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The proinflammatory cytokine tumor necrosis factor α (TNFα) inhibits hematopoietic stem cell (HSC) expansion, interferes with HSC self-renewal and compromises the ability of HSC to reconstitute hematopoiesis. We have investigated mechanisms by which TNFα suppresses hematopoiesis using the genomic instability syndrome Fanconi anemia mouse model deficient for the complementation-group-C gene (Fancc). Examination of senescence makers, such as senescence-associated β-galactosidase, HP1-γ, p53 and p16INK4A shows that TNFα induces premature senescence in bone marrow HSCs and progenitor cells as well as other tissues of Fancc–/– mice. TNFα-induced senescence correlates with the accumulation of reactive oxygen species (ROS) and oxidative DNA damage. Neutralization of TNFα or deletion of the TNF receptor in Fancc–/– mice (Fancc–/–;Tnfr1–/–) prevents excessive ROS production and hematopoietic senescence. Pretreatment of TNFα-injected Fancc–/– mice with a ROS scavenger significantly reduces oxidative base damage, DNA strand breaks and senescence. Furthermore, HSCs and progenitor cells from TNFα-treated Fancc–/– mice show increased chromosomal aberrations and have an impaired oxidative DNA-damage repair. These results indicate an intimate link between inflammatory reactive oxygen species and DNA-damage-induced premature senescence in HSCs and progenitor cells, which may play an important role in aging and anemia.
The genomic instability syndrome Fanconi anemia (FA) shares the common feature of having defective DNA damage response/repair processes with other premature aging syndromes such as Bloom (Bischof et al., 2001) and Werner (Chen et al., 2003), ataxia telangiectasia (Ito et al., 2004) and BRCA1-deficient breast cancer (Cao et al., 2003). The FA pathway has been shown to functionally interact with the proteins responsible for Bloom syndrome (BLM) (Meetei et al., 2003), ataxia telangiectasia (ATM) (Taniguchi et al., 2002)] and breast cancer (BRCA1) (Seal et al., 2003). In fact, one of the FA genes, FANCD1, turned out to be the breast cancer gene BRCA2 (Howlett et al., 2002). This suggests a biological link between the FA disease and organismal aging. The FA disease model is unique in that defects are especially profound in the rapidly proliferating hematopoietic system. Thus far, aging studies have not yet focused on hematopoietic stem cells (HSCs) in FA or these other premature aging syndromes. Nevertheless, HSCs are capable of self-renewing, have a high risk of accumulating mutations and, thus, may require fewer events to become senescent or transformed (Pelicci, 2004). In addition, HSC aging can be assessed in a quantitative fashion with well-established assays. Since senescence can cause stem cell depletion, which can conceivably lead to organismal aging, such studies may yield important insights to the mechanisms of aging.
Functional failure of HSCs can result in the decrease in number and function of HSCs and progenitor cells leading to the development of anemia, neutropenia and thrombocytopenia, and has been implicated in hematologic diseases such as bone marrow (BM)-failure diseases such as FA, myelodysplastic syndromes (MDS) and aplastic anemia (Young, 2002; Bagby, Jr, 2003; Chen, 2005). Indeed, it has been shown that FA HSCs and progenitor cells have high rates of stress-induced apoptosis and reduced repopulating ability (Haneline et al., 1998; Haneline et al., 1999; Haneline et al., 2003). The FA proteins are thus believed to play important roles in the maintenance of hematopoiesis. Consistent with the observations that cells derived from FA patients are intolerant of oxidative stress, it has been reported that FA proteins, particularly the complementation group C (FANCC) protein, play a crucial role in oxidative-stress signaling in a variety of cell types including hematopoietic cells (Kruyt et al., 1998; Cumming et al., 2001; Hadjur et al., 2001; Futaki et al., 2002; Park et al., 2004; Saadatzadeh et al., 2004; Pagano et al., 2005). More recently, cytokine hypersensitivity of FA hematopoietic cells to apoptotic cues has also been proposed as a major factor in the pathogenesis of BM failure in three FA mouse models (Fanca–/–, Fancc–/– and Fancg–/–) (Li et al., 2004; Si et al., 2006).
It is believed that cellular and organismal senescence occurs as a result of chronic exposure to extrinsic environmental factors, primarily of oxidative stress (Sohal and Weindruch, 1996). The degree of oxidative damage has been found to increase exponentially with senescence and aging in a variety of cells, and tissues of different species (Chen et al., 1995). Endogenously formed reactive oxygen species (ROS) continuously damage cellular constituents including DNA. These challenges, coupled with exogenous exposure to agents that generate ROS, are both associated with normal aging processes and linked to cardiovascular disease, age-related anemia and cancer (Maccio et al., 2005; Sablina et al., 2005; Wallace, 2005). In the aged and certain disease states, ROS produced by proinflammatory cytokines – such as tumor necrosis factor-α (TNFα) – at inflammatory sites often cause DNA damage (Goossens et al., 1999; Suematsu et al., 2003; Wheelhouse et al., 2003). However, the mechanism through which ROS mediate their effects on the aging process is unclear.
Overproduction of the proinflammatory cytokine TNFα has been implicated in pathological conditions related to anemia and aging that is characteristic of HSC function failure (Oster et al., 1989; Kitagawa et al., 1997; Young, 2000; Dufour et al., 2003). Here, we have employed the disease model of genomic instability syndrome FA to investigate the cellular mechanism by which TNFα-induced inflammatory ROS influence hematopoiesis. We demonstrate that ROS induce premature senescence in BM cells enriched for HSCs and progenitor cells through prolonged oxidative DNA-damage response and increased genomic instability.
To examine the effect of TNFα on HSC/progenitor cell activity, we injected intraperitoneally (i.p.) wild-type (WT) or Fancc–/– mice with mouse recombinant TNFα in PBS or PBS alone. Seven days later, mice were sacrificed, and peripheral blood (PB) and BM were analyzed for hematopoietic functions. TNFα-treated Fancc–/– mice exhibited mild cytopenia, as evidenced by a decrease in red cell counts, hemoglobin and hematocrit values (see supplementary material Fig. S1). Consistent with this, BM analysis of TNFα-treated Fancc–/– mice revealed a decrease in the number of cells compared with PBS-injected Fancc–/– mice (see supplementary material Table S1). In another set of experiments we analyzed hematopoietic recovery in mice injected with TNFα. We found that Fancc–/– mice recovered slowly from TNFα-induced hemo-suppression compared with WT mice (see supplementary material Fig. S2). Furthermore, Fancc–/– mice showed reduced rates of multilineage recovery (see supplementary material Table S2).
The frequencies of BM HSC/progenitor [lineage-negative, Sca-1-positive, c-Kit-positive (Lin–-Sca-1+-c-Kit+)] cells (LSK cells) were reduced by more than twofold in TNFα-treated Fancc–/– mice compared with PBS controls (Fig. 1A). We thus asked whether TNFα suppressed clonogenic progenitor activity. As shown in Fig. 1B, the total number of colonies formed by LSK cells from TNFα-treated Fancc–/– mice was more than threefold lower than that of WT mice. We also observed significantly decreased series-plating efficiency with BM LSK cells of TNFα-treated Fancc–/– mice compared with WT cells (Fig. 1C). To evaluate the in vivo effect of TNFα on HSC/progenitor cell activity, we performed BM-stem-cell transplantation. Long-term engraftment analysis shows that Fancc–/– BM HSCs were decreased in their ability to repopulate compared with WT cells (Fig. 1D), which is consistent with previous reports (Haneline et al., 1998; Haneline et al., 1999). Significantly, BM HSCs from TNFα-treated Fancc–/– mice constituted less than 10% of the peripheral blood cells at 16 weeks after transplantation compared with more than 40% reconstitution by those from TNFα-treated WT mice (Fig. 1D), indicating that TNFα impaired long-term hematopoietic reconstitution of Fancc–/– HSCs. Collectively, these results suggest that TNFα suppresses hematopoietic functions by inhibiting the self-renewal or proliferative potential of HSCs and progenitor cells.
Given that TNFα suppressed the proliferative potential of HSCs and progenitor cells, we wondered whether TNFα affected differentiation or apoptosis of these primitive cells. Fig. 2A shows that BM HSCs from TNFα-injected WT or Fancc–/– donors had the ability of multilineage reconstitution, suggesting that TNFα does not affect the differentiation of long-term repopulating HSCs. Whereas TNFα induced apoptosis in BM LSK cells of treated mice, no statistically significant difference was observed between WT and Fancc–/– mice (Fig. 2B). Thus, apoptosis was not the major consequence of TNFα-mediated inhibitory effect on the self-renewal or proliferative potential of Fancc–/– HSCs and progenitor cells. However, BM LSK cells from TNFα-injected Fancc–/– mice clearly exhibited a prolonged G2-M accumulation with decreased cells in S phase (Fig. 2C), suggesting that the G2 checkpoint is compromised in these BM HSC/progenitor cells. This result reminisces of DNA-damage-induced G2-M arrest, a hallmark of FA (Bagby, Jr, 2003; Kennedy and D'Andrea, 2005).
It has been suggested that HSC senescence plays an important role in the pathophysiology of hematologic diseases, such as aplastic anemia and MDS (Maciejewski and Risitano, 2003; Zhang et al., 2005a; Zhang et al., 2005b). To determine whether HSC senescence is a defining feature of TNFα-mediated hematopoietic suppression, we examined two established senescence markers: senescence-associated β-galactosidase (SA-β-gal) (Dimri et al., 1995) and senescence-associated heterochromatin foci (Narita et al., 2003). We found that approximately 10% of LSK cells from TNFα-injected WT mice and more than 20% of those from TNFα-injected Fancc–/– animals stained positive for SA-β-gal activity (Fig. 3A). LSK cells from TNFα-treated Fancc–/– mice also stained strongly positive for HP1-γ, indicating the formation of senescence-associated heterochromatin foci (Fig. 3B) (Narita et al., 2003). In addition, analysis of these two senescence markers in BM sections from TNFα-injected Fancc–/– mice revealed abundant positive staining in those cells (Fig. 3C) accompanied with a weak proliferation index, as identified by the proliferation marker Ki-67 (Fig. 3D). We also examined two established effectors of stress-induced senescence, p53 and p16INK4A (Randle et al., 2001; Serrano and Blasco, 2001). We observed that the BM LSK cells of TNFα-injected Fancc–/– mice stained positive for p53 and p16INK4A, whereas those from untreated mice stained essentially negative (Fig. 3E). These results suggest that senescence is a defining feature of TNFα-mediated hematopoietic suppression in Fancc–/– mice.
Telomere shorting has been proposed to be a cause of HSC senescence (Lansdorp, 2005). The telomere length of BM cells from TNFα-injected mice was comparable to that of untreated mice, regardless of genotype (see supplementary material Fig. S3A). Overexpression of the mouse telomere reverse transcriptase (mTERT) in TNFα-treated Fancc–/– BM cells enriched for HSCs and progenitor (Lin–) cells failed to abrogate senescence (see supplementary material Fig. S3B) or to restore long-term repopulating ability of these HSCs and progenitor cells (see supplementary material Fig. S3C). The production of reactive oxygen species (ROS) has been implicated as a mechanism of TNFα-induced cell death (Sakon et al., 2003; Ventura et al., 2004). To address whether ROS might be relevant for TNFα-induced hematopoietic senescence, we pretreated the TNFα-injected mice with the ROS scavenger N-acetyl-L-cysteine (NAC). Remarkably, NAC almost completely abrogated the induction of senescence-associated heterochromatin foci (HP1-γ) by TNFα in both BM LSK cells and BM sections of Fancc–/– mice (Fig. 4A), indicating that TNFα-induced senescence was mediated by oxidative stress. Examination of the effect of exogenous peroxide (H2O2; a potent ROS producer) on BM LSK cells demonstrated that, indeed, Fancc–/– HSC/progenitor cells were vulnerable to oxidative-stress-induced senescence (Fig. 4B) and inhibition of hematopoietic reconstitution (Fig. 4C). Analysis of ROS production by flow cytometry revealed that TNFα did not cause substantial ROS production in the BM of WT mice (Fig. 4D). By contrast, there was a significant ROS accumulation in the Fancc–/– BM. In vitro culture of isolated BM LSK cells in the presence of TNFα further demonstrated increased ROS production in Fancc–/– HSC/progenitor cells (see supplementary material Fig. S4).
To ascertain that it was TNFα that induced ROS production in hematopoietic cells, we took two approaches: pretreatment of mice with neutralizing anti-TNFα antibody and inactivation of TNFα signaling in Fancc–/– mice. For the latter, Fancc–/– mice were crossed with mice carrying a null mutation in the type 1 TNF receptor (Tnfr1–/– mice). Nearly complete prevention of TNFα-induced ROS accumulation (Fig. 4E) and senescence induction (Fig. 4F) was obtained with TNFα-injected mice deficient for Tnfr1. Whereas treatment of mice with anti-TNFα antibody 30 minutes after each TNFα injection did not completely neutralize serum TNFα [mean ELISA values for anti-TNFα and vehicle-injected animals were 78.6±12.4 pg/ml and 245±21.5 pg/ml, respectively, in TNFα-treated WT mice (n=6) compared with 63.9±17.2 pg/ml and 227±23.8 pg/ml, respectively, in TNFα-treated Fancc–/– mice (n=6); the basal level of serum TNFα in either WT or Fancc–/– mice was approximately 20 pg/ml], administration of the anti-TNF monoclonal antibody resulted in significant reduction of senescence in the spleen (Fig. 4F). In vitro assays further demonstrated that Fancc–/–;Tnfr1–/– BM HSC/progenitor cells were resistant to TNFα-induced ROS production and senescence (see supplementary material Fig. S5). Thus, oxidative stress is responsible for TNFα-induced hematopoietic senescence in Fancc-deficient mice.
To specifically assess TNFα-induced oxidative damage in hematopoietic organs, we immunostained for 4-hydroxy-2-nonenal (HNE), an established marker of ROS-induced tissue damage and an aldehyde product of polyunsaturated fatty acid oxidation (Cauwels et al., 2003). HNE staining was evident in the spleen of WT mice treated with TNFα (see supplementary material Fig. S6), in which we did not observe significant induction of senescence (Figs 3, ,4).4). We found further increase in HNE immunoreactivity in the treated Fancc–/– mice (see supplementary material Fig. S6). This increase in HNE immunoreactivity was largely prevented by anti-TNFα antibody or NAC treatment (see supplementary material Fig. S6). Thus, oxidative organ damage induced by TNFα, as detected by fatty acid oxidation, appears to be proportional to the level of ROS production but not the extent of induction of senescence.
To determine the effect of TNFα-induced ROS on Fancc–/– hematopoietic function, we performed two established assays: clonogenic progenitor assay and competitive hematopoietic repopulation assay using BM cells from double-knockout (Fancc–/–;Tnfr1–/–) mice. Inactivation of TNFα signaling in Fancc–/– (Fancc–/–;Tnfr1–/–) mice rescued progenitor growth (Fig. 5A) and stem cell repopulating ability (Fig. 5B). Pretreatment of TNFα-injected mice with the ROS scavenger NAC also significantly reduced the inhibitory effect of TNFα in progenitor growth and hematopoietic reconstitution. Interestingly, NAC did not further improve progenitor proliferation or hematopoietic reconstitution of BM cells from mice deficient for the Tnfr1 gene (Fig. 5A,B).
Oxidative stress can induce DNA damage that causes growth arrest and cellular senescence (Ames et al., 1993; Chen, 2000). To determine whether TNFα-generated ROS induced DNA damage in HSC/progenitor cells that appeared to have undergone ROS-dependent senescence (Fig. 4), we employed the comet assay (Fairbairn et al., 1995) using a Fpg-FLARE (fragment length analysis using repair enzymes) assay kit. This assay measures specifically oxidative DNA damage including single- and double-strand DNA breaks. There was significant accumulation of DNA damage in BM LSK cells freshly isolated from TNFα-treated WT mice and, to a much greater degree, in TNFα-treated Fancc–/– mice (Fig. 6A). Moreover, there was a significant difference between WT and Fancc–/– cells in terms of kinetics of DNA damage repair, as measured by the remaining amounts of DNA strand breaks over a period of 8 hours of TNFα treatment (see supplementary material Fig. S7). The levels of TNFα-induced DNA strand breaks remained high in Fancc–/– cells at each time point compared with those in WT cells. A similar increase in 8-oxo-deoxyguanosin (8-oxodG), an established marker of oxidative DNA damage, was also demonstrated in BM LSK cells from TNFα-injected Fancc–/– mice compared with treated WT mice (Fig. 6B). Treatment of TNFα-injected Fancc–/– mice with neutralizing anti-TNFα antibody or NAC reduced the accumulation of both DNA strand breaks and 8-oxodG (Fig. 6A,B). Thus, TNFα-induced senescence in HSC/progenitor cells appears to involve oxidative DNA damage.
Genomic instability is the cellular hallmark of FA (Bagby, Jr, 2003; Kennedy and D'Andrea, 2005), and unrepaired TNFα-induced oxidative DNA damage may increase genomic instability leading to premature senescence of HSC/progenitor cells. To test this possibility, we conducted karyotype analysis of BM Lin– (enriched for HSC/progenitor) cells from TNFα-injected WT and Fancc–/– mice. Chromosomal aberrations were rare in cells from treated WT mice, with fewer than 10% of the cells showing one aberration and almost no cells containing two or more aberrations (Table 1). By contrast, BM HSC/progenitor cells from TNFα-injected Fancc–/– mice showed significant increase in chromosomal aberrations (56% of the cells contained at least one aberration, 35% had two or more aberrations, and 23% had three or more aberrations; Table 1). FISH analysis of the chromosomal damages indicated that the TNFα-induced aberrations in Fancc–/– cells consisted mostly of chromatid breaks, gaps, chromosomal breaks, dicentric chromosomes, double minutes, chromosome fragments and fusions (Fig. 6C; Table 2), suggesting that the efficiency and accuracy of DNA strand break/repair may have been compromised in these cells.
A biochemical consequence of genomic damage is the activation of DNA-damage response markers, which include phosphorylated kinases ATM and Chk2, and phosphorylated histone H2AX (γH2AX) and p53 (Shiloh, 2003; Bartkova et al., 2005). Phosphorylation of p53 at Ser20 (p53Ser20) is considered a specific indicator of oxidative DNA damage (Shieh et al., 1999; d'Adda di Fagagna et al., 2003); whereas the formation of γH2AX foci constitutes a robust marker of DNA strand breaks (Banin et al., 1998; Celeste et al., 2003). In HSC/progenitor cells from TNFα-injected WT mice, about 15% of cells stained positive for p53Ser20 and less than 10% stained positive for γH2AX (Fig. 6D). However, a majority of the cells from TNFα-injected Fancc–/– mice were intensively stained for p53Ser20 (62%) or γH2AX (80%). Treatment of the mice with NAC reduced these damage responses to near basal levels (Fig. 6D).
The persistent high levels of oxidative DNA damage observed in HSC/progenitor cells from TNFα-injected Fancc–/– mice suggest that a deficiency in the FA pathway renders chromosomal DNA susceptible to ROS attack, thereby increasing oxidative DNA damage. Alternatively, a defect in the FA function might compromise the damage response/repair process. To distinguish between these possibilities, we treated BM cells from WT and Fancc–/– mice with H2O2, and conducted a time-course study to assess DNA-repair kinetics by examining the levels of p53Ser20 and γH2AX. Compared with WT cells, Fancc–/– cells consistently showed a significant delay in the kinetics of DNA-damage response/repair as evidenced by the much slower clearance of the markers of oxidative DNA damage (p53Ser20) and DNA strand break (γH2AX) (Fig. 6E). This suggests that Fancc–/– cells accumulated high levels of oxidative DNA damage due to impairment of DNA-damage response/repair rather than to an increase in the susceptibility of their DNA to oxidative damage.
We used the disease model of the genomic instability syndrome FA to investigate the role of TNFα-mediated inflammation in HSC senescence. We demonstrated that TNFα induced premature senescence in BM HSCs and progenitor cells as well as other tissues of mice deleted for the Fancc gene, which is required for the maintenance of genetic integrity (Bagby, Jr, 2003; Kennedy and D'Andrea, 2005). Importantly, TNFα-induced senescence of HSCs and progenitor cells correlated with the accumulation of inflammatory ROS, and oxidative DNA damage. ROS are involved in the induction of a senescent phenotype characterized by irreversible growth arrest (Chen et al., 1995; Sohal and Weindruch, 1996). The question naturally arises as to whether the inhibition of ROS can rescue HSCs and progenitor cells from TNFα-mediated senescence. Our findings revealed that pretreatment of TNFα-injected Fancc-deficient mice with antioxidant abrogated the exacerbated inflammatory phenotype, and prevented oxidative DNA damage and hematopoietic senescence. It is clear that the persistent inflammatory response, as reflected by prolonged production of inflammatory molecules, is primarily responsible for the generation of ROS and senescence, because mice deficient for both FA function and TNFα signaling (Fancc–/–;Tnfr1–/–) were resistant to TNFα-induced ROS production and senescence. Moreover, HSCs and progenitor cells from TNFα-treated Fancc-deficient mice showed increased genomic instability and an impaired repair of oxidative DNA damage. Our study thus provides a link between inflammatory ROS and HSC senescence.
Compelling evidence indicates that ROS are widely implicated in the inflammatory process (Lavrovsky et al., 2000). However, the mechanism by which ROS contribute to the proinflammatory states of the aging process is not well defined. Our results demonstrated that TNFα-induced ROS contributed to the increase in oxidative DNA damage in Fancc-deficient HSCs and progenitor cells. Inflammatory ROS have been associated with the initiation or aggravation of diverse pathological states including aging and cancers (Cutler, 2005; Wallace, 2005; Yoshida et al., 2005). We believe that in cells with compromised DNA repair capacity, the ability of inflammatory ROS to damage DNA is the mechanism through which ROS mediate their effects on the inflammatory and, thus, aging process. Indeed, studies have shown that the production of ROS by TNFα at inflammatory sites causes DNA damage (Goossens et al., 1999; Suematsu et al., 2003; Wheelhouse et al., 2003). The role of ROS in TNFα-induced inflammation and HSC senescence was validated by the use of the ROS scavenger NAC, which showed that inhibition of ROS accumulation reduced oxidative DNA damage and cellular senescence. In addition, mice treated with TNFα accumulated high levels of ROS, and administration of NAC significantly reduced the amounts of secreted pro-inflammatory cytokines (Y.Q. and Q.P., unpublished results), indicating that the inflammatory response is, at least in part, mediated by ROS. Thus, these results suggest that ROS serves as a link between inflammation and senescence.
By investigating four well-established senescence makers (SA-β-gal, HP1-γ, p53 and p16INK4A), we provided evidence that TNFα induced premature senescence in BM HSCs and progenitor cells and also other tissues of Fancc–/– mice. Recently, another senescence marker protein, SMP30, has been shown to play an important role in senescence and aging, and SPM-deficiency renders mice highly susceptible to oxidative stress (Sato et al., 2006). It would be interesting to examine the effect of TNFα on the expression pattern of this new senescence marker in HSCs and progenitor cells of Fancc–/– mice.
The persistent DNA damage response and increased genomic instability in senescent HSCs, and progenitor cells are two important features of our study. BM progenitor cells from TNFα-treated Fancc–/– mice contained high levels of DNA strand breakage and oxidative DNA damage. Moreover, studies of repair kinetics revealed much slower clearance of the oxidative DNA damage and DNA strand break markers in Fancc–/– HSCs and progenitor cells than in WT cells. Consistent with this, DNA damage response was persisted in these Fancc–/– cells. This suggests that premature senescence observed in Fancc–/– HSCs and progenitor cells may result from the prolonged activation of the oxidative DNA damage and DNA double-strand-break checkpoints. However, this prolonged checkpoint activation did not facilitate damage repair. Instead, we found dramatically increased genomic instability in Fancc–/– cells. Thus, we conclude that it is unrepaired DNA damage that results in the persistent DNA damage response in these cells.
Our study raises an important question: can inflammatory response be a link between aging and HSC function? It is long known that general inflammatory stress tends to increase with age (Chung et al., 2001). Under inflammatory conditions, HSCs must be able to produce a large number of leukocytes, which are then activated to fight against invaders or stressing agents. This, ultimately, may lead to premature exhaustion of the HSC pool. In the meantime, the HSCs become targets of the toxicity of inflammatory ROS. The ensuing consequence will depend upon the capacity of HSCs to repair ROS-induced damage, particularly DNA damage. In aged and certain disease states, inefficient repair of the oxidative damage may lead to the decrease of the HSC quality (self-renewal capacity). Hence, the toll of inflammatory stress consists in premature senescence of HSCs.
Chronic inflammation and oxidative stress are important features in the pathogenesis of BM diseases such as FA (Lavrovsky et al., 2000; Bagby, Jr, 2003; Chen, 2005). The increased oxidative stress in FA patients may be the result of an increased burden of endogenously produced oxidants as well as increased amounts of ROS generated by various inflammatory cytokines – as suggested in our study. Therefore, understanding the relationship between ROS and inflammation in the context of HSC senescence and aging in these disease states provides a unique opportunity to mechanistically comprehend, and potentially intervene in these physiologically important processes. In addition, our results suggest that, antioxidant compounds may be of therapeutic value in monitoring disease progression, and antioxidant therapy could be used to stop the initiation and propagation of inflammatory response in these diseases.
Generation of Fancc knockout mice has been described by Chen et al. (Chen et al., 1996). Fancc+/– mice were intercrossed with C57Bl/6 mice for more than ten generations to develop an inbred strain. Fancc–/– mice and their WT littermate controls were generated by interbreeding the heterozygous Fancc+/– mice. The mice were maintained on a C57BL/6 (CD45.2+) background. Fancc–/–;Tnfr1–/– double-knockout mice were generated by mating Fancc+/– with Tnfr1–/– mice (Jackson Laboratory, Bar Harbor, ME), followed by the mating of F1 heterozygous siblings. All mice were used at approximately 10-14 weeks of age. Mice were injected intraperitoneally (i.p.) with mouse recombinant TNFα (Peprotech) in PBS at 0.1 mg/kg per day for 2 consecutive days. N-acetyl-L-cysteine (NAC; Sigma) was injected i.p. at 1 mg per mouse 30 minutes before and after each TNFα injection. For anti-TNFα antibody treatment, TNFα-treated mice were injected with 20 μg of neutralizing mouse anti-TNFα antibody (R&D Systems) 30 minutes after each TNFα injection. All experimental procedures conducted in this study were approved by the Institutional Animal Care and Use Committee of Cincinnati Children's Hospital Medical Center.
BM mononuclear cells were depleted of lineage-committed cells using the magnetic-activated cell separation (MACS) cell separation columns (Miltenyi Biotec Inc.). Lin–-Sca-1+-c-Kit+ (LSK) cells were then purified by staining Lin– cells with phycoerythrin (PE)-conjucated anti-Sca-1 (Sca-1–PE) and allophycocyanin (APC)-conjucated anti-c-Kit (c-Kit–APC) antibodies (BD PharMingen) followed by cell sorting using a fluorescence-activated cell sorter (FACS) FACSCalibur (Becton Dickinson).
BM progenitor cells were cultured in a 35-mm tissue culture dish in 4 ml of semisolid medium containing 3 ml of MethoCult M 3134 (Stem Cell Technologies) and the following growth factors: 100 ng/ml stem cell factor (SCF), 10 ng/ml interleukin-3 (IL-3), 100 ng/ml granulocyte colony-stimulating factor (G-CSF) and 4 U/ml erythropoietin (Peprotech). On day 10 after plating, the colonies were counted. Colony growth results are given expressed as the mean (of triplicate plates) ± s.d.
Age-matched congenic B6.SJL-PtrcaPep3b/BoyJ (B6.BoyJ; CD45.1+) mice (Jackson Laboratories, Bar Harbor, ME) were used as transplant recipients. These mice were lethally irradiated (9.5 Gy, 110 cGy/minute, 137Cs γ-rays) and injected intravenously with 2×106 test cells (CD45.2+), mixed with 1×106 competitor cells (BoyJ; CD45.1+). Donor-derived repopulation in recipients was assessed by the proportion of leukocytes in peripheral blood that expressed the CD45.2 marker by flow cytometry. Short- and long-term engraftment and multi-lineage repopulation analysis of donor cells were performed 1 month and 4 month after transplantation, respectively.
Cells were suspended in FACS buffer (0.1% FCS in 0.02% sodium azide) and incubated with the indicated antibodies on ice for 30 minutes, followed by two washes. Data were collected on a FACSCalibur (Becton Dickinson). Labeling was done using the following antibodies: anti-Sca-1, anti-c-Kit, anti-B220, anti-CD3e, anti-CD4, anti-CD8, anti-Gr-1, anti-Mac-1 and anti-Ter119 (all from BD PharMingen, San Diego, CA).
Cells were stained with annexin V and 7-AAD using BD ApoAlert Annexin V kit (BD PharMingen) in accordance with the manufacturer's instructions. Apoptosis was analyzed by quantification of annexin-V-positive cell population by flow cytometry. For cell cycle analysis, cells were permeabilized with 0.3% Nonidet P-40 (NP-40), and then stained with propidium iodide (PI) containing 1 mg/ml RNase A, followed by FACS analysis of the G0-G1, S and G2-M populations.
Cells were cytospun onto slides and fixed in ice-cold methanol for 5 minutes at –20°C. After air drying, cells were blocked for 1 hour with 5% normal serum. Then cells were incubated with antibodies against HP1-γ (07-332; Upstate Cell Signaling Solutions, Lake Placid, NY), p53 (FL-393) and p16INK4A (F-12) (both from Santa Cruz Biotechnology) in PBS with 2% normal serum at room temperature for 1 hour. After extensive washes, cells were incubated with PE-conjugated secondary antibody (Jackson, Bar Harbor, ME). DNA was then labeled with 4,6 diamidino-2-phenylindole (DAPI; Sigma). Slides were finally mounted in mounting medium (Vector). For senescence-associated β galactosidase (SA-β-gal) assay, cells were stained using a SA-β-gal staining kit (#9860; Cell Signaling) according to manufacturer's instructions.
Cells were incubated with CM-H2DCFDA (Molecular Probe) in the dark for 15 minutes at 37°C. After washing, cells were analyzed by flow cytometry using a FACSCalibur (BD Biosciences). Data were analyzed by using the CellQuest program (BD Biosciences).
During necropsy, organs were removed, preserved in formalin and then embedded in paraffin blocks. Paraffin sections were deparaffinized, rehydrated, incubated in 0.1 mM sodium citrate (pH 6.0), washed and incubated with peroxidase blocking reagent (Vector Laboratories, VectaStain Elite ABC kit). After washing in PBS, slides were incubated with primary antibodies against HNE (11-S; Alpha Diagnostic International, San Antonio, TX) or HP1-γ (07-332; Upstate Cell Signaling Solutions, Lake Placid, NY). Following three PBS washes, slides were incubated with secondary antibody and then detected with the VectaStain Elite ABC reagents.
The retroviral vector expressing the mTERT was kindly provided by Fuyuki Ishkawa (Kyoto University, Japan). Retroviruses were prepared by the Vector Core at Cincinnati Children's Research Foundation. Retroviral supernatant was collected 36 hours, 48 hours and 60 hours after transfection. Cells were plated onto non-tissue culture 24-well plates coated with Retronectin (Takara-Shuzo, Shiga, Japan) and pre-stimulated for 2 days in Iscove's modified Dulbecco's medium (IMDM) containing 20% FCS, 100 ng/ml SCF, 20 ng/ml IL-6, and 50 ng/ml Flt-3L (Peprotech). Cells were then exposed to the retroviral supernatant for 3 hours at 37°C in the presence of 4 μg/ml Polybrene (Sigma). Cells were centrifuged at 600 g for 45 minutes. Infection was repeated twice and infection efficiency was assessed by the detection of green fluorescent protein (GFP)-positive cells by FACS.
Cells were solubilized in RIPA lysis buffer (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.5% sodium deoxycholate, 0.1% sodium dodecyl sulfate (SDS), 1% Nonidet NP-40) containing a cocktail of protease inhibitors (Calbiochem, San Diego, CA). Equal amounts of protein were separated on 10% SDS-PAGE, transferred to a nitrocellulose membrane and blotted with antibodies against p53Ser20 (Santa Cruz Biotech), γ-H2AX (Upstate Biotechnologies), and β-actin (Sigma).
Serum levels of inflammatory cytokines were measured using enzyme-linked immunoadsorbent assays (ELISA) kits (R&D Systems).
Generation of DNA strand breaks was assessed by the single-cell gel electrophoresis (comet) assay (Fairbairn et al., 1995), using a Fpg-FLARE (fragment-length analysis using repair enzymes) comet assay kit in accordance with the manufacturer's instructions (Trevigen, Gaithersberg, MO). For each experimental point at least three different cultures were analyzed, and 100 cells were evaluated for comet-tail length from each culture.
Cells were incubated in 0.1 mg/ml colcemid for 30-60 minutes and then incubated in 75 mM KCl. Chromosomes were subsequently fixed in methanol:acetic acid (3:1) and dropped onto glass slides. Metaphase chromosomes were Giemsa-stained and examined for abnormalities. Chromosome aberrations were defined using the nomenclature rules from the Committee on Standard Genetic Nomenclature for Mice.
Data were analyzed statistically using a two-tail Student's t-test. The level of statistical significance stated in the text was based on the P values. P<0.05 was considered statistically significant.
We thank Manuel Buchwald (Hospital for Sick Children, University of Toronto) for the Fancc+/– mice, Fuyuki Ishkawa (Kyoto University) for the mTERT-expressing retroviral vectors, Reena Rani for technical assistance, Jeff Bailey and Victoria Summey for bone marrow transplantation, and the Vector Core of the Cincinnati Children's Research Foundation (Cincinnati Children's Hospital Medical Center) for the preparation of retroviruses. Q.P. thanks Grover Bagby (Oregon Health Science University) for continued support. This work was supported in part by NIH grants R01 CA109641 and R01 HL076712.
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/120/9/1572/DC1