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The mammalian target of rapamycin (mTOR) is a signaling molecule that senses environmental cues, such as nutrient status and oxygen supply, to regulate cell growth, proliferation, and other functions. Unchecked, sustained mTOR activity results in defects in HSC function. Inflammatory conditions, such as autoimmune disease, are often associated with defective hematopoiesis. Here, we investigated whether hyperactivation of mTOR in HSCs contributes to hematopoietic defects in autoimmunity and inflammation. We found that in mice deficient in Foxp3 (scurfy mice), a model of autoimmunity, the development of autoimmune disease correlated with progressive bone marrow loss and impaired regenerative capacity of HSCs in competitive bone marrow transplantation. Similarly, LPS-mediated inflammation in C57BL/6 mice led to massive bone marrow cell death and impaired HSC function. Importantly, treatment with rapamycin in both models corrected bone marrow hypocellularity and partially restored hematopoietic activity. In cultured mouse bone marrow cells, treatment with either of the inflammatory cytokines IL-6 or TNF-α was sufficient to activate mTOR, while preventing mTOR activation in vivo required simultaneous inhibition of CCL2, IL-6, and TNF-α. These data strongly suggest that mTOR activation in HSCs by inflammatory cytokines underlies defective hematopoiesis in autoimmune disease and inflammation.
Mammalian target of rapamycin (mTOR) has emerged as a central regulator for cellular response to environmental cues, such as nutrition, growth factors, and oxygen supplies (1, 2). The potential involvement of mTOR in HSC function was first suggested by the observation that targeted mutation of Pten, which is a distant upstream negative regulator of mTOR, resulted in a loss of HSC function (3, 4). mTOR is implicated in Pten deficiency–mediated HSC defect, as the defects are reversed by rapamycin (3). Our recent study demonstrated that mTOR hyperactivation abrogates quiescence and function of HSCs by increasing ROS levels (5). More recently, we reported that rapamycin rejuvenates HSCs in and increases lifespan of old mice (6).
Although the consequences of mTOR activation in HSC function are now well established, the pathophysiological conditions that lead to mTOR activation in HSCs remain to be identified. In particular, it is worth considering the possibility that innate or adaptive immune activation may lead to mTOR activation in HSCs. For instance, infectious diseases, such as viral hepatitis, have long been associated with HSC defects (7). In addition, leukocytopenia is an important manifestation of systemic lupus erythematosus (8), although an HSC defect has yet to be established. These data raised an interesting issue as to whether autoimmune diseases and inflammation may cause HSC defects. Moreover, given the impact of mTOR in HSC function, it is intriguing that mTOR activation in HSCs may be responsible for the defective hematopoiesis in both autoimmune diseases and inflammation. Here we use models of autoimmune diseases and endotoxin-induced systemic inflammation to test this hypothesis.
The scurfy mice have severe autoimmune diseases and pancytopenia due to a spontaneous mutation of the forkhead box P3 (Foxp3) gene (9, 10). They therefore serve as a valuable model to determine whether and how autoimmune disease causes defective hematopoiesis. We first evaluated their bone marrow cellularity in relation to disease progression. While the 1-week-old scurfy mice that show no sign of autoimmune diseases had normal bone marrow cellularity, a progressive loss of bone marrow cells was observed in the subsequent 3 weeks when autoimmune diseases became severe. The reduction of cellularity was obvious, even after normalization of body weight (Figure (Figure1A).1A). Surprisingly, based on the stringent HSC markers, Flk2–lin–Sca-1+c-kit+CD150+CD48–CD34– (11, 12), we observed a significant increase in the number of the HSCs in the scurfy mice at 3 weeks. This increase was transient, as the number of HSCs was lower in the 4-week-old scurfy mice (Figure (Figure1,1, B and C). To understand the cellular basis for the increased HSC numbers at 3 weeks, we labeled scurfy mice and their WT littermates with BrdU. At 24 hours after BrdU injection, both scurfy and WT mice had about 60% of bone marrow cells labeled with BrdU. During the same period, about 50% of Lin–Sca1+ckit+ (LSK) cells and 40% of HSCs in WT mice were BrdU+. The considerably higher levels of cycling HSCs in young mice, in comparison to what was described in adult mice by others (3) and us (5), are consistent with a previous analysis of HSC cycles (13). In the scurfy mice, about 80% of LSK cells and HSCs were BrdU+ (Figure (Figure1C).1C). Despite an enlargement of the stem cell compartment at 3 weeks, in vitro colony formation unit (CFU) analysis revealed substantial reductions in the progenitor cell activities for all lineages of blood cells (Supplemental Figure 1; supplemental material available online with this article; doi: 10.1172/JCI43873DS1).
To determine the regenerative capacity of HSCs in scurfy mice, we carried out competitive bone marrow transplantation. We transplanted 5 × 105 bone marrow cells from scurfy or control mice, in conjunction with an equal number of recipient-type bone marrow cells, into lethally irradiated CD45.1 C57BL/6 recipients (Figure (Figure1D).1D). At given time points after transplantation, the percentage of donor-type cells was analyzed using the CD45.2 congenic markers (Figure (Figure1E).1E). As shown in Figure Figure1F,1F, bone marrow cells from 7-day-old scurfy mice had a comparable reconstitution capacity as those of control mice. However, despite a significant increase of HSC frequency, bone marrow from 3-week-old scurfy mice exhibited significantly reduced HSC activity. The defects were more pronounced in bone marrow from 4-week-old scurfy mice. Defects in production of both B cells and myeloid cells were pronounced at both 4 and 12 weeks after transplantation (Figure (Figure1G).1G). At 12 weeks, it is clear that scurfy bone marrow was substantially defective in T cell reconstitution, although the impact cannot be evaluated at 4 weeks, due to slow T cell reconstitution. Thus, the HSC defects were acquired in the scurfy mice and were progressively associated with the development of autoimmune diseases. Since the defective HSC function was observed in the scurfy mice even when the HSC compartment was enlarged (Figure (Figure1F,1F, middle panel), the defective hematopoiesis was not due to physical elimination of HSCs by the autoreactive T cells or by relocation of HSCs to other compartments. This contention is supported by the fact that, despite the presence of T cells in the bone marrow from the scurfy mice, the cotransplanted recipient-type HSCs were not destroyed (Figure (Figure1,1, E–G). Since Foxp3 is not expressed in HSCs (Supplemental Figure 2), the HSC defect is unlikely a direct consequence of Foxp3 mutation. Since the Sca-1 is an activation marker of bone marrow cells (14), we checked whether the increased HSCs in the scurfy mice at 3 weeks merely reflected more activation in the bone marrow cells. As shown in Supplemental Figure 3, the increase in HSC number in the bone marrow was largely unaffected when Sca-1 was dropped as part of the HSC markers.
To characterize the reduction of stem cells and progenitor numbers in 4-week-old scurfy bone marrow, we compared the percentage and number of short-term HSCs (ST-HSCs), Flk2–lin–Sca1+ckit+ (FLSK) cells, multipotent progenitors (MPPs), common lymphoid progenitors (CLPs), and myeloid progenitors (MPs) in the bone marrow and HSCs and MPPs in the spleen. As shown in Figure Figure2,2, A–C, and Supplemental Figure 4, a reduction of HSCs was associated with an increase of ST-HSCs. The numbers of FLSK cells, MPPs, CLPs, and MPs were not increased in the bone marrow. Significant increases of FLSK cells and HSCs were observed in the spleen (Figure (Figure2,2, D and E). Therefore, both increased mobilization and alteration of differentiation of HSCs likely contributed to the reduced HSCs and progenitors in the 4-week-old bone marrow.
We then considered the possibility that the innate immune response may cause HSC defects. To test this hypothesis, we tested whether the broad hematopoietic defects can be induced by LPS, a prototype pathogen-associated molecular pattern (PAMP) that interacts with TLR4 and triggers inflammatory response (15). As shown in Figure Figure3A,3A, we injected C57BL/6 mice with lethal doses of LPS and analyzed the complete blood cell count (CBC), bone marrow cellularity, and HSC function. Significant reductions of all lineages of blood cells were observed at 1 or 10 weeks after LPS treatment (Figure (Figure3B).3B). In addition, a massive reduction of bone marrow cellularity was observed after LPS treatment (Figure (Figure3C3C and Supplemental Figure 5). This reduction was due to the massive cell death of bone marrow cells in the LPS-treated mice (Figure (Figure3D). 3D).
We took 2 approaches to determine whether LPS induced HSC defects. First, we mixed bone marrow or HSCs from either vehicle- (PBS-) or LPS-treated mice with recipient-type bone marrow. Although the HSCs in the PBS group were as efficient as the recipient-type bone marrow in hematopoiesis, those from the LPS group were much less functional (Figure (Figure3E).3E). The defects were observed in multiple lineages and persisted over the 15 weeks studied (Figure (Figure3F).3F). Since the defects were long lasting and occurred in T, B, and myeloid cells, they likely reflect a defective HSC function. Second, to directly demonstrate the HSC defects, we used 50 FACS-sorted HSCs from the 2 groups to compete with 105 recipient-type bone marrow cells. As shown in Figure Figure3G,3G, the purified HSCs from the LPS-treated mice were significantly less potent in hematopoiesis. Again, the defects were manifested in the numbers of total leukocytes as well as T, B, and myeloid cells.
LPS may either directly inactivate HSCs or do so by inducing inflammatory cytokines. As shown in Figure Figure4A,4A, high levels of IL-6, TNF-α, or CCL2 were found 2 hours after LPS treatment. In contrast, no induction of IFN-γ, IL-10, and IL12p40 was observed. The elevation was not long lasting, as the cytokine levels at 72 hours after LPS treatment largely returned to baseline. To address the potential role for the inflammatory cytokines, we used antibodies to block the effect of IL-6 and TNF-α. In addition, we used mice with a targeted mutation of the Ccr2 gene that encodes the dominant CCL2 receptor to test the impact of CCL2 (16). Because blocking individual cytokines had no appreciable effect in preventing bone marrow hypocellularity (Supplemental Figure 5), we treated Ccr2-deficient mice with a combination of anti–IL-6 and anti–TNF-α mAbs in order to block all 3 cytokines simultaneously (Figure (Figure4B).4B). This combination largely reversed the massive loss of bone marrow cellularity (Figure (Figure4C).4C). Furthermore, LPS treatment caused a large increase in the number of cells with HSC phenotypes on day 3 (Figure (Figure4D).4D). This increase was not due to increased bone marrow activation marker Sca-1 (Supplemental Figure 6). However, most of the HSCs had undergone apoptosis, as revealed by their staining to Annexin V and permeability to the nuclear dye DAPI (Figure (Figure4E).4E). By day 7, the number of HSCs in the LPS-treated mice was lower than that in the PBS-treated mice (Figure (Figure4D).4D). The rise and fall of HSC numbers was largely eliminated in the anti–TNF-α/IL-6–treated Ccr2-deficient mice (Figure (Figure4D).4D). Likewise, apoptosis of HSCs was abrogated by blocking the 3 cytokines (Figure (Figure4E). 4E).
To test the function of HSCs, we carried out competitive bone marrow transplantation (Figure (Figure4B).4B). As shown in Figure Figure4F,4F, blocking the 3 cytokines reversed the defects in bone marrow cells from the LPS-treated mice. The impact was observed at all time points tested (Figure (Figure4F)4F) and found in T and B cell lineages (Supplemental Figure 7).
Given our previous studies on the impact of mTOR activation on HSC function (5, 6), we wondered whether the inflammatory cytokines may inactivate HSCs by stimulating mTOR. To test this possibility, we treated cKit+ bone marrow cells from WT mice with either IL-6 or TNF-α. The dose used (5 ng/ml) was no higher than what was observed in the LPS-treated mice (Figure (Figure3A).3A). As shown in Figure Figure5A,5A, treatment for 30 minutes with either cytokine substantially induced phosphorylation of S6 kinase (S6K), a downstream target of mTOR. To confirm that mTOR activation occurred in HSCs, we analyzed the pS6 levels in the LSK cells and HSCs using flow cytometry. As shown in Figure Figure5B,5B, both cytokines induced mTOR activation in both LSK cells and HSCs. However, the percentage of pS6+ cells was higher in HSCs than that in LSK cells. These data demonstrated that either IL-6 or TNF-α was sufficient to activate mTOR in HSCs.
To determine potential contribution of mTOR activation to HSC defects in LPS-treated mice, we tested whether mTOR activation in HSCs was induced by LPS in vivo. At 2 hours after LPS treatment, a substantial increase of pmTOR (Figure (Figure6,6, A and B) and pS6 (Figure (Figure6,6, C and D) was observed in total bone marrow cells, including LSK cells and HSCs. Based on mean fluorescence intensity, the increase in total bone marrow cells (<1-fold increase) was substantially less than that in either LSK cells or HSCs (2- to 3-fold increase in pmTOR and 7- to 8-fold increase in pS6). Importantly, the inflammatory cytokines IL-6, TNF-α, and CCL2 played a critical role in mTOR activation, as the induction of pS6 was abrogated when the 3 cytokines were simultaneously blocked (Figure (Figure6,6, E and F).
To determine the role for mTOR activation in LPS-induced hematopoietic defects, we injected rapamycin in conjunction with LPS, starting 1 day prior to LPS treatment (Figure (Figure7A).7A). Although this treatment did not reduce the production of IL-6, TNF-α, and CCL2 (Figure (Figure7B),7B), rapamycin essentially abrogated the LPS-induced bone marrow hypocellularity (Figure (Figure7C).7C). Corresponding to restoration of cellularity, rapamycin also prevented the massive apoptosis in bone marrow cells (Figure (Figure6D).6D). We then used competitive bone marrow transplantation to determine whether rapamycin restored the hematopoietic activity of bone marrow from LPS-treated mice (Figure (Figure7A).7A). As shown in Figure Figure7E,7E, rapamycin partially abrogated the inhibition of hematopoietic activity by LPS, as measured by either total leukocytes or B cells at either 4 or 10 weeks after transplantation. Since T cell development is usually slower, the effect of rapamycin was observed at 10 weeks only. Although a significant effect was observed for myeloid lineage at 4 weeks, the trend observed at 10 weeks is not statistically significant.
Since the levels of inflammatory cytokines were also very high in the scurfy mice (Supplemental Figure 8), we tested whether the HSC defects in the mice described in Figure Figure11 were also due to the hyperactive mTOR pathway. Again, we determined the mTOR status in the HSCs using flow cytometric analysis of phosphorylation of mTOR and S6. As shown in Figure Figure8A8A and Supplemental Figure 9, the mean fluorescence intensities of both pmTOR and pS6 were significantly increased in the scurfy HSCs in comparison with those of the WT HSCs. Consistent with our previous observation, mTOR activation was associated with increased expression of senescence marker gene cyclin-dependent kinase inhibitor 2A (p16ink4a) (Supplemental Figure 10).
In order to determine whether mTOR activation is the underlying cause of the HSC defects in the scurfy mice, we treated 2-week-old scurfy mice with rapamycin at a daily dose of 4 mg/kg for 7 days and tested the effect of rapamycin on lifespan and hematopoiesis of the scurfy mice as well as the function of HSCs in competitive bone marrow transplantation (Figure (Figure8B).8B). As shown in Figure Figure8C,8C, 1 week of rapamycin treatment significantly extended the lifespan of scurfy mice, which is consistent with a report involving 2 immunodeficiency polyendocrinopathy, enteropathy, X-linked syndrome (IPEX) patients (17). However, this treatment inhibited neither the production of inflammatory cytokines (Supplemental Figure 8A) nor lymphoproliferation as manifested by splenomegaly (Supplemental Figure 8B) and the adenopathy (our unpublished observations). Therefore, a therapeutic benefit can be achieved without overt inhibition of inflammation and lymphoproliferation.
At 2 weeks after the treatments, the bone marrow cells were isolated and tested for HSC function. As shown in Figure Figure8D,8D, rapamycin treatment significantly increased the bone marrow cellularity. In addition to the total numbers of leukocytes, a remarkable reduction of B cells was observed in the scurfy bone marrow. This is also substantially rescued by rapamycin treatment (Figure (Figure8,8, E and F). Further, the production of immature (Ter119+CD71+) and mature (Ter119+CD71–) erythroblasts was also restored by rapamycin treatment. To determine whether rapamycin restored the HSC function in the scurfy mice, the bone marrow of vehicle- or rapamycin-treated mice was mixed with an equal number of recipient-type bone marrow cells and transplanted into lethally irradiated CD45.1 recipients. The PBLs were analyzed at 3 months after bone marrow transplantation. As shown in Figure Figure8G,8G, the frequency of donor-type nucleated blood cells increased by about 6 fold in the rapamycin group. Substantial increases were found in all cell lineages tested, including T, B lymphocytes, and myeloid cells (Supplemental Figure 11). Therefore, a short-term rapamycin treatment restored the long-term HSC function of the scurfy mice in the new hosts.
It is well established that hematological abnormalities are associated with infections (18), inflammation (19), and autoimmune diseases (20). However, whether and how these pathophysiological conditions affect HSC function remain unresolved. Here, we used models for both autoimmune diseases and inflammation to demonstrate 3 intrinsic links among HSC defects and these conditions.
First, our data demonstrate that autoimmune diseases cause HSC defects. Traditionally, immunologists have focused on both innate and adaptive responses of mature leukocytes in autoimmune diseases. How the stem cells may respond in this setting has not been systematically studied. Here, we demonstrated that scurfy mice, which have spontaneous inactivation of Foxp3+ Tregs and fatal autoimmune diseases, exhibit significant HSC defects when tested in competitive bone marrow transplantation. Since Foxp3 is not expressed in HSCs (Supplemental Figure 2), it is likely that the HSC defects are secondary to autoimmune diseases.
Because of the increased burden of autoreactive T cells or autoantibodies in the scurfy mice (10, 21, 22), it may be tempting to suggest that HSCs were destroyed by self-reactive T cells or autoantibodies. However, since recipient-type bone marrow cells functioned normally when cotransplanted into irradiated host with scurfy bone marrow cells, it is unlikely either antibody-producing cells or autoreactive T cells in the scurfy bone marrow can prevent the function of normal HSCs. Furthermore, at 3 weeks, when the number of HSCs in the bone marrow was at least 10-fold higher in the scurfy mice than in WT mice, the scurfy bone marrow cells showed significant defects in hematopoiesis in transplantation assay. Therefore, hematopoietic defects are not due to physical elimination of HSCs. It is more plausible that the environment in the autoimmune mice causes functional inactivation of HSCs.
The causative relationship between autoimmune diseases and HSC defects demonstrated herein may explain frequent leucopenia in systemic lupus erythematosus (8). Moreover, AA, which is a well-established HSC disease, is believed to be due to autoimmune diseases, because the majority of the patients respond to immune suppression (23–25). It is of note that given the potential contribution of homeostatic proliferation to autoimmune diseases (26, 27), defective hematopoiesis and autoimmune diseases may form a vicious cycle to the detriment of the host. As such, breaking this cycle may have significant therapeutic implications for both autoimmune diseases and hematological defects.
Second, we demonstrated a direct link between inflammatory cytokines and HSC defects. A potential link between inflammation and HSC function has not been established. A recent study showed that LPS from Pseudomonas aeruginosa caused a block in generation of common progenitors for myeloid cells. Moreover, a germline mutation of TLR4 prevented common MP defects in LPS-treated mice (28). A different regiment of LPS treatment has lead to defects in CLPs (29). Here, we have shown that LPS caused long-lasting HSC defects by inducing IL-6, TNF-α, and Ccl2. As a consequence, reconstitutions of both the lymphoid and myeloid compartments are adversely affected.
Both infection and tissue injuries trigger production of inflammatory cytokines through recognition of either PAMPs or danger-associated molecular patterns (DAMPs) (30, 31). Host responses to DAMPs are selectively regulated by CD24-Siglec G/10 interaction (32, 33). The abnormal production of PAMPs or DAMPs or interruption of CD24-Siglec G/10 interaction may all create a proinflammatory environment. Therefore, the link between inflammatory cytokines and HSC defects, as uncovered here, may have broad significance on a variety of pathophysiological conditions. Since mTOR activation can be induced by a multitude of inflammatory cytokines, and since the increase of inflammatory cytokines is found in both experimental models, it is possible that these pathological conditions involve similar mechanisms. However, due to higher levels and the more transient nature of inflammatory cytokines, the effect of LPS is more acute.
Last, but not the least, our data demonstrated that both inflammation and autoimmune diseases induce HSC defects through mTOR activation. In both models, we observed a transient expansion and decline in HSC number, which is reminiscent of the observations made in HSCs with mutations of Pten and Tsc1, 2 upstream negative regulators of mTOR (3–5). The functional defects in HSCs are also reminiscent of the Tsc- and Pten-deficient HSCs, although the phenotype in the Pten-deficient HSCs is complicated by development of malignancy and genetic instability (3–5). The critical role of mTOR activation in HSC defects is confirmed by the effect of rapamycin. However, since all experimental conditions activate or inactivate mTOR in other cell types, one may envisage that rapamycin inhibits production of inflammatory cytokines by other cells, which in turn inactivates HSCs by mTOR-independent mechanisms. Therefore, it is formally possible that the HSC defects are due to HSC-extrinsic activation of mTOR. We consider this very unlikely, as our data indicated that rapamycin has no effect on the production of inflammatory cytokines in these settings.
We have recently reported that mTOR activation is the underlying cause of HSC senescence in the old mice (6). The stimuli that cause mTOR activation in the old mice have not been identified. Given the increased levels of inflammatory cytokines in the elderly, it is of interest to consider inflammation as a cause of HSC aging. In this regard, it is of interest that HSCs from autoimmune mice also exhibit higher levels of senescence marker p16INK4a. Taken together, our data suggest that mTOR activation underlies HSC defects associated with infection, inflammation, autoimmune diseases, and aging. This conclusion may have interesting implications for the treatment of hematological abnormalities.
C57BL/6 Ly5.1 (CD45.2) C57BL/6 Ly5.2 (CD45.1) mice were purchased from the National Cancer Institute. The scurfy mice were obtained from The Jackson Laboratory. The Ccr2–/– mice (16) were obtained from the University of Michigan Animal Core. FoxP3EGFP mice that express both FoxP3 and EGFP under the endogenous regulatory sequence of the FoxP3 locus have been described previously (34) and have been provided by T.A. Chatila (Department of Pediatrics, UCLA, Los Angeles, California, USA). All the mice were kept in the Unit of Laboratory Animal Facility at University of Michigan. All procedures involving experimental animals were approved by the University Committee on the Use and Care of Animals at the University of Michigan.
LPS from Escherichia coli 055:B5 (Sigma-Aldrich) was resuspended in 1x PBS at 2 mg/ml. Mice were treated with 0.3 mg once or twice, as indicated, by intraperitoneal injection. Rapamycin (Sigma-Aldrich) was reconstituted in absolute ethanol at 10 mg/ml and diluted in 5% Tween-80 (Sigma-Aldrich) and 5% PEG-400 (Hampton Research). Mice received 4 mg/kg rapamycin by intraperitoneal injection every other day (5). Control IgG or anti–TNF-α (35) and anti–IL-6 mAb (LS-C7955, LifeSpan Bio.) were injected intraperitoneally (100 μg/mouse) on days 0 and 3 of LPS injection.
BrdU (Sigma-Aldrich) was injected intraperitoneally into adult mice at 100 mg/kg body weight. Mice were then given 1 mg/ml BrdU in the drinking water for 24 hours before sacrifice for analysis. The BrdU staining kit (BD Biosciences) was used according to the manufacturer’s instructions.
Eight-week-old congenic recipient mice were lethally irradiated with a Cs-137 x-ray source, delivering 0.97 Gy per minute, for a combined 11.5 Gy, that were delivered 4 hours apart. Bone marrow cells of the donor type were mixed with competitive recipient-type bone marrow cells and were then transplanted into recipients by injection through the retro-orbital venous sinus. Reconstitutions were measured using flow cytometry of blood from the tail vein at the time points indicated. The red blood cells were lysed using ammonium chloride/potassium bicarbonate buffer before staining.
Bone marrow cells were flushed out from the long bones (tibias and femurs) by using a 25-gauge needle with 1x HBSS, without calcium or magnesium (Invitrogen), supplemented with 2% heat-inactivated fetal bovine serum. For flow cytometry and purification of HSCs, the immunophenotype Flk2–lin–Sca-1+c-kit+CD34–CD150+CD48- was used, and the LSK cell population was marked as lin–Sca-1+c-kit+. Lineage markers included B220, CD3, Gr-1, Mac-1, and Ter119. The anti-CD150 antibody was purchased from BioLegend, and all other antibodies were obtained from BD Biosciences. For intracellular staining, cells were first stained with indicated surface markers, then fixed with Fix buffer (BD Biosciences) for 2 hours at 4°C, followed by permeate buffer for 10 minutes at room temperature, and then refixed for 10 minutes. pmTOR antibodies (Cell Signaling Technology) were diluted at 1:100 and Alexa Fluor 488–conjugated pS6 antibodies (Cell Signaling Technology) and FITC-conjugated p16Ink4a antibodies (Santa Cruz Biotechnology Inc.) were diluted at 1:10 and incubated overnight at 4°C. FITC-conjugated secondary antibodies (Jackson ImmunoResearch Laboratories Inc.) for pmTOR staining were diluted at 1:100 and incubated for 2 hours at 4°C. The Foxp3 Staining Kit (eBioscience) was used for Foxp3 staining. Flow cytometry analysis was performed on BD LSR II (BD Biosciences). FACS sorting was performed on BD FACSDiva (BD Biosciences).
LSK cells and HSCs were purified by FACS sorting, and RNA was isolated with TRIzol (Invitrogen) and further purified using the RNeasy Kit (Invitrogen). cDNA was made from purified RNA with SuperScript III (Invitrogen). Real-time PCR was performed on the 7500 Real-Time PCR System (Applied Biosystems). The sequences of RT-PCR primers for p16Ink4a were as follows: forward, 5′-TTCTTGGTGAAGTTCGTGCGATCC-3′, and reverse, 5′-TTGAGCAGAAGAGCTGCTACGTGA-3′.
Total bone marrow cells isolated from 21-day-old WT or scurfy mice were plated into MethoCult GFM3434 (StemCell Technologies Inc.) and cultured for 12 days before being counted under a microscope (CKX31; Olympus).
c-Kit–positive cells were isolated from C57BL/6 mice bone marrows with biotin-conjugated anti–c-Kit antibody (BD Biosciences) and MACS separation column. The c-Kit–positive cells were lysed with lysis buffer (Cell Signaling) and analyzed using SDS-PAGE and Western blot with anti–phospho-S6K (Cell Signalling) and anti-actin (Abcam) antibodies.
A paired 2-tailed Student’s t test was used to compare the sorted bone marrow cells from WT or scurfy mice and to compare the peripheral blood cells or bone marrow cells from mice treated with LPS or PBS. P values of less than 0.05 were considered significant.
We thank Shenghui He and Linhua Vatan for technical help and Darla Kroft for critical reading of the manuscript. This study was supported by grants from National Institute of Health and US Department of Defense.
Conflict of interest: The authors have declared that no conflict of interest exists.
Citation for this article: J Clin Invest. 2010;120(11):4091–4101. doi:10.1172/JCI43873.
See the related Commentary beginning on page 3813.