Cutting Edge: Activation-Induced Iron Flux Controls CD4 T Cell Proliferation by Promoting Proper IL-2R Signaling and Mitochondrial Function

As an essential microelement, iron takes part in a wide variety of physiological processes, including erythropoiesis, DNA synthesis and repair, and immunity. Human disease states leading to either iron deficiency or iron overload have been shown to adversely affect the adaptive immune response (1, 2). As such, iron homeostasis is vital to the development of effective T cell–mediated immunity. In healthy individuals, the majority of bioavailable iron in circulation is bound to transferrin (Tf). These Tf–iron complexes are taken up by cells via Tf receptor 1 (TfR1)-mediated endocytosis (3), and T cell surface expression of TfR1 increases after activation (4). In addition to TfR1-mediated iron import, T cells have also been shown to take up non–Tf-bound iron (5, 6), which is known to occur through the action of several nonspecific metal ion transporters, such as divalent metal transporter 1 (DMT1) and the ZRT/IRT-like proteins 8/14 (ZIP8/14) (7–9). Once in the cytosol, iron is freed from its binding partners and enters a redox active pool known as the labile iron pool (LIP), which can then be transported to ferritin for storage (10) or used as a cofactor for other cellular proteins and enzymes. Alternatively, LIP can be directly released by the cell through ferroportin (Fpn), the only known iron exporter in mammalian cells (11).

Dysregulation of the iron homeostatic pathway can be detrimental to T cells. Early thymocyte differentiation and maturation are dependent on iron acquisition via TfR1, and the absence of TfR1 has been shown to lead to T cell developmental arrest (12, 13). Additionally, iron deficiency impairs both the activation status and proliferative capacity of peripheral T lymphocytes (14, 15). Nevertheless, the exact mechanism by which iron regulates T cell proliferation remains unclear. In this study, we show that T cell activation mobilizes intracellular iron stores, which promotes proper CD4 T cell expansion by fueling IL-2R signaling and mitochondrial function after stimulation.

Male and female C57BL/6 mice ranging from 8 to 12 wk of age were either bred in-house or purchased from The Jackson Laboratory. Mice were housed in specific pathogen-free conditions. All animal experiments were performed in accordance with the Institutional Animal Care and Use Committee of the University of Michigan.

CD4 T cells were enriched from murine splenocytes and human PBMC using a positive selection kit according to the manufacturer’s instructions (Miltenyi Biotec). T cells were activated with plate-bound anti-CD3 (5 μg/ml) and soluble anti-CD28 (1 μg/ml for murine studies; 2 μg/ml for human studies) Abs (eBioscience) for an indicated time in RPMI 1640 medium supplemented with 10% FBS, 2 mM glutamine, and penicillin/streptomycin at 37°C. Cells were treated with the indicated amounts of deferoxamine mesylate (DFO) (Sigma-Aldrich) or ferric ammonium citrate (FAC) (Sigma-Aldrich) during certain experiments. Fifty units of rIL-2 (PeproTech) was used for all IL-2 supplementation experiments. For cell proliferation, CD4 T cells were labeled with CellTrace Violet (CTV) (5 μM) (Invitrogen) in 1× PBS containing 0.1% BSA for 30 min at 37°C.

The fluorescently conjugated Abs used for surface and intracellular staining in the presence of anti-FcγR mAb (2.4G2) were as follows: anti-mouse TCR-β (H57-597) Pacific Blue/allophycocyanin, anti-mouse CD4 (GK1.5) allophycocyanin-Cy7, anti-mouse CD8 (53-6.7) PE-Cy7, anti-mouse CD71 (R17217) FITC/PerCP-Cy5.5, anti-mouse CD25 (PC61.5) PerCP-Cy5.5/PE-Cy7, anti-mouse CD69 (H1.2F3) PE-Cy7, and IL-2 (JES6-5H4) PE (all from eBioscience).

For Fpn, fixed cells were incubated with metal transporter protein Ab (rabbit anti-mouse MTP1/IREG1/Fpn) (Alpha Diagnostic) in flow cytometry buffer. Ferritin expression was measured by anti-mouse ferritin (EPR3004Y) (Abcam) staining in permeabilization buffer after fixation. To analyze STAT5 phosphorylation, cells were fixed in 80% methanol and stained with rabbit anti-mouse pSTAT5 (Tyr694) (Cell Signaling) Ab. In all stainings, AF488-conjugated anti-rabbit secondary Ab (Invitrogen) was used. For intracellular cytokine expression, stimulated CD4 T cells were restimulated with 50 ng/ml PMA (Sigma-Aldrich) and 1.5 μM ionomycin (Sigma-Aldrich) in the presence of 3 μM monensin for 4 h, followed by intracellular cytokine staining (BD Biosciences). Dead cells were excluded from the analysis based on propidium iodide (PI) (1 μg/ml) or live/dead Fixable Aqua Dead Cell Stain (Invitrogen) signal. To measure cell cycle progression, cells were fixed with 70% ethanol, then stained with 50 ng/ml PI for 15 min. Cells were acquired on a FACS Canto II (BD Bioscience), and data were analyzed using FlowJo (TreeStar software v. 10.5).

Enriched CD4 T cells were split into replicates containing 1 × 10⁶ live cells each and spun down. After removing the supernatant, the cells were analyzed for metals by inductively coupled plasma mass spectrometry (ICP-MS) as described previously (16).

To measure LIP, cells were stained with calcein-acetoxymethyl (Calcein-AM) dye (0.02 μM) (Thermo Fisher Scientific) for 10 min at 37°C and analyzed by flow cytometry. LIP was calculated based on the ratio of Calcein mean fluorescence intensity (MFI) of control versus test samples.

Total RNA was isolated from both unstimulated and stimulated CD4 T cells using the RNeasy Plus mini kit (Qiagen) according to the manufacturer’s instructions. cDNA was synthesized, and quantitative PCR was performed using SYBR Green with Applied Biosystem’s 7500HT Sequence Detection System. Fold changes were calculated from ΔCt values using the ΔΔCt method. Expression of target genes was normalized to β-actin.

Supernatants were collected from CD4 T cells stimulated with or without DFO for 3 d. ELISA assays were done in conjunction with the University of Michigan ELISA core.

Mitochondrial potential, mitochondrial mass, and mitochondrial reactive oxygen species (mtROS) were measured using tetramethylrhodamine methyl ester perchlorate (TMRM) (60 nM) (Invitrogen), MitoTracker green (30 nM) (Invitrogen), and MitoSOX (2.5 μM) (Invitrogen), respectively. Cells were then analyzed by flow cytometry.

All graphs were prepared using Prism software (Prism version 7; GraphPad Software, San Diego, CA). For comparison among multiple groups, data were analyzed using one-way ANOVA with multicomparison post hoc test. For comparison between two groups, unpaired Student t tests were used. A p value < 0.05 was considered statistically significant.

The degree to which CD4 T cells require biometals for their maintenance and function is poorly understood. We used ICP-MS to measure the levels of several metals in unstimulated CD4 T cells. Interestingly, resting CD4 T cells contain significantly higher amounts of intracellular iron than any other tested metal (Supplemental Table I). Furthermore, ICP-MS analysis revealed that CD4 T cells drastically downregulate total intracellular iron levels after activation (Fig. 1A), which corresponds with accumulation of iron in the media over time (Fig. 1B). Because ICP-MS detects both protein-bound iron and LIP, we examined whether conventional αβ T cell subsets maintain distinct levels of LIP at steady-state using Calcein-AM dye, a cell-permeable fluorescent probe that binds to free iron in the cytoplasm (Supplemental Fig. 1A). We found that CD4 T cells contain lower levels of LIP than CD8 T cells (Fig. 1C), prompting us to investigate what happens to LIP following T cell activation. The data showed that CD4 T cells steadily downregulate LIP over time after stimulation (Fig. 1D, top panel), and this reduction of LIP can be observed as early as 2 h after receiving a TCR stimulus (Supplemental Fig. 1B). Activation-induced downregulation of LIP appears to be conserved across the T cell lineage, as CD8 T cells also reduce LIP after stimulation (Fig. 1D, bottom panel). This phenomenon was found to occur in human CD4 T cells as well (Fig. 1E).

We next investigated whether other iron homeostatic processes are influenced by TCR stimulation. We began by examining changes in TfR1 and found that both mRNA and protein levels are dramatically upregulated after activation, with peak expression occurring at 2 d poststimulation (Fig. 1F, top panel; Supplemental Fig. 1C, 1D, left panels). Additionally, mRNA expression of the non–Tf-bound iron transporters ZIP8, ZIP14, and the iron-response element–containing isoform of DMT1 was increased over time after T cell activation, whereas expression of the iron-response element–null isoform of DMT1 was unchanged (Fig. 1G). We next asked whether activated T cells have increased capacity to store iron by measuring ferritin expression. Although ferritin mRNA levels decreased over time after T cell stimulation (Supplemental Fig. 1D, middle panels), ferritin protein expression increased (Fig. 1F, Supplemental Fig. 1C, middle panels). To understand whether the decrease in both total iron and LIP levels following activation is due to iron export, we measured Fpn expression. Unexpectedly, both Fpn mRNA (Supplemental Fig. 1D, right panel) and protein (Fig. 1F, bottom panel; Supplemental Fig. 1C, right panel) expression were drastically decreased following stimulation, indicating that Fpn-mediated export is not the main mechanism of iron export in activated T cells. Taken together, our results show that T cells actively take up, store, and export iron during the response to TCR stimulation.

Our data showing the downregulation of iron after activation prompted us to further investigate the role of TCR signaling strength on total iron levels. We observed that strong TCR stimuli decreased cellular iron levels dramatically, whereas weak TCR stimuli had only a negligible effect on total iron amount (Fig. 2A). LIP levels were also impacted by TCR signaling strength, with only strong TCR stimuli prompting both a dramatic decrease in LIP and efficient T cell proliferation (Fig. 2B). These data revealed that LIP levels are inversely correlated with activation-induced proliferation, and this phenomenon is dependent on the strength of the TCR stimulus (Fig. 2B). Next, we sought to examine whether CD28 signaling also contributes to the regulation of iron homeostasis. We found that CD28 signaling had only a marginal effect on the downregulation of LIP, and that TCR signaling alone was sufficient in reducing LIP levels after T cell activation (Fig. 2C).

The relationship between intracellular iron levels and T cell proliferation prompted us to examine whether modulating iron levels in culture affects CD4 T cell expansion. We found that treatment with the iron chelator DFO significantly inhibited activation-induced downregulation of both total iron levels and LIP levels (Fig. 2D, 2E). Additionally, DFO treatment inhibited proliferation in a dose-dependent manner, leading to an inverse correlation between T cell proliferation and LIP levels at either high or low concentrations of DFO (Fig. 2E). DFO treatment also reduced CD4 T cell numbers (Supplemental Fig. 2A), consistent with the observed block in T cell proliferation. Similarly, DFO treatment severely inhibited human CD4 T cell proliferation (Fig. 2F) while simultaneously stabilizing LIP levels (Fig. 2G). In addition to its effects on LIP, exposure to DFO led to a reduction in the surface expression of TfR1, ferritin, and Fpn in activated T lymphocytes (Supplemental Fig. 2B). In all, strong TCR stimuli induce rapid mobilization of iron within the cell that correlates with downstream proliferation.

The profound effect of iron availability on proliferation prompted us to investigate possible mechanisms by which iron may be controlling T cell expansion. We began by asking whether CD4 T cells are properly activated when iron availability is limited. To test this, we measured the expression of the activation markers CD25 and CD69 on DFO-treated CD4 T cells. The results showed that iron chelation reduced CD25 but not CD69 expression (Fig. 3A, Supplemental Fig. 2C). Additionally, DFO treatment had only a marginal effect on T cell growth as measured by forward scatter (Supplemental Fig. 2C, right panel).

It is known that CD25 expression is dependent on both TCR signaling and IL-2 sensing (17). Additionally, IL-2 is known to be a critical mediator of T cell survival and proliferation (18). Therefore, we next asked whether iron regulates IL-2 synthesis. We hypothesized that DFO treatment would decrease IL-2 production by activated T cells; however, CD4 T cells cultured in iron-deficient conditions secreted copious amounts of IL-2 (Fig. 3B). This was further supported by the fact that DFO treatment led to increased frequencies of IL-2⁺ CD4 T cells (Fig. 3C). The observed accumulation of IL-2 in the media after activation may be due to the continuous secretion of cytokine by cells that do not have the capability to use it. As such, we hypothesized that DFO-treated cells do not produce enough IL-2 during early TCR stimulation. However, the addition of exogenous IL-2 to the culture media was not sufficient to overcome the antiproliferative effect of DFO (Fig. 3D), as shown previously (19).

Collectively, our data indicated that insufficient iron levels during T cell activation lead to defective IL-2R signaling. To determine whether the IL-2R signaling pathway was impaired by iron chelation, we analyzed pSTAT5 levels in DFO-treated CD4 T cells, as CD25 expression is governed by STAT5 (20). The data showed that DFO reduced the amount of pSTAT5 in activated CD4 T cells (Fig. 3E, Supplemental Fig. 2D), suggesting that iron is necessary for optimal IL-2R signaling following T cell stimulation. Together, our findings show that iron modulates T cell proliferation by controlling intracellular IL-2 signaling rather than IL-2 production.

To further study the role of iron in T cell biology, we examined whether iron chelation negatively impacts cell cycle progression. DFO treatment significantly decreased the percentages of activated T cells in the S and G₂ phases of the cell cycle (Fig. 4A), corroborating previous findings that DFO is a potent S phase inhibitor in T lymphocytes (21). Iron is also known to play a role in the generation of mtROS, which have been shown to be important for T cell activation (22). Therefore, we hypothesized that iron availability during stimulation may impact mitochondrial dynamics. Indeed, iron chelation led to a sharp decrease in mitochondrial mass, mitochondrial potential, and mtROS levels over time after T cell activation (Fig. 4B, Supplemental Fig. 3A).

Last, we investigated whether the adverse effects of DFO on mitochondrial function and proliferation could be reversed by the addition of exogenous iron. The results showed that iron supplementation via FAC treatment rescued cell proliferation in the presence of DFO (Fig. 4C). Furthermore, the negative effects of DFO on cell cycle progression were abrogated by the addition of iron to the culture media (Fig. 4D, Supplemental Fig. 3B). FAC treatment also restored mitochondrial mass, mitochondrial potential, and mtROS production (Fig. 4E), further supporting a role for iron in maintaining mitochondrial biogenesis and function after stimulation.

In summary, our findings indicate that intracellular iron levels impact T cell proliferation by promoting optimal IL-2R signal transduction and mitochondrial function. T cell activation leads to a drastic loss of intracellular iron, which ultimately results in accumulation of iron in the culture media. Together, these data suggest that T cells actively avoid iron overload during the response to TCR stimuli. Cells may try to prevent iron overload by exporting iron during activation; however, we showed that Fpn levels decrease dramatically after stimulation. Our findings suggest that T cells use an Fpn-independent mechanism of iron export. T cells are known to increase heme oxygenase 1 (HO-1) expression after activation, and overexpression of HO-1 has been shown to inhibit T cell proliferation (23). Therefore, it is possible that T cells try to prevent both iron overload and HO-1 overexpression during stimulation by exporting heme-bound iron. The heme-iron exporter feline leukemia virus subgroup C receptor 1 (FLVCR1) has been shown to be essential for T cell development and peripheral maintenance (24). Therefore, T cells may increase the export of heme-iron after activation via the action of FLVCR1; however, this possibility remains to be tested.

We also found that iron chelation suppresses TCR-induced expansion and that this phenomenon was independent of the ability of the cell to produce IL-2. Instead, our results showed that IL-2R signaling in activated CD4 T cells depends on iron, as DFO treatment decreased pSTAT5 levels after stimulation. Interestingly, CD25 recycling after T cell activation occurs via the action of Tf⁺ endosomes (17), indicating that CD25 expression after stimulation may be an iron-dependent process. Although the precise steps in the IL-2R signaling cascade that are affected by iron remain unknown, our data highlight a potential role for iron in regulating intracellular signaling in activated T cells.

The role of iron in immunity has been well established, particularly through the study of macrophage biology. It is well known that macrophages aid in the recycling of iron and that this iron facilitates macrophage responses to intracellular and extracellular pathogens (25). Macrophage-mediated iron uptake has also been highlighted as a key mechanism of nutrient sequestration from tumor cells during the immune response to cancer (26). However, the role of T cell–derived iron metabolism in human disease states is relatively understudied. We found that CD4 and CD8 T cells maintain varying degrees of LIP during quiescence. Additionally, activation-induced reduction of LIP appears to be a lineage-wide phenomenon. Therefore, it is possible that different T effector cell subsets require certain levels of iron for their function. Recent work has shown that iron overload can enhance T cell inflammatory cytokine production, leading to exacerbation of autoimmune diseases like experimental autoimmune encephalomyelitis (27). Iron deficiency is also known to impair concanavalin A–induced T cell responses during acute liver inflammation (2). Therefore, the possibility that iron can modulate Th cell differentiation in the context of human disease warrants further study and may open the door for the discovery of novel therapeutic strategies for patients.

We thank Tiffany Nguyen of the Seo Laboratory for help in establishing the ICP-MS assay.

The authors have no financial conflicts of interest.