Newly Generated CD4⁺ T Cells Acquire Metabolic Quiescence after Thymic Egress

Mature naive T cells circulate through the secondary lymphoid organs (SLOs) in a quiescent state. This is characterized by small cell size, low basal metabolic activity, G₀ phase, and low or no expression of activation markers such as CD69 and CD44 (1, 2). Accumulating evidence has suggested that the quiescent status of naive T cells is an actively enforced process that involves constant subthreshold stimulation of TCR and IL-7R, as well as multiple pathways, including the forkhead box family members (FoxO1/3 and Foxp1), Krüppel-like factor 2, Schlafen2, tuberous sclerosis (Tsc)1, and autophagy-related genes Atg3/5/7 and Vps34 (1, 3–16). Deletion of at least some of these molecules revealed increased apoptotic susceptibility, acquisition of activated phenotype, and loss of full responsiveness to activating stimuli.

In contrast, higher-than-basal metabolic activity is important for T cell development in the thymus. The mitochondrial and endoplasmic reticulum (ER) volumes are high in CD4 and CD8 double-positive (DP) thymocytes that undergo positive selection and do not decline at least in CD4 single-positive (SP) thymocytes (17–19). An increase in intracellular reactive oxygen species (ROS) and a decrease in cytosolic antioxidant molecule thioredoxin Trx(1) are also observed in CD4 SP thymocytes undergoing phenotypic maturation (20). Mature SP thymocytes are more tolerant to high ROS level, as the addition of H₂O₂ or Trx reductase inhibitor increases the apoptosis of immature CD69⁺ but not mature Qa2⁺CD69⁻ SP thymocytes. Additionally, an increase in ROS level is associated with enhanced IL-2 production by mature CD4 SP thymocytes (20), and a lack of ROS in Ncf1^−/− mice affects negative selection and results in the egress of autoreactive T cells (21). These results indicate that high mitochondrial content and elevated intracellular ROS in CD4 SP thymocytes may facilitate negative selection and functional maturation in these young T cells before egress (20, 22).

Interestingly, the conditional deletion of Atg genes using CD4^cre mice results in the accumulation of mitochondria/ER and enhanced apoptosis in peripheral T cells. DP or SP thymocytes are largely not affected (6, 7). This indicates that peripheral T cells are more sensitive to changes in mitochondrial contents and metabolic activity than are CD4 SP thymocytes. It is thus reasonable to speculate that a process of metabolic reprogramming to acquire a metabolically quiescent state may occur in peripheral T cells after they leave the thymus. The initiation and regulation of this metabolic change during SP thymocytes–mature peripheral T cells transition remains unclear. However, failure in this metabolic switch predisposes T cells to apoptotic death and causes survived cells to become semiactivated with decreased immune competency (12, 14, 16, 23).

Recent thymic emigrants (RTEs) comprise a unique population of T cells that have recently completed a tightly regulated developmental program in the thymus and entered the naive T cell pool in the periphery (2, 24–28). The phenotype and function of RTEs are different from SP thymocytes as well as peripheral naive T cells (29–43). Upon activation, RTEs exhibit lower levels of proliferation, cytokine production, and aerobic glycolysis when compared with mature naive T cells (44). The SLOs and liver are important for the maturation and tolerance of RTEs in the periphery during a 2- to 3-wk period (29, 43, 45–49). Despite these findings, the basal metabolic changes during the transition from SP thymocytes to RTEs and eventually to naive T cells are not well investigated.

Mitochondria are “the powerhouses” of a cell. They also serve as signaling hubs for oxidative and Ca²⁺ signaling (50). We thus compared the mitochondria among T cell subsets, including immature (CD69⁺) and mature (CD69⁻) CD4 SP thymocytes (51), CD4⁺ RTEs, and CD4⁺CD25⁻CD44^loCD62L^hi naive T cells. A significant reduction in mitochondrial content and mitochondrial ROS (mROS) was found in RTEs and naive T cells when compared with SP thymocytes. This decrease in mitochondria required T cell egress from the thymus and occurred early after young T cells entering the circulation. The reason for mitochondrial downregulation was also investigated.

This study was carried out under strict guidance of the Ethics Committee of Peking University Health Science Center. C57BL/6 congenic mice, CD45.1 and CD45.2, were purchased from Peking University Health Science Center and Vital River Laboratory Animal Technology Company (Beijing, China). FVB-Tg (Rag2-EGFP) 1Mnz/J mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and were backcrossed to the C57BL/6 background for 14 generations (termed RAG2p-GFP in this study). Cd4-Cre mice and lymphotoxin β receptor−deficient (Ltbr^−/−) mice on the C57BL/6 background were provided by Prof. Y. Zhang (Peking University Health Science Center) and Prof. Y. Fu (University of Chicago), respectively. Tsc1^flox mice were provided by W. Zhang (Peking University Health Science Center) and backcrossed to the C57BL/6 background for 14 generations. To generate Tsc1^flox-GFP⁺-Cd4-Cre mice, Tsc1-floxed mice were crossed to Cd4-Cre transgenic mice and RAG2p-GFP transgenic mice. The mice were housed in standardized conditions with temperature maintained at 23°C on a 12-h light/12-h dark cycle. All animals were provided standard laboratory chow and water ad libitum. Acclimatization for at least 3 d to the laboratory conditions before experimental inclusion was performed. Female mice at 6–8 wk of age were used in all experiments.

To prepare CD4 SP thymocytes, single-cell suspensions of thymocytes were first incubated with FITC-conjugated CD8 Ab (BD Pharmingen, San Diego, CA) followed by anti-FITC microbeads (MACS; Miltenyi Biotec, Bergisch Gladbach, Germany). CD8⁻ thymocytes were obtained by negative selection using the autoMACS Pro Separator (Miltenyi Biotec). CD4 SP thymocytes with the phenotype of GFP⁺CD4⁺CD8⁻CD69⁺CD44^lo, GFP⁺CD4⁺CD8⁻CD69⁻Qa2⁻CD44^lo, and GFP⁺CD4⁺CD8⁻CD69⁻Qa2⁺CD44^lo were then sorted using a FACSAria II (BD Biosciences, San Diego, CA). When large amounts of SP thymocytes were needed for Western blotting, immature GFP⁺CD4⁺CD8⁻CD69⁺CD25⁻ and relatively mature GFP⁺CD4⁺CD8⁻CD69⁻CD25⁻ thymocytes were purified. For the isolation of CD4⁺ RTEs and CD4⁺ naive T cells, CD4⁺CD8⁻CD25⁻NK1.1⁻GFP⁺ (RTEs) and GFP⁻CD4⁺CD8⁻CD44^loCD62L^hiCD25⁻ (naive T) cells from mesenteric lymph nodes were purified by flow cytometry. When T cell activation was examined, purified CD69⁺ and CD69⁻ CD4 SP thymocytes, CD4⁺ RTEs, and naive T cells were cultured with plate-bound anti-CD3 (2 μg/ml) and soluble anti-CD28 (1 μg/ml) for 3 d. The cells were then stained for surface CD25 and intracellular IL-2 and analyzed by flow cytometry.

Anti-CD8 (3.155) was prepared from a hybridoma obtained from the American Type Culture Collection (Manassas, VA). Abs against CD4 (GK1.5), CD8 (53-6.7), CD69 (H1.2.F3), NK1.1 (PK136), CD62L (MEL-14), CD44 (IM7), and CD25 (3C7) were purchased from BD Pharmingen. Alexa Fluor 647–conjugated anti-mouse Qa2 was purchased from BioLegend (San Diego, CA).

To stain the mitochondria, lymphocytes were incubated with MitoTracker Green (20 nM; Invitrogen, Grand Island, NY) at 37°C for 30 min. Cellular ROS level was measured by incubation with 5-(and-6)-carboxy-2′,7′-dichlorodihydrofluorescein diacetate (10 μM; Invitrogen) at 37°C for 30 min. mROS production was measured by MitoSOX Red (2 μM; Invitrogen) at 37°C for 10 min. Mitochondrial membrane potential was measured by tetramethyl rhodamine methyl ester (TMRM; 20 nM; Invitrogen) or MitoTracker Deep Red (20 nM; Invitrogen) at 37°C for 30 or 10 min. The cells were then washed and stained with Abs against CD4, CD8, and Qa2 for thymocytes or CD4, CD8, CD62L, and CD44 Abs for RTE and naive T cells. The cells were collected on a FACSAria II (BD Biosciences) and analyzed using FlowJo software.

For analysis of the oxygen consumption rate (OCR; in picomolles per minute) and extracellular acidification rate (ECAR; in milli-pH units per minute), the Seahorse XF-24 metabolic extracellular flux analyzer was used (Seahorse Bioscience, North Billerica, MA). Sorted CD4⁺ T cell subsets were plated onto 24-well XF microplates (1.0 × 10⁶ cells per well) coated with poly-l-lysine to enhance T cell attachment. Perturbation profiling of the use of metabolic pathways by CD4⁺ T cells was done by the addition of oligomycin (1 μmol/l), carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (1 μmol/l; Sigma-Aldrich, St. Louis, MO), and rotenone (1 μmol/l; Sigma-Aldrich) and antimycin A (1 μmol/l; Sigma-Aldrich). Metabolic parameters were calculated with Seahorse Wave software.

Freshly isolated GFP⁺CD4⁺CD8⁻CD69⁻Qa2⁻CD25⁻ and GFP⁺CD4⁺CD8⁻CD69⁻Qa2⁺CD25⁻ SP thymocytes, CD4⁺ RTEs, and CD4⁺ naive T cells from mesenteric lymph nodes were immediately fixed in a 4% glutaraldehyde in 0.1 M sodium cacodylate buffer and processed as described previously (52). The images were taken from the transmission electron microscopy at Qinghua University and were analyzed using ImageJ software.

RNA was purified from various T cell subsets using TRIzol (Invitrogen). The purity of RNA was verified spectrophotometrically at 260/280 nm. The RNA samples (2 μg) were reversed transcribed into cDNAs using the FastQuant reverse transcription kit (Tiangen, Beijing, China) according to the manufacturer’s instructions. Quantitative real-time PCR was performed using FastStart Universal SYBR Green Master mix (Roche, Basel, Switzerland) on an iCycler real-time PCR system (Bio-Rad Laboratories, Hemel Hempstead, U.K.), with each sample in triplicate. The following PCR primers were used: voltage-dependent anion channel (VDAC), forward, 5′-GGCTACGGCTTTGGCTTAAT-3′, reverse, 5′-CCCTCTTGTACCCTGTCTTGA-3′; β-actin, forward, 5′-TGGAATCCTGTGGCATCCATGAAAC-3′, reverse, 5′-TAAAACGCAGCTCAGTAACAGTCCG-3′; 12S rRNA, forward, 5′-ACCGCGGTCATACGATTAAC-3′, reverse, 5′-CCCAGTTTGGGTCTTAGCTG-3′; 18S rRNA, forward, 5′-CGCGGTTCTATTTTGTTGGT-3′, reverse, 5′-AGTCGGCATCGTTTATGGTC-3′; Cytb, forward, 5′-ATTCCTTCATGTCGGACGAG-3′, reverse, 5′-ACTGAGAAGCCCCCTCAAAT-3′; cytochrome c oxidase 1 (Cox1), forward, 5′-GCCCCAGATATAGCATTCCC-3′, reverse, 5′-GTTCATCCTGTTCCTGCTCC-3′; NADH dehydrogenase (ND)1, forward, 5′-ACCATTTGCAGACGCCATAA-3′, reverse, 5′-TGAAATTGTTTGGGCTACGG-3′. The quantification was based on ΔΔCT calculations and was normalized to β-actin as loading controls. To determine mitochondrial DNA (mtDNA) content, quantitative PCR was performed as previously described (16, 17) using primers specific for the mtDNA gene and normalized to a genomic control.

Purified T cell subsets were washed with ice-cold PBS and then lysed in 50 mM Tris (pH 7.4) containing 1% Triton X-100, 150 mM NaCl, 1% sodium deoxycholate, 0.1% SDS, and a mixture of protease inhibitors and phosphatase inhibitors (Roche). The whole-cell lysates were subjected to 6–15% SDS-polyacrylamide gels and transferred to a Hybond ECL nitrocellulose filter membrane (Whatman, GE Healthcare Life Sciences, Pittsburgh, PA). The membranes were subsequently blocked for 1 h at room temperature with TBS (pH 7.4) containing 0.1% Tween 20 and 5% dried skimmed milk. The primary Abs used for hybridization at 4°C overnight were anti–peroxisome proliferator-activated receptor γ coactivator-1α (PGC-1α), nuclear respiratory factor (NRF)1, and Rieske iron-sulfur protein (RISP) from Abcam (Cambridge, MA), GABPα and mitochondrial transcription factor A (TFAM) from Santa Cruz Biotechnology (Santa Cruz, CA), LaminB1 and β-tubulin from Bioworld Technology (St. Louis Park, MN), NADH-ubiquinone oxidoreductase 1α subcomplex subunit 9 (NDUFA9) and complex II 70 kDa Fp subunit from Invitrogen (Carlsbad, CA), and VDAC, mechanistic target of rapamycin (mTOR), p-mTOR (Ser²⁴⁴⁸), S6K, p-S6K, 4E-BP1, p–4E-BP1, AMP-activated protein kinase (AMPK)α, p-AMPKα (Thr¹⁷²), p-ACC (Ser⁷⁹), p–Unc-51–like kinase (ULK1; Ser⁷⁵⁷), FoxO1, p-FoxO1 (Thr²⁴), Akt, p-Akt (Ser⁴⁷³), OPA1, p-DRP1 (Ser⁶¹⁶), p-ERK1/2 (Thr²⁰²/Tyr²⁰⁴), p-p38 (Thr¹⁸⁰/Tyr¹⁸²), p-JNK1/2 (Thr¹⁸³/Tyr¹⁸⁵), p–apoptosis signal–regulating kinase 1 (ASK1; Thr⁸⁴⁵), and β-actin from Cell Signaling Technology (Danvers, MA). Anti-Trx1 was from Redox Bio Science (Kyoto, Japan). The membranes were then washed with TBS (pH 7.4) containing 0.1% Tween 20 for three times for 10 min each time and hybridized with HRP-conjugated goat anti-rabbit, goat anti-mouse, or donkey anti-goat secondary Ab at room temperature for 1.5 h. The immunoreactive bands were detected by chemiluminescence using ECL detection reagents SuperSignal West Pico chemiluminescent substrate (Thermo Fisher Scientific, Waltham, MA) and exposed to ImageQuant LAS500 (GE Healthcare Life Sciences). The bands were quantified using ImageJ software. The cellular fractionation was performed as previously described (53).

Purified CD4⁺CD8⁻CD69⁻Qa2⁺CD44^lo (2.0 × 10⁶) thymocytes from CD45.1⁺ C57BL/6 mice were adoptively transferred into syngeneic CD45.2⁺ wild-type (WT) or Ltbr^−/− mice (with splenectomy 1 wk before the adoptive transfer) via tail vein injection. After the indicated periods of time, lymphocytes from blood, mesenteric lymph nodes, and spleen of the recipient mice were collected and stained with MitoTracker Green or MitoSOX Red. The mean fluorescence intensities of MitoTracker Green and MitoSOX Red in CD4⁺CD8⁻CD69⁻Qa2⁺CD44^lo thymocytes (CD45.1⁺ C57BL/6 mice) freshly obtained at each harvesting time point were used as controls.

In the adoptive transfer experiment with drug administration, the sorted CD4⁺CD8⁻CD69⁻ thymocytes were treated with MitoTEMPO (1 μM) or chloroquine (CQ; 5 μM) for 2 h before the transfer. The recipients were also given MitoTEMPO (0.7 mg/kg) or CQ (60 mg/kg) every day for 7 d before the transfer and 7 d after transfer. The control cells and recipients were treated with PBS. The circulating lymphocytes from the recipient mice were collected at 7 d after transfer and stained with MitoTracker Green and CD45.1.

For immunofluorescence assays, cytospins were prepared using a Shandon Cytospin (Thermo Fisher Scientific) with purified T cell subsets on microscope slides and then fixed in 4% paraformaldehyde for 30 min at room temperature. Fixed cells were permeabilized and blocked with 0.3% Triton X-100 and 5% BSA in PBS for 1 h and incubated with anti-LAMP1 (Abcam) and anti-Tom20 (Santa Cruz Biotechnology) at 4°C overnight. After washing, the cells were incubated with tetramethylrhodamine isothiocyanate–conjugated anti-rat Ab and Cy5-conjugated anti-rabbit Ab (Abcam) for 1 h protected from light at room temperature. Nuclei were stained with Hoechst 33342. The samples were examined with a Leica SP8 confocal microscope (Leica Microsystems IR) fitted with a ×63 objective, and images were processed and assembled using Leica software. The brightness and contrast were adjusted in Adobe Photoshop CS.

FTY720 (Sigma-Aldrich) was reconstituted with DMSO and diluted in PBS before use. FTY720 (1 mg/kg body weight) or DMSO (control) was i.p. injected into RAG2p-GFP transgenic mice every day for 5 consecutive days or every other day for a week.

An UltiMate 3000 UHPLC (Dionex, Waltham, MA) coupled with Q Exactive (Thermo Fisher Scientific, Carlsbad, CA) was applied in positive and negative switch mode. An Atlantis T3 column (2.1 × 100 mm; Waters) was used for liquid chromatography separation. Five millimolars ammonium acetate and acetonitrile was used for mobile phase A and B. The detailed parameters are as follows: spray voltage, 3.5 kV for positive and 2.5 kV for negative; capillary temperature, 275°C for positive and 320°C for negative; sheath gas flow rate (arbitrary units), 35; auxiliary gas flow rate (arbitrary units), 8; mass range (m/z), 70–1050 for positive and 80–1200 for negative; full mass spectroscopy resolution, 70,000.

The statistical analysis of the results was performed using GraphPad Prism 5 software (San Diego, CA). An unpaired or two-tailed paired Student t test was used to evaluate the significance of the differences between two groups. Data are presented as mean ± SD. A p value < 0.05 was considered significant (*p < 0.05, **p < 0.01, ***p < 0.005).

To study the mitochondrial contents in T cells, CD4 SP thymocytes were divided into CD69⁺Qa2⁻ immature thymocytes and CD69⁻Qa2⁻ and CD69⁻Qa2⁺ mature thymocytes (Fig. 1A). The Qa2⁺ subset of CD4 SP was previously identified as the most mature SP thymocytes that have acquired the ability to egress from thymic corticomedullary junctions (24, 54). To simplify the comparisons, CD69⁺ immature and CD69⁻ mature SP thymocytes were also isolated in some cases (2). The peripheral T cells were purified from SLOs of RAG2p-GFP transgenic mice, including GFP⁺CD4⁺CD8⁻CD44^loCD62L^hiCD25⁻ RTEs and GFP⁻CD4⁺CD8⁻CD44^loCD62L^hiCD25⁻ mature naive T cells (Fig. 1A) (29, 47). The RAG2p-GFP mice allow an easy identification of RTEs from unmanipulated mice, making the ex vivo metabolic analysis of T cells more reliable. Upon activation, CD25 upregulation and IL-2 production could be found in all of the T cell subsets. The levels of CD25 and IL-2 were similar between RTEs and naive T cells but were lower in SP thymocytes when compared with the peripheral ones (Supplemental Fig. 1A, 1B). This confirms that RTEs are functionally more mature than their precursors in the thymus.

The cell size, one of the indicators of metabolic status, was first compared among various T cell subsets. As shown in Fig. 1B and 1C, the cell size of RTEs was similar to naive T cells but was significantly smaller than SP thymocytes. We then compared the mitochondrial contents among various T cell subsets. Compared to SP thymocytes, peripheral naive T cells had ∼1.8-fold weaker staining of MitoTracker Green, a mitochondrial-selective fluorescent label commonly used for mitochondrial measurement (Fig. 1B, 1C). RTEs were not included in this analysis as GFP and MitoTracker Green had similar emission wavelengths. We thus purified various CD4⁺ T cell subsets and quantified mitochondria by measuring the amount of mtDNA relative to nuclear DNA. As shown in Fig. 1D, ∼2-fold less mtDNA was found in RTEs than in SP thymocytes, including ND1 in respiratory chain complex I, cytochrome b in complex III, Cox1 in complex IV, and 12S rRNA (Fig. 1D). The mRNA and protein levels of an outer mitochondrial membrane protein, VDAC, were also measured and lower levels were found in peripheral RTEs than in SP thymocytes (Fig. 1E, 1F). Naive T cells had either similar or slightly higher levels of mtDNA and VDAC than did RTEs (Fig. 1D–F). These results suggest that the mitochondrial content is reduced when SP thymocytes become RTEs and is maintained at low levels in mature naive T cells.

We further examined the mitochondria by transmission electron microscopy (TEM). Consistent with mtDNA analysis, RTEs revealed a nearly 2-fold reduction in mitochondrial amounts and mitochondrial area when compared with CD69⁻ SP thymocytes (Fig. 2). The cytoplasmic area and the relative area occupied by mitochondria were also significantly smaller in RTEs than in mature SP thymocytes (Fig. 2B). The areas of the nucleus were similar among various T cell subsets. Similar TEM measurements were obtained when RTEs and naive T cells or CD69⁻Qa2⁻ and CD69⁻Qa2⁺ SP thymocytes were compared (Fig. 2). These data again indicate a downregulation of mitochondrial content after SP thymocytes mature into RTEs in the periphery.

To investigate whether the reduction of mitochondrial content is accompanied with a decline of mitochondrial function, we compared mitochondrial membrane potential in different T cell subsets as it relates to mitochondrial capacity to generate ATP by oxidative phosphorylation (55). As shown in Fig. 3A, RTEs in the periphery had lower mitochondrial membrane potential than did SPs in the thymus, and naive T cells had the lowest mitochondrial membrane potential among the four T cell subsets.

We next measured mROS in T cell subsets. mROS is generated from the flow of electrons down the mitochondrial electron transport chain and can participate in cytoplasmic signaling processes (56, 57). The amount of mROS thus reflects the activity of mitochondrial oxidative metabolism. A decline of mROS also helps maintain the activity of reduction-oxidation (redox)–sensitive protein tyrosine phosphatases and prevents cells from premature activation. As shown in Fig. 3B, most Qa2⁻ and Qa2⁺ SP thymocytes were mROS^hi whereas ∼51.14 ± 15.12% of RTEs became mROS^lo. The percentage of mROS^lo cells increased to 85.58 ± 10.05% in mature naive T cells. Similar to mROS reduction, the intracellular ROS measured by carboxy-H₂DCDFA staining also showed lower levels in peripheral naive T cells than in SP thymocytes (Fig. 3C, GFP⁺ RTEs were not detected owing to fluorescence similarity).

We then performed Western blotting to directly compare the expression of several subunits in respiratory chain complex. As shown in Fig. 3D, NDUFA9 in complex I, Fp70 in complex II, and RISP in complex III all showed similar protein levels among SP thymocytes, peripheral RTEs, and naive T cells. This implicates that the reduction of mROS in RTEs may not be due to an altered electron transport chain composition.

Superoxide dismutase (SOD)₂ is responsible for the dismutation of O₂⁻ to H₂O₂/O₂ and inhibition of oxidative signals in mitochondria. Loss of SOD₂ causes an increase in mROS, a decrease in the mitochondrial membrane potential, and impaired oxidative phosphorylation (58). As shown in Fig. 3E, the protein level of SOD₂ was higher in RTEs than in mature SP thymocytes. Naive T cells had an even higher SOD₂ level (Fig. 3E). Thus, the upregulation of this antioxidant defense enzyme may contribute to mROS downregulation in peripheral T cells.

We further compared mitochondrial function and cellular metabolism by measuring OCR and ECAR in these T cell subsets. Without cell activation, SP thymocytes, RTEs, and naive T cells showed largely similar levels of mitochondrial respiration, spare respiratory capacity, and glycolytic function (Fig. 3F). Although not reaching statistical significance, the highest level of basal and maximal respiration was found in immature CD69⁺ SP thymoyctes whereas the lowest level was found in naive T cells. The metabolites were also measured by liquid chromatography–tandem mass spectrometry. Again, similar levels of metabolites or intermediates involved in the fatty acid β-oxidation were found among these T cell subsets (Supplemental Fig. 1C). However, a reduction in the levels of free carnitine, pivaloylcarnitine, tetradecanoylcarnitine, and tetradecenoylcarnitine could be detected in RTEs and naive T cells when compared with CD69⁻CD4 SP thymocytes. Additionally, lower expression of glucose transporter 1 was found in RTEs and naive T cells than in CD4 SP thymocytes (Supplemental Fig. 1D). Thus, in contrast to the continuous downregulation of mitochondrial membrane potential and mROS during SP thymocyte–RTE–naive T cell transition, the mitochondrial respiration and glycolytic function of SP thymocytes and peripheral RTEs/naive T cells were quite similar. This suggests that the changes of mitochondrial contents, mitochondrial membrane potential, and mROS during the SP–RTE–naive T transition may not be significant enough to alter the overall mitochondrial function at steady-state. However, this fine tuning of mitochondria may equip peripheral T cells, in particular naive T cells, with the ability to fully upregulate their aerobic glycolysis upon activation (44).

To investigate whether the reduction of mitochondrial content and mROS occurs in RTEs after thymic egress or in a small subset of RTE precursors (SP thymocytes) before thymic egress, we used FTY720, a sphingosine 1-phosphate analog, to block thymic egress (59, 60). After 5 d of FTY720 administration, the percentage of CD4 SP thymocytes was increased by 2-fold whereas very few GFP⁺ T cells were found in the peripheral blood (Fig. 4A, Supplemental Fig. 1E). The percentage of GFP^lo CD4 SP thymocytes was ∼5% in control mice and 20% in FTY720-treated mice, indicating a successful blockage of thymic egress and an enrichment of RTE precursors (Qa2⁺ cells) in the GFP^lo population in the thymus (Fig. 4A). As shown in Fig. 4B, Qa2⁺GFP^lo RTE precursors in FTY720-treated mice had similar levels of mROS when compared with Qa2⁺GFP^hi newly generated SP thymocytes. In contrast, lower mROS levels were found in RTEs and naive T cells in SLOs (Fig. 4B). This suggests that the downregulation of mROS and likely mitochondrial content requires thymic egress.

As the reduction of mROS occurs after young T cells have left the thymus, we examined whether peripheral lymphoid organs were required for this downregulation. To this end, RTE precursors (Qa2⁺CD69⁻ CD4 SP thymocytes) were adoptively transferred into WT mice and splenectomized Ltbr^−/− mice that lack organized SLOs (61, 62). The use of RTE precursors in the transfer experiment was to mimic the process of young T cells exiting from the thymus and entering the peripheral T cell pool. The changes of mROS and mitochondrial content of donor T cells were monitored in the recipients within a week. In WT recipients, a significant decrease in mROS was first observed in donor T cells in the peripheral blood at 1 d after transfer (Fig. 4C). The reduction of mROS in donor T cells in mesenteric lymph nodes and spleen was detectable at 3 d after transfer (Fig. 4C). The mitochondrial content by MitoTracker Green staining was reduced in donor T cells in blood, mesenteric lymph nodes, and spleen as early as 3 d after transfer (Fig. 4D). Low levels of mROS and mitochondrial mass were maintained until 7 d after adoptive transfer. In Ltbr^−/− recipients, the donor T cells had slightly more severe mROS reduction in blood at 1 d after transfer (Fig. 4E). The decrease of mitochondrial content in the circulating donor cells was similar in Ltbr^−/− and WT recipients (Fig. 4E). This suggests that young T cells that just left the thymus could quickly downregulate their mitochondrial volume and mROS in the circulation and do not rely on cell trafficking through SLOs.

To investigate whether the first pass of T cells in the blood is enough to induce mitochondrial downregulation, we treated the recipient mice with FTY720 every other day to inhibit mature T cell egress from SLOs (Fig. 4F). Two hours after the second dosing of FTY720, the mice received RTE precursors (Qa2⁺CD69⁻ CD4 SP thymocytes) via i.v. injection. Within 2 h of adoptive transfer, very few donor T cells could be found in the circulation of FTY720-treated mice (Fig. 4F). Both FTY720- and DMSO-treated recipients had similar ratios of donor T cells in the lymph nodes and spleen. As shown in Fig. 4G, the donor T cells in both types of recipients exhibit similar downregulation of mitochondria and mROS. These results suggest that the induction of mitochondrial downregulation in young T cells may occur during their first pass in peripheral blood, right after their egress from the thymus.

The homeostasis of mitochondria is constantly cross-regulated and coordinated by mitochondrial biogenesis, mitophagy, and mitochondrial dynamics (63, 64). To study the reasons for mitochondrial reduction after T cell egress, the protein expression of mitochondrial biogenesis-related molecules, including PGC-1α, NRF family members NRF1 and NRF2 (GABPα), and NRF-regulated TFAM, were first examined by Western blotting and no significant differences were found among CD4 SP thymocyte subsets, RTEs, and naive T cells (Fig. 5A, 5B). The levels of OPA1 and total and phosphorylated DRP1, essential molecules involved in mitochondrial fusion and fission, were also similar among these T cell subsets (Fig. 5B). Our results suggest that mitochondrial biogenesis and dynamics may not be the main contributors to mitochondrial reduction in RTEs.

We next examined the mitophagy levels in various T cell subsets. From TEM results, we found more autophagic structures and more mitophagosomes (abnormal mitochondria in autolysosome-like vesicles) in RTEs and naive T cells than in CD4 SP thymocytes (Fig. 5C). The Cyto-ID staining and flow cytometry analysis confirmed that naive T cells had a higher level of autophagy than did SP thymocytes (Fig. 5D). The confocal microscopy also showed an increased colocalization of lysosomal marker LAMP1 and mitochondrial import receptor Tom20 in RTEs and naive T cells when compared with SP thymocytes (Fig. 5E). We further performed the adoptive transfer experiment in which donor cells (RTE precursors) and recipients were both treated with an autophagy inhibitor, CQ. As shown in Fig. 5F, the reduction of mitochondria and mROS in circulating donor T cells was suppressed in the presence of CQ. These data indicate that enhanced autophagic clearance of mitochondria contributes to mitochondrial reduction in RTEs and naive T cells.

Multiple signaling molecules, such as mTOR, FoxO1, and MAPKs, play important roles in regulating autophagy (65–67). These signaling molecules are also essential sensors in integrating environmental cues and transducing them into programmed cellular responses (68–72). We thus investigated whether environmental differences between the thymus and periphery led to the activation of these molecules or their related signaling pathways.

Compared to SP thymocytes, peripheral RTEs and naive T cells showed lower levels of phosphorylation of mTOR (Ser²⁴⁴⁸) and its downstream targets S6K and 4E-BP1 (Fig. 6A). The levels of total mTOR and 4E-BP1 were similar among various T cell subsets. mTOR negatively regulates autophagy by phosphorylating ULK1 at Ser⁷⁵⁷ (73). Consistent with reduced mTOR phosphorylation in RTEs and naive T cells, the phosphorylation of ULK1 was also decreased in these T cells (Fig. 6B). mTOR activity can be negatively regulated by AMPK, a key sensor of cellular energy homeostasis. However, we did not observe differences in the phosphorylation of AMPK and its downstream signaling molecule ACC among various T cell subsets (Fig. 6B). The activity of mTOR was also negatively controlled by Tsc (12). We then used TSC1^fl/flCd4-Cre mice to examine whether increased mTORC1 activity led to the accumulation of mitochondria in RTEs and naive T cells. Compared to WT cells, Tsc1-deficient RTEs and naive T cells had larger cell size, higher ROS/mROS, and lower mitochondrial membrane potential (Supplemental Fig. 2A, 2B). However, the downregulation of mROS during SP–RTE–naive T transition was not completely suppressed in Tsc1-deficient T cells (Supplemental Fig. 2A, 2B). These data suggest that the changes in mTORC1 activity may not be the only factor involved in mitochondrial downregulation during SP–RTE–naive T cell transition.

We next determined whether FoxO1 was altered during SP–RTE transition. Compared to immature CD69⁺ SP thymocytes, the total FoxO1 protein level was significantly elevated in mature CD69⁻ SP thymocytes (Fig. 6C), consistent with the previous finding that FoxO1 upregulation during SP thymocyte maturation is essential in T cell quiescence (10, 15). RTEs and naive T cells had similar levels of total FoxO1 as mature CD69⁻ SP thymocytes (Fig. 6C). The phosphorylation of FoxO1 (Thr²⁴) and its regulator Akt (Ser⁴⁷³) was also increased in CD69⁻ SP thymocytes and remained high in RTEs and naive T cells (Fig. 6C). FoxO1 phosphorylation by Akt leads to the translocation of FoxO1 from the nucleus to the cytosol (74). We also found increased cytosolic localization of FoxO1 in mature CD69⁻ SP thymocytes and peripheral T cells (Fig. 6D). The levels of total, phosphorylated, and cytosolic FoxO1 showed no significant differences between CD69⁻ SP thymocytes and peripheral T cells, suggesting that FoxO1 may not be specifically involved in mitochondrial downregulation in RTEs and naive T cells.

Three groups of MAPKs have been reported to regulate autophagy, including ERK1 or ERK2, JNK1 or JNK2, and p38 MAPK (75). Compared to CD69⁺ CD4 SP thymocytes, more phosphorylation of JNK and p38 was found in CD69⁻ SP thymocytes (Fig. 6E), agreeing well with the previous reports that these two MAPKs play important roles in negative selection of SP thymocytes (76–78). Notably, the activation of JNK and p38 was further enhanced in CD4⁺ RTEs and naive T cells (Fig. 6E). RTEs and naive T cells also had elevated ERK phosphorylation when compared with CD4 SP thymocytes (Fig. 6E). These data suggest that MAPKs were activated during SP–RTE transition.

ASK1 is an MAPK kinase kinase that selectively activates JNK and p38. It can be activated by oxidative stress, ER stress, inflammatory cytokines, and LPS (79–81). The oxidative stress and intracellular redox balance also regulate ERK activity, as ERK and its upstream activator, Ras GTPases, contain redox-sensitive cysteines (82, 83). We have previously shown that during positive selection, the Trx system is important in protecting postselected DP thymocytes from ROS-induced activation of ASK1, JNK, and p38 and subsequent cell apoptosis (19). We thus determined the levels of Trx1 and ASK1 in SP thymocytes and peripheral T cells. As shown in Fig. 6F, significantly lower levels of Trx1 but higher levels of phosphorylated ASK1 were found in RTEs and naive T cells when compared with CD69⁺ and CD69⁻ CD4 SP thymocytes. These results suggest that cellular oxidation is increased after T cells enter the periphery, likely resulting in the activation of MAPKs.

To further determine whether cellular oxidation contributes to mitochondrial downregulation in RTEs and naive T cells, we used MitoTEMPO, a mitochondrial superoxide and alkyl radical scavenger, to treat donor T cells (CD4⁺CD8⁻CD69⁻ thymocytes) as well as the recipients. Seven days after adoptive transfer, the circulating donor T cells in the MitoTEMPO-treated group had higher levels of mitochondrial content than did those in PBS-treated one (Fig. 6G), indicating that oxidative stress during SP–RTE transition contributes to mitochondrial downregulation in RTEs.

Mature naive T cells circulate through peripheral lymphoid organs in a quiescent state (1). They rely primarily on catabolic metabolism and derive most of their ATP from oxidative phosphorylation, particularly fatty acid β-oxidation. Quiescent T cells also use autophagy to break down intracellular components to supply molecules for oxidative phosphorylation (84). We found that young CD4⁺ T cells (RTEs) downregulate their metabolic levels after emigrating from the thymus, being characterized by reduced cell size and cytosolic area, as well as decreased mitochondrial mass and mROS. Similar mitochondria and mROS downregulation could be also found in CD8⁺ RTEs (Supplemental Fig. 2C–G). This downregulation of mitochondria and mROS during SP thymocyte–RTE–naive T cell transition likely facilitates the quiescence and functional maturation of newly generated T cells.

The decline of mitochondrial mass and mROS does not occur during SP maturation in the thymus, even though T cells at this stage downregulate their surface CD69 and CD24 expression and upregulate essential quiescence regulators FoxO1 and Krüppel-like factor 2 (2). As RTE precursors in the thymus are resistant to high ROS-induced cell apoptosis and their IL-2 production is reduced when treated with ROS scavenger N-acetylcysteine, this delayed mitochondria and ROS downregulation may facilitate a full spectrum of negative selection (21) and the functional maturation of SP thymocytes (20). It may also explain the findings that conditional deletion of Atg3/5/7 and Vps34 (4–7) with Cd4-Cre did not lead to enhanced cell apoptosis and accumulation of mitochondrial and cellular ROS in SP thymocytes.

Our results further demonstrate that mitochondrial and mROS downregulation occurs in thymic emigrants, weeks before they become mature naive T cells. It is thus reasonable to think that the accumulation of mitochondria shown in Atg3/5/7- and Vps34-deficient naive T cells may be also found in RTEs (4–7). Additionally, the mitochondria biogenesis, fission, and fusion were not altered whereas autophagic clearance of mitochondria was upregulated in RTEs. This also agrees well with the findings that defects in autophagy lead to increased mitochondrial mass in peripheral T cells.

We also found that the reduction of mitochondrial mass and mROS requires T cell egress from the thymus. The first pass of thymic emigrants in the circulation may be enough to initiate this metabolic downregulation. The education within the SLOs is not essential. This indicates that factors in the circulation contribute to the induction of mitochondrial downregulation. One of the environmental changes that young T cells encounter when they migrate from the thymus to the circulation is an increased oxygen tension (5–13 kPa in blood versus 1.3 kPa in the thymus) (85, 86). Elevation of intracellular ROS may also occur when emigrating T cells respond to higher concentration of sphingosine 1-phosphate in the circulation (87, 88). As RTE precursors in the thymus have relatively higher levels of mitochondrial content and mROS, a sudden increase in extracellular oxygen tension or intracellular ROS level during their entry into the circulation may lead to oxidative stress. A decrease in Trx1 may further sensitize RTEs to respond to oxidative signals. Our data of increased phosphorylation of oxidative stress sensors ASK1, JNK, p38, and ERK in RTEs strongly support this possibility. The increase in mitochondrial content in circulating T cells after MitoTEMPO treatment also suggests that oxidative stress–induced activation of MAPKs may subsequently contribute to the autophagic clearance of mitochondria and reduction of ROS in young T cells (75).

Tsc1/mTOR plays an important role in regulating metabolism, T cell quiescence, and protecting cells from oxidative stress or ER stress (12, 15, 16, 89–92). The deletion of Tsc1, a stringent modulator of mTOR signaling, in T cells led to excessive mTORC1 activation, increased cell apoptosis, and altered metabolism and mitochondrial dynamics in peripheral T cells. Our data showed that RTEs and naive T cells had decreased phosphorylation of mTOR and its substrates S6K1 and 4E-BP1 when compared with SP thymocytes; the persistent mTORC1 activation by Tsc1 deletion leads to accumulation of mitochondria in not only naive T cells but also thymic emigrants. These data support the previous reports that a downregulation of mTORC1 activity in peripheral T cells is important. However, the mitochondrial content and mROS of Tsc1^−/− RTEs were still lower than in Tsc1^−/− SP thymocytes (Supplemental Fig. 2A) (12, 16, 93), implicating that signals in addition to mTOR may be involved in reducing mitochondria in young T cells.

Taken together, we demonstrate a mitochondrial downregulation process during SP thymocyte–RTE–naive T cell transition. Reaching this metabolic quiescence is likely one of the important processes during the functional maturation of RTEs. Defects in cell functions could be seen in T cells with a not-so-low level of mitochondria, mROS, and mTOR activity. For instance, mature CD4 SP thymocytes with higher levels of mitochondrial content/mROS have lower levels of Ag-induced proliferation and cytokine production when compared with peripheral CD4⁺ RTEs (25). Tsc1-deficient T cells with higher mTORC1 activity and mitochondrial accumulation have impaired T cell expansion and effector cell differentiation in vivo (12). Additionally, loss of T cell quiescence, in particular defects in mitophagy, results in the improper activation of multiclonal T cells, which may lead to autoimmunity. Thus, the downregulation of mitochondrial mass and mROS in young T cells immediately after thymic egress may also facilitate the peripheral tolerance of autoreactive T cells that escape from negative selection in the thymus. Deciphering the molecular mechanisms that regulate this mitochondrial downregulation in young T cells will increase our understanding of T cell maturation and how it becomes dysregulated under pathological conditions and during aging.

We thank Prof. Yangxin Fu (University of Chicago) and Prof. Yu Zhang (Peking University Health Science Center) for providing Ltbr^−/− and RAG2p-GFP mice. We thank Prof. Yu Zhang and Prof. Xian Wang for critical comments, helpful discussions, and critical reagents, Dr. Ying Li (Tsinghua University) for TEM analysis, and Dr. Qihua He (Peking University) for confocal microscopy.

The authors have no financial conflicts of interest.