Gene targeting RhoA reveals its essential role in coordinating mitochondrial function and thymocyte development

Abstract

Thymocyte development is regulated by complex signaling pathways. How these signaling cascades are coordinated remains elusive. RhoA of the Rho family small GTPases plays an important role in actin cytoskeleton organization, cell adhesion, migration, proliferation, and survival. Nonetheless, the physiological function of RhoA in thymocyte development is not clear. By characterizing a conditional gene targeting mouse model bearing T cell deletion of RhoA, we show that RhoA critically regulates thymocyte development by coordinating multiple developmental events. RhoA gene disruption caused a strong developmental block at the pre-TCR checkpoint and during positive selection. Ablation of RhoA led to reduced DNA synthesis in CD4⁻CD8⁻, CD4⁺CD8⁻, and CD4⁻CD8⁺ thymocytes but not in CD4⁺CD8⁺ thymocytes. Instead, RhoA-deficient CD4⁺CD8⁺ thymocytes showed an impaired mitosis. Furthermore, we found that abrogation of RhoA led to an increased apoptosis in all thymocyte subpopulations. Importantly, we show that the increased apoptosis was resulted from reduced pre-TCR expression and increased production of reactive oxygen species (ROS) which may be due to an enhanced mitochondrial function, as manifested by increased oxidative phosphorylation, glycolysis, mitochondrial membrane potential, and mitochondrial biogenesis in RhoA-deficient thymocytes. Restoration of pre-TCR expression or treatment of RhoA-deficient mice with a ROS scavenger NAC partially restored thymocyte development. These results suggest that RhoA is required for thymocyte development and indicate for the first time that fine-tuning of ROS production by RhoA, through a delicate control of metabolic circuit, may contribute to thymopoiesis.

Introduction

T cell development in thymus proceeds through a series of differentiation stages (1–4). The most immature populations in thymus comprise CD4⁻CD8⁻ double-negative (DN) thymocytes. DN thymocytes can be subdivided into four developmental stages: CD44⁺CD25⁻ (DN1), CD44⁺CD25⁺ (DN2), CD44⁻CD25⁺ (DN3), and CD44⁻CD25⁻ (DN4). DN thymocyte development is signified by TCRβ gene rearrangements and β-selection. TCRβ gene rearrangements are known as V(D)J recombination which begin at DN2 stage, continue and complete at DN3 stage. β-selection occurs at DN3 stage and is a process in which thymocytes successfully expressing pre-TCR complex consisting of rearranged TCRβ and preTα are rescued from cell death and allowed to proliferate and differentiate to DN4 and subsequent CD4⁺CD8⁺ double-positive (DP) cells. At DP stage, thymocytes undergo two concurrent developmental processes to mature to CD4⁺ or CD8⁺ single-positive (SP) thymocytes. The first process is called repertoire selection including positive and negative selection. During repertoire selection, thymocytes that express TCRαβ of low affinity to self-peptide/MHC complexes are instructed to undergo positive selection and maturation to SP cells, whereas thymocytes that express TCRαβ of high affinity to self-peptide/MHC complexes and thus are potentially autoreactive are eliminated by negative selection. The second process is termed lineage commitment, during which thymocytes that recognize self-peptide/MHC class I become CD8⁺ SP cells and thymocytes that recognize self-peptide/MHC class II become CD4⁺ SP cells.

RhoA is an intracellular signal transducer of the Rho family small GTPases that cycles between an inactive GDP-bound form and an active GTP-bound form under tight regulation (5, 6). Mostly by overexpression of dominant active or negative mutants, RhoA has been shown to modulate actin cytoskeleton organization, cell adhesion, migration, proliferation, and survival (7–11). In T cells, overexpression of the dominant mutants suggest that RhoA plays a role in T cell polarization, thymocyte adhesion, and thymic egress (12–17). Furthermore, inactivation of RhoA by C3 transferase in transgenic mice caused thymocyte developmental blocks (18–19). However, these approaches are hampered by the nonspecific nature of the dominant RhoA mutants or C3 transferase, as they may affect other Rho GTPases (20–24). Indeed, distinct cell functions of RhoA have been observed in studies of RhoA knockout mouse models. For example, contrary to the conventional view that RhoA is essential for actin cytoskeleton rearrangement and cell adhesion, RhoA-deficient primary mouse embryonic fibroblasts (MEFs) display normal actin stress fiber and focal adhesion complex formation (25). It is therefore highly desirable to use a gene targeting strategy to assess the physiological functions of RhoA in T cells.

In this study, we have examined the physiological contribution of RhoA in thymocyte development by characterizing a T cell-specific RhoA conditional knockout mouse model. We demonstrate that RhoA is required for thymocyte development by coordinating DN thymocyte β-selection, DP thymocyte positive selection, SP thymocyte lineage commitment, thymocyte proliferation and survival, and importantly, mitochondrial function. Our data suggest that RhoA couples metabolic homeostasis to thymocyte development.

Materials and Methods

Mouse gene targeting

Conditionally targeted RhoA^flox/flox mice were generated as previously described (25). The floxed allele contains loxP sites flanking exon 3 of the RhoA allele. To delete RhoA in vivo in the T cell lineage, RhoA^flox/flox mice were mated with mice expressing Cre recombinase under the control of a CD2 or Lck proximal promoter (Jackson Laboratory). RhoA^flox/flox;CD2-Cre mice (hereafter referred to as RhoA^−/−) were crossed with p14 mice expressing transgenic TCRVα2Vβ8 to generate RhoA^−/−;p14TCR Tg compound mice. Mice used for experiments ranged in age from four to eight weeks. Animals were housed under specific pathogen-free conditions in the animal facility at Cincinnati Children's Hospital Research Foundation in compliance with the Cincinnati Children’s Hospital Medical Center Animal Care and Use Committee protocols.

Flow cytometry analysis by cell-surface and intracellular staining

Single cell suspensions were prepared from thymus. Cells were incubated with various combinations of fluorophore-conjugated antibodies against the following cell-surface markers (BD Biosciences) at room temperature for 20 min: Thy1.2, CD28, c-Kit, CD4, CD8, CD44, CD25, CD69, TCRβ, IL-7Rα, HSA, CD62L, and TCR Vα2. Immunolabeled cells were analyzed by flow cytometry.

Intracellular TCRβ staining was performed as described previously (26). Briefly, cells were incubated with fluorophore-conjugated antibodies to cell-surface markers (anti-CD4, anti-CD8, anti-Thy1.2, anti-CD44, and anti-CD25, anti-CD28), as well as with a saturating amount of anti-TCRβ to block cell-surface TCRβ. Cells were then fixed at room temperature for 20 minutes with 1% paraformaldehyde. After washing in PBS, the cells were incubated with 10 mM glycine for 10 minutes to quench autofluorescence. Cells were then permeabilized using 0.5% saponin, 5% FCS, and 10 mM Hepes (pH 7.4) at room temperature for 10 minutes and stained with fluorophore-conjugated antibody to TCRβ at room temperature for 45 minutes. Intracellular staining of Bcl2 was carried out similarly, except that the Bcl2 antibody was revealed with secondary fluorophore-conjugated goat anti–mouse IgG (Jackson ImmunoResearch).

PCR genotyping to detect RhoA gene deletion

Thymocyte subsets were sorted by flow cytometry and genomic DNA from the sorted cells was isolated and analyzed by PCR using primers 5′-TCTCTGCACTGAGGGAGTTAGG-3′ (forward) and 5′-GTACATACAGGGAATGGAAAC AAGG-3′ (reverse) to detect floxed allele and primers 5′-GCACTGAGGGAGTTAGG −3′ (forward) and 5′-CTACACTAGCTGGGCAC −3′ (reverse) to detect knockout allele. For detecting floxed allele, PCR was carried out for 33 cycles with a 30-second annealing time at 58°C. For detecting knockout allele, PCR was carried out for 30 cycles with a 30-second annealing time at 60°C. PCRs were analyzed by agarose gel electrophoresis. The reaction gave rise to a 633-bp fragment for floxed allele and a 667-bp fragment for knockout allele.

Cell apoptosis analysis

Freshly isolated thymocytes were incubated with fluorophore-conjugated anti-CD4, anti-CD8, anti-CD44, and anti-CD25 antibodies for 20 min. Cells were washed, incubated with fluorophore-conjugated Annexin V (BD Biosciences) for 20 min and then analyzed by flow cytometry on a FACSCanto system using FACSDiVa software (BD Biosciences).

In vivo BrdU incorporation assay

Mice were injected intraperitoneally with 500µg BrdU. Twelve hours after injection, thymocytes were isolated, immunolabeled with fluorophore-conjugated antibodies against CD4, CD8, CD44, and CD25, fixed, permeabilized, and then incubated with fluorophore-conjugated anti-BrdU antibody, according to the manufacturer’s protocol (BD Biosciences). Immunolabeled cells were then analyzed by flow cytometry analysis.

In vitro proliferation assay

DP thymocytes were sorted by flow cytometry, plated on 48-well plates with or without anti-CD3/-CD28 antibodies, and cultured for 3 days. Cell growth rates were assayed by a nonradioactive cell proliferation assay kit (Promega).

Immunoblotting

Total thymocytes or flow cytometry-sorted DP thymocytes were lysed and protein content was normalized by Bradford assay. Lysates were separated by 10% SDS-polyacrylamide gel electrophoresis. The expression or activation (phosphorylation) of RhoA, ZAP70, ERK, and JNK was probed by Western blot using corresponding antibodies (Cell Signaling Technology).

Transcript expression analysis

RNA was isolated from thymocytes using RNeasy Micro Kit (QIAGEN) and converted to cDNA using a High Capacity cDNA Reverse Transcription Kit (Applied Biosystems Inc). Real-time PCR was performed with SYBR Green dye or Taqman assay on a 7900HT Real-Time machine (Applied Biosystems Inc). Data were analyzed using SDS 2.3 software (Applied Biosystems Inc) and normalized to GAPDH.

The primer sequences were as following: Nrf1: TATGGCGGAAGTAATGAAAGACG (forward), CAACGTAAGCTCTGCCTTGTT (reverse); Atp5I: GAGAAGGCACCGTCGATGG (forward), ACACTCTGAATAGCTGTAGGGAT (reverse); Cox5a: GCCGCTGTCTGTTCCATTC (forward), GCATCAATGTCTGGCTTGTTGAA (reverse); Ndufa2: TTGCGTGAGATTCGCGTTCA (forward), ATTCGCGGATCAGAATGG GC (reverse); HK2 F: TGATCGCCTGCTTATTCACGG (forward), AACCGCCTAGAAATCTCCAGA (reverse); slc2a: CAGTTCGGCTATAACACTGGTG (forward), GCCCCCGACAGAGAAGATG (reverse); PDK1: GGACTTCGGGTCAGTGAATGC (forward), TCCTGAGAAGATTGTCGGGGA (reverse); Pgm1: CAGAACCCTTTAACCTCTGAGTC (forward), CGAGAAATCCCTGCTCCCATAG (reverse). The Taqman primer/probe sets for Notch 3, Hes1, Notch1, PreTα, Deltex1 were Mm00435270_ m1, Mm01342805, Mm00435249_m1, Mm00492291_m1, Mm01281478_m1, respectively.

V(D)J recombination

TCRβ gene rearrangement was detected as described previously (27). In brief, genomic DNA was isolated from flow cytometry-sorted DN3 cells and amplified by PCR. PCR for Dβ2-Jβ2, Vβ5-Jβ2, Vβ8-Jβ2, Vβ11 -Jβ2, and eF-1α was performed for 31 cycles with a 1-minute annealing time at 63°C. PCR for Dβ1.1-Jβ1.7 and Dβ2.1-Jβ2.7 was first performed for 20 cycles with external primers. A total of 0.5 µl from the first amplification was used for a second PCR for an additional 31 cycles with nested internal primers.

Primer sequences for detecting Dβ2-Jβ2, Vβ5-Jβ2, Vβ8-Jβ2 and Vβ11 -Jβ2 recombination, and for eF-1α were as following: Dβ2: GTAGGCACCTGTGGGGAAGAAACT (forward), Vβ5: CCCAGCAGATTCT CTCAGTCCAACAG (forward), Vβ8: GCATGGGCTGAGGCTGATCCATTA (forward), Vβ11: TGCTG GTGTCATCCAAACACCTAG (forward), Jβ2: TGAGAGCTGTCTCCTACTA TCGATT (reverse), eF-1α: CTGCTGAGATGGGAAA GGGCT (forward), TTCAGGATAATCACCTGA GCA (reverse) (28). Primer sequences for detecting Dβ1.1-Jβ1.7 and Dβ2.1-Jβ2.7 recombination were as following: external primers: Dβ1.1ext: GAGGAGCAGCTTATCTGG TG (forward), Jβ1.7ext: AAGGGA CGACTCTGTCTTAC (reverse); Dβ2.1ext: TAGGCAACCTGTGGG GAAGAA AC (forward), Jβ2.7ext: TGAGAGCTGTCTCC TACTAT C (reverse). Nested internal primers: Dβ1.1int: GGTAGACCTATGGG AGGGC (forward), Jβ1.7int: ACCATGGTCATCCAACACA G (reverse); Dβ2.1int: GTATCACGATGTA ACATTGTG (forward), Jβ2.7int: GGAAGCGAGAGATGT GAATC (reverse) (29).

High-speed cell imaging analysis in flow using Imagestream^X

The nuclear contents and the distribution of F-actin and β-tubulin within DP thymocytes were studied by Imagestream^X (Amnis), which combines flow cytometry and microscopy (60×/numerical aperture 0.9 objective lens) capabilities. Thymocytes were harvested and cultured with anti-CD3/-CD28 antibodies for 3 days. The cells were fixed with 4% formaldehyde. Formaldehyde was removed by centrifugation, and the cell pellet was cooled on ice for at least 15 minutes, before permeabilization by consecutive suspensions in ice-cold 50% acetone, 100% acetone, and again 50% acetone solution. Cells were then incubated with fluorophore-conjugated antibodies to CD4 and CD8 and either fluorophore-conjugated anti–β-tubulin antibody (Cell Signaling) or phalloidin (Invitrogen). The nuclear stain Draq5 or DAPI (Cell Signaling Technology) was added at a concentration of 40µM and the samples were processed by Imagestream^X. Approximately 10000 events per sample were collected and analyzed with the associated Image Data Exploration and Analysis software (IDEAS; Amnis).

Metabolic assays

For measurement of oxygen consumption rate (OCR), 10⁶ freshly isolated thymocytes were resuspended in XF assay medium (pH 7.4) containing 2 mM GlutaMax, 1 mM sodium pyruvate and supplemented with 25 mM glucose, plated onto Seahorse Bioscience XF24 cell culture plates using Cell Tak (BD Bioscience), and incubated without CO₂ at 37 °C. Respiration was measured using the Seahorse XF24 Analyzer (Seahorse Biosciences) under basal condition and in the presence of the mitochondrial inhibitor oligomycin (0.6 µM) and mitochondrial uncoupler FCCP (1 µM). For measurement of extracellular acidification rate (ECAR), 10⁶ freshly isolated thymocytes were resuspended in assay medium (pH 7.4) (Sigma-Aldrich) containing 2 mM L-Glutamin and 2.5 mM Glucose, plated onto XF24 cell culture plates, and incubated as for OCR. Extracellular acidification was measured using the Seahorse XF24 Analyzer under basal condition and in the presence of the glycolysis substrate Glucose (10 mM), mitochondrial inhibitor oligomycin (0.6 µM) and glycolysis inhibitor 2 Deoxy-D-glucose (100 mM) (30, 31).

For measurements of ROS, mitochondrion numbers and mitochondrial membrane potential, thymocytes stained for CD4 and CD8 were incubated with 5 µM DCFDA, 100 nM Mitotracker Green and 50 nM DilC-5 (Invitrogen), respectively, accordingly to the manufacturer’s protocols. The cells were then analyzed by flow cytometry (31).

For mitochondrial DNA assay, thymocytes were lyzed and homogenized and mitochondrial DNA was then purified with a Mitochondrial DNA isolation Kit (Biovision), accordingly to the manufacturer’s manual. The purified mitochondrial DNA was quantified by quantitative real-time PCR with SYBR Green dye. Mitochondrial DNA content was represented by mitochondrial cyclo-oxygenase (Cox) 2 normalized to nuclear intron of β-globin. The primer sequences were: Cox2: GCCGACTAAATCAAGCAACA (forward), CAATGGGCATAAAGCTATGG (reverse); β-globin: GAAGCGATTCTAGGGAGCAG (forward), GGAGCAGC GATTCTGAGTAGA (reverse) (32). For ATP assay, thymocytes were lyzed with a cytosol extraction buffer supplied in the Mitochondrial DNA isolation Kit, homogenized and then ATP was determined by an ATP Determination Kit (Molecular Probes), accordingly to the manufacturer’s protocol.

Statistical Analysis

Data are expressed as the mean ± standard deviation (SD). Data were analyzed by a Student’s unpaired t-test with a two-tailed distribution. Significance was accepted at p<0.05.

Results

Defective thymocyte development in the absence of RhoA

To delete RhoA gene in T cell lineage, RhoA^flox/flox mice were crossbred with CD2-Cre transgenic mice. The RhoA^flox/flox;CD2-Cre (RhoA^−/−) mice had a drastic reduction in total thymocyte cellularity (Fig. 1A). The proportion of thymocyte subsets were altered with increased frequency of DN2 and DN3 but decreased frequency of DN4 and DP thymocytes (Fig. 1B, C). The numbers of all thymocyte subsets including DN1-DN4, DP, CD4⁺ SP and CD8⁺ SP thymocytes were markedly decreased in mutant mice (Fig. 1B, C). Analysis of deletion of RhoA gene and RhoA protein found that residual thymocytes in mutant mice were partially depleted of RhoA, suggesting that there is strong selection against the loss of RhoA (Fig. 1D). Because CD2-Cre may also mediate gene deletion in B cell lineage, to achieve T cell specific deletion we crossed RhoA^flox/flox mice with Lck-Cre transgenic mice. We found that RhoA deletion by Lck-Cre also caused a decrease in thymocyte numbers (Fig. 1E). However, the decrease appears to be less profound compared to that by CD2-Cre-mediated RhoA deletion. This might be explained by relatively late deletion of RhoA by Lck-Cre. Indeed, CD2-Cre is known to delete genes earlier than Lck-Cre (26), and CD2-Cre was able to delete RhoA in DN1 cells but Lck-Cre failed to do so (Fig. 1F). As a result, RhoA deletion by CD2-Cre led to an accumulation of early T cell precursors (ETPs) among DN1 cells, whereas Lck-Cre-mediated RhoA deletion had no effect on this early thymocyte subset (Fig. 1G). Collectively, these data suggest that RhoA is required for T cell development in the thymus and RhoA has a T cell intrinsic role in T cell development.

Figure 1. RhoA deficiency causes thymocyte hypocellularity.

(A) The numbers of total thymocytes from RhoA^flox/flox (wild type (WT)) and RhoA^flox/flox;CD2-Cre (RhoA^−/−) mice. (B) Left: representative flow cytometry plots of thymocyte subsets. Percentage of cells in each corresponding quadrant is indicated; Right: quantification of the numbers of thymocyte subsets. DN: CD4CD8 double-negative, DP: CD4⁺CD8⁺ double-positive, SP: single-positve. (C) Left: representative flow cytometry plots of thymocyte subsets gated from DN cells. Percentage of cells in each corresponding quadrant is indicated; Right: quantification of the numbers of thymocyte subsets within DN cells. DN1: CD44⁺CD25, DN2: CD44⁺CD25⁺, DN3: CD44CD25⁺, DN4: CD44CD25. (D) Upper: RhoA protein expression. Thymocytes from WT and RhoA^−/− mice were detected for RhoA expression by Western blot. Lower: efficiency of RhoA gene deletion. DNA from flow cytometry-sorted cells was used as PCR template to determine the floxed and knockout (KO) alleles of RhoA. Data are representative of 3 mice per genotype. (E) Quantification of the numbers of thymocyte subsets in RhoA^flox/flox;Lck-Cre and RhoA^flox/flox;LCK-Cre⁺ mice. (F) Comparison of RhoA gene deletion in DN1 cells from RhoA^flox/flox;LCK-Cre⁺ mice with that from RhoA^−/− mice. DNA from flow cytometry-sorted DN1 cells was used as PCR template to determine the floxed and KO alleles of RhoA. (G) Comparison of the frequency of early T cell precursors (ETPs) in RhoA^flox/flox;LCK-Cre⁺ mice with that in RhoA^−/− mice. ETPs were identified by flow cytometry as CD4CD8Thy1.2⁺CD44⁺c-Kit⁺CD25 cells. n=5 mice per genotype for A-C and E. For G, n=4 WT and 7 RhoA^−/− mice; and n=9 mice for both RhoA^flox/flox;LCK-Cre and RhoA^flox/flox;LCK-Cre⁺*P <0.05; **P <0.01. Error bars represent SD.

Defective β-selection in the absence of RhoA

RhoA deficiency resulted in a reduced cellularity of all DN subsets and an increased percentage of DN3 cells at the expense of DN4 and DP cells, suggesting that the developmental transition from DN3 to DN4 and then DP cells was blocked in RhoA^−/− mice. DN3 transition to DN4 and DP stage requires successful TCRβ gene rearrangements, namely V(D)J recombination, and β-selection (1–4). We found that Vβ11-Jβ2 and Vβ5-Jβ2 recombination was moderately impaired upon RhoA deletion, whereas Vβ8-Jβ2, Dβ2-Jβ2, Dβ2.1-Jβ2.7, and Dβ1.1-Jβ1.7 rearrangements were completely intact (Fig. 2A). These results suggest that RhoA deficiency doesn’t cause gross defect in TCRβ gene rearrangements. In contrast, RhoA^−/− mice showed fewer DN3 cells, particularly DN3b cells, that express intracellular (Ic) and surface TCRβ (Fig. 2B, C). A substantial reduction in the frequency of IcTCRβ⁺ DN4 cells and DN4 cells expressing surface TCRβ was also detected in RhoA^−/− mice (Fig. 2B, C). Concomitantly, the percentage of IcTCRβ⁻ DN4 cells was increased in the absence of RhoA (Fig. 2B). However, the total numbers of both IcTCRβ⁺ and IcTCRβ⁻ DN4 cells were decreased in RhoA^−/− mice (Fig. 2B). Furthermore, RhoA deficiency caused a reduced expression of preTα (Fig. 2D). Together, these data suggest that RhoA regulates β-selection by governing expression and/or transport of TCRβ and preTα, but not TCRβ gene rearrangements.

Figure 2. RhoA deficiency causes a defect in β-selection.

(A) Genomic DNA PCR analysis of V(D)Jβ recombination in flow cytometry-sorted 50,000 DN3 cells. The application of eF-1α was used as input control. Data are representative of 3 mice per genotype. (B) Flow cytometry analysis of intracellular TCRβ (IcTCRβ) protein expression. Left: representative flow cytometry plots of IcTCRβ⁺ and IcTCRβ cells gated from DN3a (CD4CD8Thy1.2⁺CD44CD25⁺CD28), DN3b (CD4CD8Thy1.2⁺CD44CD25⁺CD28⁺), or DN4 (CD4CD8Thy1.2⁺CD44CD25CD28⁺) cells. Percentage of IcTCRβ⁺ and IcTCRβ cells is indicated above bracketed lines. Right: mean percentage and numbers of IcTCRβ⁺ and IcTCRβ cells. (C) Flow cytometry analysis of cell-surface protein expression of TCRβ. TCRβ⁺ cells were gated from DN3 (CD4CD8Thy1.2⁺CD44CD25⁺) or DN4 (CD4CD8Thy1.2⁺CD44CD25) cells. Mean percentage of DN3 and DN4 cells expressing TCRβ is shown. (D) mRNA expression of preTα in DN3 (CD4CD8Thy1.2⁺CD44CD25⁺) cells. The data are presented as fold expression relative to one WT mouse. n=4 WT and 7–8 RhoA^−/− mice for B–D. *P <0.05; **P <0.01. Error bars represent SD.

Impaired proliferation and survival in DN cells in the absence of RhoA

Successfully rearranged TCRβ forms complex with preTα and CD3 into pre-TCR on the cell surface of DN3 cells. These DN3 cells are rescued from cell death and committed to proliferate and differentiate to DN4 and DP cells (33). Measurement of cell proliferation by BrdU incorporation demonstrated that in wild-type (WT) mice, there was a significant increase of cell division in DN4 cells comparing with DN3 cells, reflecting successful β-selection. However, such an increased proliferation from DN3 to DN4 was abolished in the absence of RhoA (Fig. 3A). Moreover, RhoA deficiency diminished the proliferation rate of DN4 cells without affecting that of DN3 cells (Fig. 3A). Nonetheless, Annexin V staining revealed that both DN3 and DN4 cells from the mutant were more susceptible to apoptosis compared to WT counterparts (Fig. 3B). These data provide further evidence that RhoA is required for β-selection.

Figure 3. RhoA deficiency causes an impaired proliferation and/or survival in DN3 and DN4 thymocytes that can be rescued by restoration of pre-TCR expression.

(A) Flow cytometry analysis of BrdU incorporation. BrdU⁺ cells were gated from DN3 or DN4 cells. Mean percentage of BrdU⁺ cells is shown. (B) Flow cytometry analysis of apoptosis. Annexin V⁺ cells were gated from DN3 or DN4 cells. Mean frequency of Annexin V⁺ cells is shown. (C) mRNA expression of Notch signaling molecules in DN4 cells. The data are presented as fold expression relative to one WT mouse. (D) Flow cytometry analysis of protein expression of cell-surface IL-7Rα and intracellular Bcl-2 in DN3, IcTCRβ⁺ DN4 and IcTCRβ DN4 cells. Mean fluorescence intensity (MFI) is shown. (E) Flow cytometry analysis of intracellular TCRβ (IcTCRβ) protein expression in DN3 and DN4 cells from WT;p14TCR transgenic (Tg) and RhoA^−/−;p14TCR Tg mice. Mean percentage of IcTCRβ⁺ cells is shown. (F) Flow cytometry analysis of apoptosis in DN3 or DN4 cells from the non-transgenic and transgenic mice indicated. Mean frequency of Annexin V⁺ cells is shown. (G) Flow cytometry analysis of the frequency of DN3 and DN4 cells in the non-transgenic and transgenic mice indicated. n=3–6 mice per genotype. The results from a representative experiment of two (A, C, D) or three (B) independent experiments are shown. **P <0.01. Error bars represent SD.

Cell survival at DN thymocytes is regulated by IL-7R and Notch signaling (26). Unexpectedly, pro-survival Notch1 and its downstream targets Notch3, Hes1, and Deltex1 were elevated in RhoA-deficient DN cells (Fig. 3C), possibly reflecting a compensatory effect of increase cell apoptosis, whereas the expression levels of IL-7R and its signaling transducer Bcl2 appeared normal (Fig. 3D).

We then examined if the increased apoptosis in RhoA^−/− DN cells was caused by defective pre-TCR expression. To this end, we restored pre-TCR expression in RhoA^−/− DN cells by crossing RhoA^−/− mice with p14TCR transgenic mice (Fig. 3E). We found that the restoration of pre-TCR expression in resultant RhoA^−/−;p14TCR transgenic mice partially rescued DN cell survival and distribution (Fig. 3F, G). Together, these data suggest that RhoA regulates DN cell survival during β-selection independent of IL-7R and Notch signaling, but dependent on pre-TCR expression.

Defective positive selection and lineage commitment in the absence of RhoA

RhoA deficiency led to reduced cellularity of the CD4⁺ SP and CD8⁺ SP thymocytes. The decreased numbers of SP thymocytes may not only result from defective β-selection, but also from impaired positive selection and/or lineage commitment. To examine positive selection in RhoA^−/− mice, we analyzed DP, CD4⁺ SP and CD8⁺ SP thymocytes for the surface expression of CD69, upregulation of which is a critical marker of successful positive selection (34). We found that RhoA^−/− mice had more CD69^hiCD4⁺CD8⁺, but less CD69^hiCD4⁺ and CD69^hiCD8⁺, thymocytes than the WT control mice (Fig. 4A). To determine at which specific point DP thymocyte development was blocked in RhoA^−/− mice, we analyzed the expression of TCRβ together with CD69 in total thymocytes (1, 34), by FACS. As shown in Fig. 4B, RhoA^−/− mice had fewer TCR^loCD69⁻ thymocytes (preselection DP thymocytes), likely due to defective β-selection. However, the proportion of TCR^intCD69⁺ cells (mainly DP thymocytes at the initial stage of positive selection) was increased in the absence of RhoA, whereas TCR^hiCD69⁺ (immature SP thymocytes) and TCR^hiCD69⁻ cells (mature SP thymocytes) were lower in RhoA^−/− mice. These data suggest that RhoA-deficient thymocytes were blocked at the initial stage of positive selection. The detrimental effect of RhoA deficiency on positive selection was reaffirmed in RhoA^−/−;p14TCR transgenic mice. p14TCR mice express TCRVα2Vβ8 that specifically recognizes MHC class I molecule-restricted lymphocytic choriomeningitis virus (LCMV) epitope gp33–41, and thus are dominated with monoclonal CD8⁺ T cells (35). Because the majority of DP thymocytes recognize self-ligands enabling positive selection, p14TCR mice are biased towards positive selection. RhoA deficiency in p14TCR mice resulted in a marked decrease in CD8⁺ SP thymocytes and lower expression of Vα2 TCR on CD8⁺ SP thymocytes (Figure 4C). Taken together, these results indicate that RhoA is required for thymocyte positive selection.

Figure 4. RhoA deficiency causes an impaired positive selection.

(A) Flow cytometry analysis of CD69⁺ cells gated from DP, CD4⁺ SP or CD8⁺ SP thymocytes. Mean frequency of CD69⁺ cells is shown. (B) Flow cytometry analysis of CD69 and TCRβ expression in total thymocytes. (C) Flow cytometry analysis of the frequency of CD8⁺ SP and TCRVα^hiCD8⁺ SP thymocytes in WT;p14TCR transgenic (Tg) and RhoA^−/−;p14TCR Tg mice. (D) Flow cytometry analysis of CD4⁺CD8^int cells gated from TCR^hiCD69⁺ population. Mean frequency of CD4⁺CD8^int cells is shown. (E) Flow cytometry analysis of HSACD62L^hi cells gated from CD4⁺ SP or CD8⁺ SP thymocytes. Mean frequency of HSACD62L^hi cells is shown. n=3 mice per genotype. The results from a representative experiment of two independent experiments are shown. *P <0.05; **P < 0.01. Error bars represent SD.

After the initiation of positive selection, DP thymocytes downregulate CD4 and CD8, passing through a CD4⁺CD8^int transitional stage before committing to mature CD4⁺ SP and CD8⁺ SP thymocytes (1). We found that RhoA^−/− mice had fewer CD4⁺CD8^int cells (Figure 4D). However, the frequency of mature (HSA⁻CD62L^hi) CD4⁺ SP and CD8⁺ SP thymocytes were not altered by RhoA deficiency (Figure 4E), suggesting that RhoA deficiency causes defects in early, but not late, stage of lineage commitment. Thus, RhoA is not only required for thymocyte positive selection, but also for lineage commitment.

Impaired survival and proliferation in DP and CD4⁺ and CD8⁺ SP thymocytes in the absence of RhoA

Positive selection is accompanied by cell survival and proliferation (36). It is thus logical to postulate that an impaired survival and proliferation could contribute to the decreased DP, CD4⁺ SP and CD8⁺ SP thymocytes in RhoA^−/− mice. Indeed, Annexin V staining revealed that there was an increased apoptosis in DP, CD4⁺ SP and CD8⁺ SP mutant thymocytes (Fig. 5A). Interestingly, although RhoA^−/− CD4⁺ SP and CD8⁺ SP thymocytes had less BrdU incorporation, DP thymocytes from RhoA^−/− mice had comparable BrdU incorporation to that of WT counterparts (Fig. 5B), suggesting that the S phase (DNA synthesis) of cell cycle is impaired in CD4⁺ SP and CD8⁺ SP, but not in DP, thymocytes. Because RhoA has been shown to play an important role in the M phase (mitosis) of cell cycle (25, 37, 38), we examined whether mitosis was defective in RhoA-deficient DP thymocytes. Analysis of nuclear contents by using Imagestream^x imaging flow cytometer (Amnis) found that RhoA deficiency resulted in a significant elevation of multinucleated cells in DP thymocytes, when the cells were cultured with anti-CD3/-CD28 antibodies (Fig. 5C, D, E). Both F-actin and β-tubulin assemblies, two critical structural components of mitosis (39, 40), were abolished in RhoA-deficient multinucleated cells (Fig. 5D, E). RhoA^−/− DP thymocytes bearing single nucleus also showed an impaired β-tubulin staining, compared to WT counterparts (Fig. 5F). These data suggest that RhoA is critical for regulating mitosis of DP thymocytes through controlling actin and tubulin cytoskeleton. Consistent with defective mitosis, RhoA-deficient DP thymocytes proliferated slower than WT control cells in response to TCR crosslinking with anti-CD3/-CD28 antibodies (Fig. 5G). Because survival and proliferation activities depend on TCR signaling in developing DP thymocytes, we analyzed intracellular signaling events in sorted DP thymocytes from RhoA^−/− and WT mice. As shown in Fig. 5H, the phosphorylation of ZAP70, ERK and JNK was impaired in the absence of RhoA, suggesting that RhoA is essential for TCR signaling.

Figure 5. RhoA deficiency causes an impaired survival and/or proliferation in DP and CD4⁺ SP and CD8⁺ SP thymocytes.

(A) Flow cytometry analysis of apoptosis. Annexin V⁺ cells were gated from DP, CD4⁺ SP or CD8⁺ SP cells. Mean frequency of Annexin V⁺ cells is shown. (B) Flow cytometry analysis of BrdU incorporation. BrdU⁺ cells were gated from DP, CD4⁺ SP or CD8⁺ SP cells. Mean percentage of BrdU⁺ cells is shown. (C) Imagestream analysis of nuclear contents in DP thymocytes cultured with anti-CD3/-CD28 antibodies. Mean frequency of multinucleated cells is shown. (D and E) Imagestream analysis of F-actin (D) and β-tubulin (E) distribution in DP thymocytes cultured with anti-CD3/-CD28 antibodies. Images shown in each panel are from samples pooled from at least 3 mice of same genotypes. (F) Quantification of mean fluorescence of F-actin and β-tubulin staining in DP thymocytes bearing single nucleus. (G) Proliferation rate of DP thymocytes cultured with anti-CD3/-CD28 antibodies. Fold growth relative to WT cells cultured without anti-CD3/-CD28 is shown. (H) Western blot analysis of TCR signaling events in DP thymocytes. The results are from samples pooled from at least 3 mice of same genotypes and represent two independent experiments. For A and B, n=3 mice per genotype and the results are from a representative experiment of two (A) or three (B) independent experiments. For C, n=4 WT and 6 RhoA^−/− mice. For G, n=3 mice per genotype. *P <0.05; *P <0.01. Error bars represent SD.

Altered mitochondrial function in RhoA-deficient thymocytes

Cell survival and proliferation are regulated by mitochondrial energy metabolism (41, 42). In thymocytes, it has been indicated that Notch promotes DN cell survival by regulating glucose metabolism (33). Because RhoA deficiency caused defects in thymocyte survival and proliferation, we hypothesized that depletion of RhoA may affect mitochondrial function. To substantiate this hypothesis, we first measured OCR, an indicator of oxidative phosphorylation (OXPHOS), and ECAR, an indicator of aerobic glycolysis, by Seahorse XF Cell Mito Stress Test and XF Glycolysis Test, respectively. To our surprise, RhoA-deficient thymocytes showed an increase in both basal and maximal, electron transport chain (ETC) accelerator FCCP-induced respiration (Fig. 6A), which coincided with an upregulation of a number of mitochondrial genes involved in OXPHOS, including Nrf1, Atp5I, Ndufa2, and Cox5a (Fig. 6B) (43). Consistent with the increased OXPHOS, RhoA^−/− thymocytes exhibited higher ECAR/glycolysis and increased expression of Hk2, Slc2a, Pdk1, and Pgm1, all of which have been shown to be important for glycolysis (Fig. 6C, D) (43). We subsequently measured mitochondrion mass and mitochondrial membrane potential by flow cytometry with Mito Tracker Green and DILC5, respectively. RhoA deficiency led to an increase in mitochondrion contents and mitochondrial membrane potential in all thymocyte subsets (DN, DP, CD4⁺ SP and CD8⁺ SP) (Fig. 6E, F). Further, mitochondrial DNA copy number was found increased by more than 200-fold in the absence of RhoA (Fig. 6G). However, ATP production in RhoA-deficient thymocytes was reduced (Fig. 6H).

Figure 6. RhoA deficiency causes an enhanced mitochondrial function in thymocytes.

(A) Oxygen consumption rate (OCR) in total thymocytes under basal condition and in response to oligomycin and FCCP. (B) mRNA expression of genes involved in oxidative phosphorylation in total thymocytes. The data are presented as fold expression relative to one WT sample. (C) Extracellular acidification rate (ECAR) in total thymocytes under basal condition and in response to glucose, oligomycin and 2-DG. (D) mRNA expression of genes involved in glycolysis in total thymocytes. The data are presented as fold expression relative to one WT sample. (E) Quantification of mitochondrion numbers/mass by flow cytometry analysis of Mitotracker Green staining in total thymocytes and thymocyte subsets. Mean fluorescence intensity (MFI) is shown. (F) Mitochondrial membrane potential assayed by flow cytometry of DilC-5 staining in total thymocytes and thymocyte subsets. MFI is shown. (G) Mitochondrial DNA contents in total thymocytes. Mitochondrial DNA was represented by mitochondrial Cox 2 normalized to nuclear β-globin. The data are presented as mitochondrial DNA relative to one WT mouse. (H) ATP levels in total thymocytes. In A-D, the results are averaged from three samples per genotype with each sample pooled from at least 3 mice of same genotypes. In E-H, n=5 mice per genotype. *P <0.05; **P <0.01. Error bars represent SD.

Because mitochondrial oxidation may generate reactive oxygen species (ROS) to suppress ATP production (44, 45), we reasoned that the decline in ATP production in RhoA^−/− thymocytes may be due to the increased ROS. Indeed, flow cytometry analysis of ROS levels found that consistent with the increased OXPHOS, ROS were significantly elevated in all RhoA^−/− thymocyte subsets (Fig. 7A). ROS can be deleterious to cells (42, 44). To test whether the increased ROS are responsible for the defective thymocyte development in RhoA^−/− mice, we treated the mice with antioxidant N-acetylcysteine (NAC) to scavenge ROS. NAC repressed ROS production in RhoA^−/− thymocyte subsets to the comparable levels to that in NAC-treated WT thymocytes (Fig. 7B). The increase in cell apoptosis in RhoA^−/− thymocytes was partially reversed by NAC treatment (Fig. 7C). NAC also partially reversed the increase in the frequency of DN thymocytes and the decrease in the frequency of DP thymocytes in RhoA^−/− mice (Fig. 7D). In addition, the hypocellularity in total thymocytes and thymocyte subpopulations in RhoA^−/− mice was partially rescued by NAC treatment (Fig. 7E). Together, these data suggest that RhoA represses mitochondrial metabolism and ROS production, contributing to its regulation of thymocyte development.

Figure 7. RhoA promotes thymocyte development partially through repressing ROS hyper-production.

(A) RhoA^−/− thymocytes show increased levels of ROS revealed by flow cytometry analysis of DCFDA staining in total thymocytes and thymocytes subsets. Mean fluorescence intensity (MFI) is shown. (B) ROS scavenger NAC treatment reverses the ROS levels in RhoA^−/− thymocytes. WT and RhoA^−/− mice were injected i.p. with 15 µg/g of body weight NAC every other day for 14 days. Ten days after last NAC injection, the mice will sacrificed and thymocytes were harvested, stained with CD4, CD8 and DCFDA, and analyzed by flow cytometry. (C) NAC treatment partially rescues cell apoptosis in RhoA^−/− thymocytes. Mice were treated with NAC as described in B. Apoptosis in thymocyte subsets was assayed by flow cytometry analysis of Annexin V⁺ cells. (D, E) NAC treatment partially reverses the changes in frequency of DN and DP thymocytes (D) and partially rescues thymocyte hypocellularity (E) in RhoA^−/− mice. Mice were treated with NAC as described in B. The frequency and numbers of thymocyte subsets were revealed by flow cytometry analysis. In A, n=5 mice per genotype. In B-E, n=3 mice per genotype. The results from a representative experiment of two independent experiments are shown. *P <0.05; **P <0.01. Error bars represent SD.

Discussion

In this study, we report that RhoA is essential for thymocyte development, with RhoA deletion leading to multiple defects during thymopoiesis. RhoA regulates a complex mechanism governing β-selection, positive selection, cell survival and proliferation, and pre-TCR and TCR signaling. Importantly, RhoA integrates energy metabolism in this process.

During thymocyte development, pre-TCR checkpoint is crucial for DN cell transition to DP cells (26). DN cells that successfully express pre-TCR are prevented from programmed cell death and are committed to proliferate and differentiate to DP cells (26, 33). We show that RhoA^−/− DN3 cells expressed reduced intracellular and cell surface TCRβ. Consistent with this, we found fewer IcTCRβ⁺ DN4 cells in RhoA^−/− mice. RhoA^−/− DN cells also had less preTα expression. These data suggest that RhoA is crucial for pre-TCR expression. By expressing transgenic TCR to restore pre-TCR expression in RhoA^−/− DN cells, we demonstrate that RhoA-regulated pre-TCR expression contributes to DN cell survival. Furthermore, because DN4 cells that do not express TCRβ are usually eliminated by apoptosis, the increased apoptosis in RhoA^−/− DN4 cells likely reflects the increased frequency of IcTCRβ⁻ DN4 cells in the mutant mice.

Survival and proliferation of thymocytes at the pre-TCR checkpoint are not only regulated by pre-TCR, but also by IL-7R and Notch (26, 46). The expressions of IL-7R and its signaling transducer Bcl2 remained unchanged in RhoA-deficient DN cells, whereas Notch1 and a few Notch1 targets were upregulated in the absence of RhoA. The intact IL-7R signaling may reflect an overlapping role of RhoA with the closely related RhoB and/or RhoC, while the increased Notch signaling may be a compensatory effect of the increased apoptosis. However, we cannot exclude the possibility that the elevated Notch signaling is intrinsic to RhoA deletion, i.e. RhoA directly negatively regulates Notch signaling. Indeed, Rac1/Rac2 and Cdc42 of the Rho GTPase family have been shown to inhibit Notch signaling during mouse T cell development and Drosophila wing growth/ development, respectively (26).

It is interesting that RhoA regulates proliferation of DP and SP cells through different mechanisms. RhoA deficiency resulted in a reduced BrdU incorporation in CD4⁺ SP and CD8⁺ SP thymocytes. In contrast, the levels of BrdU incorporation in RhoA^−/− DP thymocytes were indistinguishable to that in WT control cells. Nonetheless, RhoA^−/− DP thymocytes contained more multinucleated cells compared to WT counterparts. Therefore, RhoA appears to promote mitosis in DP thymocytes, but DNA synthesis in CD4⁺ SP and CD8⁺ SP thymocytes. The role of RhoA in mitosis of DP thymocytes seems to be broadly conserved because RhoA is also involved in mitosis of a number of other cells including keratinocytes, mouse embryonic fibroblasts, and hematopoietic progenitor cells (25, 37, 38).

It has been shown that inactivation of RhoA in C3 transferase transgenic mice blocked thymocyte development (18, 19). However, compared to the C3 mice that showed no survival defect in DN4 cells (19), RhoA^−/− mice exhibited increased apoptosis in this population. Moreover, whereas C3 mice showed normal positive selection (18), RhoA^−/− mice were impaired at this checkpoint. These differences are likely due to the nonspecific nature of C3 transferase which inactivates RhoA as well as RhoB and RhoC (24).

Metabolic pathways have been appreciated to be essential for naïve T cell activation and effector T cell differentiation (43, 45, 47, 48). However, it remains poorly understood whether and how metabolic programs are regulated in thymocyte development. An intriguing finding from our study is that RhoA appears to serve as a coordinator of mitochondrial metabolic programs with thymocytes development. RhoA deficiency in thymocytes resulted in an increased mitochondrial function. Although it suggests that RhoA represses mitochondrial metabolic programs, we cannot rule out the possibility that the enhanced mitochondrial function in RhoA^−/− thymocytes reflects, at least in part, a compensatory effect of the increase apoptosis.

It is known that mitochondria are a major source of ROS, where they are produced as byproducts of OXPHOS (42, 49). In line with the increased OXPHOS, ROS production was elevated in RhoA-deficient thymocytes. Excessive ROS can lead to broad cellular damage. For example, sustained ROS inhibit peripheral T cell proliferation and induce apoptosis in thymocytes (50, 51). We therefore reasoned that the increased ROS levels in RhoA-deficient thymocytes might contribute to the increased apoptosis and dampened thymocyte development. To test this, we treated RhoA^−/− mice with ROS scavenger NAC and found that removal of excessive ROS by NAC partially restored cell survival and the frequency and numbers of thymocytes. These results suggest that RhoA regulates thymocyte survival and development not only through governing pre-TCR expression, but also oxidative stress. Because a low level of ROS can act as signaling intermediates that in general are beneficial to cells (50, 52), our findings indicate for the first time that a fine-tuned ROS production regulated by RhoA through a delicate control of metabolic program, is important for thymocyte development. In sum, our study demonstrates that RhoA coordinates multiple developmental events to regulate thymopoiesis, providing a missing link between mitochondrial metabolism and thymocyte development.