Osteoclasts adapt to physioxia perturbation through DNA demethylation
2021; Springer Nature; Volume: 22; Issue: 12 Linguagem: Inglês
10.15252/embr.202153035
ISSN1469-3178
AutoresKeizo Nishikawa, Shigeto Seno, Toshitada Yoshihara, Ayako Narazaki, Yuki Sugiura, Reito Shimizu, Junichi Kikuta, Reiko Sakaguchi, Norio Suzuki, Norihiko Takeda, Hiroaki Semba, Masamichi Yamamoto, Daisuke Okuzaki, Daisuke Motooka, Yasuhiro Kobayashi, Makoto Suematsu, Haruhiko Koseki, Hideo Matsuda, Masayuki Yamamoto, Seiji Tobita, Yasuo Mori, Masaru Ishii,
Tópico(s)MicroRNA in disease regulation
ResumoArticle18 October 2021Open Access Transparent process Osteoclasts adapt to physioxia perturbation through DNA demethylation Keizo Nishikawa Corresponding Author Keizo Nishikawa [email protected] orcid.org/0000-0002-9725-1556 Laboratory of Cell Biology and Metabolic Biochemistry, Department of Medical Life Systems, Graduate School of Life and Medical Sciences, Doshisha University, Kyotanabe, Japan Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Search for more papers by this author Shigeto Seno Shigeto Seno orcid.org/0000-0003-3861-6444 Department of Bioinformatic Engineering, Graduate School of Information Science and Technology, Osaka University, Osaka, Japan Search for more papers by this author Toshitada Yoshihara Toshitada Yoshihara Department of Chemistry and Chemical Biology, Gunma University, Kiryu, Japan Search for more papers by this author Ayako Narazaki Ayako Narazaki Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Search for more papers by this author Yuki Sugiura Yuki Sugiura Department of Biochemistry, Keio University, Tokyo, Japan Search for more papers by this author Reito Shimizu Reito Shimizu Laboratory of Cell Biology and Metabolic Biochemistry, Department of Medical Life Systems, Graduate School of Life and Medical Sciences, Doshisha University, Kyotanabe, Japan Search for more papers by this author Junichi Kikuta Junichi Kikuta Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Laboratory of Bioimaging and Drug Discovery, National Institutes of Biomedical Innovation, Health and Nutrition, Ibaraki, Japan Search for more papers by this author Reiko Sakaguchi Reiko Sakaguchi WPI-Research Initiative-Institute for Integrated Cell-Material Science, Kyoto University, Kyoto, Japan Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan Search for more papers by this author Norio Suzuki Norio Suzuki Division of Oxygen Biology, Tohoku University Graduate School of Medicine, Sendai, Japan Search for more papers by this author Norihiko Takeda Norihiko Takeda Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan Search for more papers by this author Hiroaki Semba Hiroaki Semba Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan Department of Cardiovascular Medicine/Basic Research, The Cardiovascular Institute, Tokyo, Japan Search for more papers by this author Masamichi Yamamoto Masamichi Yamamoto Department of Artificial Kidneys, Graduate School of Medicine, Kyoto University, Kyoto, Japan Search for more papers by this author Daisuke Okuzaki Daisuke Okuzaki orcid.org/0000-0002-4552-783X Single Cell Genomics, Human Immunology, WPI Immunology Frontier Research Center, Osaka University, Suita, Japan Search for more papers by this author Daisuke Motooka Daisuke Motooka Genome Information Research Center, Research Institute for Microbial Diseases, Osaka University, Suita, Japan Search for more papers by this author Yasuhiro Kobayashi Yasuhiro Kobayashi Institute for Oral Science, Matsumoto Dental University, Shiojiri, Japan Search for more papers by this author Makoto Suematsu Makoto Suematsu Department of Biochemistry, Keio University, Tokyo, Japan Search for more papers by this author Haruhiko Koseki Haruhiko Koseki orcid.org/0000-0001-8424-5854 Developmental Genetics Group, Center for Integrative Medical Sciences, RIKEN, Yokohama, Japan Search for more papers by this author Hideo Matsuda Hideo Matsuda orcid.org/0000-0002-4701-3779 Department of Bioinformatic Engineering, Graduate School of Information Science and Technology, Osaka University, Osaka, Japan Search for more papers by this author Masayuki Yamamoto Masayuki Yamamoto orcid.org/0000-0002-9073-9436 Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Japan Search for more papers by this author Seiji Tobita Seiji Tobita Department of Chemistry and Chemical Biology, Gunma University, Kiryu, Japan Search for more papers by this author Yasuo Mori Corresponding Author Yasuo Mori [email protected] orcid.org/0000-0001-7180-1938 WPI-Research Initiative-Institute for Integrated Cell-Material Science, Kyoto University, Kyoto, Japan Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan Search for more papers by this author Masaru Ishii Corresponding Author Masaru Ishii [email protected] orcid.org/0000-0002-4215-007X Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Laboratory of Bioimaging and Drug Discovery, National Institutes of Biomedical Innovation, Health and Nutrition, Ibaraki, Japan Search for more papers by this author Keizo Nishikawa Corresponding Author Keizo Nishikawa [email protected] orcid.org/0000-0002-9725-1556 Laboratory of Cell Biology and Metabolic Biochemistry, Department of Medical Life Systems, Graduate School of Life and Medical Sciences, Doshisha University, Kyotanabe, Japan Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Search for more papers by this author Shigeto Seno Shigeto Seno orcid.org/0000-0003-3861-6444 Department of Bioinformatic Engineering, Graduate School of Information Science and Technology, Osaka University, Osaka, Japan Search for more papers by this author Toshitada Yoshihara Toshitada Yoshihara Department of Chemistry and Chemical Biology, Gunma University, Kiryu, Japan Search for more papers by this author Ayako Narazaki Ayako Narazaki Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Search for more papers by this author Yuki Sugiura Yuki Sugiura Department of Biochemistry, Keio University, Tokyo, Japan Search for more papers by this author Reito Shimizu Reito Shimizu Laboratory of Cell Biology and Metabolic Biochemistry, Department of Medical Life Systems, Graduate School of Life and Medical Sciences, Doshisha University, Kyotanabe, Japan Search for more papers by this author Junichi Kikuta Junichi Kikuta Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Laboratory of Bioimaging and Drug Discovery, National Institutes of Biomedical Innovation, Health and Nutrition, Ibaraki, Japan Search for more papers by this author Reiko Sakaguchi Reiko Sakaguchi WPI-Research Initiative-Institute for Integrated Cell-Material Science, Kyoto University, Kyoto, Japan Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan Search for more papers by this author Norio Suzuki Norio Suzuki Division of Oxygen Biology, Tohoku University Graduate School of Medicine, Sendai, Japan Search for more papers by this author Norihiko Takeda Norihiko Takeda Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan Search for more papers by this author Hiroaki Semba Hiroaki Semba Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan Department of Cardiovascular Medicine/Basic Research, The Cardiovascular Institute, Tokyo, Japan Search for more papers by this author Masamichi Yamamoto Masamichi Yamamoto Department of Artificial Kidneys, Graduate School of Medicine, Kyoto University, Kyoto, Japan Search for more papers by this author Daisuke Okuzaki Daisuke Okuzaki orcid.org/0000-0002-4552-783X Single Cell Genomics, Human Immunology, WPI Immunology Frontier Research Center, Osaka University, Suita, Japan Search for more papers by this author Daisuke Motooka Daisuke Motooka Genome Information Research Center, Research Institute for Microbial Diseases, Osaka University, Suita, Japan Search for more papers by this author Yasuhiro Kobayashi Yasuhiro Kobayashi Institute for Oral Science, Matsumoto Dental University, Shiojiri, Japan Search for more papers by this author Makoto Suematsu Makoto Suematsu Department of Biochemistry, Keio University, Tokyo, Japan Search for more papers by this author Haruhiko Koseki Haruhiko Koseki orcid.org/0000-0001-8424-5854 Developmental Genetics Group, Center for Integrative Medical Sciences, RIKEN, Yokohama, Japan Search for more papers by this author Hideo Matsuda Hideo Matsuda orcid.org/0000-0002-4701-3779 Department of Bioinformatic Engineering, Graduate School of Information Science and Technology, Osaka University, Osaka, Japan Search for more papers by this author Masayuki Yamamoto Masayuki Yamamoto orcid.org/0000-0002-9073-9436 Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Japan Search for more papers by this author Seiji Tobita Seiji Tobita Department of Chemistry and Chemical Biology, Gunma University, Kiryu, Japan Search for more papers by this author Yasuo Mori Corresponding Author Yasuo Mori [email protected] orcid.org/0000-0001-7180-1938 WPI-Research Initiative-Institute for Integrated Cell-Material Science, Kyoto University, Kyoto, Japan Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan Search for more papers by this author Masaru Ishii Corresponding Author Masaru Ishii [email protected] orcid.org/0000-0002-4215-007X Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan Laboratory of Bioimaging and Drug Discovery, National Institutes of Biomedical Innovation, Health and Nutrition, Ibaraki, Japan Search for more papers by this author Author Information Keizo Nishikawa *,1,2,3, Shigeto Seno4, Toshitada Yoshihara5, Ayako Narazaki3, Yuki Sugiura6, Reito Shimizu1, Junichi Kikuta2,3,7, Reiko Sakaguchi8,9, Norio Suzuki10, Norihiko Takeda11, Hiroaki Semba11,12, Masamichi Yamamoto13, Daisuke Okuzaki14, Daisuke Motooka15, Yasuhiro Kobayashi16, Makoto Suematsu6, Haruhiko Koseki17, Hideo Matsuda4, Masayuki Yamamoto18, Seiji Tobita5, Yasuo Mori *,8,9 and Masaru Ishii *,2,3,7 1Laboratory of Cell Biology and Metabolic Biochemistry, Department of Medical Life Systems, Graduate School of Life and Medical Sciences, Doshisha University, Kyotanabe, Japan 2Department of Immunology and Cell Biology, WPI-Immunology Frontier Research Center, Osaka University, Suita, Japan 3Graduate School of Medicine/Frontier Biosciences, Osaka University, Suita, Japan 4Department of Bioinformatic Engineering, Graduate School of Information Science and Technology, Osaka University, Osaka, Japan 5Department of Chemistry and Chemical Biology, Gunma University, Kiryu, Japan 6Department of Biochemistry, Keio University, Tokyo, Japan 7Laboratory of Bioimaging and Drug Discovery, National Institutes of Biomedical Innovation, Health and Nutrition, Ibaraki, Japan 8WPI-Research Initiative-Institute for Integrated Cell-Material Science, Kyoto University, Kyoto, Japan 9Department of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan 10Division of Oxygen Biology, Tohoku University Graduate School of Medicine, Sendai, Japan 11Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan 12Department of Cardiovascular Medicine/Basic Research, The Cardiovascular Institute, Tokyo, Japan 13Department of Artificial Kidneys, Graduate School of Medicine, Kyoto University, Kyoto, Japan 14Single Cell Genomics, Human Immunology, WPI Immunology Frontier Research Center, Osaka University, Suita, Japan 15Genome Information Research Center, Research Institute for Microbial Diseases, Osaka University, Suita, Japan 16Institute for Oral Science, Matsumoto Dental University, Shiojiri, Japan 17Developmental Genetics Group, Center for Integrative Medical Sciences, RIKEN, Yokohama, Japan 18Department of Medical Biochemistry, Tohoku University Graduate School of Medicine, Sendai, Japan *Corresponding author. Tel: +81 774 65 6868; E-mail: [email protected] *Corresponding author. Tel: +81 75 383 2761; E-mail: [email protected] *Corresponding author. Tel: +81 6 6879 3880; E-mail: [email protected] EMBO Reports (2021)22:e53035https://doi.org/10.15252/embr.202153035 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Oxygen plays an important role in diverse biological processes. However, since quantitation of the partial pressure of cellular oxygen in vivo is challenging, the extent of oxygen perturbation in situ and its cellular response remains underexplored. Using two-photon phosphorescence lifetime imaging microscopy, we determine the physiological range of oxygen tension in osteoclasts of live mice. We find that oxygen tension ranges from 17.4 to 36.4 mmHg, under hypoxic and normoxic conditions, respectively. Physiological normoxia thus corresponds to 5% and hypoxia to 2% oxygen in osteoclasts. Hypoxia in this range severely limits osteoclastogenesis, independent of energy metabolism and hypoxia-inducible factor activity. We observe that hypoxia decreases ten-eleven translocation (TET) activity. Tet2/3 cooperatively induces Prdm1 expression via oxygen-dependent DNA demethylation, which in turn activates NFATc1 required for osteoclastogenesis. Taken together, our results reveal that TET enzymes, acting as functional oxygen sensors, regulate osteoclastogenesis within the physiological range of oxygen tension, thus opening new avenues for research on in vivo response to oxygen perturbation. Synopsis Within the physiological range of oxygen tension, ten-eleven translocation (TET) enzymes act as functional oxygen sensors involved in osteoclastogenesis. Oxygen tension ranges from 17.4 to 36.4 mmHg in osteoclasts of live mice. Hypoxia in this range decreases TET activity of osteoclasts but does not affect energy metabolism and HIF activity. Tet2/3 regulate osteoclastogenesis by controlling Prdm1 expression via oxygen-dependent DNA demethylation. Introduction Oxygen plays an essential role in serving as the final electron acceptor for aerobic respiration and is also a substrate in reactions that generate metabolites and structural macromolecules. Since a reduction in oxygen availability (hypoxia) has physiological and pathophysiological implications (Semenza, 2002; Palazon et al, 2014), aerobic organisms such as animals and land plants adopt alternative solutions to direct their primary hypoxia responses (Hammarlund et al, 2020). A major breakthrough in understanding such hypoxia responses in animals came through the discovery of the prolyl hydroxylase (PHD)/hypoxia-inducible factor (HIF) axis (Wang & Semenza, 1993; Maxwell et al, 1999; Ohh et al, 2000; Jaakkola et al, 2001). PHD enzymes are members of the 2-oxoglutarate (2-OG)-dependent dioxygenase enzyme family and act as oxygen sensors to catalyze the oxidation of specific prolyl residues in HIF-α proteins that enable their recognition by ubiquitin ligase complexes and their subsequent degradation by the proteasome. A decrease in oxygen availability results in the attenuation of PHD-mediated hydroxylation to enable HIF-α stabilization. The stabilized HIF-α can then heterodimerize with aryl hydrocarbon receptor nuclear translocator (ARNT) to trigger a genetic program that controls angiogenesis, cell growth, and a switch to a glycolytic cell metabolism, thereby enabling cellular adaptation to hypoxia (Semenza, 2002; Wilson et al, 2020). Moreover, recent research on the hypoxia response machinery revealed the importance of other 2-OG-dependent dioxygenases whose activities also depend on oxygen availability. The DNA demethylases—ten-eleven translocation (TET) methylcytosine dioxygenases—catalyze the oxidation of 5-methylcytosine, but their activity is inhibited in severe hypoxic conditions of the tumor microenvironment, resulting in DNA hypermethylation (Laukka et al, 2016; Thienpont et al, 2016). The Jumonji-C (JmJC) domain-containing histone lysine demethylases (KDMs) also belong to the dioxygenase family of enzymes, which catalyze the oxidation of methyl groups on histones. Consequently, the hypoxia-induced decrease in the activity of KDMs leads to an altered histone methylation status, with implications for tumorigenesis (Batie et al, 2019; Chakraborty et al, 2019). These observations have raised intriguing questions as to which enzyme(s) play a pivotal role in oxygen sensing. On the basis of the Michaelis–Menten constant (KM) for oxygen, since each of the dioxygenases has a different affinity for oxygen [e.g., KDM5A and KDM6A have a KM of 117 and 260 mmHg, respectively (Batie et al, 2019; Chakraborty et al, 2019), TET2 has a KM of 39 mmHg (Laukka et al, 2016), and PHD1 has a KM ranging from 87 to 299 mmHg (Wilson et al, 2020)], the degree of changes in the reactions catalyzed by the dioxygenases largely depends on the factors that define hypoxia, which are in turn dependent on the tissue and intracellular concentration of oxygen in a given cell type. However, the extent of oxygen tension that a specific cell experiences in vivo still remains ambiguous. Bone homeostasis depends on the intimate coupling of bone resorption by osteoclasts and bone formation by osteoblasts (Takayanagi, 2007; Lorenzo et al, 2008). Bone remodeling imbalance arising because of increased osteoclast activity and/or decreased osteoblast activity leads to bone-wasting states in diseases such as osteoporosis, rheumatoid arthritis, and periodontitis (Boyle et al, 2003; Teitelbaum & Ross, 2003; Zaidi, 2007). Glycolysis is the major metabolic pathway that regulates osteoblast differentiation, which is coordinated by the HIF-α proteins (Guntur et al, 2014; Regan et al, 2014; Dirckx et al, 2018). Indeed, hypoxia affects osteogenesis at a variety of levels ranging from direct action to indirect regulation via the stimulation of angiogenesis (Wang et al, 2007; Ramasamy et al, 2014; Xie et al, 2014). Thus, hypoxia is thought to provide a favorable environment for osteoblastogenesis (Lee et al, 2017). In contrast, oxidative metabolism is an oxygen-dependent energy-producing process that is essential for osteoclast differentiation (Ishii et al, 2009; Nishikawa et al, 2015). Of note, since bone resorption driven by the massive secretion of acids and specialized proteinases is an energy-consuming process, osteoclasts require substantial oxygen to support their energy demands with respect to both differentiation and function. Thus, oxygen is likely to be an important environmental factor for osteoclast regulation; however, the in vivo oxygen tension in osteoclasts and the effect of its perturbation on osteoclastogenesis remain unclear. Several methods to assess oxygen perturbation in different tissues are available. Immunohistochemistry for pimonidazole adducts and HIF-α accumulation is widely used for analyzing hypoxia within tissue (Varia et al, 1998; Ramasamy et al, 2014), but it is neither quantitative nor indicative of oxygen tension. Electrochemical electrodes are used to measure oxygen tension in real time; e.g., the tip of micro-electrode is positioned directly in the target tissue, and the electrode in the gas analyzer measures oxygen using the collected blood. However, these invasive procedures are not suitable for the measurement of oxygen tension in hard tissue. On the other hand, non-invasive methods to quantitatively assess oxygen tension in vivo have been developed based on phosphorescence quenching (Vanderkooi et al, 1987; Rumsey et al, 1988) and magnetic resonance techniques including magnetic resonance imaging (MRI), nuclear magnetic resonance (NMR), and electron paramagnetic resonance (EPR) (Roussakis et al, 2015). These methods have both advantages and limitations in terms of applicable targets, spatial resolution, tissue permeability, convenience, and reversibility. Of these methods, optical imaging utilizing a phosphorescence probe has great advantages in determining the spatio-temporal dynamics of oxygen in both soft and hard tissues (Spencer et al, 2014; Hirakawa et al, 2015; Yoshihara et al, 2015). Here, using two-photon phosphorescence lifetime imaging microscopy with a cell-penetrating phosphorescent probe that we originally developed, we first succeeded in determining the physiological range of oxygen tension in the bone marrow at the single-cell resolution. Furthermore, we show that TET enzymes play a pivotal role in oxygen sensing and are involved in osteoclastogenesis via an epigenetic regulation mechanism within the physiological range of oxygen tension. Results Definition of physiological normoxia and hypoxia for osteoclasts in the local bone marrow environment of live mice Phosphorescence quenching by oxygen is a standard method for the sequential monitoring and non-invasive measurement of oxygen concentration (Vanderkooi et al, 1987; Rumsey et al, 1988). We recently developed a method to assess intracellular oxygen tension using a cell-penetrating phosphorescent probe (BTPDM1) based on the Ir (III) complex and a bifurcated fiber system (Hirakawa et al, 2015). The current study used a two-photon laser scanning microscope (TPLSM) equipped with time-correlated single-photon counting (TCSPC) system, to assess combined intensity-lifetime imaging of phosphorescence in live mice. This technique was named two-photon phosphorescence lifetime imaging microscopy (2PLIM). Prior to imaging, BTPDM1 was injected into TcirgGFP/+ mice, their osteoclast lineage cells of which express green fluorescence protein (GFP) (Sun-Wada et al, 2009). GFP fluorescence enabled osteoclasts to be visualized while the phosphorescence lifetime was simultaneously measured using the TCSPC system (Fig 1A and B). The phosphorescence lifetime of mice osteoclasts during air inhalation was 2.98 ± 0.02 μs (Fig 1C). In order to calculate oxygen tension of osteoclasts via phosphorescence lifetime, we prepared a calibration curve showing the relationship between phosphorescence lifetime and oxygen concentration (Fig EV1A and B). We cultured primary osteoclasts treated with BTPDM1 and measured phosphorescence lifetime under various oxygen concentrations. The results indicated that the in vivo oxygen tension of osteoclasts was 36.4 ± 0.7 mmHg (4.8 ± 0.1%; Figs 1D and EV1C). Next, in order to determine the physiological range of oxygen tension in osteoclasts, we measured the oxygen tension of osteoclasts in mice during hypoxic air inhalation (approximately 14% oxygen to reduce arterial oxygen saturation [spO2] to 55%). As a result, phosphorescence lifetime of osteoclasts in these mice was prolonged to 3.62 ± 0.03 μs, which in turn was estimated as 17.4 ± 0.6 mmHg (2.3 ± 0.1%) of oxygen tension (Figs 1E and EV1C). These results enabled us to introduce a physiological context to oxygen tension of osteoclasts in vivo, i.e., ca. 5% as physiological normoxia (referred to as physioxia) in the mice whose spO2 is 98% during air inhalation and ca. 2% as physiological hypoxia in the mice whose spO2 reduced to 55% during inhalation of hypoxic air containing 14% oxygen. Figure 1. Measurement of passive physiological range of oxygen tension (pO2) of osteoclasts A. Schematic diagram of the two-photon intravital imaging and 2PLIM. B. Representative intravital image of calvarial bone marrow of Tcirg1EGFP/+ male mice treated with BTPDM1 showing osteoclasts (left, EGFP fluorescence) and 2PLIM image for pO2 changes (right, phosphorescence lifetime of BTPDM1). Scale bar, 20 μm. C. Phosphorescence lifetime in each osteoclast of calvarial bone marrow of mice upon exposure to ambient air (n = 91 from six mice). Data denote mean ± s.e.m. D, E. Change in pO2 of osteoclasts in the mice upon exposure to various concentrations of oxygen from 21% to 14% pO2. Magnified PLIM images of osteoclasts under different peripheral oxygen saturations (SpO2, D). The pO2 of each osteoclast in these mice was plotted (E, n = 47 from three mice for each SpO2). Scale bar, 20 μm. Data denote mean ± s.e.m. **P < 0.01 (ANOVA). F. Effect of physioxia (5% pO2) or physiological hypoxia (2% pO2) on osteoclastogenesis. TRAP-stained cells (left panel) and the number of TRAP-positive cells with more than three nuclei (right). Scale bar, 100 μm. Data denote mean ± s.e.m. **P < 0.01 (n = 8 biological replicates; t-test). Download figure Download PowerPoint Click here to expand this figure. Figure EV1. Reciprocal plot of phosphorescence lifetime and oxygen concentration A. PLIM images of in vitro-cultured osteoclasts under different conditions of oxygen concentration. Scale, 50 μm. B. The phosphorescence quenching due to dissolved oxygen in solution can be examined by the Stern–Volmer equation. The approximate line was constructed by a straight-line approximation, and an approximation formula and the coefficient of determination are shown. C. Phosphorescence lifetime in each osteoclast of the calvarial bone marrow of mice upon inhalation of normal (21% pO2) and hypoxic (14% pO2) air. The phosphorescence lifetime of each osteoclast was plotted (right, n = 47 from three mice for each SpO2). Data denote mean ± s.e.m. **P < 0.01 (ANOVA). Download figure Download PowerPoint Studies that have been conducted so far on the effect of hypoxia on osteoclast differentiation indicate that oxygen is a negative regulator of osteoclast differentiation (Arnett et al, 2003; Fukuoka et al, 2005; Murata et al, 2017). Osteoclast differentiation was evaluated in vitro by counting multinucleated cells (MNCs) positive for the osteoclast marker, tartrate-resistant acid phosphatase (TRAP), following stimulation of bone marrow-derived monocyte/macrophage precursor cells (BMMs) with receptor activator of nuclear factor-κB ligand (RANKL), and in the presence of macrophage colony-stimulating factor (M-CSF)(Nishikawa et al, 2010; Hayashi et al, 2012). We confirmed that the formation of TRAP-positive MNCs was significantly enhanced when they were cultured in hypoxic conditions of 10% oxygen, compared with that under atmospheric air conditions (Appendix Fig S1A). However, our 2PLIM revealed that these culture conditions do not reflect in vivo oxygen levels. In order to mimic in vivo oxygen tension, we investigated the effect of hypoxia on osteoclast differentiation by performing an in vitro experiment either in 5% oxygen (physioxia) or 2% oxygen (hypoxia). The experiment produced results contrary to those of previous results. The formation of TRAP-positive MNCs was severely impaired under 2% oxygen compared with that under 5% oxygen (Fig 1F and Appendix Fig S1B). Nevertheless, neither the number of proliferative BMMs nor the apoptotic activities of BMMs were affected in both 5% and 2% oxygen (Appendix Fig S1C and D). These results suggest that oxygen is a positive regulator of osteoclast differentiation. The effect of hypoxia on bone metabolism was studied in vivo by studying an experimental mouse hypoxia models generated under acute (oxygen inhalation at a low concentration) conditions. Mice exposed to hypoxia displayed high bone mass phenotypes with low numbers of osteoclasts, while no obvious difference in the number of osteoblasts was observed (Fig 2A–C). Although the hypoxia signaling pathway is involved in the promotion of osteoblastogenesis (Wang et al, 2007), our experimental conditions, wherein the mice were exposed to 14% oxygen for 10 days, did not affect osteoblastogenesis. Taken together, these results suggest that hypoxia causes high bone mass due to defective osteoclast differentiation. Figure 2. Effect of oxygen deprivation on bone metabolism A. μCT analysis of the femurs of 10-week-old male mice upon exposure to low oxygen concentration (14% pO2, n = 5) or ambient air (n = 7), and of 10-week-old Hif1aRank–/–; Hif2aRank–/– male mice upon exposure to low oxygen concentration (14% pO2, n = 6) or ambient air (n = 5) for 10 days (axial view of the metaphyseal region). Scale, 0.5 mm. B. Histological analysis of the proximal tibias of 10-week-old male mice upon exposure to low oxygen concentration (14% pO2) or ambient air for 10 days, and of 10-week-old Hif1aRank–/–; Hif2aRank–/– male mice upon exposure to low oxygen concentration (14% pO2) or ambient air (toluidine blue staining [arrow, osteoclasts]). Scale, 100 μm. C. Parameters for osteoclasts and osteoblasts during bone morphometric analysis of 10-week-old male mice under low oxygen concentration (n = 3) or ambient air (n = 5), and 10-week-old Hif1aRank–/–; Hif2aRank–/– male under low oxygen concentration (n = 3) or ambient air (n = 5) for 10 days. Data denote mean ± s.e.m. *P < 0.05; NS, not significant (t-test). Download figure Download PowerPoint HIF-α is dispensable for the osteoclast response to physioxia perturbation The above-stated findings raised the necessity for elucidating the mechanism(s) underlying oxygen-induced regulation o
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