CD4+CD8+ double-positive (DP) T cells represent a minor subpopulation of T lymphocytes found in the periphery of adult rats. In this study, we show that peripheral DP T cells appear among the first T cells that colonize the peripheral lymphoid organs during fetal life, and represent ∼40% of peripheral T cells during the perinatal period. Later their proportion decreases to reach the low values seen in adulthood. Most DP T cells are small size lymphocytes that do not exhibit an activated phenotype, and their proliferative rate is similar to that of the other peripheral T cell subpopulations. Only 30–40% of DP T cells expresses CD8β chain, the remaining cells expressing CD8αα homodimers. However, both DP T cell subsets have an intrathymic origin since they appear in the recent thymic emigrant population after injection of FITC intrathymically. Functionally, although DP T cells are resistant to undergo apoptosis in response to glucocorticoids, they show poor proliferative responses upon CD3/TCR stimulation due to their inability to produce IL-2. A fraction of DP T cells are not actively synthesizing the CD8 coreceptor, and they gradually differentiate to the CD4 cell lineage in reaggregation cultures. Transfer of DP T lymphocytes into thymectomized SCID mice demonstrates that these cells undergo post-thymic maturation in the peripheral lymphoid organs and that their CD4 cell progeny is fully immunocompetent, as judged by its ability to survive and expand in peripheral lymphoid organs, to proliferate in response to CD3 ligation, and to produce IL-2 upon stimulation.

Most peripheral T lymphocytes fall into two main subsets: MHC class II-restricted CD4+CD8 helper cells and MHC class I-restricted CD4CD8+ cytotoxic cells. It is broadly accepted, therefore, that the expression of the CD4 and CD8 molecules by those mature T cells is mutually exclusive and that only the immature T lymphocytes express both Ags simultaneously during the intrathymic maturation process. However, T cells bearing a CD4+CD8+ double-positive (DP)3 phenotype can be found in small numbers in the peripheral blood of healthy humans (1, 2, 3) and in the secondary lymphoid organs and peripheral blood of rats and mice (4, 5, 6). Unlike humans and rodents, a prominent DP T lymphocyte population is detected in the periphery of swine, monkeys, and some strains of chickens (7, 8, 9, 10). It has been reported that the proportion of peripheral DP lymphocytes increases during the neonatal period in mice and humans (2, 11, 12, 13, 14), in certain disease conditions such as bacterial and viral infections (3, 15, 16, 17), autoimmune disorders (3, 18, 19, 20), tumors (3, 21), and after transplantation (22, 23), in aged individuals (7, 10, 24), and spontaneously in some healthy adult humans (25, 26, 27).

The phenotype and function of peripheral DP T cells is a conflicting matter. Some authors consider that the DP T cell subset is comprised of mature lymphocytes with a naive phenotype (4, 14), whereas others have described the expression of memory T cell markers in peripheral DP cells (7, 10, 28). An intermediate phenotype between naive and memory has been also reported in some human DP T cell populations (26). Functionally, DP lymphocytes from swine, chickens, and humans show helper activities (7, 9, 27, 29), whereas those from monkeys seem to have dual functions which overlap with CD4+CD8 and CD4CD8+ T cells (30). Furthermore, some reports have pointed out that peripheral DP T lymphocytes are only partially immunocompetent (11, 27).

The origin of peripheral DP T cells is also unclear. They could represent DP thymocytes that have prematurely escaped from the thymus (4, 11, 14), or CD4+CD8 T cells that re-express CD8 after activation or exposure to lymphokines such as IL-4 (7, 31, 32).

In the present report, we study the evolution of rat peripheral DP T cells during fetal, postnatal, and adult life, and analyze their phenotype, origin, and functionality. We show that rat peripheral DP T lymphocytes have an intrathymic origin and that they represent a partially immunocompetent thymocyte subpopulation that is exported to the periphery, where they conclude their maturation to become functionally competent CD4+CD8 T cells.

Wistar Hannover rats were maintained in our animal facilities. Rat fetuses at days 19–21 of gestation were obtained from timed pregnancies. The day of finding a vaginal plug was designated day 0 of gestation. C.B17 SCID mice were purchased from Harlan Iberica (Barcelona, Spain) and maintained in specific pathogen-free breeding facilities.

Mouse or hamster anti-rat mAbs of the following specificities were obtained from BD PharMingen (San Diego, CA) and Serotec (Oxford, U.K.) and used in the current study: CD4 (OX35), CD8α (OX8), CD8β (341), CD25 (OX39), CD28 (JJ319), CD45RC (OX22), CD49d/VLA-4 (Mrα41), CD53 (OX44), CD62L/L-selectin (HRL1), CD71/transferrin receptor (OX26), CD134 (OX40), NKR-P1A (10/78), MHC II/RT1B (OX6), MHC II/RT1D (OX17), TCRαβ (R73), TCRγδ (V65), TCRVβ8.2 (R78), TCRVβ8.5 (B73), TCRVβ10 (G101), and TCRVβ16 (His42). Three-color flow cytometric analyses were performed with Abs labeled with FITC, PE, CyChrome, or PerCP. In some experiments, biotinylated Abs that were revealed with streptavidin-CyChrome (BD PharMingen) were used as third Ab. In the four-color analyses, unconjugated Abs were revealed with allophycocyanin-conjugated F(ab′)2 of goat anti-mouse IgG (Caltag Laboratories, Burlingame, CA). Flow cytometric analyses were performed as described previously (33). Stained cells were analyzed in a FACSCalibur flow cytometer (BD Biosciences, San Jose, CA) from the Servicio Común de Investigación (Faculty of Biology, Complutense University of Madrid. Madrid, Spain). The data were analyzed using PC-Lysis and CellQuest research softwares (BD Biosciences).

To isolate thymus and lymph node T cell subpopulations, cell suspensions were stained with anti-CD4 and anti-CD8 mAbs and purified by magnetic sorting using VarioMACS (Miltenyi Biotec, Bergisch Gladbach, Germany) in conjunction with an anti-FITC Multisort kit and anti-PE Microbeads (Miltenyi Biotec) according to the manufacturer’s instructions.

To determine the proportion of proliferating cells among peripheral T lymphocytes, cells were stained with either anti-CD4 and anti-TCRαβ, anti-CD8 and anti-TCRαβ, or anti-CD4 and anti-CD8 mAbs and then fixed with 30% ethanol and incubated with 7-amino actinomycin D as previously described (34). The proportion of dead cells after dexamethasone or anti-CD3 treatments was estimated by propidium iodide staining. Analyses were conducted in FACScan and FACSCalibur flow cytometers (BD Biosciences) using PC-Lysis and CellFit research softwares.

As previously described (35), neonatal rats were intrathymically injected with 5 μl of a solution of FITC (0.5 mg/ml; Sigma-Aldrich, Madrid, Spain) and adult rats with 20 μl of FITC (1 mg/ml). Control animals received the same amount of FITC dropped into the mediastinal cavity. After 14 h, the spleen, blood, and peripheral and mesenteric lymph nodes were harvested and stained as described above.

In some experiments, 20–24 h after intrathymic injection of FITC, rats were anesthetized and the thymic lobes were removed by suction. Seven days after thymectomy, the phenotype of FITC+ cells in the peripheral tissues was analyzed.

Peripheral T cell subpopulations were purified as described above and used to test the proliferative responses to plate-bound anti-CD3 (0.1–1 μg/ml), Con A (1–10 μg/ml), and recombinant human IL-2 (1–10 U/ml). Cultures (1 × 105 cells/well) were pulsed with 10 μM 5-bromo-2′-deoxyuridine (BrdU) for 12 h, and a specific kit from Boehringer Mannheim (BrdU Labeling and Detection kit III; Boehringer Mannheim, Mannheim, Germany) was used to measure BrdU incorporation into newly synthesized DNA (33). IL-2 production was estimated in the supernatants from those cultures using an ELISA kit specific for rat IL-2 (R&D Systems, Minneapolis, MN).

Conditions for pronase stripping and coreceptor re-expression followed the protocol developed by Suzuki et al. (36), with slight modifications as we described previously (35).

The basic procedures for HOS cultures have been previously described (35, 37). Thymic stromal cells were obtained from thymic fragments cultured with 1.35 mM 2-deoxyguanosine (Sigma-Aldrich España) for 7 days, trypsinized (0.25% trypsin in 0.02% EDTA; Sigma-Aldrich España), and depleted from residual thymocytes by treatment with anti-TCRαβ and anti-TCRγδ mAbs bound to sheep anti-mouse Ig-coated magnetic beads (Dynal, Oslo, Norway). Thymic stromal cells and magnetically purified peripheral DP lymphocytes were mixed at a ratio 1:2. The mixture of cells was inoculated into wells of a 96-well V-bottom plate in 0.2 ml of RPMI 1640 medium (Life Technologies, Grand Island, NY), supplemented with 10% FCS (Harlan Sera Lab, Leicestershire, U.K.), l-glutamine (2 mM), sodium pyruvate (1 mM), 2-ME (5 × 10−5 M), streptomycin (100 μg/ml), and penicillin (100 U/ml; all from Life Technologies). The plates were centrifuged at 500 × g for 3 min and placed into a plastic bag, then the air inside was replaced for a gas mixture (70% O2, 25% N2, and 5% CO2). The plastic bag was incubated at 37°C.

Peripheral DP lymphocytes (2 × 106) isolated from pronase-treated lymph node cell suspensions and lymph node CD4+ T cells were injected i.v. into thymectomized SCID mice. The animals were sacrificed at different time points, and their spleen, lymph node, and blood cells were used to analyze the expression of rat CD4 and CD8 Ags. An anti-TCRαβ mAb, which does not cross-react with the mouse molecule, was used to identify rat cells.

As others and we have described previously (4, 5), a low percentage (<3%) of DP T lymphocytes appears in the peripheral blood and secondary lymphoid organs of adult rats. However, higher proportions of DP T cells could be detected during fetal and postnatal life (Fig. 1,A). First peripheral DP T cells were detected on fetal day 20 (Fig. 1,B), when T cells start to colonize the peripheral lymphoid organs (35). During the perinatal period, the proportion of this cell subset rapidly increased, representing 30–40% of peripheral T cells (Fig. 1,B). In the following days, the percentage of DP cells gradually decreased to reach the low adult values by the fourth week of life (Fig. 1,B). Nevertheless, the absolute numbers of DP T cells gradually increased from the end of fetal life until the adult stage in the peripheral blood and all the secondary lymphoid organs analyzed (spleen and mesenteric and peripheral lymph nodes) (Fig. 1 C).

FIGURE 1.

Evolution of the percentages and absolute numbers of peripheral DP T cells during development. A, Rat lymph node cells from different developmental stages were incubated with anti-CD4, anti-CD8, and anti-TCRαβ mAbs. Dot plots show CD4 vs CD8 expression on gated TCRαβ+ cells. F, fetal; PN, postnatal. B, The proportion of CD4+CD8+ cells among TCRαβ+ lymphocytes from peripheral (PLN) and mesenteric (MLN) lymph nodes, spleen, and blood was estimated by flow cytometry in fetal, postnatal (P), and adult (A) developmental stages. C, The percentages of CD4+CD8+TCRαβ+ cells from lymph nodes, spleen, and blood were estimated by flow cytometry and used to calculate the total number of peripheral DP T cells during development. D, Expression of TCRαβ, TCRγδ, NKR-P1A, and CD8β on gated CD4+CD8α , CD4+CD8α+, and CD4CD8α+ T cell subsets from postnatal lymph nodes. Data are representative of 5–10 independent experiments. Similar results were obtained with cells from spleen and peripheral blood and from different developmental stages.

FIGURE 1.

Evolution of the percentages and absolute numbers of peripheral DP T cells during development. A, Rat lymph node cells from different developmental stages were incubated with anti-CD4, anti-CD8, and anti-TCRαβ mAbs. Dot plots show CD4 vs CD8 expression on gated TCRαβ+ cells. F, fetal; PN, postnatal. B, The proportion of CD4+CD8+ cells among TCRαβ+ lymphocytes from peripheral (PLN) and mesenteric (MLN) lymph nodes, spleen, and blood was estimated by flow cytometry in fetal, postnatal (P), and adult (A) developmental stages. C, The percentages of CD4+CD8+TCRαβ+ cells from lymph nodes, spleen, and blood were estimated by flow cytometry and used to calculate the total number of peripheral DP T cells during development. D, Expression of TCRαβ, TCRγδ, NKR-P1A, and CD8β on gated CD4+CD8α , CD4+CD8α+, and CD4CD8α+ T cell subsets from postnatal lymph nodes. Data are representative of 5–10 independent experiments. Similar results were obtained with cells from spleen and peripheral blood and from different developmental stages.

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We examined comparatively the phenotype of peripheral DP T cells and that of peripheral CD4 and CD8 single-positive (SP) lymphocytes. All DP T cells expressed TCRαβ with the same high intensity that peripheral SP T cells did (Fig. 1,D). No expression of TCRγδ and the NK cell marker NKR-P1A could be seen among DP T lymphocytes (Fig. 1,D). Interestingly, DP T cells were heterogeneous in the expression of the CD8β chain, and 60–70% of this cell subset corresponded to CD4+CD8α+β lymphocytes (Fig. 1,D), which suggested that the coreceptor was being expressed as an homodimer αα (38). Most peripheral T lymphocytes, including DP cells, expressed CD45RC, CD53, CD28, VLA-4, and L-selectin (Fig. 2,A). A high frequency (30–45%) of DP T cells stained for the activation marker CD25, whereas <10% of SP T cells were CD25+ (Fig. 2,A). However, the expression of other activation markers (CD71, CD134, and MHC class II) could be only detected in a low proportion of DP cells very similar to that found in the CD4 SP cell subset (Fig. 2,A). Accordingly, DP cell subset included small size lymphocytes like SP T cell subpopulations (FSC mean: CD4 402; DP 397; CD8 403). Furthermore, the proliferation index of DP T cells, although changed throughout development, was always very similar to that of SP T cells (Fig. 2 B), arguing against an activated condition of peripheral DP T cells.

FIGURE 2.

Phenotype of peripheral DP T cells. A, The expression of different Ags on gated CD4+CD8α , CD4+CD8α+, and CD4CD8α+ T cell subsets from mesenteric and peripheral lymph nodes is shown. Similar results were obtained with cells from spleen and peripheral blood, as well as from fetal, postnatal, and adult stages. B, Lymph node cells from different developmental stages were stained with anti-CD4, anti-CD8α, or anti-TCRαβ mAbs and incubated with 7-amino actinomycin D. The proportion of CD4, DP, and CD8 T lymphocytes in S+G2+M phases of the cell cycle was calculated. Similar results were obtained using splenocytes and peripheral blood cells. C, Lymph node cells were stained with anti-CD4, anti-CD8α, anti-CD8β, and mAbs specific for different TCRVβ segments. The proportion of TCRVβ+ cells in the CD4+CD8α CD8β, CD4+CD8α+CD8β, CD4+CD8α+CD8β+, and CD4CD8α+CD8β+ T lymphocyte subsets was estimated by flow cytometry.

FIGURE 2.

Phenotype of peripheral DP T cells. A, The expression of different Ags on gated CD4+CD8α , CD4+CD8α+, and CD4CD8α+ T cell subsets from mesenteric and peripheral lymph nodes is shown. Similar results were obtained with cells from spleen and peripheral blood, as well as from fetal, postnatal, and adult stages. B, Lymph node cells from different developmental stages were stained with anti-CD4, anti-CD8α, or anti-TCRαβ mAbs and incubated with 7-amino actinomycin D. The proportion of CD4, DP, and CD8 T lymphocytes in S+G2+M phases of the cell cycle was calculated. Similar results were obtained using splenocytes and peripheral blood cells. C, Lymph node cells were stained with anti-CD4, anti-CD8α, anti-CD8β, and mAbs specific for different TCRVβ segments. The proportion of TCRVβ+ cells in the CD4+CD8α CD8β, CD4+CD8α+CD8β, CD4+CD8α+CD8β+, and CD4CD8α+CD8β+ T lymphocyte subsets was estimated by flow cytometry.

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The comparison of TCRVβ usage among CD4 SP, CD8 SP, and DP T cell subsets showed that the expression of Vβ8.5 and Vβ16 was very similar in all peripheral T cells (Fig. 2,C). In contrast, the proportion of Vβ8.2- and Vβ10-expressing cells among CD4 SP and DP lymphocytes was higher than in the CD8 SP T cell subset (Fig. 2 C), suggesting a possible relationship between peripheral DP and CD4 SP lymphocytes.

No major differences in the above-described phenotypic characteristics were detected when DP T cells from different peripheral lymphoid organs (spleen, mesenteric and peripheral lymph nodes, and peripheral blood) or different developmental stages (fetal, postnatal, and adult) were compared (data not shown). No differences were observed either when comparing the phenotype of the two DP cell subsets defined according to CD8β expression (data not shown).

According to the expression of the CD8β chain, DP T cells could be subdivided into two subpopulations, one bearing the CD8αβ heterodimer (CD4+CD8α+β+; DPαβ) and the other presumably expressing CD8αα homodimers (CD4+CD8α+β; DPαα). These results suggested that DPαβ cells were of thymic origin, whereas DPαα cells could represent extrathymically derived T cells (39). To better determine the origin of peripheral DP cells, we analyzed the phenotype of the recent thymic emigrants (RTEs). For this purpose, neonatal and adult rats were intrathymically injected with FITC, and 14 h later FITC-labeled cells appearing in lymph nodes, spleen, and peripheral blood were analyzed. Both neonatal and adult RTEs were mostly TCRαβhigh (>98%) and included a consistent population of CD4+CD8α+ T cells, which represented 30–40% of the neonatal RTEs and <5% of the adult RTES (Fig. 3,A). Remarkably, only 30–40% of the neonatal and adult CD4+CD8α+ RTEs expressed the CD8β chain (Fig. 3 A) and 60–70% of the DP RTEs were CD4+CD8α+β, implying that both peripheral DP T cell subsets derive from the thymus.

FIGURE 3.

Intrathymic origin of peripheral DP T cells. A, Postnatal and adult rats were intrathymically injected with FITC and, after 14 h, lymph node cells were stained with anti-CD4, anti-CD8α, and anti-CD8β mAbs. The proportion of FITC+ cells is shown in the leftdot plots. Dot plots in the middle show CD4 vs CD8 expression among FITC+ RTEs. Histograms represent the expression of CD8β within the CD4+CD8α+ cell subset of RTEs. The percentage of TCRαβhigh cells among FITC+ RTEs was always ≥97%. Similar results were observed in spleen and peripheral blood. B, Thymocytes were stained with anti-CD4, anti-CD8α, anti-CD8β, and anti-TCRαβ mAbs. The left histogram shows the expression of TCRαβ in the total thymocyte population and the proportion of TCRαβhigh thymocytes. The dot plot in the middle represents the CD4 vs CD8α expression among TCRαβhigh thymocytes. The right histogram shows the expression of CD8β within the CD4+CD8α+TCRαβhigh thymocyte subpopulation.

FIGURE 3.

Intrathymic origin of peripheral DP T cells. A, Postnatal and adult rats were intrathymically injected with FITC and, after 14 h, lymph node cells were stained with anti-CD4, anti-CD8α, and anti-CD8β mAbs. The proportion of FITC+ cells is shown in the leftdot plots. Dot plots in the middle show CD4 vs CD8 expression among FITC+ RTEs. Histograms represent the expression of CD8β within the CD4+CD8α+ cell subset of RTEs. The percentage of TCRαβhigh cells among FITC+ RTEs was always ≥97%. Similar results were observed in spleen and peripheral blood. B, Thymocytes were stained with anti-CD4, anti-CD8α, anti-CD8β, and anti-TCRαβ mAbs. The left histogram shows the expression of TCRαβ in the total thymocyte population and the proportion of TCRαβhigh thymocytes. The dot plot in the middle represents the CD4 vs CD8α expression among TCRαβhigh thymocytes. The right histogram shows the expression of CD8β within the CD4+CD8α+TCRαβhigh thymocyte subpopulation.

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The intrathymic origin of both DPαα and DPαβ cells was further confirmed by the occurrence of these cell subpopulations in the thymus. The analysis of the expression of CD4, CD8α, and CD8β in the neonatal and adult TCRαβhigh thymocyte subsets revealed that ∼70% of DP TCRαβhigh thymocytes coexpressed CD8α and CD8β, whereas the remaining 30% of DP TCRαβhigh cells only expressed the CD8α chain (Fig. 3 B).

Therefore, these results show that both peripheral DP T cell subsets, DPαα and DPαβ, come from thymocyte subpopulations which would be exported by the thymus.

Once established the relationship between peripheral DP lymphocytes and some thymocyte subpopulations, we analyzed the functional characteristics of these DP T cells to know their maturational state. First we studied the sensitivity of the peripheral T cell subsets and immature DP thymocytes to undergo apoptosis in response to corticosteroids. As previously reported (40), DP thymocytes were extremely sensitive to glucocorticoid-mediated cell death, whereas all peripheral T cell subsets, including DP cells, were comparatively resistant to glucocorticoid-induced apoptosis, even with the highest concentrations of dexamethasone used (Fig. 4 A). These results demonstrate that peripheral DP cells have already acquired the resistance to glucocorticoid-induced death, a feature of the positively selected lymphocytes (41).

FIGURE 4.

Functional features of peripheral DP T cells. A, The resistance of DP thymocytes and different peripheral T cell subsets to undergo apoptosis in response to dexamethasone was assayed by exposing the T cell subpopulations to the indicated doses of dexamethasone for 20 h. The proportion of dead cells was estimated by propidium iodide staining. B, The proliferative responses of peripheral T cell subsets to Con A and plate-bound anti-CD3 Abs were estimated after 48 h of culture. BrdU was added for the last 12 h and the BrdU incorporation to newly synthesized DNA was measured as described in Materials and Methods. C, The proportion of dead cells in anti-CD3-stimulated cultures of either DP thymocytes or distinct peripheral T cell subsets was measured by propidium iodide staining. D, The proliferative response to different doses of plate-bound anti-CD3 Abs in the presence of IL-2 was analyzed after 48 h of culture. BrdU incorporation was estimated by a BrdU-labeling kit. E, The supernatants from anti-CD3-stimulated cultures using either CD4, DP, or CD8 T cell subsets were assayed for IL-2 production using an ELISA-specific kit.

FIGURE 4.

Functional features of peripheral DP T cells. A, The resistance of DP thymocytes and different peripheral T cell subsets to undergo apoptosis in response to dexamethasone was assayed by exposing the T cell subpopulations to the indicated doses of dexamethasone for 20 h. The proportion of dead cells was estimated by propidium iodide staining. B, The proliferative responses of peripheral T cell subsets to Con A and plate-bound anti-CD3 Abs were estimated after 48 h of culture. BrdU was added for the last 12 h and the BrdU incorporation to newly synthesized DNA was measured as described in Materials and Methods. C, The proportion of dead cells in anti-CD3-stimulated cultures of either DP thymocytes or distinct peripheral T cell subsets was measured by propidium iodide staining. D, The proliferative response to different doses of plate-bound anti-CD3 Abs in the presence of IL-2 was analyzed after 48 h of culture. BrdU incorporation was estimated by a BrdU-labeling kit. E, The supernatants from anti-CD3-stimulated cultures using either CD4, DP, or CD8 T cell subsets were assayed for IL-2 production using an ELISA-specific kit.

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We next analyzed the proliferative response of DP T cells to different stimulators, including Con A and immobilized anti-CD3 mAbs. As shown in Fig. 4,B, peripheral DP cells exhibited a poor proliferative response to any of the concentrations of Con A or anti-CD3 mAbs used, whereas CD4 SP and CD8 SP T lymphocytes proliferated vigorously in response to the same stimuli. The reduced ability of the DP T cells to expand upon stimulation prompted us to examine whether these cells would undergo apoptosis after TCR stimulation, as it has been described for DP thymocytes (42). Propidium iodide staining was used to quantify cell death after stimulation with anti-CD3 mAbs. The results of this analysis showed that DP T cells, as well as SP T cells, were resistant to CD3-induced apoptosis (Fig. 4,C). By contrast, DP thymocytes were highly susceptible (Fig. 4 C). Therefore, a high incidence of apoptosis was not the explanation to the low responsiveness to CD3 stimulation exhibited by peripheral DP T cells.

Interestingly, all peripheral T cell subpopulations showed similar proliferative responses upon IL-2 stimulation (Fig. 4,D). Then we analyzed the effect of IL-2 addition to anti-CD3-stimulated cultures. In the presence of IL-2, at any of the concentrations used, DP T cells could proliferate in response to CD3 ligation at the same level as CD4 SP and CD8 SP T cells (Fig. 4,D). These data strongly suggested that the unresponsiveness status of DP T cells could be due to a defect in IL-2 production. To support this view, we measured the amount of IL-2 in the supernatants from anti-CD3-stimulated cultures. The results showed, confirming our hypothesis, a minimal production of IL-2 by stimulated DP T cells in comparison to both CD4 and CD8 SP T cell subpopulations (Fig. 4 E).

On the other hand, no significant differences were observed when comparing the functional characteristics of postnatal and adult DP T cells (data not shown).

The results shown above demonstrate that rat DP T lymphocytes represent a peripheral T cell subset which has not reached its total functional capabilities and originates from immature thymocyte subpopulations. To know their evolution and role in the periphery, we first analyzed the possible relationship between the DP T cells and the other peripheral T cell subsets. Some of the data from the phenotypic study, like the expression of TCRVβ fragments and some other molecules such as CD71, CD134, MHC class II, and CD62L, suggested a possible relationship between DP and CD4 SP T cells. In addition, although the level of expression of CD4 in DP T cells was comparable to that in CD4 SP T cells, the CD8 expression levels were variable, which may imply the existence of a down-regulation process of CD8α and CD8β molecules in correlation to a differentiation process. Thus, we examined whether all peripheral T cells were actively expressing both CD4 and CD8 Ags, since thymocytes have been found to retain coreceptor proteins on the cell surface for several hours even after active biosynthesis of the molecule was terminated. For this purpose, DP lymphocytes were isolated and assessed for active coreceptor synthesis using the pronase-stripping/coreceptor re-expression assay devised by Suzuki et al. (36). Interestingly, 20–25% of DP T cells were able to re-express only the CD4 coreceptor after pronase stripping, indicating that they have already terminated the synthesis of CD8 coreceptor (Fig. 5 A).

FIGURE 5.

Peripheral DP T cells become CD4 SP T cells. A, Magnetically sorted DP T lymphocytes were subjected to coreceptor stripping by pronase treatment, and the re-expressed CD4 and CD8 molecules were analyzed after the resting period. B and C, Generation of CD4 SP T cells after culturing peripheral CD4+CD8α+CD8β+/− (B) and CD4+CD8α+CD8β+ (C) lymphocytes with thymic stromal cells (ratio 2:1) in HOS-reaggregation cultures for different time periods. Dot plots represent CD4 vs CD8α expression on gated TCRαβ+ cells, and histograms (C) show CD8β expression on the indicated CD4- and DP-gated cells. D, Postnatal rats were intrathymically injected with FITC and thymectomized 24 h later. CD4 vs CD8α expression on FITC+ RTEs was analyzed 14 h after the intrathymic labeling (iT initial) and 7 days after the intrathymic labeling and subsequent thymectomy (iT plus TX 7 days).

FIGURE 5.

Peripheral DP T cells become CD4 SP T cells. A, Magnetically sorted DP T lymphocytes were subjected to coreceptor stripping by pronase treatment, and the re-expressed CD4 and CD8 molecules were analyzed after the resting period. B and C, Generation of CD4 SP T cells after culturing peripheral CD4+CD8α+CD8β+/− (B) and CD4+CD8α+CD8β+ (C) lymphocytes with thymic stromal cells (ratio 2:1) in HOS-reaggregation cultures for different time periods. Dot plots represent CD4 vs CD8α expression on gated TCRαβ+ cells, and histograms (C) show CD8β expression on the indicated CD4- and DP-gated cells. D, Postnatal rats were intrathymically injected with FITC and thymectomized 24 h later. CD4 vs CD8α expression on FITC+ RTEs was analyzed 14 h after the intrathymic labeling (iT initial) and 7 days after the intrathymic labeling and subsequent thymectomy (iT plus TX 7 days).

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Once we demonstrated that a fraction of DP T cells are in fact CD4 SP T cells, we studied the ability of peripheral DP lymphocytes to differentiate to the CD4 cell lineage. We conducted HOS reaggregation cultures with peripheral DP T cells and thymic stromal cells, an in vitro assay in which we have previously shown that DP cells differentiate into SP cells (35). The DP T cell population used in these experiments was first isolated according to CD4 and CD8α expression and then included DPαα and DPαβ cells. Since the majority of DP T cells experienced a gradual maturation to CD4 SP T cells throughout the culture period (Fig. 5,B), this implies that both DP cell subpopulations are able to give rise to CD4 SP T cells. In another set of reaggregation cultures, we used DPαβ T cells, isolated according to CD4 and CD8β expression, and after 4 days, both DPαα and CD4 SP cells appeared in the cultures (Fig. 5 C). All of these results demonstrate that peripheral DP T cells are able to become CD4 SP T cells, presumably according to a sequential differentiation pathway which includes the following steps: DPαβ→DPαα→CD4.

Subsequently, we studied whether the maturational transition from DP to CD4 SP cells seen in vitro could also occur in an in vivo model. To address this issue, we analyzed the evolution of a peripheral T cell population since its migration from the thymus. Neonatal rats were then injected intrathymically with FITC to label thymocytes and thymectomized 24 h later to prevent continued emigration of FITC+ thymocytes. As shown in Fig. 5 D, 7 days after the intrathymic injection, the FITC+ peripheral T cell population contained a decreased proportion of DP cells in conjunction with an increased CD4+ cell subset, whereas the CD8+ cell subpopulation remained virtually unchanged. Because CD4 and DP lymphocytes showed similar proportions of apoptotic and cycling cells (data not shown), the decreased numbers of DP cells indicate that the differentiation of DP lymphocytes into CD4 SP cells also occurs in vivo in the peripheral organs.

We extended the study of this differentiation process by injecting i.v. purified rat DP T cells into SCID mice. To avoid, in the injected cell population, the presence of those DP T cells that have already initiated the differentiation to the CD4 cell lineage, peripheral T cells were subjected to coreceptor stripping by pronase treatment and subsequent re-expression culture, previously to the isolation of the DP T lymphocytes. In this way, DP cells that had already terminated CD8 synthesis exhibited a CD4+ SP phenotype after the re-expression culture, and they were not isolated in the DP cell population. Likewise, to prevent a possible thymic influence in the differentiation process, SCID mice were thymectomized before transferring the DP T cells. Under these conditions, rat DP T lymphocytes experienced a gradual differentiation to the CD4 cell lineage throughout a 50-day period (Fig. 6). This process could be observed in all of the secondary lymphoid organs studied, as well as in peripheral blood (Fig. 6), indicating that DP T cells properly home to lymphoid organs.

FIGURE 6.

Peripheral DP T cells develop into CD4 T cells in the absence of thymus. SCID mice were thymectomized and i.v. injected with DP T cells isolated from pronase-treated lymph node cell suspensions. Rat CD4 vs CD8α expression was analyzed on gated rat TCRαβ+ cells (histograms) present in different peripheral lymphoid tissues and at different days postinjection. PLN, Peripheral lymph node; MLN, mesenteric lymph node.

FIGURE 6.

Peripheral DP T cells develop into CD4 T cells in the absence of thymus. SCID mice were thymectomized and i.v. injected with DP T cells isolated from pronase-treated lymph node cell suspensions. Rat CD4 vs CD8α expression was analyzed on gated rat TCRαβ+ cells (histograms) present in different peripheral lymphoid tissues and at different days postinjection. PLN, Peripheral lymph node; MLN, mesenteric lymph node.

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The phenotypic analysis of the CD4 progeny of i.v. injected DP T cells showed that the expression of both CD8β and CD25 Ags had been down-regulated in parallel to the expression of the CD8α chain (Fig. 7,A). Thus, CD4 SP cells derived from DP cells showed an identical phenotype to that exhibited by fresh CD4 SP T cells (Figs. 1,D and 2A), whereas CD4+CD8low cells expressed an intermediate phenotype (Fig. 7 A).

FIGURE 7.

The progeny of i.v. injected DP T cells is functionally immunocompetent. A, The expression of CD8β and CD25 on rat CD4+CD8α+, CD4+CD8αint, and CD4+CD8α T cell subsets was analyzed 50 days after the injection of rat DP T cells into thymectomized SCID mice. B, Evolution of the total cell number of the rat T cell subsets found in lymph nodes, spleen, and peripheral blood from thymectomized SCID mice i.v. injected with rat peripheral DP T cells. C–E, Rat CD4DP and DP T cells were obtained 50 days after the injection of peripheral DP T cells into thymectomized SCID mice. CD4CD4 T cells were isolated from SCID mice injected with peripheral CD4 T cells. The three T cell subsets were assayed for their proliferative responses to plate-bound anti-CD3 Abs (C) and IL-2 (D) and their ability to produce IL-2 upon anti-CD3 stimulation (E).

FIGURE 7.

The progeny of i.v. injected DP T cells is functionally immunocompetent. A, The expression of CD8β and CD25 on rat CD4+CD8α+, CD4+CD8αint, and CD4+CD8α T cell subsets was analyzed 50 days after the injection of rat DP T cells into thymectomized SCID mice. B, Evolution of the total cell number of the rat T cell subsets found in lymph nodes, spleen, and peripheral blood from thymectomized SCID mice i.v. injected with rat peripheral DP T cells. C–E, Rat CD4DP and DP T cells were obtained 50 days after the injection of peripheral DP T cells into thymectomized SCID mice. CD4CD4 T cells were isolated from SCID mice injected with peripheral CD4 T cells. The three T cell subsets were assayed for their proliferative responses to plate-bound anti-CD3 Abs (C) and IL-2 (D) and their ability to produce IL-2 upon anti-CD3 stimulation (E).

Close modal

The numbers of CD4 SP T cells generated after transfer of DP cells also increased progressively, so that 4 × 104 CD4 SP cells detected on day 1 post-transfer yielded 10 × 106 cells after 50 days (Fig. 7 B), indicating that rat donor cells not only populated host lymphoid organs, but also survived and proliferated there.

Despite the evidence that peripheral DP T cells can develop into phenotypically mature, long-lived CD4 SP T cells, it was critical to demonstrate that these cells were immunocompetent. We therefore tested the peripheral progeny of injected DP cells for their ability to proliferate in response to CD3 ligation. We isolated rat CD4 SP cells derived from SCID mice i.v. injected with either rat peripheral DP T cells (CD4DP cells) or peripheral CD4+CD8 T cells (CD4CD4 cells), as well as rat peripheral DP T cells, and comparatively measured their proliferative responses to immobilized anti-CD3 Abs. As shown in Fig. 7,C, CD4DP cells proliferated in response to anti-CD3 stimulation as vigorously as CD4CD4 cells, whereas their DP precursors exhibited a poor response. Additionally, the three lymphocyte populations showed a similarly high response to IL-2 (Fig. 7,D) and when the IL-2 production in the anti-CD3-stimulated cultures was measured, similar concentrations of IL-2 were found to be produced by stimulated CD4CD4 and CD4DP cells, in contrast to the minimal production by DP T cells (Fig. 7 E). Then, the gain of function of CD4DP cells with respect to their DP precursors runs parallel to the acquisition of their IL-2 production ability.

These results unequivocally establish that peripheral DP T lymphocytes develop into functional CD4 SP progeny after i.v. injection and demonstrate a lack of requirement for the thymic microenvironment in the final maturation of some CD4+CD8 T cells.

In normal adult rats, a small subpopulation of T cells coexpressing CD4 and CD8 coreceptors appears in the periphery in addition to the typical helper CD4 SP and cytotoxic CD8 SP T cells (4, 5). In this study, we show that these peripheral DP cells are already present among the first T cells colonizing the peripheral lymphoid organs during fetal life and represent 30–40% of peripheral T lymphocytes during the perinatal period. Later during neonatal life, the proportion of DP T cells gradually decreases to reach the low values seen in adulthood, although in absolute terms the numbers of DP T cells gradually increase from the end of fetal life until the adult stage in all of the peripheral lymphoid organs analyzed. Similar to rats, the proportion of peripheral DP T cells in adult mice and humans is very low (1, 2, 3, 6) and is transiently increased around birth time (2, 11, 12, 13, 14). In contrast, adult swine, monkeys, and some chickens have prominent populations of peripheral DP T cells (7, 8, 9, 10) whose proportions gradually increase with age (7, 10, 43).

The phenotype of the rat DP T cells described in our study is very similar to that reported by Kenny et al. (4). One of the most outstanding phenotypic features of DP T lymphocytes is the heterogeneous expression of CD8β chain that allows subdividing them into cells expressing the CD8αβ heterodimer and cells that presumably express CD8αα homodimers. The proportions of these DP T cell subpopulations in the different peripheral lymphoid organs remain constant throughout life, and no major differences can be observed when comparing their phenotypes. Traditionally, it has been considered that the expression of CD8β requires intrathymic T cell maturation, while CD8αα homodimer-expressing cells arise extrathymically (39). However, we demonstrate that both rat DP T cell subpopulations, DPαβ and DPαα, are of thymic origin. They both appear among RTEs after intrathymic injection of FITC and derive from TCRαβhigh thymocyte subpopulations clearly identifiable at any developmental stage. Supporting the intrathymic origin of peripheral DP lymphocytes, Kenny et al. (4) described that more than half of rat DP T cells expressed Thy-1, a cell marker for rat RTEs (44). Likewise, thymectomy eliminates partial or totally the number of peripheral DP T cells in rats (Ref. 4) and our unpublished results) and humans (18), but not in swine (45). Bonomo et al. (11) also demonstrated by intrathymic injection of FITC the thymic origin of peripheral DP lymphocytes in neonatal mice, and Res et al. (14) concluded that human cord blood DP T cells are derived from a mature DP thymocyte subset that can emigrate from the thymus.

In contrast, peripheral DP lymphocytes from swine, chickens, and monkeys, as well as some of those appearing in adult humans are not considered as thymus-derived cells but as CD4+CD8 T cells that re-express CD8 after activation. This is due to the lack of expression of CD8β and CD1 molecules and/or because they show functional and phenotypic properties of memory T cells (8, 9, 10, 28, 45). It is unlikely, however, that rat DP T cells represent activated T lymphocytes. They exhibit a naive CD45RChigh phenotype and a small lymphocyte morphology, they do not show a higher proliferative rate than SP T cell subsets and, although a high proportion of those cells express CD25, only a few of them express other cell markers like CD134, CD71, or MHC Ags that are up-regulated upon T cell activation in the rat (46, 47, 48). Additionally, the proportion of CD25+ cells detected among peripheral DP T cells is very similar to that found among DP RTEs as soon as 3 h after intrathymic FITC injection (data not shown), which argues against the acquisition of this cell marker after activation in the periphery.

Although the resistance to undergo apoptosis in response to corticosteroids indicates that rat peripheral DP T cells have been already submitted to a positive selection signal (41), the poor proliferative response to Con A or anti-CD3 mAbs, but not to IL-2, points out that these cells are not fully immunocompetent. That deficient response to CD3/TCR stimulation is not caused by the lack of responding cells, since peripheral DP T cells are as resistant as SP T cells to CD3-induced apoptosis, but by the inability of stimulated DP lymphocytes to produce IL-2. Therefore, rat DP T cells constitute a peripheral T cell subset that has not reached the full functional maturity, similar to that found by Kay et al. (27) in the peripheral blood of healthy humans. These peripheral DP lymphocytes share many similarities with functionally immature human CD4+CD8α+CD8β+/−TCRαβhigh (49) and mouse CD4+CD8αβlowTCRαβhigh thymocyte subsets (50) which have been positively selected but have not acquired a fully developed helper function, since they respond poorly to different proliferative stimuli and cannot produce high levels of cytokines upon activation. Hyporesponsive DP T cells have not been described in swine or monkeys (30, 45), in which, on the contrary, these lymphocytes seem to represent a primed T cell subset.

The next question to be elucidated is the physiological relevance of the DP T lymphocyte subset in the periphery. Given that CD25 is expressed by a high proportion of rat DP T cells, Kenny et al. (4) have recently postulated that they could represent regulatory T cells involved in preventing autoimmunity (51, 52), instead of thymocyte subpopulations which have not completed their functional maturation and have been released by the thymus, as we demonstrate in the current study. However, the same authors discarded that possibility because CD25 does not seem to be a definitive marker for regulatory T cells in the rat (53). In agreement with them, we show that DP T cells do not share many of the typical features described for the regulatory CD4+CD25+ T cells: 1) <50% of DP T cells appearing in the different developmental stages express CD25; 2) they do proliferate in response to exogenous IL-2 alone as vigorously as peripheral SP T cells; and 3) when they mature and down-regulate CD8αβ expression, they also down-regulate the expression of CD25 and acquire the ability to respond to stimulation through the TCR and to produce IL-2 at the same levels as SP T cells. Furthermore, the proliferative responses of CD4+CD25 T cells to Con A or soluble anti-CD3 mAbs were not suppressed by rat DP T cells when mixed in various ratios (our unpublished results).

Previously, Bonomo et al. (11) demonstrated that some immature DP T cells prematurely escaped from the mouse thymus and proposed that those cells may complete their process of maturation in the peripheral lymphoid tissues. In agreement with this, our results clearly demonstrate that rat peripheral DP T lymphocytes gradually differentiate into CD4 SP T cells, presumably following a sequence of differentiation which includes the stages CD4+CD8α+CD8β+→ CD4+CD8α+CD8β→ CD4+CD8αCD8β. A similar cell stage sequence has been described by Vanhecke et al. (49) during the intrathymic differentiation of human DP thymocytes to CD4 SP cells.

The CD4 progeny originated from DP T cells is fully functional, as judged by its ability to survive and expand in peripheral lymphoid organs, to proliferate in response to CD3 ligation, and to produce IL-2 upon stimulation. Therefore, we can conclude that rat peripheral DP lymphocytes are partially immunocompetent T cells that belong to the CD4 cell lineage and undergo post-thymic phenotypic and functional maturation in the peripheral lymphoid tissues. Similar post-thymic differentiation processes affecting CD4+ T cells have been previously reported in mice and rats (44, 54, 55) and have been shown to be independent of the presence of the thymus (44, 56), a fact also confirmed by our results. The nature of the signal(s) required for post-thymic development of some CD4+ T cells is unknown at present. Peripheral MHC class II expression has been shown to be necessary for the long-term survival and optimal expansion of CD4+ T cells (57, 58, 59); however, the involvement of other signals mediated by cytokines, costimulatory molecules, and/or extracellular matrix components cannot be discarded.

Finally, in view of all of these results, it seems evident that important species-specific differences among peripheral DP T cells exist. Nevertheless, before assuming that DP lymphocytes could represent different T cell subsets in distinct animal species, additional studies should revise some issues as the possible different functionality of DP T lymphocytes in young and adult individuals, which could reflect a different composition of the DP T cell subset throughout life in the same species. Likewise, the lack of expression of CD8β and CD1 molecules has been the only feature to consider DP T cells as extrathymically derived cells in some species. However, other experimental approaches seem to be necessary to definitively discard the intrathymic origin of DP T cells in those species, still more when it has been described in the existence of CD4+CD8α+CD8βCD1TCRαβhigh thymocytes which are able to migrate from the thymus (14, 49).

We thank Adriana Bonomo for invaluable help with the intrathymic injections of neonatal rats.

1

This work was supported by grants from the Ministerio de Educación y Cultura (PB97-0332), Fondo de Investigaciones Sanitarias (FIS98/0041), Comunidad de Madrid (08.3/0014/1997, 08.3/0027/1998, and 08.3/0041/2000), and Ministerio de Ciencia y Tecnología (PM99-0060).

3

Abbreviations used in this paper: DP, double positive; HOS, high oxygen submersion; RTE, recent thymic emigrant; SP, single positive; BrdU, 5-bromo-2′-deoxyuridine.

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