Impaired CD8 T cell memory and CD4 T cell primary responses in IL-7Rα mutant mice

Generation of IL-7Rα^449F mice

To analyze the role of signaling effectors downstream of IL-7Rα on in vivo T lymphocyte function, we used homologous recombination with a replacement vector to generate the IL-7Rα^449F knock-in mice (Fig. 1 A; and Fig. S1 and Supplemental Materials and methods available at http://www.jem.org/cgi/content/org/jem.20061871/DC1). The targeted IL-7Rα locus retained the endogenous promoter and upstream exons; however, exons 6–8 were replaced with cDNA encoding the transmembrane and cytoplasmic domains, with disruption of the YXXM motif by the Y449F substitution (Fig. 1 A). The phosphorylated IL-7Rα YXXM motif has been shown to be the binding site for the Src homology 2 domain of PI3 kinase and activation of downstream pathways (30–32). The Neo^R selection marker was deleted by Cre-mediated recombination to eliminate potential promoter effects. Initial analysis of mutant IL-7Rα protein expression from thymocyte lysates indicated patterns comparable to WT thymocytes with mature polypeptides (90 kD) (Fig. 1 B). Most importantly, we determined that the Y449F knock-in mutation did not affect surface expression of IL-7Rα on peripheral CD4 and CD8 T cells (Fig. 1 C). Furthermore, signaling analysis of WT and IL-7Rα^449F–cultured T cell blasts showed that TCR activation of the Erk and p38 MAPK pathways was unaffected, but that IL-7 activation of STAT5 requires IL-7Rα Y449 (Fig. 1 D). Interestingly, IL-7 by itself does not induce phosphorylation of Akt (Fig. 1 D) or S6 kinase (unpublished data) in primary murine T cells at the time points tested.

Development of early hematopoietic lineages in IL-7Rα^449F mice

To perform a comprehensive analysis of hematopoietic development in IL-7Rα^449F mice, we investigated the effect of loss of IL-7Rα Y449 signaling during development of early hematopoietic lineages using flow cytometry. We found the total number of bone marrow hematopoietic stem cells (HSCs) and derivatives were unaffected by the mutation (Fig. 2 A). Once again, we determined that the Y449F knock-in mutation does not affect surface expression of IL-7Rα on HSCs (Fig. 2 B). However, the earliest thymic T cell progenitors were severely impaired in both IL-7Rα mutants as indicated by a 20-fold decrease in the total number of early thymic progenitors (ETPs; characterized as Lin⁻CD3⁻CD4⁻CD8⁻CD25⁻CD117⁺ [33]) in IL-7Rα^449F and IL-7Rα^−/− mice in comparison to WT (Fig. 2 C).

IL-7Rα^449F thymocytes bypass a developmental block

Complete disruption of IL-7Rα results in decreased survival of early T lymphocyte progenitors causing a severe cytopenia that precludes detailed analysis of its role in peripheral T cells (1, 2, 26). In contrast, IL-7Rα^449F mice showed only a fourfold decrease in thymic cellularity compared with WT despite the severe ETP defect (Fig. 3 A, right), suggesting a role for IL-7Rα Y449-independent signaling in thymocyte development. In agreement with this hypothesis, the distribution of thymocyte subsets in IL-7Rα^449F mice was indistinguishable from WT (Fig. 3 A, left). To further examine thymocyte development, WT, IL-7Rα^449F, and IL-7Rα^−/− thymocytes were analyzed by expression of CD44 and CD25 in the CD4⁻CD8⁻ DN compartment using flow cytometry. At all DN stages (DNI, CD4⁻CD8⁻CD44⁺CD25⁻; DNII, CD4⁻CD8⁻CD44⁺CD25⁺; DNIII, CD4⁻CD8⁻CD44⁻ CD25⁺; DNIV, CD4⁻CD8⁻CD44⁻CD25⁻), the defects in IL-7Rα^449F were less severe than in IL-7Rα^−/− mice (Fig. 3 B, left). In addition, IL-7Rα^449F mice had significantly higher numbers of DP thymocytes than IL-7Rα^−/− mice (Fig. 3 A). This could be explained by the observation that IL-7Rα^449F and WT mice showed similar production of DP thymocytes from DNIV precursors,whereas IL-7Rα^−/− DNIV cells encountered a severe developmental block at this stage (Fig. 3 B, right). This indicates an IL-7Rα Y449–independent compensatory mechanism that contributes to restoration of normal thymocyte development and partial rescue of total T cell numbers. The role of IL-7Rα signaling in positive selection was examined by determining the frequency of TCR-β⁺ DP thymocytes that had downregulated heat stable antigen (HSA) (Fig. 3 C) or upregulated the maturation marker CD69 (unpublished data). These analyses showed that neither complete deletion nor mutation of IL-7Rα resulted in impaired positive selection, and we conclude that IL-7Rα is nonessential in this process.

Commitment to the γδ T cell lineage is IL-7Rα dependent (34) and occurs late in the DNII to DNIII stage (35). We assessed γδ development by flow cytometry by staining total thymocytes with anti-γδ antibody and a lineage cocktail. Our data showed that γδ development is Y449 dependent because their numbers were markedly diminished (Fig. 3 D). Collectively, our data shows that IL-7Rα Y449 signals play critical roles in both αβ and γδ T cell development.

To examine the basis of restored cellularity in IL-7Rα^449F mice, cell turnover was determined by BrdU incorporation to monitor uptake by rapidly cycling DN thymocytes. We found IL-7Rα^449F DNIV cells that lack surface expression of the mutated IL-7Rα (unpublished data) have markedly decreased BrdU uptake, whereas the DNIII and DP stages undergo more rapid cycling compared with WT (Fig. 3 E). Collectively, the data suggest that IL-7Rα–driven cell division in DN and DP thymocytes is Y449 independent.

To examine the severity of the IL-7Rα^449F mutation on developmental potential, competitive repopulation experiments were performed. This analysis revealed that IL-7Rα^449F thymocytes were less robust than their WT counterparts. Similar to what was seen in whole mice, when either WT or IL-7Rα^449F whole bone marrow were delivered separately, IL-7Rα^449F recipient thymic development resulted in fourfold lower cellularity than WT recipients (unpublished data). However, when WT (CD90.1⁺) and IL-7Rα^449F (CD90.2⁺) whole bone marrow were delivered at a 1:1 ratio, the thymic output was comprised of <5% IL-7Rα^449F–derived cells, although subset distribution was still normal (Fig. 3 F). Collectively, our data shows that knock-in disruption of IL-7Rα Y449 signals causes stage-specific defects, particularly dramatic at the ETP stage, but that these are bypassed to generate normal thymocyte subsets.

IL-7Rα Y449-independent signals are involved in peripheral T cell development

The severe developmental defect in IL-7Rα^−/− thymocytes (1) causes a paucity of T cells in the periphery that confounds further analysis of the role of IL-7 in T cell function (3). When peripheral T cells were examined in IL-7Rα^449F mice, we found that the relative subset distribution is similar between IL-7Rα^449F and WT mice, whereas IL-7Rα^−/− mice had a severe disruption of CD4 and CD8 T cell development (Fig. 4 A). Quantification of peripheral lymphocytes revealed that IL-7Rα^449F mice had significantly lower total cell numbers than WT mice (CD4 T cells: 2.5 ± 0.3; CD8 T cells: 3.8 ± 0.6; B cells: 3.7± 1.0 fold decrease, n = 3) but that these subsets were higher than in IL-7Rα^−/− mice, thus indicating that lymphocyte development is partially IL-7Rα Y449 independent (Fig. 4 B). Consistent with the expectation that this targeted mutation only affects IL-7Rα–mediated events, development of NK cells (NK1.1⁺CD3⁻) was unaffected in the spleens of both IL-7Rα^449F and IL-7Rα^−/− mice (Fig. 4 B). TCR-β and CD3ɛ expression levels on WT, IL-7Rα^449F, and IL-7Rα^−/− splenic T cells were analyzed by flow cytometry and found to be comparable between WT and IL-7Rα^449F mice, whereas those from IL-7Rα^−/− mice were severely deficient (Fig. 4 C). This shows that IL-7Rα Y449 signaling is dispensable for peripheral T cell maturation as assessed by antigen receptor expression.

To eliminate the possibility of altered expression of other γc receptor family members in IL-7Rα^449F mice accounting for differences in peripheral T cell function, surface expression levels of cytokine receptor chains γc and IL-2/15Rβ were examined. Our data showed only marginal differences in IL-7Rα and γc expression between WT and IL-7Rα^449F CD4 and CD8 T cells, although the frequency of IL-2/15Rβ–expressing cells was increased in both IL-7Rα^449F and IL-7Rα^−/− T cells (Fig. S2, available at http://www.jem.org/cgi/content/org/jem.20061871/DC1).

Homeostatic proliferation is IL-7Rα Y449 dependent

Because IL-7Rα^449F mice are able to support T cell development, we sought to address what aspects of T cell function were impaired or retained. IL-7 has been shown to be involved in homeostatic proliferation of naive CD4 and CD8 T cells (9, 13, 14). To address the role of IL-7Rα Y449 in homeostatic proliferation, IL-7Rα^449F and WT (CD45.2⁺) T cells were labeled with the mitotic tracker CFSE and transferred into either normal or irradiated recipient host mice (CD45.1⁺). In stark contrast to WT T cells, IL-7Rα^449F T cells were unable to proliferate when transferred into acutely lymphopenic hosts as measured by CFSE dilution (Fig. 5). Indeed the deficiency was as severe as that in IL-7Rα^−/− cells. A small pool of CD8 T cells undergo limited rounds of division, but these appear IL-7Rα independent because it was also observed with IL-7Rα^−/− cells (Fig. 5). Our data strongly argue that homeostatic proliferation of both CD4 and CD8 T cells is dependent on signals initiated by IL-7Rα Y449.

IL-7Rα Y449 is essential for CD4 primary immune responses

To determine the role of IL-7Rα Y449 in generating a functional polyclonal T cell immune response in vivo, WT and IL-7Rα^449F mice were infected with a recombinant Listeria monocytogenes strain expressing an immunodominant foreign, MHC class I (K^b)–restricted peptide, SIYRYYGL (rLM-SIY) (Priatel, J., L. Zenewicz, H. Shen, and H. Teh, personal communication). Primary responses to rLM-SIY were monitored at day 7, and antigen-specific T cells were enumerated by IFN-γ production in CD4 or CD8 T cells after in vitro stimulation with immunodominant MHC class II (listeriolysin O [LLO]_190-201)– and MHC class I (SIY)–restricted peptides, respectively. Surprisingly, IL-7Rα^449F CD4 T cells failed to generate a detectable primary immune response after infection (Fig. 6 A). In contrast, IL-7Rα^449F CD8 T cells mounted a robust response that was equivalent to WT controls in total cell numbers (Fig. 6 A), independent of lower initial T cell numbers in IL-7Rα^449F mice. In vitro anti-CD3/anti-CD28 TCR stimulation assays showed that IL-7Rα^449F CD4 and CD8 T cells had impaired proliferation response to low, but not high, levels of TCR stimulation (Fig. 6 B) and this may account for the defective CD4 primary response.

To determine whether or not the decrease in B cells or an altered splenic compartment could account for the lack of CD4 T cell primary response in IL-7Rα^449F mice, equal numbers of WT Thy1.1 (CD90.1⁺) T cells were adoptively transferred into WT and IL-7Rα^449F CD90.2⁺ hosts. The mice were infected with rLM-SIY and responses measured at day 7 after infection by determining IFN-γ expression upon in vitro stimulation with the indicated peptides. The fact that WT CD90.1⁺ CD4 T cells responded strongly in IL-7Rα^449F hosts (Fig. 6 C) confirmed that the IL-7Rα^449F CD4 defect was T cell intrinsic and not a result of inefficient antigen presentation. These data show that CD4, but not CD8 T cells, require IL-7Rα Y449–mediated signals to generate primary responses to rLM-SIY.

IL-7Rα Y449 is essential for maintenance of CD8 T cell memory

Previous reports have suggested that IL-7Rα is critical for the generation and/or maintenance of T cell memory. The fact that IL-7Rα^449F mice mount a strong primary CD8 T cell response to the MHC class I–restricted SIY peptide enabled us to investigate whether the formation of CD8 T cell memory is dependent on IL-7Rα Y449. WT and IL-7Rα^449F mice were infected with rLM-SIY and analyzed 45 d later for antigen-specific memory CD8 T cells by expression of IFN-γ in response to peptide. Despite generating equivalent numbers of CD8 T cell effectors as WT at day 7, IL-7Rα^449F mice showed an eightfold reduction in memory CD8 T cell numbers compared with WT mice (Fig. 7 A). Not surprisingly, the severely impaired primary CD4 T cell response in IL-7Rα^449F mice resulted in very few LLO-reactive memory CD4 T cells. This experiment indicates an essential role for IL-7Rα Y449 signaling in regulation of CD8 memory T cells.

To test the fitness of IL-7Rα^449F memory T cells, WT and IL-7Rα^449F mice previously immunized with rLM-SIY were homologously rechallenged at 100 d after infection. 3 d after reinfection, the IL-7Rα^449F mice showed an increase in the number of SIY-specific CD8 T cells but failed to reach the same level as achieved by WT controls (Fig. 7 B).

Because IL-7Rα^449F T cells have impaired proliferation at low dose stimulus of the TCR, the possibility existed that the number of IL-7Rα^449F memory CD8 T cells was underestimated by assessing their ability to produce IFN-γ upon peptide challenge. To confirm our findings of decreased CD8 memory T cells, an H2-K^b MHC I–Ig fusion protein conjugated to PE (MHC–Ig dimer) was loaded with SIY peptide and used for detection of antigen-specific cells. As shown in Fig. 7 C, the number of IL-7Rα^449F memory CD8 T cells remains significantly lower (2.5-fold) than WT at day 60, although the decrease is not quite as dramatic as that shown by IFN-γ induction. This suggests that antigen-specific IL-7Rα^449F memory CD8 T cells have compromised TCR-mediated cytokine induction. Strikingly, when memory T cells were assayed at day 100 in a parallel experiment, the WT CD8 memory T cell numbers remained constant, whereas IL-7Rα^449F CD8 memory T cell numbers continued to decrease (Fig. 7 C). These data demonstrate a requirement for IL-7Rα Y449 signals in maintenance and long term survival of memory CD8 T cells.

To address whether or not the lack of a primary CD4 T helper cell expansion in the IL-7Rα^449F mice was affecting the generation of an efficient memory CD8 T cell compartment, we performed adoptive transfer experiments wherein WT CD90.1⁺ T cells were transferred into IL-7Rα^449F (CD90.2⁺) hosts. This allows WT and IL-7Rα^449F CD8 T cells to encounter antigen in the same environment and receive the same signals from activated WT CD4 T helper cells. WT and IL-7Rα^449F antigen-specific CD8 T cells were enumerated using MHC–Ig dimer staining at day 7 and 45 after rLM-SIY infection. To account for differences in transplant versus host cell numbers within spleens, we normalized the number of antigen-specific CD8 memory T cells against the number of antigen-specific primary CD8 T cells and represented this as a percentage. The normal range of effector T cells that survive the contraction period to become memory cells is between 5 and 10%. We saw that >15% of WT but <5% of IL-7Rα^449F effector CD8 T cells were maintained at memory stage at day 45 after infection (Fig. 7 D), refuting the hypothesis that lack of CD4 T cell help accounts for the defect in IL-7Rα^449F CD8 T cell memory.

We therefore looked at possible cell intrinsic disruptions in IL-7Rα^449F CD8 memory T cells. Memory T cells require a balance of cell survival and low-level proliferation for long-term maintenance. Previous studies have demonstrated a role for IL-7 in survival of memory CD8 T cells by regulating levels of the survival factor Bcl-2 (9, 36). Using intracellular staining and flow cytometry, we found that the levels of Bcl-2 in WT and IL-7Rα^449F CD8⁺ IFN-γ⁺ memory T cells were equivalent after rechallenge (Fig. 7 E) and at day 45 after infection (not depicted), and therefore cannot account for the maintenance defect. IL-15 has been demonstrated to regulate proliferation of memory CD8 T cells (6–8), but it remained possible that the IL-7Rα^449F memory CD8 T cell pool was decreasing over time because of decreased proliferation. In agreement with previous studies that cycling of memory cells is regulated by IL-15 (37), we found that the cycling of IL-7Rα^449F memory CD8 T cells was not disrupted as shown by BrdU incorporation (Fig. 7 F). Thus, the defect in IL-7Rα^449F memory CD8 T cells does not result from impaired cycling or from a lack of Bcl-2 survival signals.

STAT5, but not Bcl-2, is IL-7Rα Y449 dependent

To more fully address the requirement of IL-7Rα Y449 in activation of known IL-7 signaling pathways, WT and IL-7Rα^449F thymocytes and splenocytes were stimulated ex vivo with purified, recombinant IL-7 and analyzed by flow cytometry for phosphorylation of STAT5 and up-regulation of Bcl-2. This analysis showed that STAT5 activation was entirely IL-7Rα Y449 dependent in all IL-7–responsive subsets (thymic DN, CD4 and CD8, and splenic CD4 and CD8 T cells) (Fig. 8 A). Surprisingly, two prosurvival members of the Bcl-2 family, Bcl-2 and Bcl-xL, were IL-7Rα Y449 independent (Fig. 8 B and unpublished data). Analysis of Bcl-2 regulation showed that IL-7Rα^449F DN thymocytes had decreased levels of Bcl-2 after 24 h ex vivo culture compared with WT and that they were severely impaired in their ability to up-regulate Bcl-2 in response to IL-7 (Fig. 8 B). However, the remaining thymocyte subsets and peripheral T cells showed significant induction of Bcl-2 expression at the same time points (Fig. 8 B) leading to the conclusion that IL-7–mediated Bcl-2 regulation is mostly Y449 independent. Our findings support the hypothesis that IL-7–induced Bcl-2 is important during the DN stages of development to impart survival signals that allow differentiation events to occur (26, 27), but importantly, refutes the notion that Bcl-2 is sufficient in IL-7Rα–mediated events in peripheral T cells, particularly maintenance of CD8 memory T cells.

Mice.

Animals were housed at the University of British Columbia, Microbiology and Immunology Department Animal Facility in accordance with University of British Columbia Animal Care and Biosafety Committee certificates. C57BL/6, B6.SJL-Ptprc^a Pepc^b (BoyJ), B6.PL-Thy1^a/CyJ (Thy1.1⁺), and IL-7Rα^−/− mice were obtained from The Jackson Laboratory.

IL-7Rα^449F mice generation.

IL-7Rα^449F mice were generated by homologous recombination as detailed in Supplemental Materials and methods (available at http://www.jem.org/cgi/content/org/jem.20061871/DC1). A replacement vector was generated in pKSloxPNT (51) (provided by Robert Farese, Gladstone Institute of Cardiovascular Disease, University of California, San Francisco, CA) that substituted exons 6–8 encoding the transmembrane and cytoplasmic domains of IL-7Rα with a partial cDNA (nucleotides 707–1,398) encoding a Y-to-F mutation at residue 449 and an SV40 polyadenylation signal. The targeted locus would retain regulation by the endogenous promoter. RF8 ES cells (129/SvJae origin) (52) were electroporated with linearized vector, selected, and targeted ES clones were identified by genomic Southern analysis (see Supplemental Materials and methods). Two out of three targeted clones were transiently transfected with MC-Cre (provided by Shinya Yamanaka, Institute for Frontier Medical Sciences, Kyoto University, Kyoto, Japan) to delete the loxP-flanked Neo^R cassette and subclones verified as described (see Supplemental Materials and methods). IL-7Rα^+/449F founders were generated by standard procedures and backcrossed at least six generations with C57BL/6 mice. PCR genotyping was performed with primers A (5′ TCTTCCTGAACAGCCAG), B (5′ TCTCTTCTGTGAGCTACGG), and C (5′ CCCTGAACCTGAAACATAAA) for 30 cycles each of 30 s at 94°C, 30 s at 60°C, and 1 min at 72°C. Primers A and B amplify a 489-bp fragment from the WT allele, whereas primers A and C amplify a 595-bp fragment from the knock-in allele.

Antibodies.

FITC, PE, CyC, APC, Alexa 647, APC Cy7, PE Cy7, and biotinylated antibodies were obtained from BD Biosciences (anti-γδ TCR, NK1.1, B220, Gr-1, Mac-1, CD3ɛ, CD4, CD8α, CD25, CD44, CD69, CD90.1, CD90.2, c-kit, IL-2/15Rβ, γc, HSA, Sca-1, TCR-β, Bcl-2, phospho-STAT5, BrdU, and H-2Kb–Ig recombinant fusion protein, and mouse DimerXI) or eBioscience (anti-CD4, CD8α, CD45.2, and IL-7Rα). Unlabeled antibodies were obtained from Cell Signaling Technology (anti-Akt, phospho-Akt Ser473, phospho-STAT5, phospho-Erk, Erk, and p38 MAPK), BD Biosciences (anti-phospho p38 MAPK), Santa Cruz Biotechnology, Inc. (anti–IL-7Rα), and Sigma-Aldrich (anti–α-tubulin). Rabbit polyclonal anti-STAT5A and STAT5B were a gift from Dr. Lothar Hennighausen (National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD). Fluorescent secondary antibodies used in the Odyssey Infrared Imaging System (LiCOR) were obtained from Rockland Immunochemicals (goat anti–rabbit IgG IRDye800) and Invitrogen (goat anti–mouse IgG Alexa 680). Horseradish peroxidase–conjugated secondary antibodies used in chemiluminescent immunoblotting were from GE Healthcare.

Flow cytometry.

Surface staining for stem cell and lymphocyte characterization was performed according to standard procedures. For analysis of phospho-STAT5 activation, cells were acid stripped (10 mM Na citrate, pH 4, 140 mM NaCl) and starved for 4 h in serum-free RPMI + 1% BSA. Cells were stimulated by addition of 25 ng/ml recombinant murine IL-7 (Chemicon) for 10 min at 37°C in serum-free media. Cells were fixed for 10 min in 1% PFA at RT and permeabilized in 100% ice-cold methanol. Cells were then stained for detection of phospho-STAT5, CD4, and CD8. For detection of Bcl-2 up-regulation, cells were cytokine stripped and serum starved as described for phospho-STAT5, and stimulated with IL-7 for 20 h at 37°C. Cells were fixed and permeablized for 1 h at RT in 1% PFA, 0.1% Tween-20 and stained for surface markers and intracellular Bcl-2. Samples were collected on either a FACSCalibur or LSRII (BD Biosciences), and data were analyzed with FlowJo software (Tree Star).

Cell cycle analysis.

Two doses of 1 mg BrdU were delivered i.p. to WT and IL-7Rα^449F mice at 4-h intervals. 20 h after the second BrdU dose, thymi were harvested and stained for surface markers (for DN, α-CD25, CD44, and lineage cocktail α-CD3, CD4, CD8, γδ TCR, Gr-1, Mac1 B220; for DP, α-CD4, CD8). Cells were then treated according to manufacturer's instructions for detection of BrdU (BD Biosciences).

Western blotting.

Samples were lysed in lysis buffer (10 mM Tris-HCl, pH 7.5, 5 mM EDTA, 1% Triton X-100), and cleared lysates were quantified and normalized for total protein level. For immunoblot analysis of IL-7Rα expression, total thymocyte lysates were resolved by SDS-PAGE, and proteins were electrophoretically transferred to nitrocellulose. A polyclonal anti–IL-7Rα from Santa Cruz Biotechnology, Inc. (sc-662) was used for detection and visualized with donkey anti–rabbit HRP and SuperSignal West Pico ECL reagents (Pierce Chemical Co.). For analysis of signaling pathways, WT and IL-7Rα^449F CD8 T cells were cultured for 48 h at 10⁶ cells/ml in the presence of 2 μg/ml ConA in complete media, then maintained in 20 U/ml IL-2 for 3 d, and finally, cytokine stripped and serum starved for 4 h. Cells were stimulated with 25 ng/ml IL-7, 1 μg soluble α-CD3, or 10 nM IL-2. Western blots were probed for detection of phosphorylated and total levels of Akt, p38 MAPK, Erk, and STAT5 using the LiCor Odyssey imaging system.

Homeostatic proliferation.

Spleens were collected from mice and prepared as single cell suspensions. Red blood cells were lysed using ACK lysis buffer (155 mM NH₄Cl, 10 mM KHCO₃, 100 μM EDTA). Lymphocytes were B cell depleted using sheep anti–mouse IgG DynaBeads (Invitrogen). Purified cells were CFSE labeled as previously described (53). T cell subsets were quantified by flow cytometry, and 5 × 10⁶ T cells were transferred i.v. to CD45.1 congenic mice that either had or had not received a sublethal dose of irradiation the previous day (600 rads). After 7 d, spleens of recipient mice were harvested and stained for CD45.2, CD4, and CD8α.

Competitive repopulation.

Bone marrow cells were isolated from the femurs of Thy1.1 and IL-7Rα^449F mice. After red blood cell lysis, 2 × 10⁶ whole bone marrow cells from Thy1.1 (CD45.2⁺, CD90.1⁺) and IL-7Rα^449F (CD45.2⁺, CD90.2⁺) donors were adoptively transferred into lethally irradiated (1200 rads) BoyJ (CD45.1⁺) congenic recipients. Recipient mice received Biosol (neomycin sulfate) at 2 mg/ml in their water the night before irradiation and for the next 2 wk. At 4 wk after injection, recipient thymuses were harvested and stained for CD45.2, CD90.1, CD90.2, CD4, and CD8α.

In vitro TCR stimulation assay.

Peripheral T cells were purified from the spleens of WT and IL-7Rα^449F mice using α-CD4 and α-CD8 MACS beads (Miltenyi Biotec) according to manufacturer's instructions. Purified, CFSE-labeled CD4 and CD8 WT and IL-7Rα^449F T cells were cultured in vitro for 3 d in RPMI + 10% FBS in the presence of 1 μg α-CD28 and the indicated doses of plate-bound α-CD3. Cells were harvested, stained for CD4 and CD8, and dilution of CFSE was monitored by cytometry.

L. monocytogenes infection and response assays.

Naive C57BL/6 and IL-7Rα^449F mice were injected intravenously with 10⁵ CFU or previously immunized mice infected with 10⁶ CFU of rLM-SIY in PBS. At appropriate time points after infection, spleens were collected, homogenized to single cell suspension, red blood cells lysed, and the cells restimulated in vitro with epitope-specific peptides for 5 h at 37°C in the presence of 1 μl/10⁶ cells Golgi plug (BD Biosciences). The CD4-specific LLO peptide was added at 5 μM and the CD8-specific SIY peptide at 1 μM in Iscove's media + 10% FCS. After stimulation, cells were fixed and permeabilized in 2% PFA, 0.2% Tween-20 for 20 min at RT. Cells were then stained for CD4, CD8α, and IFN-γ. Antigen-specific cells were designated as CD4⁺ or CD8⁺ cells that expressed IFN-γ after in vitro challenge with antigenic peptides. SIY peptide loading of the BD Biosciences MHC I–Ig dimer fusion protein was performed according to manufacturer's instructions with a 40-M excess of peptide. For flow cytometric analysis of SIY-specific CD8 T cells using peptide loaded dimer, 0.5 μg dimer/2 × 10⁶ cells was used.

For adoptive transfer experiments, WT T cells were purified from Thy1.1 mice by B cell depletion, and 10⁷ T cells were injected intravenously into C57BL/6 or IL-7Rα^449F recipients. The next day, recipient mice were infected with 10⁵ CFU of rLM-SIY, and analysis was performed as outlined in the previous paragraph (with addition of anti-CD90.1) at 7 d after infection. Analysis of Bcl-2 levels and BrdU incorporation was performed with or without peptide stimulation and without IL-7 stimulation. Bcl-2 intracellular staining was performed as described previously. For cell cycle analysis, mice were given a single 2-mg dose of BrdU i.p. and then fed BrdU in their water for 8 d (0.8 mg/ml). Staining was performed according to manufacturer's instructions (BD Sciences).

Statistical analysis.

All data are presented as mean ± SEM and analyzed by Student's t test. Significance was set at p-value of <0.05.

Online supplemental material.

Fig. S1 shows genomic Southern blot analysis of ES clones with knock-in targeted IL-7Rα loci. Fig. S2 shows that IL-7Rα^449F peripheral T cells have largely unperturbed γc and IL-2/15Rβ receptor subunits. Supplemental Materials and methods provides details of the knock-in targeting of the IL-7Rα locus. Online supplemental material available at http://www.jem.org/cgi/content/org/jem.20061871/DC1.