TRECs). TRECs, episomal circles of DNA that are produced as a byproduct of the recombination of TCR genes (23), are found in mature thymocytes and naive T cells recently released from normal thymus. Proliferation of T cells in the periphery leads to dilution of TRECs, because these DNA episomes do not replicate during cell division. TRECs have therefore been used to identify naive T cells and recent thymic emigrants (23). We performed RT-PCR analysis on both input HPCs and lymphocytes produced in the construct. Figure 2F shows the absence of TRECs in the HPC population (AC133+ input cells) and the presence of TRECs in lymphocytes produced in the construct. These findings indicate that a subset of HPCs underwent rearrangement of the TCR genes during coculture with skin cells, as would be expected during normal T cell development.
Newly produced T cells express a diverse TCR repertoire. To examine the TCR diversity of cells produced in the thymus construct, newly produced T cells were subjected to spectratyping via TCR-CDR3 length analysis. This technique allows identification of Vβ usage as well as diversity within each Vβ family. The number of peaks within a Vβ family is an index of the complexity of the TCR repertoire utilizing that Vβ. Two spectratypes are shown (Figure 3), both derived from the same HPC inoculum but differentiated in the presence of skin cells from 2 different donors. T cells produced in skin cell cultures from the first donor were significantly more diverse (Figure 3A). T cells of all 26 Vβ subfamilies tested were represented in the small population of newly generated T cells analyzed. Fully 85% of these Vβ families (22 of 26) displayed significant TCR diversity. T cells produced in cultures from the second skin donor were less diverse, suggesting that skin donors may vary in their ability to support diverse T cell differentiation (Figure 3B).
Spectratype analysis of T cells produced in skin cell cultures. T cells generated in skin cell cultures were subjected to TCR-CDR3 length analysis. Diversity within each Vβ family is signified by multiple peaks. Precursor cells from 1 bone marrow donor were matured in skin cell cultures from 2 different skin donors. T cells produced in cultures from the first skin donor (A; 8 × 105 cells analyzed) were more diverse and had a different T cell repertoire than T cells produced in cultures from the second skin donor (B; 5 × 105 cells analyzed).
Epithelial cells mediate positive selection in the thymus, thereby determining the TCR repertoire of developing thymocytes (22). If epithelial cells in skin cell cultures are mediating positive selection, different skin donors would be expected to produce different patterns of oligoclonality. We have found that T cells developing from the same HPC inoculum in cultures using different skin cell donors do have differing repertoire patterns. For example, the first skin donor we tested produced clonal populations in subfamilies Vβ8, Vβ19, Vβ23, and Vβ24, and the second produced completely clonal populations in a distinct subset (Vβ7, Vβ21, and Vβ16; Figure 3, A and B). Also, the greatest diversity was seen in largely nonoverlapping subsets of Vβ subfamilies: Vβ4, Vβ5, Vβ6, and Vβ9 versus Vβ9, Vβ11, Vβ22, and Vβ23. Thus, not only does the ability to support diverse T cell production vary among donors, but T cells from different Vβ subfamilies are preferentially produced from different skin cell donors.
Newly produced T cells are mature and functional. Mature and functional T cells are distinguished by their ability to proliferate, express the activation antigen CD69, and produce cytokines in response to stimulation through the TCR-CD3 complex. To determine whether the T cells produced in our system were mature and functional, we evaluated the response of these cells to T cell mitogens and alloantigens. T cells produced in the thymus construct proliferated in response to treatment with phytohemagglutinin (Figure 4, A and B). Proliferation was comparable to that seen using PBLs (construct-derived T cells, 28% ± 2.6%, versus PBLs, 34% ± 7.9%, mean ± SD, n = 3; PBL data not shown). Newly produced CD4+ and CD8+ single-positive T cells stimulated with concanavalin A expressed robust levels of the early activation marker CD69 (Figure 4, C–F). CD69 upregulation of newly produced T cells was also comparable to results obtained using PBLs (construct-derived T cells, 82% ± 2.6%, versus PBLs, 89% ± 2.9%; PBL data not shown). A subset of these activated cells also produced IL-2, TNF-α, and IFN-γ, as demonstrated by intracellular flow cytometry (Figure 4, G–J). Significant proliferation was also seen in response to allogeneic stimulator cells in mixed leukocyte reactions (MLRs; Figure 4K). Newly generated T cells proliferated in response to allogeneic stimulator cells at levels comparable to those seen in mature T cells isolated from peripheral blood, providing additional evidence for the functional maturity of these cells.
T cells produced in skin cell cultures are mature and functional. (A and B) Proliferation of T cells in response to (A) control medium and (B) phytohemagglutinin (PHA). (C–F) Expression of CD69 activation antigen in response to (C) control medium and (D–F) concanavalin A (Con A) by (C and D) total T cells and (E) CD4+ and (F) CD8+ subsets. (G and H) Production of TNF-α by CD3+ cells in response to (G) control medium and (H) concanavalin A treatment. (I and J) Production of (I) IFN-γ and (J) IL-2 in response to concanavalin A treatment. Treatment with control medium is not shown for IFN-γ and IL-2 samples but was identical to TNF-α control shown in G. The percentage of positive cells is shown. (K) Proliferation of T cells generated in skin cell cultures (T cells) in response to allogeneic PBMCs. Proliferation was assayed by BrdU incorporation on day 6 of the MLR. BrdU incorporation was detected via flow cytometry with gating on CD3+ T cells. PBMCs were derived from allogeneic, unrelated donors [PBMCs (A), donor A; PBMCs (B), donor B]. Error bars represent the SD of 3 measurements.
Delta-like Notch ligands and T cell production. The Notch ligand delta-like 1 has been shown to enhance T cell development via maintenance and expansion of progenitor cells, induction of lymphoid differentiation at the expense of myeloid differentiation, and the biasing of lymphocyte production toward T and away from the B lineage (24–26). OP9 monolayers expressing delta-like 1 support the full development of murine CD8+ T cells, but support human T cell development only to the double-positive stage (27, 28).
In order to examine skin cells for the presence of delta-like ligands, we stained human skin and cultured skin cells with an antibody that recognizes epitopes common to human delta-like 1–4. Suprabasal keratinocytes expressed delta-like Notch ligands (Figure 5A), and more than 50% of keratinocytes freshly disaggregated from skin expressed high levels of delta-like ligands (Figure 5B). Keratinocytes grown in low-density cultures were a mix of basaloid, small proliferating cells that did not express delta-like ligands and larger, differentiating cells that expressed high levels of delta-like molecules (Figure 5C). Culture of keratinocytes under high-density conditions that encouraged differentiation and stratification induced expression of delta-like ligands. Greater than 50% of keratinocytes grown in high-density cultures expressed delta-like ligands by day 6 after confluence of the cultures (Figure 5D). These cells maintained delta-like expression when subsequently plated on matrices (data not shown). In contrast, fibroblasts in the dermis of human skin and fibroblasts grown in culture did not express delta-like ligands (Figure 5, A and B).
Expression of delta-like Notch ligands in human keratinocytes. (A) Sections of normal human skin stained for delta-like Notch ligands demonstrated abundant staining of suprabasal epidermal keratinocytes but no staining of dermal fibroblasts. Magnification: ×10. (B) Keratinocytes freshly isolated from skin expressed delta-like ligands, but fibroblasts lacked expression. SSC-H, side scatter. (C) Cultured keratinocytes contained a mixture of large differentiating cells that expressed high levels of delta-like Notch ligands (green) and smaller, basaloid cells that were negative for delta-like ligand expression. Magnification: ×4. (D) Keratinocytes cultured under high-density conditions that encouraged differentiation expressed delta-like ligands.
T cells undergo negative selection in the construct, rendering them tolerant to self MHC. Negative selection during intrathymic development leads to clonal deletion of autoreactive T cells, a process mediated most efficiently by bone marrow–derived DCs (22). In our skin cell cultures, we observed that DCs were produced in parallel with T cells. Many of these cells, distinguished by high HLA-DR expression and low CD14 expression, were adherent to the surface of fibroblasts in the constructs (Figure 1, E and F). Thus, developing T cells likely come into contact with autologous DCs during development in the skin cell cultures. We performed a series of MLRs to determine whether these DCs could induce tolerance in developing T cells. DCs, either autologous or allogeneic, were produced from different bone marrow donors and used as stimulator cells in the MLRs.
DCs were produced by culturing HPCs for 21 days in the presence of matrices colonized only with dermal fibroblasts. These cultures produced a combination of monocytes (CD14hiHLA-DRlo) and DCs (CD14loHLA-DRhi), in agreement with a previous report that dermal fibroblasts support the differentiation of monocytes and DCs from HPCs (29). These cells were cultured for 5 days in GM-CSF and IL-4, then matured for 48 hours with LPS. Output cells were at least 80% mature myeloid-lineage DCs (CD14loHLA-DRhiCD86hi). DCs were irradiated and added to cultures of T cells generated in skin cell cultures from autologous and allogeneic bone marrow samples. T cells produced in the skin construct proliferated in response to allogeneic DCs but failed to respond to autologous DCs (Figure 6). PBMCs proliferated strongly in response to DCs from both donors. These results show that T cells developing in the skin cell construct do not proliferate in response to self MHC and other antigens carried by the DCs, providing evidence that negative selection occurs within the skin cell culture microenvironment.
Newly generated T cells are tolerant to challenge with autologous, but not allogeneic, DCs. T cells produced in skin cell cultures from HPCs of bone marrow donor A were exposed to donor A–derived DCs during the process of T cell development. Donor A–derived lymphocytes were therefore examined for their ability to respond to DCs derived from the same donor (donor A) and a second, unrelated donor (donor B) in the MLR. Responses of PBMCs to the DCs of both donors are included to demonstrate the immunogenicity of both DC populations. Proliferation was assayed by BrdU incorporation on day 6 of the MLR. BrdU incorporation was detected via flow cytometry with gating on CD3+ T cells. PBMCs were derived from a third, unrelated donor. Error bars represent the SD of 3 measurements. A duplicate set of experiments produced similar results.
Expression of the AIRE gene and autoantigens. The thymus also has the ability to eliminate T cells reactive to proteins normally expressed only in specific peripheral organs. Expression of the AIRE transcription factor induces a subset of medullary epithelial cells to express high levels of proteins normally limited to specialized tissues, and this expression has been shown to be crucial for the removal of autoreactive T cells (30, 31).
We tested skin cell cultures for expression of AIRE by real-time RT-PCR and found that combined HPC/skin cell cultures expressed AIRE, although at significantly lower levels than in the thymus (Figure 7, A and B). Expression was fairly constant during the 28-day culture period (Figure 7B). DCs in the thymus also express AIRE and a panel of autoantigens (32, 33). In order to determine whether AIRE was expressed by skin or hematopoietic cells, we next tested cultures of keratinocytes and fibroblasts grown on 3-dimensional matrices in the absence of HPCs. We found that mixed cultures of fibroblasts and keratinocytes also expressed AIRE (Figure 7C). Lymphotoxin-α (LTα) induces the expression of the AIRE gene both in intact mouse thymus and in thymic epithelial cell lines (34). LTα mediates its effect by binding to the lymphotoxin-β receptor (LTβR), and this receptor is upregulated by IFN-γ (35). Keratinocytes express LTα and upregulate LTβR after treatment with IFN-γ) (36). We observed a modest increase in AIRE expression when cocultures of keratinocytes and fibroblasts were pretreated with IFN-γ and stimulated with recombinant LTα (Figure 7C). To determine whether fibroblasts or keratinocytes expressed AIRE, we tested cells that were grown separately on 3-dimensional matrices. AIRE is expressed by several types of epithelial cells in mice, including Fallopian tube, kidney, and large airway epithelia (37). We were therefore surprised to find that AIRE was expressed by dermal fibroblasts, not by keratinocytes (Figure 3C). To our knowledge, this is the first report of AIRE expression in cells from the skin.
Skin cell cultures express autoantigens and transcription factors important for thymic and T cell development. (A and B) Skin cell/HPC cultures express AIRE. Equal amounts of mRNA derived from skin cell/HPC cultures and from normal human thymus were analyzed by real-time RT-PCR for AIRE expression in the presence (+) or absence (–) of RT. RNA from skin cell–colonized matrices was isolated on days 0, 7, 14, 21, and 28 after the addition of HPCs. Signal for each sample was first normalized to cyclophilin A expression and then normalized to thymus expression. mw, molecular weight markers (100-kb ladder). (C) Fibroblasts in skin cell cultures express AIRE. AIRE expression was assayed by real-time RT-PCR in keratinocyte (Ker) and fibroblast (Fib) cocultures without HPC and in cultures of keratinocytes or fibroblasts alone. Combined cultures of skin cells were assayed after treatment with control medium (Con), IFN-γ, or IFN-γ followed by LTα stimulation. Signal for each sample was normalized to cyclophilin A and thymus expression. (D and E) Skin cell cocultures with and without HPC express the autoantigens MBP, PLP, S100β and thyroglobulin by quantitative real-time RT-PCR. RNA from skin cell/HPC cocultures harvested on day 14, matrices colonized with keratinocytes and fibroblasts alone, and thymus was analyzed in the presence (+) or absence (–) of RT. Signal for all samples was normalized to cyclophilin A and thymus expression. No product was detected in samples without RT. (F) Skin cell/HPC cultures express Foxn1 and Hoxa3, but do not express Pax1. Day-28 skin cell/HPC cultures were analyzed by real-time RT-PCR. Samples without RT had no signal in all samples. Expression was normalized to cyclophilin A, and expression of each factor was then normalized to thymus expression.
We observed relatively low levels of AIRE expression in skin cell cultures. To determine whether this expression induces autoantigen expression, we tested skin cell/HPC cultures by quantitative real-time RT-PCR for a panel of noncutaneous autoantigens which included myelin basic protein (MBP), myelin phospholipid protein (PLP), S100β, and thyroglobulin. These 4 autoantigens were detected in both human thymus and in day-14 combined skin cell/HPC cultures (Figure 7, D and E). By day 14, DCs were likely present in skin cell/HPC cultures. Thymic DCs are known to express AIRE and a subset of autoantigens (38). To determine whether skin cells alone expressed autoantigens, we tested cocultures of fibroblasts and keratinocytes in the absence of HPCs. We found that these cultures expressed lower, but detectable, levels of all 4 autoantigens (Figure 7E).
Skin cell cultures express the thymic transcription factors Hoxa3 and Foxn1. Mutations in several transcription factors, including Foxn1, Hoxa3, and Pax1, result in impaired development or function of the thymus. Foxn1 is critical for the differentiation and survival of thymic epithelial cells. Foxn1 deficiency induces the nude phenotype in mice, rats, and humans, characterized by hairlessness and congenital athymia (39). Nude mice have a rudimentary thymus that lacks all types of thymic epithelial cells and cannot support T cell differentiation. We found significant expression levels of Foxn1 in skin cell/HPC cultures by real-time RT-PCR (Figure 7F). This result is consistent with the fact that Foxn1 is also critical for differentiation of epithelial cells in the skin (40). _Foxn1_-deficient mice have abnormal keratinocyte maturation and produce fragile hairs with altered keratin composition. Transient Foxn1 expression has recently been shown induce a transcriptional program leading to the differentiation of epidermal keratinocytes (41).
Hoxa3, a member of the Hox family of transcription factors, is critical for the development of structures in the pharyngeal region. _Hoxa3_-deficient mice are athymic and also lack parathyroid glands (42). We found significant levels of Hoxa3 expression in skin cell/HPC cultures (Figure 7F). This is consistent with the finding that Hoxa3 also plays a role in keratinocyte biology. Hoxa-3 is expressed by keratinocytes and promotes migration of these cells during wound healing (43).
Deficiency of the Pax1 transcription factor has more subtle effects on thymic function than Foxn1 and Hoxa3 deficits. _Pax1_-deficient mice have a hypoplastic thymus that is less efficient at supporting T cell development (44). In contrast to our findings with Foxn1 and Hoxa3, we found no evidence of Pax1 expression in skin cell/HPC cultures by real-time RT-PCR (Figure 7F).