Study subjects were recruited from the University of North Carolina, Chapel Hill (UNC-CH) and the NC Children’s Hospital. CB samples from full-term infants were collected immediately after birth or obtained from the Carolinas Cord Blood Bank. Direct capillary or venous blood samples (0.5–3.0 mL) from infants age 2 weeks to 14 months were collected by nursing or phlebotomy personnel during routine physician visits. The infants consisted of similar numbers of males and females. The exact number of infants per age group is listed in Table 1; we had 70 CB samples, 35 samples from infants between the ages of 1 and 9 months and 64 samples from 1 year olds (10–14 months). The large number of infants (n = 169) was required because limited blood volumes (0.5–2.0 mL) from infants age 2 weeks to 1 year restricted the number of assays and parameters that could be analyzed. The infant participants were recruited from the North Carolina population that consists of about 46% Caucasians, 46% African-Americans, and 8% other groups, and within the ethnic categories, 10% represent Hispanics.

Exclusion criteria included HIV-positive status of the mother, known in utero infections, treatment of the mother with immunosuppressive drugs, diagnosis of mother or child with immunosuppressive disorder, life-threatening malformations of the infant or life expectancy <6 months. Infant blood samples were also excluded if the infant had a bleeding disorder or had a chronic infection. The Virology, Immunology, and Microbiology Core of the UNC Center for AIDS Research provided blood samples from healthy adults. Age, sex, and race of the adult donors were unknown. The study was approved by the UNC-CH Institutional Review Board, and informed parental consent was obtained. Institutional guidelines strictly adhere to the World Medical Association’s Declaration of Helsinki.

Cord blood from full-term infants was collected into CB collection bags containing CPD anticoagulant, whereas all other blood samples were collected into EDTA-containing blood tubes. Blood samples were processed within 24 h. Whole blood was used for Phosflow™ analysis of JAK2 and STAT proteins; for all other assays, peripheral blood mononuclear cells (PBMCs) were isolated by gradient density separation using Lymphocyte Separation Medium (MP Biomedicals, OH) as described (55, 56).

CD4⁺ T cells were enriched by magnetic bead separation using the CD4⁺ T cell enrichment Kit from Stemcell Technologies (Vancouver, BC, Canada) according to the manufacturer’s instructions. The purity of CD4⁺ T cell populations was confirmed by flow cytometry and exceeded 96%.

Peripheral blood mononuclear cells were stained with CD3 (SP34-2)-Pacific Blue, CD4 (L200)-PerCPCy5.5, CD8 (RPA-T8)-Alexa Fluor700, CD45RA (5H9)-FITC, and CCR7 (3D12)-PeCy7 to determine naïve [T(N): CD45RA⁺CCR7⁺], central memory [T(CM): CD45RA⁻CCR7⁺], and effector/effector memory cell [T(Eff/EM), CD45RA^±CCR7⁻] T cell populations. All antibodies, unless noted otherwise, were purchased from BD Biosciences (San Jose, CA, USA). The expression of cytokine receptors on CD4⁺T cells was quantitated using PE-labeled antibodies: IFN-γR1 (CD119-GIR-208), IFN-γR2 (2HUB-159, Biolegend, San Diego, CA, USA), IL-4Rα (hIL4R-M57), IL-2Rα (CD25-clone-4E3, Miltenyi Biotec, Auburn, CA, USA), or IL-2Rβ (CD122-clone-Mikβ). Cytokine receptor expression is reported as percentage of CD4⁺ T cell populations or quantitated on a per cell basis using a standard curve generated with Quantibrite PE beads (BD Biosciences). The staining protocol was performed at 4°C. Samples were acquired on a LSRII instrument (BD Biosciences) by collecting a minimum of 300,000 events. The flow cytometry data were analyzed using FlowJo Software (Tree Star, Ashland, OR, USA), version 9.8. Antibodies were titrated prior to use and gating was based on Fluorescence Minus One (FMO) and isotype antibody controls.

Phosflow™ analysis was performed to measure the activation of JAK2 after IFN-γ stimulation or the activation of STAT proteins according to the manufacturer’s protocols (BD Biosciences) using Perm Buffer III (CD4⁺ T cells) or Perm Buffer IV (NK cells). Although most JAK and STAT molecules contain multiple tyrosine and/or serine phosphorylation sites, the analysis was limited to a single phosphorylation site. Briefly, 300–400 µL of whole blood was treated for 10 min with 10 µg/mL of human recombinant IFN-γ (Miltenyi Biotec) and samples were then stained with antibodies to total JAK2 or with antibodies to phosphorylated JAK2 (Tyr1008; clone D4A8, Cell Signaling Technology, Danvers, MA, USA). Phospho-JAK2 (pJAK2) was detected with anti-rabbit IgG (H + L), F(ab’)2 Fragment (Alexa Fluor 647, Cell Signaling Technology). To detect the phosphorylation of specific STAT proteins after stimulation with IFN-γ IL-4, IL-2, or IL-7, whole blood was incubated for 15 min with 10 µg/mL of recombinant IFN-γ, 0.1 µg/mL of recombinant IL-4 (R&D, Minneapolis, MN, USA) or 0.1 µg/mL human IL-2 or IL-7 (Miltenyi Biotec), respectively, and subsequently stained for phosphorylated STAT-1 (Tyr 701; clone 58D6, Cell Signaling Technology), STAT6 (Y641; clone pY641) or STAT5 (Y694; clone pY694, both BD Biosciences), respectively. Non-phosphorylated STAT1, STAT6, or STAT5 proteins were measured in parallel. Unstimulated whole blood served as negative control. STAT4 phosphorylation (pSTAT4 Y693) was measured using the same protocol, except that PBMCs were stimulated for 45 min with IL-12 (75 ng/mL) in serum-free RPMI 1640 media. To distinguish CD4⁺ T cells with different antigen experience, blood samples were costained with CD3 (SP34-2)-PeCy7, CD4 (L200)-PerCPCy5.5, and CD27 (L128)-PE, and naïve, central memory, and effector/effector memory populations were defined as being CD27⁺, CD27^low, or CD27⁻, respectively (57). Cytokine concentrations and incubation times were determined in preliminary experiments (data not shown). Samples were acquired on a LSRII instrument analyzed with FlowJo Software. Cells positive for JAK2 or STAT phosphorylation are reported as percentage of CD4⁺ T cells after subtracting the values from the untreated samples. Antibodies were titrated prior to use and gating was based on FMO and isotype antibody controls. In addition, a subset of samples was analyzed on the Amnis^® ImageStreamX instrument according to the manufacturer’s protocol.

The phosphorylation of JAK3 in response to stimulation with IL-4 or IL-2 was measured by Western blot. CD4⁺ T cells were enriched as described above and either stimulated immediately with IL-4 or IL-2 or preactivated for 2 days prior to cytokine stimulation. For the latter, CD4⁺ T cells were stimulated at a concentration of 2–4 × 10⁶ cells/mL with plate-bound anti-CD3 (10 µg/mL; clone SP34-2), precoated overnight in a 24-well plate, and cultured at 37°C and 5% CO₂. Controls were set up in parallel with media only. After 48 h, to activate JAK3, CD4⁺ T cells were stimulated for 5 min with 0.1 µg/mL recombinant human IL-4 or IL-2 at 37°C in a water-bath followed by a 5-min centrifugation at 300 × g (total stimulation time: 10 min). Cells were lysed with 100 µL of Blue Loading Pack (Cell Signaling Technology) per 4 × 10⁶ cells. Protein concentrations were determined using the Pierce 660 nm Assay (Thermo Scientific, Rockford, IL, USA). Samples were loaded onto 8% Novex^® Tris-Glycine gels (Novex^®, Life Technologies, NY, USA). The XCell SureLock™ Mini Cell was used for electrophoresis in Novex^® Tris-Glycine SDS running buffer and for subsequent transfer using Novex^® Tris-Glycine transfer buffer. Transfer PVDF membranes were obtained from Bio-Rad Laboratories (Hercules, CA, USA). All detection antibodies and reagents were obtained from Cell Signaling Technology unless noted otherwise. To determine JAK3 phosphorylation, the membranes were incubated with phospho-JAK3 antibody (Tyr980/981; clone D44E3; 1:1,000 dilution) in 1× TBST overnight, stripped using Restore Plus Western blot (Pierce/Thermo Scientific, Rockville, IL, USA) and re-probed for non-phosphorylated JAK3 (clone D1H3; 1:1,000 dilution:) and beta (β)-Actin (clone 13E5; 1:50,000 dilution) as loading control. The membranes were then incubated with secondary anti-Rabbit IgG HRP-linked antibody at a dilution of 1:2,000 and developed using the Amersham™ ECL Prime Western Blotting Detection Reagent (GE Healthcare Life Sciences, Pittsburgh, PA, USA). Images were captured using the Bio-Rad ChemiDoc MP System and densitometric measurements were obtained using the Image Lab Software Version 5.2.1. Protein sizes were verified based on the NuPage HiMark Prestained Protein Standard (Life Technologies, Grand Island, NY, USA).

Data were analyzed using GraphPad Prism Software (Version 6, 2014; La Jolla, CA, USA). Continuous measurements were compared between independent age groups using non-parametric Mann–Whitney tests. Statistical comparisons were only performed if the sample size was n ≥ 4 in both age groups. Associations between continuous variables was estimated using non-parametric Spearman rank correlation. To visualize the age-dependent changes in STAT activation in response to IL-4, IFN-γ, and IL-2, a LOESS trend curve was generated with a smooth parameter of 0.5. Only statistically significant p-values are shown in the figures. A 0.05 significance level was used with no adjustment for multiple hypothesis testing.

Cytokine receptor expression was evaluated both by quantifying the number of receptors per CD4⁺ T cell and by determining the percentage of CD4⁺ T cells expressing each cytokine receptor. Two main comparisons were performed. First, we determined whether there was a significant change in cytokine receptor expression from birth to 1 year of age. Second, we tested whether the expression of cytokine receptors differed between 1-year-old infants and adults. These comparisons were conducted both for total CD4⁺ T cells and for CD4⁺ T cell subsets. The analysis of CD4⁺ T cells according to their phenotypic characterization as naïve [T(N); CD45RA⁺CCR7⁺], central memory [T(CM); CD45RA⁻CCR7⁺], or effector memory/effector [T(EM/Eff); CD45RA^±CCR7⁻] CD4⁺ T cells was important because the majority of infant peripheral blood CD4⁺ T cells represent naïve CD4⁺ T cells and the relative frequencies of these CD4⁺ T cell population change with age and pathogen exposure. Thus, this analysis was necessary to accurately evaluate age-related differences in infant CD4⁺ T cells from birth to 1 year, and equally critical for the comparison of infant and adult CD4⁺ T cell parameters.

Cord blood CD4⁺ T cells expressed significantly more IL-4Rα molecules per CD4⁺ T cell than adults (Table 2). Although there was an age-dependent decline in median IL-4Rα numbers/CD4⁺ T cell comparing birth to 1 year, these differences were not statistically significant. The percentage of IL-4Rα⁺CD4⁺ T cells also did not vary substantially with age (Table 3). To initiate signaling upon IL-4 binding, the IL-4Rα chain dimerizes with the common γ-chain (γc); we did, however, not measure the common γ-chain, because previous studies have already shown that it is expressed at lower levels in infants (58).

The IFN-γ receptor consists of the IFN-γR1, promoting IFN-γ binding, and the IFN-γR2 that mediates signaling to the JAK kinases. The number of IFN-γR1 molecules per CD4⁺ T cell did not significantly differ between birth and 1 year (Table 2). CD4⁺ T cells in CB and in blood from 1-year-old infants expressed significantly higher numbers of IFN-γR1/CD4⁺ T cell than adults (Table 2; Figure 1A). In contrast, the percentage of CD4⁺ T cells expressing IFN-γR1 increased from birth to 1 year, and even at 1 year, the percentage of IFN-γR1⁺CD4⁺ T cells was significantly lower compared to adults (Table 3, Figure 1A). This difference was especially pronounced in the subset of effector memory/effector CD4⁺ T cells (Table 3; Figure 1B). We did not observe significant changes in total expression levels of IFN-γR2 (Table 2), and although the frequencies of naïve and effector memory/effector IFN-γR2 CD4⁺ T cells seemed to decrease in the first year, the biological significance of the changes is questionable given their very low frequencies (Table 3). Note that the relatively low frequencies of IFN-γR1 or IFN-γR2 CD4⁺ T cells in infants were not the result of poor staining, as IFN-γR-positive B cells could be easily enumerated in the same samples (Figure 1C). The analysis of the ratio of relative expression levels of IL-4Rα to IFN-γR1 or IL-4Rα to IFN-γR2 on infant CD4⁺ T cells did not reveal changes that would support a switch from a more Th2-biased response at birth versus a more balanced Th2/Th1 response at 1 year.

An analysis of the IL-2R was included because T cell expansion is a critical step in T cell differentiation. The IL-2R consists of three distinct chains, the α, β, and γ-chain. The IL-2Rα chain (or CD25) is of special importance as it is a marker of regulatory T cells (Tregs), which occur in higher frequencies in infant compared to adult blood, but CD25 is also expressed on activated T cells that are more frequent in adult peripheral blood. Both, absolute numbers of IL-2Rα per CD4⁺ T cell and the percentage of IL-2Rα⁺CD4⁺ T cells were higher at 1 year of age than at birth (Tables 2 and 3; Figure 1D). At 1 year, the number of IL-2Rα molecules per CD4⁺ T cells exceeded that of adult CD4⁺ T cells (Table 2), whereas the percentage of IL-2Rα⁺CD4⁺ T cells was significantly lower compared to adults (Table 3). Whether or not these results reflected the dichotomy of the CD25 molecule as a marker of Treg or activated CD4⁺ T cells was not a focus of the study. However, soluble CD25 (sCD25) plasma levels were higher at 1 year compared to CB (Figure 1D), and there was a significant positive correlation between plasma sCD25 levels and the percentage of IL-2Rα⁺CD4⁺ T cells (Figure 1D). Although these results were indicative of increasing activated CD4⁺ T cells with age, higher sCD25 plasma levels (Figure 1D) and IL-2Rα⁺CD4⁺ T cells (Table 3) were already increased by 1–3 months and did not further increase from 3 to 14 months.

The expression levels of IL-2Rβ did not differ between infant CD4⁺ T cells at birth and or at 1 year, or between CD4⁺ T cells of 1-year-old infants and adults. We did not measure the expression of the common γ-chain (see above).

The results highlight that naïve, central memory, and effector memory CD4⁺ T cells show age-dependent differences in cytokine receptor expression that are likely to influence the functional capacity of these CD4⁺ T cell subsets.

As the Th2-bias of infant CD4⁺ T cells is well documented, we hypothesized that CB CD4⁺ T cells would activate the JAK/STAT signaling pathway in response to IL-4 as effectively as adult CD4⁺ T cells. Upon binding of IL-4 to its receptor, JAK1 and JAK3 will be activated by engaging the IL-4Rα or the common γ chain, respectively. As the common γ chain is also part of the IL-2 receptor, we chose to measure JAK3 activation to maximize the number of parameters that could be analyzed considering the small sample volumes available. The limited blood volumes for 10- to 14-month-old infants also required that we pooled PBMCs from multiple infants (pool 1: n = 9; pool 2: n = 5) to enrich CD4⁺ T cells for Western blot analysis.

The protein levels of total JAK3 were comparable between CD4⁺ T cells isolated from CB, 1-year olds or adults (Figure 2A). With the caveat that exact frequencies of pJAK3⁺CD4⁺ T cells could not be quantified by Western blot, CD4⁺ T cells isolated from CB or 1-year-old infants were equally able to activate pJAK3 in response to IL-4 (Figures 2B,C). Thus, densitometry analysis showed no statistically significant differences in the increase of median pJAK3 densities in IL-4 stimulated compared to unstimulated CD4⁺ T cells in CB (19-fold; range: 4.9- to 25.4-fold), samples of 1 year olds (13-fold; range: 10.7- to 15.4-fold), and adults (2.5-fold; range: 2.2- to 11.6-fold).

The effective activation of pJAK3 by IL-4 in CB was confirmed by the fact that the downstream IL-4-specifc transcription factor STAT6 was activated in 64% (median value) of CB CD4⁺ T cells, although subject-to-subject variation at birth was high (range: 14–99%; Figure 2D). Despite the seemingly potent induction of the JAK/STAT signaling pathway by IL-4 in CB, median frequencies of pSTAT6⁺CD4⁺ T cells were lower compared to adults (93%; range: 60–98%). The activation of pSTAT6 significantly increased with age (Figure 2D), and by 4–6 months of age, pSTAT6 activation was comparable to 1-year-old infants and to adults (Figure 2D). The age-dependent increase in IL-4-induced pSTAT6 activation observed in the total CD4⁺ T cell populations appeared to be mainly reflective of changes in naive CD4⁺ T cells (Figure 2D). Although memory CD4⁺ T cells also responded with higher pSTAT6 activation by 4–6 months and at 1 year, pSTAT6 activation remained below [T(CM)] or within the lower range [T(EM/Eff)] of adult memory CD4⁺ T cells (Figure 2D). However, as naïve cells represented the majority of infant CD4⁺ T cells, the reduced pSTAT6 activation by infant memory cells was masked in the total CD4⁺ T cell population.

To test for the translocation of activated pSTAT6 to the nucleus, a critical event for downstream gene induction, we utilized Amnis^® ImageStreamX analysis. CB and adult CD4⁺ T cells showed similar expression of unphosphorylated STAT6 (Figure 3). Upon IL-4-stimulation, activated pSTAT6 was clearly detectable in the nucleus of CB CD4⁺ T cells, but the fluorescence intensity was slightly stronger in CD4⁺ T cells of an 8-month-old infant and in adult CD4⁺ T cells (Figure 3).

In contrast to JAK3 activation by IL-4, only a small fraction of CB CD4⁺ T cells induced JAK2 phosphorylation in response to IFN-γ stimulation (Figure 4A). The histograms in Figure 4B are representative for CB and adult CD4⁺ T cells and demonstrate that unstimulated CD4⁺ T cells in both age groups have similar low levels of pJAK2 and similar background level staining in FMO controls, but clearly differ in the levels of pJAK2 after IFN-γ stimulation. However, by 1 year of age, pJAK2 activation was even higher than in adult CD4⁺ T cells (Figure 4A). To more specifically determine when infant CD4⁺ T cell acquired improved IFN-γ signaling capacity, we measured STAT1 phosphorylation in CD4⁺ T cells from birth to 1 year. As expected, the increase in pSTAT1 activation paralleled the observed increase in pJAK2⁺ CD4⁺ T cells (Figure 4C). In CB, only 1.3% (median; range: 0.0–10.7%) CD4⁺ T cells activated pSTAT1 upon IFN-γ stimulation (Figure 4D). Activation of pSTAT1 was already increased by 1–3 months of age (median: 26.1%) and by 4–6 months pSTAT1⁺CD4⁺ T cell frequencies were detected in about 40% CD4⁺ T cells (Figure 4D); representative histograms are shown in Figure S1 in Supplementary Material. Although there was a significant positive correlation between increasing infant age and higher frequencies of pSTAT1⁺CD4⁺ T cells, this relationship did not appear to be linear (Figure 4D). In fact, IFN-γ-induced pSTAT1 activation seemed to plateau at about 6 months of age (Figure 4D; Table S1 and Figure S2 in Supplementary Material). As was shown for total STAT6 expression, unphosphorylated STAT1 could be detected in both CB and adult CD4⁺ T cells (Figure 4E). However, consistent with the age-dependent increase in pSTAT1 activation, activated pSTAT1 was localized mainly in the cytoplasm in CB CD4⁺ T cells, whereas in samples from 6-month-old infants pSTAT1 was clearly detectable in the nucleus (Figure 4E).

Compared to only a 1.4-fold increase in median frequencies of pSTAT6⁺CD4⁺ T cells from birth (63.8%) to 1 year (90.6%), median frequencies of pSTAT1⁺CD4⁺ T cells increased 30-fold in the first year (birth: 1.3%; 1 year: 38.0%) (Figure 5A). Furthermore, in samples for which sufficient blood volume was available to test both IL-4 and IFN-γ-induced JAK/STAT signaling, we observed a more than 10-fold increase in the ratio of pSTAT1:pSTAT6 from 0.04 (median; range: 0.0–0.2) at birth to 0.5 (range: 0.4–0.9) at 1 year (Figure 5B). Whether or not these findings have direct implications for CD4⁺ Th cell differentiation, was outside the scope of the current study. To further probe CB CD4⁺ T cells for reduced signaling function in response to Th1-promoting cytokines, we tested whether STAT4 phosphorylation after in vitro IL-12 stimulation was also reduced. Indeed, only 0.42% (median value; range: 0.0–4.2%) CB CD4⁺ T cells responded with STAT4 phosphorylation (Figure 6A). Despite higher IL-12 signaling ability of CD4⁺ T cells in 1-year-old infants, median frequencies of pSTAT4⁺CD4⁺ T cells still accounted for <1% in 10- to 14-month-old infants (range: 0.1–1.7%) and remained significantly lower compared to pSTAT4⁺ frequencies in adult CD4⁺ T cells (median: 3.3%; range: 0.2–18.2%) (Figure 6A). Consistent with this result, few CD4⁺ T cells with nuclear pSTAT4 localization could be detected in 1-year-old infants (Figure 6B). In addition, it appeared that the nuclear translocation might have been less efficient compared to nuclear localization of pSTAT6 or pSTAT1 in CD4⁺ T cells of 1-year-old infants stimulated with IL-4 or IFN-γ, respectively (Figure 6B). However, the tools for a more quantitative protein analysis of the Amnis^®ImageStreamX data were not available.

Considering that the IL-12Rβ2 chain is not expressed on naive CD4⁺ T cells (59, 60), we compared in pSTAT4⁺CD4⁺ T cell frequencies in memory CD4⁺ T cells of different age groups. Despite the low frequencies of CD4⁺ T cells that can be phenotypically characterized as T(CM) or T(EM/Eff) CD4⁺ T cells in CB, about 1 and 4%, respectively, could activate pSTAT4. By 1 year, the functional capacity of T(EM/Eff) CD4⁺ T cells to signal in response to IL12 was equivalent to adult T(EM/Eff) CD4⁺ T cells (Figures 6C,D), but the low frequencies of differentiated infant CD4⁺ T cells likely prevented the detection of this response within the total CD4⁺ T cell population (Figure 6A). Despite higher frequencies of memory CD4⁺ T cells in adults, IL-12Rβ2 expression is relatively low within the total CD4⁺ T cell population (Figure S3 in Supplementary Material). In contrast, NK cells (CD3⁻CD14⁻CD19⁻CD56⁺), even in infants, show more pronounced expression of IL-12Rβ2 (Figure S3 in Supplementary Material), with IL-12Rβ2 expression being highest on CD16⁺ NK cell subsets (Figure S3 in Supplementary Material).

IL-2 is a key cytokine promoting the proliferation and expansion of CD4⁺ T cells. As pointed out above, IL-2 signals through the common γ chain of the IL-2R and activates the JAK3 kinase. The activation of pJAK3 by IL-2 appeared to be comparable in CD4⁺ T cells from CB, blood of 1-year-old infants and adults (Figure 7A; see Figure 2A for controls). However, because we could not quantitate the exact number of pJAK3⁺CD4⁺ T cells by Western blot (Figure 7A), we measured the activation of the IL-2 specific transcription factor STAT5, downstream of JAK3. This analysis showed that frequencies of pSTAT5⁺CD4⁺ T cells were positively correlated (p < 0.0001) with increasing age between birth and 1 year (Figure 7B). However, even at 1 year, median pSTAT5⁺CD4⁺ T cell frequencies (36%; range: 25–51%) remained significantly lower compared adults (median: 69%; range: 44–80%) (Figure 6B).

To determine whether this effect was specific for IL-2, we tested whether JAK/STAT signaling to the related common γ-chain cytokine IL-7, that also promotes proliferation, was impaired in infants as well. Compared to <1% median frequencies of pSTAT5⁺ CD4⁺ T cells activated by IL-2 in CB, IL-7 induced about 19% (range: 0.3–73.2%) pSTAT5⁺CD4⁺ T cells (Figure 7C). IL-7-induced frequencies of pSTAT5⁺CD4⁺ T cells increased to 72% (range: 62.1–87.3%) by 4–6 months and reached 90% (range: 45.2–95.5%) by 1 year, levels comparable to those achieved in adults (median: 93.3%; range: 83.0–97.6%) (Figure 7C). The differential signaling capacity of infant CD4⁺ T cells to IL-2 or IL-7 was further supported by image analysis of blood samples from 6-month-old infants (Figure 7D). Although we could consistently detect pSTAT5 in the nucleus of samples stimulated with IL-7, activation and nuclear translocation of pSTAT5 in response to IL-2 was variable (Figure 7E).

The difference in IL-2 versus IL-7 signaling was mainly due to differential activation of naïve CD4⁺ T cells (Figures 8A,B). At 1 year, pSTAT5⁺ frequencies within the naïve CD4⁺ T cell population were still significantly lower (p < 0.0001) compared to adults after IL-2 stimulation, whereas IL-7-induced pSTAT5⁺ CD4⁺ T(N) cell frequencies were comparable between 1-year olds and adults (Figure 8). In T(CM) and T(EM/Eff) CD4⁺ T cells of 1-year-old infants, the activation of pSTAT5 remained below adult levels after both IL-2 and IL-7 stimulation (Figures 8A,B). Consistent with the fact that CD4⁺ T cell differentiation into memory T cells is associated with increased expression of CD25, the IL-2Rα, we noted an age-dependent positive correlation between IL-2-induced pSTAT5 activation and frequencies IL-2Rα⁺CD4⁺ T cells (Figure 8C).