
==== Front
Mol Metab
Mol Metab
Molecular Metabolism
2212-8778
Elsevier

S2212-8778(24)00143-1
10.1016/j.molmet.2024.102012
102012
Original Article
Mitochondrial protein deacetylation by SIRT3 in osteoclasts promotes bone resorption with aging in female mice
Richardson Kimberly K. 128
Adam Gareeballah Osman 128
Ling Wen 128
Warren Aaron 12
Marques-Carvalho Adriana 12
Thostenson Jeff D. 13
Krager Kimberly 4
Aykin-Burns Nukhet 4
Byrum Stephanie D. 57
Almeida Maria 126
Kim Ha-Neui hkim@uams.edu
12⁎9
1 Center for Musculoskeletal Disease Research, USA
2 Division of Endocrinology, Department of Internal Medicine, USA
3 Department of Biostatistics, USA
4 Division of Radiation Health, Department of Pharmaceutical Sciences, USA
5 Department of Biochemistry and Molecular Biology, USA
6 Department of Orthopedic Surgery, University of Arkansas for Medical Sciences, USA
7 Arkansas Children's Research Institute, Little Rock, AR, USA
⁎ Corresponding author. University of Arkansas for Medical Sciences, Division of Endocrinology and Metabolism, 4301 W. Markham St. #587, Little Rock, 72205-7199, USA. hkim@uams.edu
8 Kimberly K. Richardson, Gareeballah Osman Adam and Wen Ling contributed equally to this work.

9 Lead contact.

16 8 2024
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© 2024 The Author(s)
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https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Objectives

The mitochondrial deacetylase sirtuin-3 (SIRT3) is necessary for the increased bone resorption and enhanced function of mitochondria in osteoclasts that occur with advancing age; how SIRT3 drives bone resorption remains elusive.

Methods

To determine the role of SIRT3 in osteoclast mitochondria, we used mice with conditional loss of Sirt3 in osteoclast lineage and mice with germline deletion of either Sirt3 or its known target Pink1.

Results

SIRT3 stimulates mitochondrial quality in osteoclasts in a PINK1-independent manner, promoting mitochondrial activity and osteoclast maturation and function, thereby contributing to bone loss in female but not male mice. Quantitative analyses of global proteomes and acetylomes revealed that deletion of Sirt3 dramatically increased acetylation of osteoclast mitochondrial proteins, particularly ATPase inhibitory factor 1 (ATPIF1), an essential protein for mitophagy. Inhibition of mitophagy via mdivi-1 recapitulated the effect of deletion of Sirt3 or Atpif1 in osteoclast formation and mitochondrial function.

Conclusions

Decreasing mitophagic flux in osteoclasts may be a promising pharmacotherapeutic approach to treat osteoporosis in older adults.

Highlights

• Targeted deletion of Sirt3 in osteoclast-lineage cells attenuates skeletal aging in female but not male mice.

• SIRT3 is required for maintenance of healthy mitochondria in osteoclast cells of aged mice.

• Pharmacological inhibition of mitophagy is sufficient to mimic the effects of conditional loss of Sirt3 on osteoclastogenesis.

• PINK1 is dispensable for osteoclast maturation and skeletal aging in mice.

• SIRT3-target proteins that may be responsible for the stimulation of bone resorption.

Keywords

SIRT3
Mitochondria
Osteoclasts
Aging
Acetylation
Proteomics
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pmc1 Introduction

Osteoclasts are giant, multinucleated cells that are responsible for the resorption of bone tissue, a task that requires the secretion of protons and collagenolytic enzymes that dissolve the bone matrix. During adulthood, bone resorption coupled to bone formation rejuvenates the skeleton and is required to maintain bone strength [1]. With aging, the amount of bone resorbed by osteoclasts is not fully restored with new bone deposited by osteoblasts and this excessive bone resorption leads to loss of bone mass and increases fracture risk [2]. Nonetheless, the mechanisms causing the elevated bone resorption in aged bone remain unclear.

Osteoclasts differentiate from hematopoietic stem cell precursors of the myeloid lineages in response to receptor activator of NF-κΒ ligand (RANKL), the indispensable cytokine for bone resorption. The most important source of RANKL for adult bone remodeling are cells of the mesenchymal lineage in particular osteocytes. Nonetheless, other cytokines produced by a variety of cells of both hematopoietic and mesenchymal lineage can also stimulate bone resorption. The binding of RANKL to its membrane receptor RANK activates multiple signaling pathways to promote differentiation, cell fusion, bone resorptive activity, and survival [[3], [4], [5]]. RANKL also stimulates mitochondrial function in osteoclasts, most likely in preparation for the highly energetic task of resorbing bone [3,4]. Indeed, during differentiation from mononuclear macrophage precursors to multinucleated osteoclast, the number and size of mitochondria increase [[6], [7], [8], [9]]. Mitochondria biogenesis, function, and quality control are critical for osteoclasts differentiation and resorptive activity under physiologic and pathophysiologic conditions [[8], [9], [10], [11], [12], [13], [14]].

The health of mitochondria is regulated by processes that include fusion and fission of the tubular mitochondrial network, mitochondrial biogenesis, and the elimination of unwanted mitochondria by mitophagy (i.e., mitochondria-specific autophagy) [15,16]. In mammalian cells, there are four major types of mitophagy: PINK1/PARKIN-mediated mitophagy, receptor-mediated mitophagy, lipid-receptor mediated mitophagy, and piecemeal mitophagy [17]. While PINK1/PARKIN mainly regulates ubiquitin-dependent mitophagy, receptor-mediated mitophagy can be activated in different cellular contexts by numerous autophagy receptors constitutively located at the outer mitochondrial membrane, including Bnip3 and Nix. We still do not understand the molecular mechanisms that control mitochondrial quality and metabolic function in osteoclasts or how changes to these mechanisms result in bone disease.

We and others have shown that the NAD-dependent mitochondrial protein deacetylase Sirtuin-3 (SIRT3) is required for the increase in bone resorption with estrogen deficiency and aging [18,19]. In many tissues, SIRT3 promotes mitochondrial quality control and attenuates the development of multiple age-related metabolic diseases [[20], [21], [22], [23]]. Some of these effects have been attributed to the deacetylation of specific mitochondrial proteins [24,25]. Indeed, altered lysine acetylation of mitochondrial proteins is recognized as an important metabolic regulatory mechanism [24,25]. We have previously proposed that the pro-resorptive effects of Sirt3 are due to its role in promoting mitophagy and, thereby, maintaining the proper functioning of mitochondria [18]. However, our previous work was performed with mice with global loss of Sirt3. Therefore, we cannot rule out the possibility that the skeletal effects are secondary to changes in hormones or other metabolic derangements. Furthermore, the mitochondrial proteins and specific lysine residues that are targeted by Sirt3 have not been examined in bone cells.

To investigate whether SIRT3 in osteoclasts contributes to skeletal aging, we generated a mouse model in which Sirt3 was specifically deleted in the myeloid lineage, including pre-osteoclasts and osteoclasts. Using quantitative proteomics, we also identified lysine acetylation targets of SIRT3 in osteoclast mitochondria.

2 Results

2.1 Targeted deletion of Sirt3 in osteoclast-lineage cells attenuates age-associated bone loss in female mice

We first examined the mRNA expression and enzymatic activity of SIRT3 in cultured osteoclasts obtained from C57BL/6 wild-type mice at 6 or 16 months of age. Notably, we found that Sirt3 activity but not mRNA or protein expression was dramatically increased in aged mice (Figure 1A–C). To investigate the role of SIRT3 in osteoclast-lineage cells, we generated mice with conditional deletion of Sirt3 in the myeloid lineage including pre-osteoclasts and osteoclasts. To this end, Sirt3f/f mice were crossed with mice expressing the Cre recombinase under the control of Lysozyme M promoter. For our subsequent work, we used Sirt3 conditional knockout mice (Sirt3ΔLysM) and littermate controls (Sirt3f/f). Sirt3ΔLysM mice appeared normal at birth and were indistinguishable from littermate controls (Sirt3f/f) with respect to body size (data not shown), weight, and femoral length during adulthood (Figures S1A and S1B).Figure 1 Age-related bone loss requires osteoclast SIRT3 in female mice. (A) mRNA expression, (B) protein levels, and (C) enzymatic activity of SIRT3 in cultures of bone marrow-derived macrophages (BMMs) from 6- or 16-month-old female C57BL/6 mice treated with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. (D–J) A cohort of female Sirt3ΔLysM mice and Sirt3fl/fl littermate controls were aged to 16 months. Another cohort of female mice of the same genotypes euthanized at 6 months of age served as young controls. (D) Sequential bone mass density (BMD) measurements in the aging cohort by dual-energy x-ray absorptiometry (n = 19–20 animals/group). (E–F) Imaging and quantification of femoral bones from female Sirt3ΔLysM mice and wild-type littermates by micro-CT after sacrifice (n = 12–20 animals/group). Scale bar: 100 μm. (G–H) Imaging and quantification of cortical porosity at the distal metaphysis of the femur by micro-CT (n = 17–19 animals/group). Scale bar: 100 μm. (I–J) Imaging and quantification of vertebral bones from female Sirt3ΔLysM mice and wild-type littermates by micro-CT after sacrifice (n = 12–20 animals/group). Scale bar: 100 μm. Line and error bars represent mean ± SD. P values were determined using (A–C) Student's t-test or (F–J) 2-way ANOVA. Interaction terms generated by 2-way ANOVA are shown below each graph. (D) P values were determined using ∗, Student's t-test or #, repeated-measures ANOVA between Sirt3ΔLysM mice and Sirt3fl/fl littermate controls in the same age.

Figure 1

To confirm efficient and specific deletion of Sirt3 in Sirt3ΔLysM mice, we cultured bone marrow-derived macrophages (BMMs) and bone marrow-derived stromal cells isolated from female mice. In macrophages from Sirt3ΔLysM mice, the mRNA levels and activity of Sirt3 were about 91% and 54% lower, respectively, compared to littermate controls (Figures S2A and S2B). In contrast, stromal cells, did not show a significant reduction in Sirt3 mRNA (Figure S2A). Other tissues, such as brain and liver showed no changes in Sirt3 levels between the two genotypes (Figure S2A). While SIRT3 is the most abundant sirtuin in mitochondria, an increase in the other mitochondrial sirtuins (SIRT4 and SIRT5) may compensate for the decrease in Sirt3. We did not detect any changes in the expression of Sirt4 and Sirt5 in BMMs lacking Sirt3 (Figure S2C).

We next aged Sirt3ΔLysM and littermate control mice up to 16 months. Serial DXA bone mineral density (BMD) analysis revealed that female control mice attained peak bone mass at about 6 months and exhibited age-related bone loss in the femur beginning at 14 months of age and in the spine at 12 months of age (Figure 1D). These findings are in line with previous studies describing skeletal aging in C57BL/6 female mice [26]. BMD in Sirt3ΔLysM mice was indistinguishable from controls up to 12 month of age. However, conditional deletion of Sirt3 attenuated the loss of both femur and spine BMD with aging, although this effect more pronounced in the femur (Figure 1D). Analysis of the bone microarchitecture after sacrifice using micro-CT, confirmed that femoral cortical bone (Figure 1E–F and S3A–S3B) and spinal trabecular bone (Figure 1H–I and S3C–S3F) were indistinguishable between the two genotypes at 6 months. As expected, control mice had lower femoral cortical thickness at 16 than at 6 months of age (Figure 1E,F). While bones enlarge with aging, as revealed by an increase in both total and medullary area of the femur (Figure S3C and S3D), a disproportionate increase in medullary area relative to the increase in total area causes cortical thinning. The age associated loss of cortical bone was greatly attenuated in Sirt3ΔLysM mice (Figure 1E,F). Aging also causes an increase in intracortical bone remodeling and cortical porosity [27]. The cortical porosity that develops with age seen in control mice was prevented in Sirt3ΔLysM mice (Figure 1G,H). Control mice also exhibited a decrease in trabecular bone volume and 3D-BMD at the spine between 6 and 16 months of age (Figure 1I–J and S3F). This low bone mass was associated with a decrease in trabecular number and increase in trabecular spacing, whereas no changes were detected in trabecular thickness (Figures S3C–S3E). Female Sirt3ΔLysM mice also lost trabecular bone in the spine with age but to a lesser extent than control mice (25% vs 41%) (Figure 1I,J). In contrast to the female mice, the conditional deletion of Sirt3 in osteoclasts of male mice had no effect on the age-related loss of cortical and trabecular bone mass measured by DXA (Figure S4A) or micro-CT (Figures S4B–S4H). Taken together, these results indicate that targeted deletion of Sirt3 in osteoclast-lineage cells attenuates skeletal aging in female but not male mice.

2.2 Deletion of Sirt3 in osteoclast-lineage cells suppresses bone resorption in aged female mice

The preservation of endocortical bone in the 16-month-old female Sirt3ΔLysM mice suggested that the Sirt3 knockout mice were protected from the increase in endocortical bone resorption that occurs with advancing age [27,28]. Histomorphometric quantification of osteoclasts revealed that 16-month-old female Sirt3ΔLysM mice showed no differences in osteoclast or osteoblast number compared to control mice (Figure 2A–C), as seen in Sirt3 null mice [18]; however, mice lacking Sirt3 in osteoclast-lineage cells had lower serum levels of C-terminal telopeptide of type 1 collagen (CTx), a bone resorption marker (Figure 2E) and lower expression of the active osteoclast-specific markers Tartrate-resistant acid phosphatase (TRAP or TRAPase), also called acid phosphatase 5 (Acp5), DC-stamp, and Integrin B3 (Itgb3) (Figure 2D). These results indicate that osteoclast function is decreased at the endocortical surface in the absence of SIRT3 in the osteoclast-lineage cells, and this allows for the preservation of bone. The conditional deletion of Sirt3 did not affect the serum levels of N-terminal propeptide of type I procollagen (P1NP), a bone formation marker (Figure 2E). In contrast to the female mice, the conditional deletion of Sirt3 in osteoclasts of male mice had no effect on the serum levels of CTx (Figure S4I).Figure 2 Conditional deletion of Sirt3 in LysM-Cre–targeted cells decreases bone resorption in aged female mice. (A–C) Number of osteoclasts (N.Oc/B.Pm) (A), osteoclast surface (Oc.S/BS) (B) and number of osteoblasts (N.Ob/B.Pm) (C) per endocortical bone surface of nondecalcified femur sections stained for TRAPase activity from 16-month-old female Sirt3ΔLysM mice and wild-type littermates (n = 17–20 animals/group). (D) qPCR of mRNA isolated from L1 vertebrae (n = 6 animals/group) obtained from 16-month-old female mice. (E) Serum concentration of a collagen degradation product (CTx) and N-terminal propeptide of type I procollagen (P1NP) in 16-month-old female Sirt3ΔLysM mice and wild-type littermates by ELISA (n = 17–20 animals/group). (F–H) BMMs were isolated from 16-month-old female Sirt3ΔLysM mice and wild-type littermates and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 5 days. (F) Representative pictures (left) and number (right) of TRAP+ multinucleated osteoclasts generated from BMMs (triplicates of pooled cultures). Scale bar: 500 μm. (G) Representative pictures (left) and resorbed areas (right) of Von Kossa–stained bone biomaterial surface (triplicates). Scale bar: 500 μm. The resorbed areas appear white, and the areas of the mineralized surface that were not resorbed appear black. (H) Osteoclast marker levels in mRNA of cultured osteoclasts measured by qPCR (triplicate cultures). Line and error bars represent mean ± SD. P values were determined using Student's t-test. All measures were performed in cultured BMMs pooled from 4 to 5 mice/group and repeated at least twice.

Figure 2

BMM cultures from female Sirt3ΔLysM mice formed smaller osteoclasts and had less resorbed area indicating decreased osteoclast maturation and function (Figure 2F,G), similar to the findings obtained when using cells from Sirt3-knockout mice [18]. The expression of osteoclast differentiation markers was also decreased in cultured osteoclasts from aged Sirt3ΔLysM mice (Figure 2H).

2.3 Targeted deletion of Sirt3 in osteoclast-lineage cells causes mitochondrial dysfunction

Osteoclasts from aged Sirt3-null mice exhibit impaired mitochondrial respiration and mitophagy [18]. Here we examined whether these effects are due to cell intrinsic actions of Sirt3 by quantifying mitochondrial activity in cultures of BMMs from 16-month-old female Sirt3ΔLysM mice and age-matched controls. Mitochondrial respiration and ATP-linked respiration were strongly diminished with conditional deletion of Sirt3 (Figure 3A,B). Deletion of Sirt3 also decreased the oxygen consumption rate associated with proton leak (Figure S5A), indicating a disruption of electron flow through the electron transport chain. Maximum respiration (i.e., maximal electron transport chain activity), reserve respiratory capacity (i.e., the difference between maximum oxygen consumption rate and basal oxygen consumption rate), and nonmitochondrial respiration were also significantly diminished with deletion of Sirt3 (Figures S5B–S5D).Figure 3 Conditional deletion of Sirt3 in LysM-Cre–targeted cells decreases osteoclast mitophagy in aged female mice. BMMs were isolated from 16-month-old female Sirt3ΔLysM mice and wild-type littermates and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. (A–B) Mitochondrial and ATP-linked respirations per cell, in osteoclasts, were measured by Seahorse (n = 8–14 wells/group). (C) MitoSOX to evaluate mitochondrial ROS production (n = 6 wells/group). (D) Representative pictures of osteoclasts stained with JC-1 (left), and quantification of mitochondrial membrane potential (right) (triplicates of pooled cultures). Scale bar: 500 μm. (E) Ultrastructural appearance of mitochondria of osteoclasts by TEM (left) and the number of cristae formation per mitochondria section (right) (triplicates of pooled cultures). (F) ATP levels (expressed as relative light units, RLU). (G–H) Representative mitochondrial protein levels by Western blot (left) and expression levels as the indicated ratio (right) (triplicates of pooled cultures). (I) Representative PINK1 acetylation levels by Western blot (left) and expression levels as the indicated ratio (right) (triplicate cultures). (J) Tfam levels in mRNA of cultured osteoclasts measured by qPCR (triplicate cultures). Line and error bars represent mean ± SD. P values were determined using Student's t-test. All measures were performed in cultured BMMs pooled from 4 to 5 mice/group and repeated at least twice.

Figure 3

We next determined whether deletion of Sirt3 in osteoclast-lineage cells is sufficient to induce mitochondrial dysfunction. To do so, we quantified mitochondrial ROS and mitochondrial membrane potential and found that deletion of Sirt3 increased ROS and decreased mitochondrial membrane potential (Figure 3C,D). These features are commonly observed when damaged mitochondria accumulate due to lack of mitophagy [[29], [30], [31]]. Abnormal mitochondria morphology is a well-established determinant of mitochondrial dysfunction [32]. Using transmission electron microscopy, we found that osteoclasts from female Sirt3ΔLysM mice had smaller mitochondria with diminished cristae structure compared to those from littermate controls (Figure 3E). Consistent with this, deletion of Sirt3 decreased ATP levels (Figures 3F).

We next examined whether targeted deletion of Sirt3 in osteoclast-lineage cells could replicate the effects of global deletion of Sirt3 on mitochondrial dynamics and mitophagy. Optic atrophy 1 (Opa1), an essential GTPase responsible for fusion of the mitochondrial inner membrane [33], promotes mitochondrial quality control via fusion-dependent mechanisms, as well as stimulation of mitophagy via fusion-independent mechanisms [34,35]. Protein levels of Opa1 were decreased in the Sirt3 conditional knockout osteoclasts (Figure 3F), as were the levels of Mitofusin 2 (Figure 3G). As seen in Sirt3 null mice [18], markers for receptor-mediated mitophagy such as Bnip3 and Nix were reduced (Figure 3H), and acetylation of the ubiquitin-mediated mitophagy marker PINK1 was increased (Figure 3I). In addition, conditional deletion of Sirt3 had a mild effect on mRNA levels of Tfam (Figure 3J), the transcription factor that contributes to mitochondria biogenesis [36]. Taken together, these results suggest that SIRT3 is required for maintenance of healthy mitochondria in osteoclast cells of aged female mice.

2.4 Inhibition of mitochondrial division and mitophagy replicates the effects of Sirt3 deletion in osteoclasts

To further examine the contribution of mitochondria dynamics and mitophagy in osteoclasts, we used mdivi-1, a pharmacological inhibitor of mitochondrial fission and mitophagy [[37], [38], [39], [40], [41], [42], [43]], in BMMs from 24-month-old female C57BL/6 mice. Mdivi-1 decreased the protein levels of Bnip3 and Nix (Figure 4A). This effect was associated with decreased mitochondrial respiration (Figure 4B–G). In the presence of mdivi-1, BMMs formed smaller osteoclasts that resorbed much less bone when cultured on Osteo Assay surface plates, and this effect was dose dependent (Figure 4H,I). To test how a block on mitophagy affects RANKL-induced osteoclastogenesis, we added mdivi-1 into cultures under 2 conditions: (1) mdivi-1 was present in culture throughout a 5-day period and (2) mdivi-1 was added on day 3.5 and kept in culture for the remainder of the 5-day experiment. We found that the amount of resorption was identical regardless of whether cultures were exposed to mdivi-1 for 5 days or mdivi-1 was added on day 3.5 (Figure 4I). These observations indicate that during the RANKL-induced osteoclastogenesis, mdivi-1 acts on late stage of osteoclast differentiation, not on early osteoclast progenitors. Consistent with this, addition of mdivi-1 decreased expression of late/terminal osteoclast differentiation markers in cultures of osteoclasts from aged mice (Figure 4J). These results indicate that pharmacological inhibition of mitophagy is sufficient to mimic the effects of conditional loss of Sirt3 on osteoclastogenesis in female mice.Figure 4 Inhibition of mitophagy by mdivi-1 mimics the effects of Sirt3 deletion in aged osteoclasts. Osteoclasts developed in cultures of BMMs from 24-month-old female C57BL/6 mice treated with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 (A–G) or 5 days (H–J) in the presence of mdivi-1 (10 μM). (A) Representative mitochondrial protein levels by Western blot (left) and expression levels as the indicated ratio (right) (triplicate cultures). (B–G) Different fractions of mitochondrial and nonmitochondrial respirations per cell, in osteoclasts, measured by Seahorse (n = 14–16 wells/group). (H) Representative pictures (left) and number (right) of TRAP+ multinucleated osteoclasts (triplicate cultures). Scale bar: 500 μm. (I) Representative pictures (left) and resorbed areas (right) of Von Kossa–stained bone biomaterial surface (quadruplicate cultures). Scale bar: 500 μm. (J) Osteoclast marker levels in mRNA of cultured osteoclasts measured by qPCR (triplicate cultures). Line and error bars represent mean ± SD. P values were determined using Student's t-test (A–G) or 1-way ANOVA (H–J). All in vitro assays were performed in cultured BMMs pooled from more than 3 mice and repeated at least twice.

Figure 4

2.5 The age-associated bone loss occurs independently of PINK1 in female mice

SIRT3 induction of PINK1/PARKIN mitophagy contributes to the health of several tissues including liver, heart, and eye [[44], [45], [46]]. This evidence, together with our current findings, suggest that PINK1 mediates the stimulation of mitophagy by SIRT3 in osteoclasts during skeletal aging. To address this hypothesis, Pink1 knockout and littermate wild-type female mice were aged up to 16 months. Consistent with previous studies [47], Pink1 knockout mice appeared normal at birth and were indistinguishable from wild-type mice with respect to body weight and femoral length (Figures S6A and S6B). Both Pink1 knockout and wild-type mice lost bone mass with age (Figure 5A–G). Pink1 knockout mice lost cortical bone at the femur with age (16.1%) to a larger extent than wild-type control mice (9.9%) (p-int = 0.03). This difference was due to higher but not significant cortical bone mass in 6-month-old Pink1 null mice compared to controls of the same age (Figure 5B). Likewise, deletion of Pink1 had no effect on the RANKL-induced osteoclast formation and the expression of osteoclast markers in cultured BMMs of aged mice (Figure 5H,I). Furthermore, the protein levels of the mitophagy markers Bnip3 and Nix were unaffected by deletion of Pink1 in cultured osteoclasts (Figure 5J). Overall mitochondrial and non-mitochondrial respiration were not changed by deletion of Pink1 (Figures 5K–L and S7A–S7D). These results suggest that PINK1 is dispensable for osteoclast maturation and skeletal aging in female mice.Figure 5 Deletion of Pink1 does not affect age-related bone loss. A cohort of female Pink1 knockout mice and littermate controls were aged to 16 months. Another cohort of female mice of the same genotypes euthanized at 6 months of age served as young controls. (A–B) Imaging and quantification of femoral bones from female Pink1 knockout mice and littermate controls by micro-CT after sacrifice (n = 7–20 animals/group). (C–G) Imaging and quantification of vertebral bones from female Pink1 knockout mice and littermate controls by micro-CT after sacrifice (n = 7–20 animals/group). (H–I) BMMs were isolated from 16-month-old female Pink1 knockout mice and littermate controls and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 4 (H–I) or 3 days (J–L). (H) Pink1 and osteoclast marker levels in mRNA of cultured osteoclasts measured by qPCR (triplicate cultures). (I) Number of TRAP+ multinucleated osteoclasts (quadruplicate cultures). (J) Representative mitophagy protein levels by Western blot. (K–L) Mitochondrial and ATP-linked respirations per cell, in osteoclasts, were measured by Seahorse (n = 12–15 wells/group). Line and error bars represent mean ± SD. P values were determined using 2-way ANOVA (B–G) or Student's t-test (H–I and K–L). Interaction terms generated by 2-way ANOVA are shown below each graph. All in vitro measures were performed in cultured BMMs pooled from 4 to 5 mice/group and repeated at least twice.

Figure 5

2.6 Osteoclasts that lack Sirt3 exhibit hyperacetylation of mitochondrial proteins

We next searched for targets of SIRT3 that could mediate its effects on mitochondria and mitophagy. First, by immunoblotting with a pan acetyl-lysine antibody, we examined the total protein acetylation in both mitochondria and cytosol isolated from the cultured BMMs of 24-month-old female C57BL/6 mice. Lysine acetylation was higher in mitochondria versus cytoplasmic fractions (Figure 6A). Mitochondrial protein acetylation was downregulated during osteoclastogenesis as shown in BMM cultures with or without RANKL (Figure 6B). As expected, deletion of Sirt3 increased protein acetylation as determined in cultured osteoclasts from aged female Sirt3ΔLysM mice (Figure 6C). There was also a slightly higher level of protein acetylation in bone tissue from Sirt3ΔLysM mice (Figure 6D).Figure 6 Deletion of Sirt3 induces hyperacetylation of mitochondrial proteins in aged osteoclasts. (A–C) BMMs were isolated from 24-month-old female C57BL/6 mice (A–B) or 16-month-old female Sirt3ΔLysM mice and littermate controls (C) and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. Representative blots showing the acetylation status of mitochondrial proteins in whole cell lysates by Western blot (left) and expression levels as the indicated ratio (right) (triplicate cultures). (D) Western blot analysis of protein isolated from femoral bone shafts (n = 4 animals/group) obtained from 16-month-old female mice. All Western blot represents a minimum of 3 independent experiments. (E–J) BMMs were isolated from 16-month-old female Sirt3 null mice and littermate controls and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. (E–G) Quantitative analysis of the global proteome was performed with label-free tandem mass spectrometry (triplicate cultures). The fold change in total proteins is presented in volcano plots for the effects of RANKL treatment or genotype (E). Top 5 highly up- or down-regulated pathways (RANKL vs Vehicle) (F–G). (H–J) A comprehensive acetylome analysis was performed with 3 samples per group as described in the flow chart (H). The fold change in hyperacetylated proteins is presented in volcano plots for the effects of RANKL treatment or genotype (I). Top 10 hyper-acetylated proteins (Sirt3 KO vs WT). Asterisk indicates published sites (J). Line and error bars represent mean ± SD. P values were determined using Student's t-test.

Figure 6

Mass spectrometry was used to analyze the global proteome of macrophages and osteoclasts cultures from 16-month-old Sirt3 null and control mice. Of a total of 4,400 identified proteins, 387 were mitochondrial proteins. As expected, macrophages and osteoclasts exhibited major differences among the proteins identified (Figure S8) with RANKL greatly increasing DC-Stamp, Atp6v0d2, Itgb3, CathK, and TRAP, known RANKL target proteins (Figures 6E and S9A). Some of mitophagy regulators previously identified including Cluh [48,49], Aco2 [50], and Sdha [51] were significantly increased in osteoclasts compared to BMMs (Figure 6E). Gene Ontology analysis also indicated that RANKL altered several metabolic pathways including oxidative phosphorylation and Parkinson's disease directly related to mitochondrial complexes (Figure 6F). In turn, deletion of Sirt3 decreased proteins and pathways related to mitochondria function, including NDUF and COX family as well as oxidative phosphorylation and Parkinson's disease (Figures S10 and 6G). The overall fold change in protein levels was low when comparing cells with and without Sirt3 (Figure 6E), suggesting that post translational modifications (acetylation) may be responsible for the mitochondrial dysfunction in osteoclasts of Sirt3 knockout mice.

Lysine acetylome analysis (Figure 6H) identified a total of 567 acetylated peptides with a fold change of >2.5 and a P value of <0.05. In line with the RANKL activation of SIRT3-mediated mitophagy in osteoclasts, lysine acetylation levels of mitophagy regulators, Senp3 [52] and Galectin-1 [53], decreased by about 8.9- or 4.45-fold during osteoclastogenesis, respectively (Figure 6I). While RANKL enhanced 69 acetyl-lysine peptides in wild-type cells, deletion of Sirt3 resulted in hyperacetylation of 305 peptides, 18 of which were mitochondrial proteins with significant hyperacetylation at multiple sites (Figure 6I–J). Some of these proteins have been previously identified as substrates of SIRT3 including ATP5O [22]. The largest fold changes in lysine acetylation were observed in proteins linked to ATP synthase (Figure 6I–J), the last complex of the electron transport chain. In particular, ATPIF1 showed the highest levels of acetylation (3.9- to 213-fold) at 5 lysine residues (K49, K52, K62, K83, and K103). We confirmed the increase in lysine acetylation of ATPIF1 by coimmunoprecipitation in osteoclast lysates from Sirt3 knockout female mice (Figure S11A). Interestingly, the physical interaction between SIRT3 and ATPIF1 increases with age in osteoclasts (Figure S11B). We generated ATPIF1 mutant constructs with lysine residues corresponding to acetylation sites mutated to arginine to prevent acetylation (Figure S11C). Notably, activation of ATPIF1 deacetylation at lysine 49 in RAW 264.7 cells augmented the formation of giant osteoclasts, akin to those observed in cultured osteoclasts from aged mice. This suggests that SIRT3-mediated deacetylation of ATPIF1 is indispensable for osteoclastogenesis during skeletal aging.

We next examined the relevance of the top 5 protein targets of SIRT3 to osteoclastogenesis. To this end, each of ATPIF1, ATP5O, HMGCL, HSPE1, and MTHFD1L were knockdown in BMMs from female wild-type mice using shRNA. Silencing of Atpif1 and Hspe1 impaired, Atp5o and Hmgcl had no effect, and Mthfd1l increased osteoclast maturation (Figures 7A and S12). Because in some cells ATPIF1 is important for mitophagy [[54], [55], [56]], we focused on this protein. In line with an important role for this protein in osteoclasts, levels of ATPIF1 increased by about 2-fold during osteoclastogenesis (Figure 7B). We confirmed that knockdown of Atpif1 decreased resorption and osteoclast markers (Figure 7C). These effects were associated with a decrease in basal, ATP-linked respiration and proton leak (Figure 7D–F). The levels of Bnip3 and Nix were also decreased, indicating defective mitophagy (Figures 7G). Knockdown of Hspe1, a chaperonin related to mitochondrial unfolded protein responses [57], did not affect the expression of mitophagy markers in osteoclasts (not shown).Figure 7 SIRT3-induced ATPIF1 deacetylation activates osteoclast mitophagy in aged mice. (A–E, not B) BMMs from 24-month-old female C57BL/6 mice transduced with lentiviral vectors expressing control sh-RNA or sh-RNAs targeting the hyperacetylated proteins and cultured with RANKL for 5 days (A and C) or 3 days (D–G). (A) Representative pictures (left) and number (right) of TRAP+ multinucleated osteoclasts and Von Kossa–stained bone biomaterial surface. (triplicate cultures). Scale bar: 500 μm. (B) BMMs were isolated from 24-month-old female C57BL/6 mice and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. Western blot analysis of expression of ATPIF1. (triplicate cultures). (C) Levels of mRNA of an osteoclast marker in cultured osteoclasts measured by qPCR. (triplicate cultures). (D–F) Different fractions of mitochondrial respiration per cell measured with Seahorse (n = 10–14 wells/group). (G) Western blot analysis of expression of mitophagy marker. (triplicate cultures). Line and error bars represent mean ± SD. P values were determined using Student's t-test. All in vitro assays were performed in cultured BMMs pooled from more than 3 mice and repeated at least twice.

Figure 7

Together, these results demonstrate that Sirt3 deficiency significantly changes the acetylome of osteoclasts, and indicate the SIRT3-target proteins that may be responsible for the stimulation of bone resorption.

3 Discussion

Loss of bone mass is one of the most common features of human aging, but the mechanisms responsible remain unclear. Herein, we provide evidence that SIRT3 in osteoclasts promotes bone resorption and loss of bone mass in female but not male mice, thereby contributing to skeletal aging. Skeletal aging is characterized by a decrease in bone formation in both cortical and trabecular bone. In long bones, an increase in osteoclast number is also a major contributor to cortical thinning and intracortical porosity with aging [27,58]. In contrast, osteoclast number does not increase with aging in trabecular bone. Deletion of Sirt3 in osteoclasts does not alter osteoclast number but decreases bone resorption. This, along with the findings that the loss of both trabecular and cortical bone was attenuated by Sirt3 deletion, strongly suggests that an increase in osteoclast resorptive activity, independent of changes in osteoclast number, contributes to skeletal aging. While the reasons for the sexual dimorphic role of SIRT3 remain unclear, it is possible that an increase in osteoclast activity with aging occurs predominantly in females. Support for this idea comes from evidence that the magnitude of intracortical bone remodeling with aging is higher in female than male mice [27]. Although the Sirt3 gene was expected to be specifically deleted in osteoclast lineage cells, there could be off-target expression of Cre in estrogen-producing cells. Alternatively, the loss of Sirt3 in osteoclasts may lead to changes in certain secreted factors that could affect estrogen levels, potentially contributing to the bone phenotype in females. We measured estrogen levels in 16-month-old female Sirt3ΔLysM mice and their WT littermate controls and found that the conditional deletion of Sirt3 in osteoclasts had no effect on estrogen levels (Figure S13), indicating that the sexually dimorphic effects on the skeletal phenotype were not due to differences in estrogen levels in female mice.

When comparing our current finding with the previous results obtained when we aged Sirt3 knockout mice [18], we noticed that the bone protective effect conferred by Sirt3 deletion was of higher magnitude in the Sirt3 knockout mice. Indeed, in the previous study the loss of cortical bone in male mice was attenuated by Sirt3 deletion, in contrast to the lack of an effect in the present study. Likewise, no loss of endocortical bone was seen in female Sirt3 knockout mice up to 16 month of age while in the Sirt3ΔLysM female mice the loss of endocortical bone was attenuated. These findings suggest that in addition to macrophages and osteoclast, SIRT3 actions in other cell types contribute to the pro-resorptive effects seen in Sirt3 knockout mice. Further genetic studies are required to elucidate whether SIRT3 expression within the bone microenvironment such as in mesenchymal, endothelial, or hematopoietic lineage cells could be promoting bone resorption in the aged skeleton.

Our previous findings, along with the present results, show that SIRT3 promotes mitochondrial homeostasis in osteoclasts. Mitochondria are a central node of the effects of RANKL on osteoclast maturation and bone resorption [[6], [7], [8], [9]]. Additionally, we demonstrate that the suppression of SIRT3-mediated mitochondrial homeostasis increases oxidative stress in aged osteoclast precursors. Mitochondrial ROS in osteoclasts contributes to physiological bone resorption [13]. We have previously identified signaling pathways used by RANKL to enable osteoclasts to accumulate ROS [13,14]. However, osteoclasts from aged Sirt3ΔLysM mice exhibit much higher mitochondrial ROS production than cells from wild-type mice (Figure 3). This is associated with a decrease in mitochondrial membrane potential and a disorganized morphology of mitochondria. Excessive levels of ROS cause numerous cellular derangements and cell death [59,60]. Similar to our findings, the global deletion of SIRT3 increases oxidative stress in aged hematopoietic stem cells [61]. However, while the increase in ROS due to SIRT3 deletion enhances the number of hematopoietic stem cells in old mice, the excessive ROS in osteoclasts from Sirt3ΔLysM mice is associated with decreased bone resorptive activity. These different outcomes are most likely related to different harmful versus beneficial thresholds for ROS in these cell types. The increase in ROS with age in bone [26] might explain why the skeletal effects of SIRT3 are observed in old but not in young mice. The higher levels of ROS with aging might elicit mitochondrial protective mechanisms and antioxidant systems, such as those provided by SIRT3, which are not necessary in young animals when the levels of ROS are lower. Support for this idea is provided by findings that SIRT3 stimulates mitochondrial oxidative metabolism in response to fasting and membrane depolarization [22,23].

Similar to our findings in osteoclasts, deletion of Sirt3 is associated with accumulation of damaged mitochondria in different cell types [[62], [63], [64]], and some studies suggest that mitophagy is reduced in cells that lack Sirt3 [[65], [66], [67]]. Damaged or dysfunctional mitochondria are degraded via different forms of mitophagy including ubiquitin-mediated mitophagy via the PINK1and PARKIN pathway and receptor-mediated mitophagy mediated by BNIP3 and NIX. Several studies have reported that BNIP3 and NIX also act in PINK1/PARKIN-mediated mitophagy [68]. In osteoclasts lacking Sirt3, we found increased acetylation of PINK1 and decreased protein levels of BNIP3 and NIX. These findings, along with the increase in ROS a well-established inducer of mitophagy, and the presence of damaged mitochondria suggest that mitophagy is compromised in the absence of Sirt3. We found that acetylated PINK1 accumulates in Sirt3-deficient osteoclasts and that this is associated with reduced levels of BNIP3 and NIX [18] (Figure 3), suggesting that deacetylation of PINK1 by SIRT3 promotes mitophagy. Similar changes in PINK1 were observed in Sirt3-deficient cardiomyocytes [69]. Nonetheless, a functional role for PINK1 has been seen in brain and heart but not pancreas and skeletal muscle [47,70]. Similar to SIRT3, the actions of PINK1 become apparent under stress conditions, such as chronic mitotoxicity or aging [[71], [72], [73]]. Nonetheless, we found that skeletal aging was similar between control and Pink1 knockout mice. These results indicate that PINK1 is not a critical mediator of the effects of Sirt3 in osteoclasts and skeletal aging. It remains possible that other mitophagy pathways independent of PINK1 are involved in osteoclast mitochondrial quality control and the excessive bone resorption that occurs with aging. Although we propose that SIRT3 regulates osteoclast maturation and function through mitophagy (primarily by measuring protein levels of its markers), there is still a need to better understand the direct role of SIRT3 in mitochondrial quality control. One of the long-standing limitations in the field is the lack of specific markers or direct measurements for mitophagy. Recently, researchers have developed mitophagy-specific reporter mice to elucidate the role of mitochondrial proteins in different tissues [[74], [75], [76]]. Using this mouse model to study skeletal aging and the contribution of osteoclast mitophagy to pathophysiological conditions would be worthwhile.

Proteomics studies using tissues from Sirt3 knockout mice have elucidated the array of acetylation and sirtuin interactions in mitochondria [77,78]. Our results from the SIRT3 acetylation targets in osteoclasts are in line with these previous studies and implicate oxidative phosphorylation and mitochondrial translation as the major pathways targeted by SIRT3. These findings suggest that the decrease in oxygen consumption rate seen in Sirt3 deficient osteoclasts result from direct inhibition of electron transport chain proteins. Our in vitro functionality assay revealed that knockdown of Atpif1 and Hspe1, two out of the top five deacetylation targets of Sirt3, recapitulated the effects of Sirt3 deletion in inhibiting osteoclast maturation. We focused our attention on ATPIF1 because of its role in multiple aspects of mitochondria function including mitophagy. Via the regulation of ATP synthase activity, ATPIF1 controls both the bioenergetics and structure of mitochondria [[79], [80], [81]]. ATPIF1 also promotes cell survival by inhibiting key events of intrinsic apoptosis including decreased mitochondrial fission, permeabilization, and cytochrome c release [82,83]. Under stress conditions, ATPIF1 blocks ATPase activity and maintains high cellular ATP levels to promote mitophagy. We found that knockdown of Atpif1 in BMMs suppressed markers of mitophagy in vitro (Figure 7). Together, these results suggest that ATPIF1 is a major mediator of the actions of SIRT3 in osteoclasts. Interestingly, deletion of SIRT3 prevents RANKL-induced ATP levels in aged mice (Figure 3). Based on the demonstrated role of ATPIF1 in blocking ATPase activity and maintaining high cellular ATP levels [[54], [55], [56]], especially under stress conditions, it is possible that the SIRT3-ATPIF1 signaling pathway can directly affect energy metabolism through ATP production in osteoclasts. Moreover, the levels of ATPIF1 increased with osteoclast differentiation (Figure 7). ATPIF1 gene promoter analysis and ChiP-seq data have revealed interactions with transcription factors such as c-Myc, c-Fos and NFκB [80,84]. These signaling pathways are activated by RANKL [85,86]. In line with this, we found that mitochondrial respiratory chain inhibitors, such as rotenone [12], antimycin A, or oligomycin (Figure S14), inhibit osteoclast formation. Future studies using mouse models and genetic manipulations of ATPIF1 should elucidate the role of this protein or ATP production in bone resorption and in mediating SIRT3 actions. For example, to test the hypothesis that the deacetylation of ATPIF1 contributes to mitophagy and the resorptive activity of osteoclasts, it would be worthwhile to generate new mouse models in which endogenous ATPIF1 is replaced with acetylation mutant proteins in cells expressing LysM-Cre.

In conclusion, we show that SIRT3-mediated lysine deacetylation of a variety of mitochondrial proteins, including ATPIF1 is of critical importance to mitochondrial quality control and the increases in bone resorption with aging. Many lines of evidence obtained using Sirt3 knockout mice with physiologic aging or using models of disease indicate that SIRT3 delays aging-associated disease in many tissues including neuroinflammation [87], hepatic steatosis [78], glucose intolerance [77], and dysregulated nutrient sensing [88]. This has led to the idea that stimulation of SIRT3 could represent an anti-aging strategy. Our previous and current findings that within bone cells SIRT3 has a predominant role in osteoclasts and, thereby, contributes to bone loss with aging indicates that SIRT3 activation could be detrimental to the aged bone. To the best of our knowledge, the signaling pathways that operate downstream of SIRT3 and their contributions to the increased bone resorption that occurs with aging, have not been examined. Our mitochondria protein hyperacetylation atlas provides a key resource for understanding osteoclast mitochondria metabolism. The elucidation of the mechanisms that modulate mitochondria homeostasis in bone cells may guide the development of therapies to attenuate skeletal aging.

4 Materials and methods

4.1 Animal experiments

The Sirt3 global knockout mice were generated and maintained as previously described [18] in our facility using pairs of mice heterozygous for the Sirt3 null allele. Pink1 knockout mice were obtained from the Jackson Laboratory (stock no. 017946). Mice with conditional deletion of Sirt3 in the myeloid lineage were generated by a two-step breeding strategy. Male homozygous LysM-Cre transgenic mice (The Jackson Laboratory; stock no. 004781) were crossed with Sirt3 floxed (f/f) mice (C57BL/6 genetic background) (The Jackson Laboratory; stock no. 031201) to generate mice heterozygous for the Sirt3 floxed allele and the Cre allele. Male Sirt3f/+;ΔLysM and female Sirt3f/f mice were intercrossed to generate Sirt3f/f and Sirt3ΔLysM mice. Offspring were genotyped by PCR using the following primer sequences: Sirt3-flox primer #1: 5′ CTGGCTTTGGGTTTAAGCAG 3′ and primer #2: 5′ GGAGGCTGAGGCTAAAGA GC 3′. To determine whether SIRT3 in osteoclasts plays a role in age-related bone loss, mice were aged up to 16 months. Six-month-old mice of the same genotypes were used as young adult controls.

Offspring from all genotypes were tail-clipped for DNA extraction at the time of sacrifice or weaning (21 days) and then group-housed with same-sex littermates. Genomic DNA extracted from tail samples was used for PCR-based genotyping following the protocols from the Jackson Laboratory. All mice used in this study were housed under standard laboratory conditions with a 12-hour dark, 12-hour light cycle, a constant temperature of 23 °C, and humidity of 48%. A standard rodent diet (Envigo, Teklad 22/5) containing 22% protein, 1.13% calcium, and 0.94% phosphorus was provided to mice ad libitum. For the aging study, at 8 months of age, the mice were switched to Teklad 2014 rodent diet (Envigo) containing 14% protein and 4% fat, and mice were given acidified water ad libitum. Investigators were blinded to study groups during animal handling and endpoint measurements. All procedures were approved by the Institutional Animal Care and Use Committees of the University of Arkansas for Medical Sciences and the Central Arkansas Veterans Healthcare System.

4.2 DXA and micro-CT

Bone mineral density (BMD) measurements were performed by dual-energy X-ray absorptiometry (DXA) using a PIXImus densitometer (GE Lunar) on mice sedated with 2% isoflurane. Data were analyzed as previously described [18]. Scans of the entire left femur or lumbar spine were used for the measurement of BMD. A micro-CT40 scanner from Scanco Medical was used to measure bone microarchitecture as previously described [18,89]. For trabecular bone measurements, the vertebrae (L5) and the metaphysis of left femur were used, whereas cortical bone was measured at the left femoral diaphysis (midpoint of the bone length as determined in scout view). Prior to analysis, the bones were dissected, cleaned, fixed in Millonig's phosphate buffer (Leica Biosystems), and gradually dehydrated in 100% ethanol.

4.3 CTx, P1NP, and estrogens ELISA

Blood was collected into 1.7 mL EDTA-coated microcentrifuge tubes by retro-orbital bleeding. The blood was then kept on ice for 1 h and centrifuged at 6,150 g at 4 °C for 10 min to separate serum from cells before analysis. Circulating CTx, P1NP, and estrogens in serum were measured using a mouse RatLaps (CTx-I) EIA kit (Immunodiagnostic Systems), Rat/Mouse PINP EIA kit (Immunodiagnostic Systems), and mouse estrogen ELISA kit (Abcam), respectively, according to the manufacturer's directions.

4.4 Bone histology

Freshly dissected lumbar vertebrae (L5) and left femurs were fixed overnight in 10% Millonig's neutral-buffered formalin with 5% sucrose, followed by dehydration with ethanol and storage in 100% ethanol until embedding in methyl methacrylate (Sigma–Aldrich) and longitudinal sectioning (5 μm thickness). To measure the static indices of osteoclast number and surface, the sections were stained for tartrate-resistant acid phosphatase (TRAP) with the Leukocyte Acid Phosphatase (TRAP) Kit (Sigma–Aldrich) and counterstained with toluidine blue (Sigma–Aldrich). Briefly, deplasticized sections were stained with TRAP solution (a mixture of Fast Garnet GBC Base Solution, Sodium Nitrite Solution, Napthol AS-BI Phosphate Solution, Acetate Solution, and Tartrate Solution) for 1 h and 45 min at 37 °C. Using the OsteoMeasure Analysis System (OsteoMetrics, Inc), the following dynamic measurements were made: total perimeter (B.Pm), single label perimeter (sL.Pm), double label perimeter (dL.Pm), and mineral apposition rate (MAR). The following values were then calculated: mineralizing surface (MS/BS = [1/2sL.Pm + dL.Pm]/B.Pm × 100; %); bone formation rate (BFR/BS = MAR × MS/BS; μm2/μm/day). Histomorphometry determinations of the trabecular bone were restricted to the secondary spongiosa. One section per sample was analyzed by a histopathologist blinded to the study groups. The terminology used in this study is that which is recommended by the Histomorphometry Nomenclature Committee of the American Society for Bone and Mineral Research [90].

4.5 Cell culture

Bone marrow macrophages (BMMs) were obtained as described previously [12,13]. Briefly, total bone marrow cells were harvested from the tibiae and femora of mice. After depleting red blood cells with ACK buffer (0.01 mM EDTA, 0.011 M KHCO3, and 0.155 M NH4Cl, pH 7.3), cells were plated in α-MEM complete medium containing 10% fetal bovine serum (FBS), 1% penicillin and streptomycin (PSG), and 10 ng/mL M-CSF (R&D Systems) in 10-cm tissue culture plates overnight. Non-adherent cells were plated in Petri dishes (10 cm) and cultured with 10 mL α-MEM complete medium supplemented with Macrophage colony stimulating factor (M-CSF) (30 ng/mL) for 4 days to obtain BMMs, which were used as osteoclast precursors. To obtain mRNA and protein, cells were then plated in 6-well plates at a density of 0.3 × 106 cells per well with 2 mL of α-MEM complete medium with 30 ng/mL M-CSF and 30 ng/mL RANKL (R&D Systems) for the indicated number of days. To obtain TRAP-positive mature osteoclasts, cells were plated in 48-well plates at a density of 0.5 × 104 cells per well with the same medium described above. Cultured osteoclasts were fixed with 10% neutral-buffered formalin for 10 min and stained for TRAP. A Leukocyte Acid Phosphatase Assay Kit (Sigma–Aldrich) was used for the TRAP staining, and the manufacturer's instructions were followed. Cells were plated at minimum in triplicate for all TRAP staining assays. To examine the effects of inhibition of mitophagy and mitochondrial respiratory chain on osteoclastogenesis, BMMs were isolated from 24-month-old female C57BL/6 mice and cultured with 30 ng/mL M-CSF and 30 ng/mL RANKL in the presence or absence of 10 μM mdivi-1 (Sigma–Aldrich), 5 nM Antimycin (Sigma–Aldrich) and 0.1 μM Oligomycin (Sigma–Aldrich). Sirt3 activity was measured using a Sirt3 Activity Assay Kit (Abcam, Cambridge, United Kingdom) according to the manufacturer's directions as described previously [91]. For stromal cell cultures, total bone marrow cells were obtained as described above. Cells from 4 to 5 mice of each group were pooled and cultured with 20% FBS, 1% PSG, and 50 μg/mL ascorbic acid (Sigma–Aldrich) in 10-cm culture dishes for 7 days. Adherent bone marrow stromal cells were trypsinized and replated in 12-well tissue culture plates at 0.2 × 106 cells per well with 10% FBS, 1% PSG, 50 μg/mL of ascorbic acid, and 10 mM β-glycerophosphate (Sigma–Aldrich) for 3 days to perform qPCR assays.

4.6 Bone resorption assay

BMMs were isolated as described above and stimulated with RANKL to form osteoclasts on Osteo Assay Surface 24-well plates (Corning Life Sciences) coated with an inorganic bone biomaterial surface. The osteoclasts were removed using a 2% hypochlorite solution for 5 min, washed with distilled water, and dried at room temperature. To identify the resorbed area, Von Kossa staining was performed in the wells. In the dark, wells were treated with 150 μL per well of 5% (w/v) aqueous silver nitrate solution for 20 min. Plates were then washed for 5 min with distilled water and incubated in the dark with 150 μL per well of 5% (w/v) sodium carbonate in a 10% formalin solution. Wells were then washed twice with PBS, rinsed with distilled water, and dried in a 50 °C oven for 30 min. Four wells per group were assessed microscopically. In this assay, the resorbed areas appear white, and the areas that are not resorbed appear black.

4.7 Assessment of mitochondrial membrane potential

The relative mitochondrial membrane potential was determined using the lipophilic cationic probe 5, 5′, 6, 6′-tetrachloro-1,1′,3,3′-tetraethylbenzimdazol-carbocyanine iodide (JC-1; Molecular Probes). The cultured osteoclasts were incubated in the dark with 7.5 μM JC-1 for 30 min. Fluorescence was observed using a Nikon Eclipse 800 microscope with a 60× water immersion objective equipped with a dual filter for fluorescein and rhodamine. Green signals are indicative of the monomeric form of JC-1 (i.e., lower membrane potential), and the red signals correspond to the aggregate form (i.e., higher membrane potential). Thus, yellow signals indicate relatively normal membrane potential (i.e., combined aggregate and monomer). Average mean intensity fluorescence was calculated using the ratio of JC-1 aggregate to monomer (590 nm/530 nm) using the Nikon NIS Elements software. All images were captured with equal exposure times.

4.8 Mitochondrial superoxide production

The mitochondrial-specific probe MitoSOX Red Mitochondrial Superoxide Indicator (M36008, Thermo Fisher Scientific) was used to evaluate mitochondrial superoxide anion levels. BMMs were seeded in a 96-well black microplate and treated with RANKL for 3 days. Following treatment, the culture medium was replaced with assay medium containing 3 μM of MitoSOX. Cells were then incubated for 30 min at 37 °C. Subsequently, a kinetic assay was performed for 120 min to evaluate the MitoSOX oxidation rate, measured by fluorescence (λex/λem = 510/580 nm), using a Cytation™ 5 microplate reader. The measurements were normalized using Hoechst staining.

4.9 TEM for visualizing mitochondrial morphology and measurement of mitochondrial cross-sectional area

Cultured osteoclasts were washed three times in PBS and fixated in 2.5% glutaraldehyde in 0.1M sodium cacodylate buffer, at room temperature. Cells were post stained with 1% osmium tetroxide/0.8% K3Fe(CN)6 in buffer, followed by 1% tannic acid and 0.5% uranyl acetate, both in water. Next, cells were dehydrated in graded ethanol series followed by 100% propylene oxide and rotated overnight in 50% PO/resin before embedded in 100% resin. 50-nm-thick sections were collected on copper mesh grids and post stained with uranyl acetate and lead citrate. Images were taken at 19000× magnification on a Tecnai F20 (FEI) at 80 kV. The mitochondrial cross-sectional area was measured on images (19000× magnification) by Image J software using the previously reported method [92].

4.10 ATP levels

Intracellular ATP levels were measured using a luciferin-luciferase-based assay with the CellTiter-Glo® Luminescent Cell Viability Assay (G7570, Promega), following the manufacturer's protocol. Briefly, bone marrow macrophages were cultured in a 96-well white–wall tissue culture plate with 30 ng/mL RANKL for 3 days. The culture media were then replaced with 100 μl of assay reagent (CellTiter-Glo Buffer and CellTiter-Glo Substrate). Each extracted sample was mixed for 2 min on an orbital shaker to promote cell lysis, followed by a 10-minute incubation at room temperature. The luminescence signal was monitored using a Cytation™ 5 microplate reader.

4.11 Mitochondrial respiration and cellular bioenergetics

Bone marrow macrophages were plated in Seahorse XF96 plates and treated with 30 ng/mL RANKL for the indicated number of days to generate pre-osteoclasts with or without 10 μM mdivi-1. The medium in the wells was replaced with XF assay medium (Agilent), and the plates were kept in a non-CO2 incubator for 20 min at 37 °C. After recording 3 total cellular respiration measurements with the XF96 analyzer, 10 μg/mL oligomycin (Sigma–Aldrich) was added to inhibit mitochondrial ATP synthase and measure the decrease in the oxygen consumption rate that is linked to ATP turnover. An oxidative phosphorylation uncoupler, 5 μM FCCP (Sigma–Aldrich), was used to determine the maximal respiration potential of the cultured cells. The amount of non-mitochondrial oxygen consumption was assessed by inhibiting the electron respiratory chain activity with a 10 μM antimycin A (Sigma–Aldrich) and 10 μM rotenone cocktail (Sigma–Aldrich). These data were used to calculate the mitochondrial basal respiration, ATP-linked respiration, reserve respiratory capacity, and proton leak as previously described [91].

4.12 Quantitative RT-PCR

Total RNA was purified from cultured bone marrow macrophages, stromal cells, soft tissues, and vertebral bones (L1) with TRIzol reagent (ThermoFisher Scientific) according to the manufacturer's directions. RNA quantity and 260/280 ratio were determined using a NanoDrop instrument (Thermo Fisher Scientific). Complementary DNA (cDNA) was reverse transcribed from 1 to 2 μg of total RNA using a High-Capacity cDNA Reverse Transcription kit (Applied Biosystems) according to the manufacturer's instructions. PCR was performed using a custom TaqMan Gene Expression Assays manufactured by Applied Biosystems. The primers and probes for murine Sirt3 (Mm00452131_m1), Acp5 (Mn00475698_m1), Ctsk (Mm00484039_m1), Itgb3 (Mm00443980_m1), Tfam (Mm00447485_m1), Pink1 (Mm00550827_m1), DC-Stamp (Mm04209236_m1), Sirt4 (Mm01201915_m1), and Sirt5 (Mm00663723_m1) were manufactured by the TaqMan Gene Expression Assays service (Applied Biosystems). Relative mRNA expression levels were normalized to the house-keeping gene ribosomal protein S2 (Mm00475528_m1) using the ΔCt method [93].

4.13 Western blot analysis

Mature osteoclast-enriched femoral cortical bone was prepared by removing the ends of femurs, flushing the bone marrow by centrifugation, and removing surface cells by scraping with a scalpel. Bone fragments were immediately frozen in liquid nitrogen and pulverized. Cultured cells were washed twice with ice-cold PBS. Proteins were extracted with a buffer containing 20 mM Tris–HCL, 150 mM NaCl, 1% Triton X-100, protease inhibitor mixture, and phosphatase inhibitor cocktail (Sigma–Aldrich) on ice for 30 min. The cell lysates were centrifuged at 13,200 rpm for 15 min at 4 °C, and the supernatants were collected in new tubes. The protein concentration of cell lysates was determined using a DC Protein Assay kit (Bio-Rad). Protein samples (20–40 μg per sample) were subjected to polyacrylamide gel electrophoresis on 8%–15% SDS-PAGE gels and transferred electrophoretically onto polyvinyl difluoride membranes (Merck Millipore). The membranes were blocked in 5% fat-free milk/Tris-buffered saline for 120 min and incubated with a primary antibody followed by a secondary antibody conjugated with horseradish peroxidase. We used rabbit polyclonal antibodies for OPA1 (1:1000, Abcam, #ab42364). Rabbit monoclonal antibodies against SIRT3 (1:1000, Cell Signaling, #5490), VDAC (1:1000, Cell Signaling, #4661), Nix (1:1000, Cell Signaling, #12396) were used to detect SIRT3 and the other representative mitochondria proteins. Mouse monoclonal antibodies against Mitofusin2 (1:1000, Abcam, ab56889), Bnip3 (1:1000, Abcam, ab10433), and PINK1 (1:500, Santa Cruz Biotechnology, sc-517353) were used to detect their corresponding protein. We also used rabbit polyclonal antibodies for Ac-Lysine (1:1000, Cell Signaling, #9441) and ATPIF1 (1:1000, Cell Signaling, # 8528). Polyclonal antibodies against Ac-PINK1 were produced in rabbits in cooperation with Creative Biolabs Inc (1:1000). Ac-PINK1 antibodies bind to one of the 2 acetylated lysines or both [18]. Blots were stripped and re-probed with anti-β-actin antibody (Santa Cruz Biotechnology; sc-81178, 1:2000). Bound antibodies were detected with ECL reagents (Millipore) and imaged and quantified with a VersaDoc™ imaging system (Bio-Rad).

4.14 Examination of global proteome and lysine acetylome

Cultures of pre-osteoclasts were prepared by flushing whole bone marrow cells as described above from femurs and tibias of 16-month-old Sirt3 global or conditional knockout and littermate control mice. Samples were analyzed by the UAMS Proteomics Core using tandem mass tag with an Orbitrap Lumos mass spectrometer. Briefly, total protein samples from cultured osteoclasts were prepared for digestion using the filter-assisted sample preparation (FASP) method [94] and used for quantitative analysis of the global proteome. Approximately 100 μg of the digested peptides were desalted using C18 stop-and-go extraction tips. In a separate set of experiments, mitochondria-enriched fractions from cultured osteoclasts with equal protein content (5 mg) were trypsinized and subjected to acetyl-lysine immunocapture using the PTMScan Acetyl-Lysine Motif kit (Cell signaling technology) in cooperation with Cell Signaling Technology [[95], [96], [97]]. Peptides were purified with a 0.7-mL C18 Sep-Pak column, and eluted peptides were lyophilized. Acetylated peptides were enriched with the anti-acetyl immunoaffinity beads provided in the kit. Liquid chromatography was performed and interfaced to a Quadrupole-Orbitrap™ Mass Spectrometer (Q-Exactive™; Thermo Fisher Scientific) via nano-electrospray ionization as previously described [98]. The mass spectrometer was programmed to acquire by data-dependent acquisition and tandem mass spectra from the top 20 ions in the full scan from 350 to 1,600 m/z.

4.15 Quantitative analyses of proteomic data

For isobaric tandem mass tag (TMT) protein analysis, proteins and reporter ions were identified and quantified using MaxQuant (Max Planck Institute) with MS3 quantitation. Scaffold Q + S (Proteome Software) was used to verify MS/MS-based peptide and protein identifications. Protein identifications were accepted if they could be established with less than 1% false discovery rate and contained at least 2 identified peptides. Protein probabilities were assigned by the Protein Prophet algorithm [99] to perform reporter ion-based statistical analysis. TMT MS3 reporter ion intensity values were log2 transformed and missing values were imputed by a normal distribution for each sample using Perseus (Max Planck Institute). TMT batch effects were removed using ComBat [100] in order to correct for technical variation due to multiplexing samples across multiple TMT10plex batches. Statistical analysis was performed using Linear Models for Microarray Data (limma) with empirical Bayes (eBayes) smoothing to the standard errors [101]. Proteins with an FDR adjusted p-value <0.05 and a fold change >2 were considered to be significant.

Proteins with an FDR adjusted p value < 0.05 and a fold change >2.0 were considered significant. Significant proteins were used to identify important protein networks and pathways using the Ensemble of Gene Set Enrichment Analyses (EGSEA) Bioconductor package and Qiagen's Ingenuity Pathway Analysis [102]. Volcano plots were obtained to demonstrate the effects of Sirt3 deletion and RANKL treatment. Also, protein sets that were significantly up- or downregulated and/or hyper- or hypoacetylated were screened to detect protein–protein binding recognized by Sirt3 deletion.

4.16 Co-immunoprecipitation

Cultured osteoclasts were washed twice with ice-cold PBS and lysed on ice in a buffer containing 20 mM Tris, pH 7.5, 50 mM NaCl, 0.1% NP-40, 2 mM EDTA, and protease/phosphatase inhibitors, incubated on ice for 30 min, and then cleared by centrifugation at 13,200 rpm for 15 min at 4 °C. A total of 0.8 mg of protein was incubated with 1.5 μg of the primary antibody against ATPIF1 (Cell Signaling, # 8528) overnight at 4 °C with rotation. Then the samples were incubated with 50 μl of beads (Thermo Fisher Scientific) for 2 h at 4 °C. Samples were further fractioned by SDS-PAGE and analyzed by Western blotting as described above. Mouse monoclonal antibodies against SIRT3 (1:2000, Santa Cruz Biotechnology, sc-365175) were used to detect SIRT3 proteins.

4.17 ATPIF1 K49R mutant

ATPIF1 mutant constructs with lysine residues at acetylation sites mutated to arginine (K49R) to prevent acetylation at K49 were generated in cooperation with Biocytogen (Figure S11C). Raw 264.7 cells were seeded in 6-well plates 24 h before transfection. Cells were incubated with ATPIF1-wild type (WT) and ATPIF1 K49R Mutant plasmids for 20 min at room temperature in serum-free medium containing Lipofectamine Plus reagent (Invitrogen), according to the manufacturer's instructions. Cells were then washed and incubated in α-MEM complete media for 24 h, followed by the addition of 30 ng/mL RANKL for 5 days.

4.18 Lentiviral transduction of bone marrow macrophages

Lentiviral particles for ATPIF1 (sc-141374-v), ATP5O (sc-76010-v), HMGCL (sc-146051-v), HSPE1 (sc-40655-v), and MTHFD1L (sc-149679-v) were purchased from Santa Cruz Biotechnology, Inc for osteoclast differentiation assays. Non-targeted shRNA lentiviral particles (sc-108080) were used as a shRNA control. Whole bone marrow cells were obtained from 24-month-old female C57BL/6 mice as described above. Twenty-four hours later, non-adherent cells were submitted to a Ficoll-Hypaque gradient (Sigma–Aldrich), and cells at the interface were cultured in the presence of 30 ng/mL M-CSF, 8 μg/mL polybrene (Santa Cruz Biotechnology), and the lentiviral particles for 16 h. We further cultured the infected bone marrow macrophages with M-CSF for 24 h and then added 2 μg/mL puromycin (Santa Cruz Biotechnology) for 48 h to remove uninfected cells.

4.19 Statistics

For spine and femur BMD data in Figures 1D and S4A, repeated-measures ANOVA models were considered with main effects of genotype and month, interactions of the main effects, month as categorical or continuous variables, with a possible quadratic term. After examining significance of effects and model fit statistics, models with categorical effects of genotype, month, and their interaction were chosen for both models. These analyses were done with SAS v9.4. All other data were analyzed using GraphPad Prism 9 (GraphPad Software). Statistically significant treatment effects were detected with a 2-way ANOVA after determining that the data were normally distributed and exhibited equivalent variances. In some cases, log or rank transformations were used to obtain normally distributed data and equal variance. This was followed by pairwise comparisons using Tukey's procedure. For experiments involving a comparison of only 2 groups, a 2-tailed Student's t-test was used. P < 0.05 was considered significant.

4.20 Study approval

The Institutional Animal Care and Use Committees of the University of Arkansas for Medical Sciences and the Central Arkansas Veterans Healthcare System reviewed and approved all studies involving mice.

Data and code availability

All data reported in this paper will be shared by the lead contact upon request. This paper does not report the original code. Any additional information required to reanalyze the data reported in this work is available from the lead contact upon request.

CRediT authorship contribution statement

Kimberly K. Richardson: Methodology, Investigation, Formal analysis, Data curation. Gareeballah Osman Adam: Writing – original draft, Methodology, Investigation, Formal analysis, Data curation. Wen Ling: Methodology, Investigation, Formal analysis, Data curation. Aaron Warren: Methodology, Investigation. Adriana Marques-Carvalho: Validation, Investigation, Formal analysis, Data curation. Jeff D. Thostenson: Software, Methodology, Formal analysis, Data curation. Kimberly Krager: Methodology, Investigation, Formal analysis, Data curation. Nukhet Aykin-Burns: Writing – original draft, Validation, Resources, Methodology. Stephanie D. Byrum: Writing – original draft, Software, Resources, Methodology, Investigation, Formal analysis, Data curation. Maria Almeida: Writing – review & editing, Resources, Conceptualization. Ha-Neui Kim: Writing – review & editing, Writing – original draft, Supervision, Project administration, Investigation, Funding acquisition, Formal analysis, Conceptualization.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A Supplementary data

The following is the Supplementary data to this article:Multimedia component 1

Supplementary Figure 1. Conditional deletion of Sirt3 in LysM-Cre–targeted cells does not affect mouse development. A cohort of female Sirt3ΔLysM mice and Sirt3fl/fl littermate controls were aged to 16 months. Another cohort of female mice of the same genotypes euthanized at 6 months of age served as young controls. The graphs indicate (A) body weight and (B) femoral length in young and aged groups (n = 12–20 animals/group). Lines and error bars represent mean ± SD. P values were determined using 2-way ANOVA. Interaction terms generated by 2-way ANOVA are shown below each graph.

Supplementary Figure 2. Sirt3 was effectively deleted in LysM-Cre–targeted cells. (A) Quantitative PCR (qPCR) of mRNA isolated from bone marrow macrophages (BMMs), stromal cells, brain (n = 3 animals/group), and liver (n = 3 animals/group) obtained from 6-month-old female mice. BMMs and stromal cells were cultured from bone marrow cells as described in the methods section (triplicate cultures). (B) SIRT3 enzymatic activity in cell lysates of cultured osteoclasts measured by fluorescence (triplicate cultures). (C) mRNA levels of other mitochondrial deacetylases in BMMs were measured by qPCR (triplicate cultures). Line and error bars represent mean ± SD. P values were determined using Student’s t-test. All measures were performed in cultured BMMs or stromal cells pooled from 4–5 mice/group.

Supplementary Figure 3 (related to Figure 1). Age-related bone loss requires osteoclast SIRT3 in female mice. A cohort of male Sirt3ΔLysM mice and Sirt3fl/fl littermate controls were aged to 16 months. Another cohort of male mice of the same genotypes euthanized at 6 months of age served as young controls. (A–B) Quantification of femoral bones from female Sirt3ΔLysM mice and wild-type littermates by micro-CT after sacrifice (n = 12–20 animals/group). (C–F) Quantification of vertebral bones from female Sirt3ΔLysM mice and wild-type littermates by micro-CT after sacrifice (n = 12–20 animals/group). Line and error bars represent mean ± SD. P values were determined using 2-way ANOVA. Interaction terms generated by 2-way ANOVA are shown below each graph.

Supplementary Figure 4. Age-related bone loss does not require osteoclast SIRT3 in male mice. A cohort of male Sirt3ΔLysM mice and Sirt3fl/fl littermate controls were aged to 16 months. Another cohort of male mice of the same genotypes euthanized at 6 months of age served as young controls. (A) Sequential BMD measurements in the aging cohort by dual-energy x-ray absorptiometry (n = 20 animals/group). (B–C) Imaging and quantification of femoral bones from male Sirt3ΔLysM mice and wild-type littermates by micro-CT after sacrifice (n = 11–18 animals/group). (D–H) Quantification of vertebral bones from male Sirt3ΔLysM mice and wild-type littermates by micro-CT after sacrifice (n = 11–18 animals/group). (I) Serum concentration of a collagen degradation product (CTx) and N-terminal propeptide of type I procollagen (P1NP) in 16-month-old male Sirt3ΔLysM mice and wild-type littermates by ELISA (n = 17–18 animals/group). Line and error bars represent mean ± SD. (A) P values were determined using Student’s t-test or repeated-measures ANOVA between Sirt3ΔLysM mice and Sirt3fl/fl littermate controls in the same age. The other P values were determined using (C–H) 2-way ANOVA or (I) Student’s t-test. Interaction terms generated by 2-way ANOVA are shown below each graph.

Supplementary Figure 5 (related to Figure 3). Conditional deletion of Sirt3 in LysM-Cre–targeted cells decreases mitochondria respiration in aged female mice. BMMs were isolated from 16-month-old female Sirt3ΔLysM mice and wild-type littermates and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. (A–D) Different fractions of mitochondrial and nonmitochondrial respirations per cell, in osteoclasts, measured by Seahorse (n = 11–14 wells/group). Line and error bars represent mean ± SD. P values were determined using Student’s t-test. All measures were performed in cultured BMMs pooled from 4–5 mice/group and repeated at least twice.

Supplementary Figure 6. Deletion of Pink1 does not affect mouse development. Female Pink1 knockout mice and littermate controls were aged to (A) 6 or (B) 16 months. The graphs indicate (A) body weight and (B) femoral length (n = 7–20 animals/group). Lines and error bars represent mean ± SD. P values were determined using Student’s t-test.

Supplementary Figure 7 (related to Figure 5). Deletion of Pink1 does not affect mitochondria respiration in aged female mice. BMMs were isolated from 16-month-old female Pink1 knockout mice and littermate controls and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. (A–D) Different fractions of mitochondrial and nonmitochondrial respirations per cell, in osteoclasts, measured by Seahorse (n = 12–15 wells/group). Line and error bars represent mean ± SD. P values were determined using Student’s t-test. All in vitro measures were performed in cultured BMMs pooled from 4–5 mice/group and repeated at least twice.

Supplementary Figure 8. Principle component analysis shows intrinsic variability of proteome among different samples in the study. Quantitative analysis of global proteome was performed as indicated in Figure 6.

Supplementary Figure 9. Top 15 up- or down-regulated proteins by RANKL in the study. Quantitative analysis of global proteome was performed as indicated in Figure 6.

Supplementary Figure 10. Kyoto Encyclopedia of Genes and Genomes (KEGG) data on regulation of each mitochondria complex by Sirt3 deletion in osteoclasts. Quantitative analysis of global proteome was performed as indicated in Figure 6.

Supplementary Figure 11. Deletion of Sirt3 induces hyperacetylation of ATPIF1 in aged osteoclasts. (A) BMMs were isolated from 16-month-old female Sirt3ΔLysM mice and littermate controls and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. Acetylation level of ATPIF1 in aged osteoclasts. Endogenous ATPIF1 were immunoprecipitated from cell lysates and immunoblotted using Ac-Lysine antibodies. (B) BMMs were isolated from 6- or 24-month-old female C57BL/6 mice and were cultured with M-CSF (30 ng/mL) and RANKL (30 ng/mL) for 3 days. Endogenous ATPIF1 were immunoprecipitated from cell lysates and immunoblotted using SIRT3 antibodies. (C) Raw 264.6 cell line transduced with plasmids expressing WT or lysine acetylation mutant targeting the K49 of ATPIF1 protein and cultured with RANKL for 5 days. Lines and error bars represent mean ± SD. P values were determined using Student’s t-test.

Supplementary Figure 12. Effects of mitochondrial protein silencing in aged osteoclasts. BMMs from 24-month-old female C57BL/6 mice were transduced with lentiviral vectors expressing control sh-RNA or sh-RNAs targeting the hyperacetylated proteins and cultured as shown in Figure 7 (triplicates of pooled cultures). Representative pictures of TRAP+ multinucleated osteoclasts. Scale bar: 500 μm. The representative image for sh-Con was adopted from Figure 7.

Supplementary Figure 13. Conditional deletion of Sirt3 in LysM-Cre–targeted cells does not affect estrogen levels in females. Serum concentration of estrogen in 16-month-old female Sirt3ΔLysM mice and wild-type littermates by ELISA (n = 11–12 animals/group).

Supplementary Figure 14. Mitochondria-targeted inhibition attenuates RANKL-induced osteoclast differentiation. BMMs were cultured with Antimycin (5 nM) and Oligomycin (0.1 μM) in the presence of RANKL (30 ng/mL) for 4.5 days. (A) After culturing, cells were fixed and stained for TRAP. Scale bar: 500 μm. (B) TRAP-positive multinucleated cells containing three or more nuclei were counted as mature osteoclasts. Line and error bars represent mean ± SD. P values were determined using 1-way ANOVA.

Multimedia component 1

Data availability

Data will be made available on request.

Acknowledgments

This work was financially supported by grants R01AR080736, R01AR082418, R56AR056679, R01AG068449, R24GM137786, P20GM109005, and P20GM125503 from the 10.13039/100000002 National Institutes of Health, United States and Bone and Joint Initiative at UAMS. We thank Lee Ann MacMillan-Crow, PhD for technical support in mitochondrial assays, and J.A. Crawford, H. Wu, and S.B. Berryhill for technical assistance in bone histology and imaging. We thank the National Resource for Quantitative Proteomics, Digital Microscopy Core, Flow Cytometry Core Facility, and Division of Laboratory Animal Medicine at UAMS for their professional assistance in this work. The schematic diagrams and graphical abstract were created with BioRender.com. Editorial assistance was provided by the Science Communication Group at UAMS.

Appendix A Supplementary data to this article can be found online at https://doi.org/10.1016/j.molmet.2024.102012.
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