PHD2 Deletion in CD8+ T Cells Worsens TAC-Induced Cardiac Inflammation, Heart Failure, and Pulmonary Remodeling

Abstract

Background: Cardiac inflammation is a key driver of cardiac fibrosis and heart failure. Cytotoxic CD8⁺ T cells play important roles in modulating inflammatory responses, especially during infections and autoimmune diseases. HIFs (hypoxia-inducible factors) regulate CD8⁺ T cell function and cardiac remodeling. However, the role of HIF in CD8⁺ T cells during heart failure development remains unclear.

Methods: We generated CD8⁺ T cell-specific PHD2 (prolyl hydroxylase domain protein 2)-deficient mice (PHD2^CKO), in which HIF bioavailability is increased due to the loss of PHD2. PHD2^CKO and wild-type littermates were subjected to pressure overload via transverse aortic constriction. Cardiac function, inflammation, and CD8⁺ T cell responses were assessed. CD8⁺ T cell metabolism was analyzed using Seahorse assays to measure extracellular acidification rate and oxygen consumption rate. HIF1α and HIF2α protein levels were measured by Western blotting.

Results: Under baseline conditions, PHD2 deletion in CD8⁺ T cells had no effect on heart function or effector molecule expression. Following transverse aortic constriction, PHD2^CKO mice showed exacerbated cardiopulmonary inflammation, fibrosis, and dysfunction. These effects were associated with enhanced CD8⁺ T cell activation and cytokine production. In vitro, PHD2-deficient CD8⁺ T cells exhibited increased glycolysis (extracellular acidification rate), reduced oxidative phosphorylation (oxygen consumption rate), and elevated HIF1α-but not HIF2α-levels. Pharmacological inhibition of HIF1α, but not HIF2α, reversed these metabolic and functional changes.

Conclusions: PHD2 deletion in CD8⁺ T cells aggravates cardiopulmonary remodeling after pressure overload by enhancing CD8⁺ T cell activation and cytokine release via a HIF1α-dependent mechanism.

Keywords: cytokines; fibrosis; heart failure; hypoxia; mice.

Figure 1.. CD8⁺ T cell–specific PHD2 deficiency alters cardiac function in female and male mice following transverse aortic constriction (TAC).

(A, D) Survival was assessed by the Kaplan–Meier method with log-rank test, and body weight was recorded weekly for 5 weeks in females. (A) and male (D) mice subjected to sham or TAC surgery. (B, E) Representative M-mode echocardiographic images obtained 5 weeks post-sham or TAC surgery in female (B) and male (E) mice. (C, F) Quantification of cardiac function and hypertrophy parameters in female (C) and male (F) mice, including left ventricular ejection fraction (LVEF), fractional shortening (LVFS), left ventricular (LV) weight to tibia length (TB) ratio, and total ventricular weight to TB ratio. Data are presented as mean ± SEM; n = 11–13 per group for female mice and n = 7–15 per group for male mice. Normality was confirmed by Shapiro–Wilk test and variance homogeneity by Levene’s test. Accordingly, either Welch’s one-way ANOVA followed by the Games–Howell post hoc test or a one-way ANOVA with Tukey’s post hoc test was used for group comparisons (n = sample size).

Figure 2.. CD8⁺ T cell–specific PHD2 deficiency modulates cardiac inflammation, hypertrophy, and fibrosis.

(A) Representative immunofluorescence images showing CD45⁺ leukocyte infiltration in the left ventricle. (B) Flow cytometry gating strategy for identifying cardiac CD45⁺ immune cells. (C) Quantification of total immune cell numbers based on surface marker expression. (D) Representative histological images of wheat germ agglutinin (WGA) staining to assess cardiomyocyte size, and Sirius Red staining for interstitial fibrosis. Scale bars: 100 μm (CD45, WGA), 100 μm and 500 μm (Sirius Red). n = 10 mice per group for CD45, WGA, and Sirius Red; n = 5 per group for flow cytometry. Normality was assessed using the Shapiro–Wilk test, and homogeneity of variances was evaluated using Levene’s test. Accordingly, either Welch’s one-way ANOVA followed by the Games–Howell post hoc test or a one-way ANOVA with Tukey’s post hoc test was used for group comparisons (n = sample size).

Figure 3.. PHD2 deficiency alters cytokine production in cardiac CD8⁺ T cells.

(A) Representative flow cytometry plots showing the proportions of CD4⁺, CD8⁺, and CD4⁻CD8⁻ T cells among cardiac CD3⁺ T cells. (B–D) Quantification of IFN-γ⁺, TNF-α⁺, and IFN-γ⁺TNF-α⁺ double-positive cells within the cardiac CD8⁺ T cell population. Data are presented as mean ± SEM (n = 5 per group). Normality was confirmed by Shapiro–Wilk test and variance homogeneity by Levene’s test. Accordingly, either Welch’s one-way ANOVA followed by the Games–Howell post hoc test or a one-way ANOVA with Tukey’s post hoc test was used for group comparisons (n = sample size).

Figure 4.. Pulmonary CD8⁺ T cell activation and cytokine production in PHD2-deficient mice.

(A) Representative flow cytometry plots of pulmonary T cell subsets: CD4⁺, CD8⁺, and CD4⁻CD8⁻ T cells. (B) Flow cytometry quantification of pulmonary CD8⁺ T cell subsets: naïve (CD44⁻CD62L⁺), effector (CD44⁺CD62L⁻), and memory (CD44⁺CD62L⁺) populations. (C–E) Quantification of IFN-γ⁺, TNF-α⁺, and IFN-γ⁺TNF-α⁺ double-positive CD8⁺ T cells in the lung. Normality was assessed using the Shapiro–Wilk test, and homogeneity of variances was evaluated using Levene’s test. Depending on the results, either a nonparametric Kruskal–Wallis test with Dunn’s post hoc analysis or a one-way ANOVA followed by Tukey’s post hoc test was applied to compare the four groups. Data are presented as mean ± SEM, with sample sizes of n = 7 for sham groups and n = 6 for TAC groups (n = sample size).

Figure 5.. In vitro effects of PHD2 deficiency on CD8⁺ T cell metabolism and activation.

(A) Oxygen consumption rate (OCR) analysis in activated CD8⁺ T cells, showing basal respiration, ATP production, maximal respiration, and spare respiratory capacity. (B) Extracellular acidification rate (ECAR) analysis showing basal glycolysis, glycolytic capacity, and glycolytic reserve. (C) Representative flow cytometry plots and quantification of IFN-γ⁺TNF-α⁺ double-positive CD8⁺ T cells. (D) Representative histograms and quantification of GLUT1 (glucose transporter 1) expression in CD8⁺ T cells. n = 6 per group for Seahorse assays (OCR and ECAR); n = 3 per group for flow cytometry. Variance homogeneity was assessed using Levene’s test, and normality was evaluated with the Shapiro–Wilk test. Accordingly, either a paired t-test or the nonparametric Wilcoxon signed-rank test was used to compare paired samples (n = sample size).

Figure 6.. HIF-dependent metabolic reprogramming in PHD2-deficient CD8⁺ T cells in vitro.

(A) Western blot analysis of PHD2, HIF-1α, and HIF-2α protein levels in stimulated CD8⁺ T cells, with densitometric quantification. Variance homogeneity was assessed using Levene’s test, and normality was evaluated with the Shapiro–Wilk test. Depending on the results, either a paired t-test or the nonparametric Wilcoxon signed-rank test was used to compare paired samples. (B–C) Seahorse analysis of OCR (B) and ECAR (C), showing metabolic parameters as in Figure 5. (D) Representative flow cytometry plots and quantification of IFN-γ⁺TNF-α⁺ double-positive CD8⁺ T cells. (E) GLUT1 expression levels assessed by flow cytometry. (F) Schematic diagram summarizing the proposed mechanism by which PHD2 deficiency promotes CD8⁺ T cell metabolic activation and inflammatory function via HIF stabilization. n = 3 per group for Western blotting and flow cytometry; n = 6 per group for Seahorse assays. Normality was assessed using the Shapiro–Wilk test, and homogeneity of variances was evaluated using Levene’s test. Depending on the results, either a nonparametric Kruskal–Wallis test with Dunn’s post hoc analysis or a one-way ANOVA followed by Tukey’s post hoc test was applied to compare the four groups. Data are presented as mean ± SEM (n = sample size).