The adult mammalian central nervous system (CNS) contains a population of slowly dividing oligodendrocyte precursor cells (OPCs), i.e., adult OPCs, which supply new oligodendrocytes throughout the life of animal. While adult OPCs develop from rapidly dividing perinatal OPCs, the mechanisms underlying their quiescence remain unknown. Here, we show that perinatal rodent OPCs cultured with thyroid hormone (TH) under hypoxia become quiescent and acquire adult OPCs-like characteristics. The cyclin-dependent kinase inhibitor p15/INK4b plays crucial roles in the TH-dependent cell cycle deceleration in OPCs under hypoxia. Klf9 is a direct target of TH-dependent signaling. Under hypoxic conditions, hypoxia-inducible factors mediates runt-related transcription factor 1 activity to induce G1 arrest in OPCs through enhancing TH-dependent p15/INK4b expression. As adult OPCs display phenotypes of adult somatic stem cells in the CNS, the current results shed light on environmental requirements for the quiescence of adult somatic stem cells during their development from actively proliferating stem/progenitor cells.
Oligodendrocytes (OLs) are myelinating cells of the vertebrate central nervous system (CNS). They are derived from oligodendrocyte precursor cells (OPCs)1, which are also called NG2 glial cells or O-2A cells. In the rat optic nerve, OPCs first appear at the brain-end of the nerve on embryonic day 16 (E16) and migrate to the nerve, reaching the eye-end around the day of birth (E21)2. OPCs in the developing rat optic nerve exhibit a limited numbers of cell divisions before they terminally differentiate into OLs: the first OLs appear around birth, and their numbers rapidly increase over the following six weeks until the end of optic nerve myelination3. In parallel with this process, rapidly dividing perinatal OPCs disappear from the myelinated nerve, as slowly dividing adult OPCs gradually increase and persist in the adult nerve4,5,6,7. Whereas less than 5% of OPCs are adult OPCs in the optic nerve on postnatal day 7 (P7), almost 70% of OPCs are adult OPCs by P306. Adult OPCs constitute appoximately 5% of the cells throughout the adult CNS, where they have a crucial role in remyelination following CNS damage througout the life of animal, suggesting that adult OPCs are adult somatic stem cells8,9,10. Fate-mapping studies in transgenic mice have shown that adult OPCs develop from perinatal OPCs11. However, the molecular mechanisms underlying the perinatal-to-adult transition remain unknown12.
The developmental processes from OPCs into OLs in vivo can be reproduced in vitro 13. OPCs prepared from P7 rat optic nerve differentiate into OLs after up to 8 divisions when cultured in 20% O2, either as purified OPCs in serum-free medium containing platelet derived growth factor (PDGF), the major mitogen for OPCs14, 15, and thyroid hormone (TH)16, or as single OPCs on astrocyte monolayers in 0.5% fetal bovine serum (FBS)17. PDGF withdrawal induces premature OPC cell cycle withdrawal and differentiation into OLs18, and in the absence of TH, OPCs can continue to proliferate in culture for more than 13 months19, 20. During this long-term culture, although the cells acquire adult OPC-like morphologies and cell surface antigens, their cell cycle dose not decelerate19.
As oxygen concentration in culture might affect the timing of cell differentiation21, we examined the effects of hypoxia on the OL differentiation of perinatal OPCs22. Surprisingly, a specific O2 concentration of less than 1.5% decelerates the cell cycle of perinatal OPCs without OL differentiation and these cells acquire adult OPC-like characteristics. Furthermore, this deceleration of cell cycle is mediated by runt-related transcription factor 1 (Runx1)23 through the promotion of G1 arrest in OPCs via cyclin-dependent kinase inhibitor (CKI) p15/INK4b24, 25 induction.
Decreasing O2 biphasically regulates OPC proliferation in a TH-dependent manner
Purified P7 rat OPCs (>97% pure) were cultured at clonal density in the serum-free medium containing PDGF and TH16, 26. Concentrations of O2 in the air interface of the culture ranged between 1% and 20%. The cell numbers after culture for 10 days were significantly greater in 5.0% O2 versus 20% O2 (Fig. 1a). Between 5.0% and 2.0% of O2, the cell numbers modestly decreased. Under conditions with less than 1.5% O2, OPCs displayed notably suppressed cell numbers. Immunocytochemistry using the OPC-specific monoclonal antibody A2B5, which recognizes ganglioside GT322, 27, and the monoclonal antibody against galactocerebroside (GC), a cell surface marker of differentiated OLs28, revealed that the percentage of GC-positive (GC+) cells decreased with decreasing O2 level (Fig. 1b, upper panel). Notably, the ratio of the A2B5-positive (A2B5+) cells versus the GC+ cells reached a maximum at 1.5% O2 (Fig. 1b, lower panel), where most of the cells remained A2B5+ and GC-negative (GC−). The immunocytochemistry result (Supplementary Fig. S1) indicated that the A2B5+ cells maintained OPC-specific bipolar bodies in 1.5% O2. In OPCs, TH is known to limit the number of cell divisions to execute terminal differentiation into OLs in 20%16. Thus we tested whether the effects of TH depend on the O2 concentrations by a clonal analysis (see details in Supplementary Fig. S1). In both 20% O2 and 1.5% O2, the application of TH decreased the number of cell divisions to comparable extents of up to 8 times (Fig. 1c). Conversely, when OPCs were cultured with PDGF in the absence of TH in either 20% or 1.5% O2, cells continued to actively proliferate for over 40 days (Fig. 1d), indicating that the decrease in cell number in both 20% and 1.5% O2 depends on TH. This also suggests that the lack of increased cell numbers in 1.5% O2 does not result from hypoxia per se.
As shown in Fig. 1a, OPCs displayed notable proliferation in 3% O2. Therefore, we characterized OPC proliferation with 3% O2 (Supplementary Fig. S2a). The addition of TH did not inhibit the proliferation of OPCs during the first 10 days. We passaged the cells, re-plated them at clonal density, and cultured them for another 10 days in 3% O2. Under these conditions, the cells exhibited cycle withdrawal after up to 7 divisions (Supplementary Fig. S2b); more than 70% of them were GC+ OLs (Supplementary Fig. S2c and d). Such phenotypes observed in 3% O2 differed from those seen in 1.5% O2, and were analogous to those found in 20% O2, while the timing of OL differentiation was delayed.
Hypoxia neither induces cell death and senescence nor inhibits OL differentiation
In 20% O2, the number of GC+ OLs was significantly increased during the period between 6 and 12 days, whereas the number of the same cells was not increased during the same period in 1.5% O2 (Fig. 2a). Moreover, the failure of increase of OPCs in the presence TH was not because of decreased cell viability, as indicated by only small percentages of dead cells in 1.5% O2 (Fig. 2b and Supplementary Fig. S11). Furthermore, the lack of increases in OPCs in 1.5% O2 in the presence of TH was not because of the OPCs undergoing replicative senescence29: after the 20 days of culture in 1.5% O2, the cells retained bipolar OPC morphology without expressing senescence-associated β-galactosidase (SA-βGal)20, 30 (Fig. 2c). Although several studies have shown that hypoxia inhibits OL differentiation31, 32, hypoxia in 1.5% O2 is not sufficient to prevent the terminal differentiation of rat optic nerve OPCs into OLs. As shown in Fig. 2d, PDGF withdrawal caused the rapid differentiation of OPCs into GC+ OLs14, 15, 18; this event did not differ in 20% O2 and 1.5% O2.
Hypoxia converts perinatal OPCs into adult-like OPCs in culture
To examine roles of TH in controlling cell proliferation, freshly prepared P7 OPCs were pre-cultured in the presence of PDGF with or without TH for 2 or 15 days in 1.5% O2 and then treated with bromodeoxyuridine (BrdU) for 20 hours. OPCs undergoing the 2-day TH treatment did not suppress the active BrdU incorporation compared with those cultured without TH (Fig. 3a). On the other hand, the 15-day TH treatment suppressed BrdU incorporation, as 3.7 ± 1.2% of the cells were BrdU-positive (BrdU+) in the presence of TH, whereas 81.6 ± 5.5% were BrdU+ in the absence of TH. We also examined effects of the 40-day TH treatment of P7 OPCs in 1.5% O2; the cells were treated with BrdU for the last 5 days. Under these conditions, 1.6 ± 0.8% of the cells were still labeled with BrdU, which was consistent with a notion that adult OPCs exhibit gradually slowed proliferation but a sustained ability to incorporate BrdU in vivo 33.
We also examined the beginning of the TH-dependent cell cycle deceleration in OPCs. P7 OPCs were cultured with TH in 1.5% or 3% O2, and their cell divisions over 5 days were compared using time-lapse microscopy. OPCs treated with TH in 1.5% O2, but not those in 3% O2, displayed an elongation in the average cell division time, which specifically began on culture day 3 (Fig. 3b). Previous observations have shown that over 95% of P7 rat OPCs cultured in 20% O2 became A2B5−/GC+ OLs within 15 days34. We further observed the effects of longer culture periods on the phenotypes of the cells. Treating OPCs cultured in 1.5% O2 with TH resulted in a decreased population of A2B5+ cells and an increased population of the GC+ cells. These two subpopulations reached a plateau around at day 20 (Fig. 3c). Collectively, in the presence of TH, the hypoxic culture helps sustain undifferentiated A2B5+ cells for longer period.
Over time, the A2B5+ cells exhibiting cell cycle deceleration in 1.5% O2 acquired multi-process morphologies resembling that of adult OPCs4,5,6,7 (Fig. 3d). These multi-process cells moved slowly; according to an analysis using time-lapse video microscopy, when P7 OPCs were cultured in 1.5% O2 for 21 days, the bipolar cells not treated with TH were highly motile and proliferated rapidly (Video 1). On the other hand, the multi-process cells treated with TH exhibiting cell cycle deceleration were much less motile (Video 2); this slow movement demonstrated by the cells was similar to that shown by adult OPCs in vivo 35. And these multi-process cells were could be maintained in 1.5% O2 with TH for more than 4 months without differentiating into OLs.
Similar to perinatal OPCs22, adult OPCs are induced to differentiate either into OLs by the withdrawal of PDGF or into type-2 astrocytes by the addition of 10% FBS4, 5, 7. As shown in Fig. 3e, when P7 OPCs which were cultured in 1.5% O2 with PDGF and TH for 15 days and then in PDGF-free medium for 5 days, they acquired morphological features of OLs and expressed OL marker, myelin basic protein (MBP). On the other hand, P7 OPCs, which were treated for 15 days with PDGF and TH, followed by a 5-day treatment with 10% FBS, the cells exhibited the characteristic morphology of type-2 astrocytes22 and expressed glial fibrillary acidic protein (GFAP). In addition, using a time-lapse video microscopy, we confirmed the morphological changes from OPCs into OLs (Video 3) and into type-2 astrocytes (Video 4) under hypoxia. These data indicate that the cells with a decelerated cell cycle under hypoxia maintained the developmental bipotency of OPCs.
Adult OPCs isolated from adult rat optic nerve can be stimulated to proliferate rapidly by a combination of PDGF, neuregulin 1 (NRG1), and isobutyl methylxanthine (IBMX) in TH-free culture7. To test if this is also the case for the cell cycle-decelerated cells under hypoxia, we dissociated the cells after 15 days of hypoxic culture with TH and re-plated them at clonal density in 20% O2, with or without TH, in the presence of PDGF, NRG1, and IBMX for 7 days (Fig. 3f). The cells cultured without TH in the presence of NRG plus IBMX divided rapidly up to 6 times without differentiation.
Taken together, the phenotypes of P7 rat optic nerve OPCs treated with PDGF and TH for 15 days in 1.5% O2 in vitro are consistent with those of adult OPCs prepared from adult rat optic nerve5, 7. Based on these findings, perinatal OPCs cultured with PDGF and TH under hypoxia for over two weeks are characterized by slow proliferation and an A2B5+ phenotype with developmental bipotency, and thus are designated adult-like OPCs.
p15/INK4b induces G1 arrest in adult-like OPCs
To investigate mechanisms for the TH-dependent deceleration of the cell cycle in OPCs, total RNA was extracted from P7 OPCs cultured in 1.5% O2 with or without TH for 15 days, and gene expressions were analyzed quantitatively on microarray (Supplementary Table S1 ). Among 129 of the TH-dependent up-regulated genes, we identified the gene of p15/INK4b (Cdkn2b)24. Among 60 of the TH-dependent down-regulated genes, 48 genes (80%), including Cyclin A2 (Ccna2), Cyclin B1 (Ccnb1) and Cyclin B2 (Ccnb2), were related to the cell cycle progression. Subsequently, we confirmed the TH-dependent gene expressions of Cyclins and CKIs. RT-PCR analysis showed that TH modestly increased the mRNA levels of G1 Cyclins, Cyclin D1 and D3, and decreased the mRNA levels of G2/M Cyclins, Cyclin A2, B1 and B2 (Fig. 4a). These data suggest that the TH-dependent deceleration of cell cycle in adult-like OPCs results from G1 arrest. p15/INK4b protein induces G1 arrest via direct binding to Cdk424. TH notably increased the mRNA level of p15/INK4b, with negligible effects on mRNA levels of other CKIs tested (Fig. 4b). TH also increased the p15/INK4b protein level in OPCs in 1.5% O2 (immunocytochemistry, in Fig. 4c; western blotting, in Supplementary Fig. S3b).
To examine effects of down- or up-regulation of p15/INK4b gene expression on proliferation of adult-like OPCs, we cultured P7 OPCs without TH for 12 days in 1.5% O2 and then transfected them with anti-p15/INK4b siRNA. The siRNA inhibited the cell cycle deceleration in perinatal OPCs induced by TH in 1.5% O2 (Fig. 4d). Conversely, when a retroviral vector was used to overexpress p15/INK4b in freshly isolated P7 OPCs, which were then cultured for 7 days at clonal-density without TH in 3% O2, where adult-like OPCs are never seen, the number of cell division was reduced and the cells maintained the A2B5+ bipolar morphology (Fig. 4e). These results suggest that p15/INK4b is an effector molecule that mediates the TH-dependent cell cycle deceleration in adult-like OPCs cultured in 1.5% O2.
Klf9 is the major mediator of TH signal to induce adult-like OPCs
As the expression of p15/INK4b in OPCs was regulated transcriptionally, the mechanisms by which TH induces p15/INK4b were further examined. The microarray data in Supplementary Table S1 shows that of the genes encoding transcription factors, 10 of them (Csrp1, Dbp, Dbx2, Hif2α, Klf9, Mrf, Mro, Nkx6.2, Rev-erbA and Runx1) are transcriptionally up-regulated with TH, and 5 of them (Cbx2, Csrp2, Foxm1, Klf14 and Mybl2) are down-regulated with TH. Individual genes of transcription factors up-regulated with TH were assessed by siRNA-mediated gene silencing to examine the effects on OPC proliferation. The gene silencing of 6 transcription factors (Csrp1, Hif2α, Klf9, Nkx6.2, Rev-erbA and Runx1) inhibited the TH-dependent cell cycle deceleration in OPCs in 1.5% O2 (Fig. 5a). The efficiencies of siRNAs were confirmed by western blotting (Supplementary Fig. S4a).
To determine whether the gene expression levels of these transcription factors are under the direct control of thyroid hormone receptor alpha 136, an immediate early-response assay was performed37. P7 rat OPCs cultured without TH in 1.5% O2 for 10 days were pre-treated with cycloheximide for 6 hours, after which TH was added to the medium. The alterations in the gene expression levels of the transcription factors (Csrp1, Hif2α, Klf9, Nkx6.2, Rev-erbA and Runx1) and p15/INK4b were assessed by RT-PCR as a function of time after the TH treatment. Klf9 (Klf9) was the only one gene showing an immediate early-response to the TH stimulation (Fig. 5b). Klf9 (Krüeppel-like factor 9) is a GC-box binding transcription factor that has been considered essential for the TH-dependent differentiation into OLs in cultures with 20% O2 38. Therefore, we examined the effects of Klf9 overexpression in P7 OPCs cultured without TH in 1.5% or 3% O2 conditions for 7 days. In the absence of TH in 1.5% O2, the overexpression of Klf9 induced the cell cycle deceleration in OPCs (Fig. 5c). However, in 3% O2, no cell cycle deceleration in Klf9 overexpressing OPC was seen. Note that, in both O2 conditions, Klf9 overexpression did not accelerate OL differentiation, as the cells maintained OPC-specific A2B5+ bipolar bodies (Supplementary Fig. S5 and Supplementary Fig. S6a). Collectively, although Klf9 plays a crucial role in TH-signaling in adult-like OPCs, other factors that are induced with hypoxia must be needed for the cell cycle deceleration in OPCs.
Runx1 determines OPC cell cycle deceleration
To determine which transcription factor(s) responsible for the deceleration of cell cycle show the 1.5% O2 condition specific gene up-regulation, we then prepared total RNA from P7 rat OPCs cultured with or without TH in 1.5% O2 for 1, 4 and 15 days, as well as from the same cells cultured in 3% O2 for 15 days. The gene expression patterns of the six transcription factors were compared with those of p15/INK4b by RT-PCR (Fig. 5d ). TH-dependent gene up-regulation of p15/INK4b was undetectable on day 1, showed a modest elevation on day 4, and was finally evident on day 15 in 1.5% O2, whereas the TH-dependent up-regulation of the p15/INK4b gene was undetectable on day 15 in 3% O2 conditions. These results support the idea that p15/INK4b is the major effector molecule of the TH-dependent cell cycle deceleration in OPCs in 1.5% O2. Among six transcription factors, Runx1 was the only one that showed the same TH-dependent gene expression pattern as p15/INK4b.
Thus, we hypothesized that the transcription factor Runx1 ultimately determines the TH-dependent cell cycle deceleration in OPCs in 1.5% O2. In rodents, the gene expression of Runx1 is controlled by the dual promoters P1 and P2; these promoters generate two major protein variants (Runx1c and Runx1b)39. In hypoxic OPCs, TH activates the proximal promoter P2 and induces the expression of the Runx1b variant (Supplementary Fig. S7). Freshly prepared P7 rat OPCs were transfected with a retroviral vector for Runx1 (variant Runx1b) overexpression and cultured with or without TH in 3% O2 for 7 days. Notably, Runx1 over-expression decelerated the cell cycle of OPCs irrespective of the presence or absence of TH or hypoxia (Fig. 5e). The cells maintained OPC-specific A2B5+ bipolar morphologies (Supplementary Fig. S6b). These results suggested that the gene expression of Runx1 is crucial for the TH-dependent cell cycle deceleration in OPCs in 1.5% O2. Runx1 is known to determine the expression of p15/INK4b40. Runx1 overexpression increased p15/INK4b protein level in OPCs in 3% O2 (Fig. 5f). These results suggest that the timing of the Runx1 gene expression determines the timing of the TH-dependent cell cycle deceleration in OPCs in 1.5% O2.
Hypoxia-inducible factors induce Runx1 gene expression
We attempted to investigate the mechanism underlying the hypoxia-dependent Runx1 gene expression. Among the six TH-dependent up-regulated transcription factors, Hif2α (Epas1) is an α-subunit of the hetero dimeric transcription factor known as hypoxia-inducible factors (HIFs)41. Other subunits of HIFs, Hif1α (another α-subunit of HIFs) and Hif1β (common β-subunit of HIFs) are constitutively expressed in OPCs (Fig. 6a). In hyperoxic conditions, Hifα proteins (Hif1α or Hif2α) are hydroxylated by prolyl-hydroxylase (PHD) and degraded by proteasome. In 1.5% O2, both Hif1α and Hif2α proteins are stabilized and accumulated in nucleus of OPCs (Fig. 6b,c). The administration of chetomin42 at 2 μM or CAY1058543 at 10 μM, both of which can inhibit Hif1α and Hif2α, induced the cell death of OPCs in 1.5% O2 (Supplementary Fig. S8). These results suggest that HIFs are necessary for the survival of OPCs under hypoxia. On the other hand, the siRNA mediated gene silencing of Hif1α or Hif2α showed that inhibiting both of them inhibited the TH-dependent cell cycle deceleration in OPCs in 1.5% O2 (Fig. 6d). Conversely, the stabilization of Hifα proteins decelerated the cell cycle in OPCs. The administration of a PHD inhibitor N-(2-methoxy-2-oxoacetyl)glycine methyl ester (DMOG)44, which stabilizes Hifα proteins, to OPCs cultured with TH in 3% O2 induced the cell cycle deceleration to an extent comparable to that observed in 1.5% O2 (Fig. 6e). Furthermore, DMOG increased Runx1 gene expression in OPCs cultured with TH in 3% O2, in parallel with the up-regulation of HIFs-related genes41, such as lactate dehydrogenase A (Ldha), phosphoglycerate kinase 1 (Pgk1) and vascular endothelial growth factor A (Vegfa) (Fig. 6f). Consequently, Runx1 protein level was increased in OPCs treated with TH and DMOG in 3% O2 (Fig. 6g), suggesting that the Runx1 gene expression is under the control of transcription factor HIFs, and that the O2 sensing of OPCs mainly depends on the stabilities of Hifα proteins.
Relevance of adult-like OPCs in culture to adult OPCs in rat optic nerves
To examine the relevance of the adult-like OPCs in culture and adult OPCs in vivo, we then compared the gene expression level of the adult-like OPC-related transcription factors in freshly prepared A2B5+/GC− optic nerve OPCs from P7 rat and in those from P14 rat. Previous studies have shown that less than 5% of OPCs in P7 rat optic nerves are adult OPCs6 and more than 50% of OPCs in P14 rat optic nerves are adult OPCs45. A clonal analysis showed that approximately 50% of purified A2B5+/GC− OPCs in P14 rat optic nerves divided fewer than 3 times over 13 days of culture in 1.5% O2, regardless of the presence or absence of TH (Fig. 7a). This finding suggests that these slowly dividing P14 OPCs are committed to becoming adult OPCs. The results of qRT-PCR showed that Runx1 and Klf9 mRNA levels were more than 9-fold and 4-fold greater, respectively, in P14 OPCs than those in P7 OPC (Fig. 7b). Furthermore, we attempted to confirm the increase in the Runx1 protein level in slowly dividing P14 OPCs; FACS sorted A2B5+/GC− P14 OPCs were stained with anti-Runx1 and anti-Ki67 antibodies. Ki67 is a protein marker of proliferating cells; thus, we considered Ki-67-negative (Ki-67−) OPCs as adult OPCs. The percentage of Ki67− cells expressing Runx1 protein was significantly greater than that of Ki67+ cells (Fig. 7c), suggesting that Runx1 contributes to the development of adult OPCs in vivo.
To explore OPCs under hypoxic conditions in the developing rat optic nerve in vivo, we used pimonidazole, which is activated in hypoxic cells to form stable covalent adducts with thiol groups in proteins46. We injected pimonidazole intraperitoneally into both P7 and P14 rats and left the animals for 2 hours. Then, we purified the OPCs from their optic nerves and stained the cells for pimonidazole and Ki-67. As shown in Supplementary Fig. S9b, in both P7 and P14 OPCs, about one-third of the OPCs were pimonidazole-positive (Pimo+), suggesting that these cells were exposed to hypoxic environments in vivo. At P7 when there are relatively few adult OPCs, only 23.7 ± 4.6% of the Pimo+ cells were Ki-67− (Fig. 7d). On the other hand, at P14, 55.4 ± 2.3% of the Pimo+ cells were Ki-67−. These results suggest that hypoxic conditions in the developing rat optic nerve promote the transition of perinatal OPCs to adult OPCs.
TH-dependent cell cycle deceleration in rat cortex OPCs and mouse OPCs
OPCs purified from P7 rat cerebral cortex47 clearly showed TH-dependent cell cycle deceleration in 1.5% O2 (Fig. 8a–c). Moreover, OPCs purified from P7 mouse optic nerves48 also showed the TH-dependent cell cycle deceleration in hypoxic culture, while the hypoxic conditions at 1% O2 were necessary in the case of mouse (Fig. 8d). These results suggest that the TH-dependent cell cycle deceleration in OPCs under hypoxic condition is a general event occurring in rodents.
The current study provides evidences that, under hypoxic culture conditions at less than 1.5% O2 in the presence of PDGF and TH, actively proliferating perinatal OPCs progressively slow their proliferation and acquire adult OPC-like characteristics. The mechanisms for the TH-dependent cell cycle deceleration in OPCs involve Runx1. Furthermore, results collected in P14 rats showed that A2B5+ OPCs showing decreased proliferative signals express greater levels of Runx1 protein, suggesting pivotal roles of Runx1 in the development of adult OPCs in vivo.
Our study raised an important question as to whether the hypoxic conditions used in our culture are physiologically relevant11. The magnitude of hypoxia (equivalent 1% O2; pO2 = 7.2 mmHg at 37 °C) is known to occur in embryos—in the thymus, kidney medulla and bone marrow49, 50. The Km value of heme oxygenase-2, an O2-sensing enzyme in the CNS, is 15 mM (pO2 = 11 mmHg at 37 °C)51, which is close to our hypoxic culture (1.5% O2; pO2 = 13.8 mmHg at 37 °C). While the O2 concentrations in the CNS under normal conditions are estimated to be approximately 2–5% O2 21, 50, and a simulation study of O2 diffusion suggested that O2 concentrations in a tissue decrease by 10-fold at a distance of several cell diameters from the nearest capillary52. These previous observations led us to hypothesize that a considerable number of OPCs reside in hypoxic niches in vivo. When we injected pimonidazole into living rats, as many as 30% of the OPCs in the optic nerves were positively labeled (Supplemmentary Fig. S9b). Considering that pimonidazole preferentially labels proteins in cells at O2 concentrations less than 14 mM (pO2 = 10 mmHg at 37 °C)45, the current results suggest that the pimonidazole-labeled OPCs actually reside under hypoxic conditions in vivo that are comparable to culture condition with less than 1.5% O2. It has been shown that a hypoxic environment is necessary to maintain the quiescence of adult OPCs in vivo 32. The current results suggest that such hypoxic environment is also essential for the development of adult OPCs by mechanisms involving Runx1.
Runx1 is a member of the Runx protein family53. Members of this family can directly bind to DNA via a conserved DNA-binding motif, known as the runt domain. In the peripheral nervous system, the defect of Runx1 activity causes impaired responses to noxious stimuli54. On the other hand, little is known about Runx1 loss of function in the CNS55. However, increasing Runx1 protein expression inhibits the proliferation and promotes the maturation of not only olfactory ensheathing cells, non-myelinating axon-wrapping glial cells in the olfactory nerve56, but also microglia, although they are myeloid lineage cells, in the postnatal forebrain57. To the best of our knowledge, there have been no previous reports about Runx1 expression in oligodendrocyte lineage cells. Importantly, Runx1 is the master gene of the development of hematopoietic stem cells (HSCs)23 and is required for the differentiation of HSCs from hemogenic endothelium in embryos58, as well as for the inhibiting the proliferation of myeloid progenitors in the bone marrow after birth59. In mouse and human HSCs, the overexpression of Runx1 protein variants, Runx1b (the variant expressed in the TH-stimulated hypoxic OPCs) or Runx1c, induces HSC quiescence in vivo 60, 61. Additionally, the Hif1α protein level in HSCs has been shown to play a critical role in regulating the quiescence and the differentiation of these cells62, 63. Thus, it is not unreasonable to suggest that as in OPCs, HIFs may induce HSCs quiescence via inducing Runx1 gene expression in hypoxic niches in the bone marrow.
Our studies showed that p15/INK4b is the major effector molecule for the TH-dependent cell cycle deceleration in hypoxic OPCs, and its expression depends on the transcription factor Runx1, finally, the sensing mechanisms for environmental O2 concentrations by OPCs depend on the HIFs-mediated up-regulation of Runx1 gene expression. The current results shed light on the possible involvement of an unidentified, intrinsic cellular developmental program resembling the intracellular developmental timer that determines the timing of the OL differentiation of perinatal OPCs cultured in 20% O2 3. Several lines of observation support this hypothesis. First, both the perinatal-to-adult transition of OPCs cultured in 1.5% O2 and the OL differentiation of OPCs in 20% O2 culture depend on extracellular PDGF and TH. Second, in the experiments using P7 rat optic nerve OPCs, the timing for the transition of OPCs from perinatal to adult-like in 1.5% O2 culture was almost identical to that for the differentiation into OL in 20% O2 culture (Fig. 1c). Third, in 20% O2 culture, TH can be replaced by retinoic acid (RA) to induce the time-dependent OL differentiation16. The cell cycle deceleration in OPCs in 1.5% O2 was also induced by adding RA (Supplementary Fig. S10). Revealing the mechanisms for the intrinsic cellular developmental program that determines the timing of the TH-dependent differentiation of adult-like OPCs under hypoxia deserve further investigation. Our findings are summarized in Fig. 9.
Materials and Methods
All experiments in this study were carried out in accordance with the regulations of Keio University School of Medicine. The ethics committee of Keio University School of Medicine approved the current protocols for the animal experiments (ID; 09201-1). And the recombinant DNA experiments were carried out in accordance with the guidelines of the Ministry of Education, Culture, Sports, Science and Technology of Japan.
Preparation of rodent OPCs
Sprague-Dawley (SD) rats and C57BL/6 mice were obtained from Japan SLC Inc. (Hamamatsu, Japan). The immunopanning purification of OPCs from rat optic nerves was performed according to Barres et al.26. In our experiments, over 97% of the cells were A2B5+ OPCs. The immunopanning purification of OPCs from rat cerebral cortex was performed according to Dugas et al.47. And the immunopanning purification of OPCs from mouse optic nerves was performed according to Watkins et al.48.
Clonal density culture of OPCs
For clonal density cultures, 1,000–2,000 of purified OPCs were inoculated in poly D-lysine (PDL)-coated T25 culture flasks or slide flasks in serum-free modified BS medium16, containing 10 ng/ml PDGF, 5 ng/ml NT-3, 5 μg/ml insulin, 5 μM forskolin (20 μM for mouse OPCs)64, with or without TH (a mixture of 3,3′,5-triiodo-L-thyronine and thyroxine; 40 ng/ml of each). Cells were cultured at 37 °C in 5% CO2 and 1~20% O2 (in hypoxic culture conditions, O2 was replaced with N2).
The procedures of immunocytochemistry were described previously19. Nucleuses were stained with DAPI (1 μg/ml) or PI (2 μg/ml) containing RNase A (50 μg/ml). Samples were examined with a BZ-9000 inverted fluorescence microscope (Keyence). Anti-GFAP rabbit polyclonal antibodies (G4546, Sigma-Aldrich), anti-MBP rat monoclonal antibody (MAB386, Millipore), anti-Ki-67 rabbit polyclonal antibodies (AB9260, Millipore), anti-Hif1α rabbit polyclonal antibodies (NB100-479, Novus Biologicals), anti-Hif2α rabbit polyclonal antibodies (NB100-122, Novus Biologicals), Anti-CDKN2B (p15/INK4b) rabbit polyclonal antibodies (bs-4269R, Bioss Antibody), anti-RUNX1/AML1 rabbit polyclonal antibodies (ab23980, Abcam).
Nuclear staining dyes PI (2.5 μg/ml) and Hoechst 33342 (5 μg/ml) were added to the culture for 90 minutes. PI+/Hoechst 33342+ double-positive cells were considered dead.
SA-βGal assay was carried out obeying Debacq-Chainiaux et al.30. Cells were fixed with 2% PFA. They were rinsed and incubated in staining solution (40 mM citric acid/Na phosphate buffer (pH 6.0), 5 mM K4[Fe(CN)6], 5 mM K3[Fe(CN)6], 150 mM NaCl, 2 mM MgCl2, 1 mg/ml X-gal) for 16 hours at 37 °C. After then, cells were post-fixed with methanol. The samples were examined with an inverted microscope CKX41 (Olympus).
BrdU incorporation assay
The procedures of the BrdU incorporation assay were described previously19. In short, cells cultured with 5 μM of BrdU were fixed and permeabilized in cold 70% ethanol, and incubated in 6N HCl-1% Triton-X100 for 15 minutes at room temperature (RT). After the rinse, cells were neutralized with 0.1 M sodium borate (in PBS with 1% Triton-X100) for 10 minutes at RT. Then cells were blocked with 50% normal goat serum-1% Triton-X100 and incubated with anti-BrdU rat monoclonal antibody (OBT0030, Oxford Biotechnology), followed by Alexa Fluor 488 goat anti-rat IgG antibodies. After post-fix, the nucleuses were stained with Hoechst 33342 (5 μg/ml).
Time-lapse differential interference microscopic images of cells cultured on PDL/fibronectin (FN)-coated glass base culture dishes were recoded every 30 minutes for 5 days (120 hours) or every 15 minutes for 24 hours by using an incubator microscope LCV110 (Olympus). Editing of video images were carried out using Meta Imaging Software version 6.1 (Molecular Devices).
Gene expression DNA Chip analysis
P7 rat optic nerves OPCs were cultured with or without TH in 1.5% O2 conditions for 15 days. Preparation of total RNA from OPCs was carried out using TRIZOL Reagent (Life Technologies) obeying manufacture’s instructions. The data analysis of Agilent Rat Gene Expression Microarray (Rat GE 4 × 44 k v3, 26,930 Entrez gene RNAs, Agilent Technologies) was carried out using GeneSpring13.1 software (Agilent Technologies).
Reverse transcription-PCR (RT-PCR)
Total RNA purification was carried out using NucleoSpin RNA XS kit (Macherey-Nagel GmbH & Co.). SMARTer PCR cDNA synthesis kit (Clontech) was used for the cDNA synthesis obeying supplier’s instructions. For the preparation of PCR reaction mixture, PrimeStar GXL DNA polymerase kit (Takara-Bio) was used. The sequences of all PCR primers are shown in Supplementary Table S2. The condition of PCR was 10 seconds at 98 °C for the denaturing, 15 seconds at 60 °C for the annealing and 40 seconds at 68 °C for the extension. The total numbers of cycling were 20–35. The PCR products were analyzed on 2% agarose gel electrophoresis.
Quantitative RT-PCR (qRT-PCR)
For the isolation of total RNA from OPCs, TRIZOL Reagent (Thermo Fisher Scientific) was used. Takara PrimeScript II 1st strand cDNA synthesis kit (Takara-Bio) was used for cDNA synthesis obeying supplier’s instructions. 20 μl of qRT-PCR reaction mixture consisted of 10 μl of BIO-RAD iQ SYBR Green Supermix (BIO-RAD), 5 μl of primer mix (2 μM of each forward and reverse primers) and 5 μl of template cDNA (0.2 ng/μl). The sequences of PCR primers for qRT-PCR are shown in Supplementary Table S3. Real-time PCR was performed on a CFX96 Real-Time System (BIO-RAD). The condition of PCR was 5 seconds at 95 °C for the denaturing, 30 seconds at 60 °C for the annealing and the extension. The total numbers of cycling were 40. The resulting values were normalized to the endogenous control gene, glyceraldehyde 3-phosphate dehydrogenase (Gapdh) or β-actin (Actb). Results were presented as the relative expression to that of the control using the comparative Ct method (∆∆Ct).
siRNA mediated gene silencing
The sequences of the target sites of each siRNA are shown in Supplementary Table S4. The inhibitory efficiency on the protein expression by anti-Runx1, anti-Hif1α, anti-Hif2α, anti-p15/INK4b siRNAs is shown in Supplementary Fig. S4. Co-transfection of siRNA and green fluorescent protein (GFP) expressing reporter plasmid DNA pMaxGFP (Lonza) to OPCs was carried out using Amaxa Basic Glial Cells Nucleofector kit (Lonza) obeying supplier’s instructions. P7 rat OPCs cultured without TH in 1.5% O2 for 14 days were dissociated with trypsin. 3.3 × 105 cells were suspended with 100 μl of Nucleofector solution mixture. Then, 2 μl of siRNA mixture (1 μM each) and 2 μl of GFP-expressing reporter plasmid pMaxGFP DNA (0.5 mg/ml) were added to it. Electroporation was operated using Nucleofector II (Lonza) programmed with #O-005. Cells suspended with 9 ml of TH-free BS medium were sifted using a 40 μm cell strainer. 3 ml of cell suspensions were inoculated into each PDL-coated slide flask. Cells were cultured in 1.5% O2 for 2 hours to allow them attaching the bottom, then all medium were replaced to 3 ml of fresh BS medium with TH. Cells were cultured in 1.5% O2 for 4–5 days. The number of cells in the GFP-expressing clones was counted.
Construction of retrovirus vector plasmid DNA
Constructs of retrovirus vectors and the scheme of pINK4b-IRES-ZsGreen are shown in Supplementary Fig. S4b.
Retrovirus vector infection
Preparation of retrovirus vectors was described previously20. Virus particles were added to the 2.0 × 104 of freshly prepared P7 rat OPCs that had been cultured on PDL-coated T25 flasks with 5 ml of TH-free BS medium containing 10 μg/ml of protamine sulfate. Cells were cultured in 3% O2 for 4 hours to allow virus infection, then the cells were dissociated with trypsin and re-inoculated on PDL-coated slideflasks in clonal density and cultured for 5–7 days. Cells were fixed and stained with DAPI. The number of ZsGreen+/DAPI+ cells in each clone was counted.
P14 rat optic nerve derived anti-GC-panned cells26 were stained with AlexaFluor647-labeled A2B5 antibody (#563776, BD Biosciences). Stained cells were resuspended in staining medium with 1 μg/ml PI, and sorted by SORP FACSAria (BD Biosciences). Cells were attached to glass slides by Cytospin (Thermo Scientific) and were fixed with 4% PFA-PBS fixative. Each slide was stained with following primary and secondary antibodies: Alexa Fluor488-labeled anti-Ki67 mouse monoclonal antibody (1:100; #558616, BD Biosciences), anti-Runx1 rabbit polyclonal antibodies (1:100; ab23980, Abcam), and anti-rabbit IgG-Cy3 donkey polyclonal antibodies (1:500; 711-165-152, Jackson ImmunoResearch). Samples were incubated with primary antibodies overnight in humidified chambers at 4 °C. Secondary antibody and nuclear stain DAPI were placed on sections for 2 hours at RT. Samples were mounted with PermaFluor (Labvision), and analyzed by a laser-scanning confocal microscopy (FV-10; Olympus).
Pimonidazole labeling in vivo
In vivo labeling of pimonidazole was carried out using Hypoxyprobe-1 kit (Hypoxyprobe, Inc.). P14 or P7 rats were administrated pimonodazole (60 mg/kg) via intraperitoneal injection62. Two hours later, animals were sacrificed and optic nerves were dissected within 5 minutes. 10,000 of optic nerve OPCs were suspended with 0.2 ml of TH-free BS medium and inoculated on PDL/gelatin-coated 12 mm glass base culture dishes and were cultured in 20% O2 for 90 minutes at 37 °C to allow them attaching the bottom. Cells were fixed with 4% PFA and were examined by immunocytochemistry.
In the case of the comparisons two, the data were evaluated statistically by Student’s t test. And one-way ANOVAs tests were used for multiple comparisons. Probability (P) < 0.05 was considered statistically significant. Error bars in each graph represent standard deviations (s.d.).
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We thank Martin C. Raff for comments on the manuscript. We also thank Naoharu Takano, Takehiro Yamamoto and Yoji A. Minamishima for critical advices on the experiments. We also thank Megumi Shiota, Hitoshi Tsugawa and the staffs of Collaborative Research Resources, Keio University School of Medicine for technical assistances. This study was supported by JST ERATO Suematsu Gas Biology Project. Y. Kabe and K. Takubo are supported by AMED-CREST.