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Third Department of Medicine, Teikyo University School of Medicine (R.O., M.T., K.T.), Ichihara, Chiba 299-0111; the Division of Endocrinology, Department of Internal Medicine, University of Tokyo School of Medicine (S.F., T.F., Y.T.), Bunkyo-ku, Tokyo 112-8688; and Sankyo Co., Ltd., Pharmacology and Molecular Biology Research Laboratories (M.M.), Shinagawa-ku, Tokyo 140-8710, Japan
Address all correspondence and requests for reprints to: Ryo Okazaki, M.D., Third Department of Medicine, Teikyo University School of Medicine, 3426–3 Anesaki, Ichihara, 299-0111 Japan. E-mail: rokazaki{at}med.teikyo-u.ac.jp
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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(PPAR)
induces adipogenic differentiation of stromal cells; however, whether
this would affect osteoblast/osteoclast differentiation is unknown.
Thus, we examined the effects of the thiazolidinedione (TZD) class of
antidiabetic agents that activate PPAR on osteoblast/osteoclast
differentiation using mouse whole bone marrow cell culture. As
reported, all TZDs we tested (troglitazone,
pioglitazone, and BRL 49653) markedly
increased the number of Oil Red O-positive adipocytes and the
expression of adipsin and PPAR 2. 1
,25-Dihydroxyvitamin
D3 [1,25-(OH)2D3] did not affect
adipogenic differentiation induced by TZDs. TZDs did not affect
alkaline phosphatase activity, an early marker of osteoblastic
differentiation, despite their marked adipogenic effects. TZDs
decreased the number of tartrate-resistant acid phosphatase-positive
multinucleated osteoclast-like cells induced by
1,25-(OH)2D3 or PTH. Troglitazone
dose dependently inhibited basal and
1,25-(OH)2D3- and PTH-induced bone resorption
as assessed by pit formation assay. Interleukin-11 blocked the
induction by troglitazone of adipogenesis, but had no
effect on the inhibition of osteoclast-like cell formation. These
results indicate that TZDs are potent inhibitors of bone resorption in
vitro. Inhibitory effects of TZDs on osteoclastic bone resorption was
not osteotropic factor specific and did not appear to be related to
their adipogenic effects. Thus, TZDs may suppress bone resorption in
diabetic patients and prevent bone loss. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Peroxisome proliferator-activated receptor- (PPAR), a member of
the steroid receptor superfamily, plays a pivotal role in the
differentiation of adipocytic cells (8). PPAR activation by
15-deoxy-
12,14-PG J2 or the thiazolidinedione (TZD)
class of antidiabetic agents leads to adipogenic differentiation of
various types of cells (8, 9, 10, 11, 12). Because bone marrow tissue and bone
marrow cells express PPAR, it is plausible that TZDs, such as
troglitazone (Tro), pioglitazone (Pio), and
BRL 49653 (BRL), affect bone marrow cell differentiation.
Indeed, Gimble et al. reported that TZDs promote
adipogenesis in a mouse bone marrow stromal cell line, BMS2, and
primary mouse bone marrow cells (13). Furthermore, forced expression of
PPAR in fibroblasts or myoblasts induce transdifferentiation of
these cells into adipocytes (11, 14). These results suggest that TZDs
affect osteoblastic differentiation of bone marrow stromal cells.
Subsequently, they also may affect osteoclastic differentiation.
However, little is known about the effects of TZDs on the
differentiation of osteoblasts and osteoclasts.
TZDs are clinically useful for diabetic patients, who are commonly associated with increased bone resorption (15) and/or high risk of osteoporosis, such as postmenopausal state and aging. Thus, it is important to clarify if adipogenic actions of TZDs in bone marrow affects bone resorption and formation. We examined the effects of TZDs on osteoblastic and osteoclastic differentiation using mouse whole bone marrow cell culture and showed an inhibitory effect of TZDs on osteoclast-like cell formation and bone resorption in vitro.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Assay of tartrate-resistant acid phosphatase-positive
multinucleated cell [TRAP(+)MNC] formation
TRAP(+)MNC formation was assessed as described previously with
modification (16). Bone marrow cells were disaggregated from tibia and
femur of 6-week-old male ddY mice (Japan SLC, Shizuoka, Japan). Cells
were plated in 24-well plates in MEM supplemented with 10% FBS.
Twenty-four hours later, medium was changed, and test agents were added
to the culture. After 3 days, medium was changed again, and the cells
were cultured 3 more days in the presence of test agents. At the end of
culture, cells were fixed with 10% formalin and stained with TRAP. All
TRAP-positive cells with three or more nuclei in each well were
counted. All experiments were performed in quadruplicate. All
procedures involving animals were approved by institutional animal care
committee.
Pit formation assay on dentine slice
The pit formation assay was performed as previously described
(17) with modification. Briefly, bone marrow cells disaggregated from
hind limbs of 14-day-old ICR mice were plated on a dentine slice in
DMEM supplemented with 10% FBS. Four hours later, cells were rinsed
once with the medium and cultured with test agents for 4 days. At the
end of the culture, cells were subjected to cell viability assay, and
the dentine slices were subjected to pit formation assay. For cell
viability assay, 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyl
tetrazolium bromide (MTT) was added to the culture at a final
concentration of 25 µg/ml and incubated for 4 h. After the
incubation, media were removed, the precipitated dye was solubilized
into dimethylsulfoxide, and absorbance was measured at 540 nm. Dentine
slices were cleaned with a handy engine (Yoshida Seiko Co. Ltd, Tokyo,
Japan) for 10 sec and stained with acid hematoxylin. The total area of
the pits was measured by an image analyzer (PIAS-LA555, PIAS Co. Ltd,
Tokyo, Japan).
Alkaline phosphatase assay
Bone marrow cells were plated in 12-well plates and cultured as
described for the TRAP(+)MNC assay. At the end of culture, cells were
washed twice with ice-cold PBS and scraped in 10 mM
Tris-HCl containing 2 mM MgCl2 and 0.05%
Triton X-100, pH 8.2. The cell suspension was homogenized using Pellet
Pestle (Kontes, Vineland, NJ) on ice after two cycles of freezing and
thawing. Aliquots of supernatants were subjected to protein assay using
a Bio-Rad Laboratories, Inc. kit (Richmond, CA) according
to Bradford’s method and to alkaline phosphatase (ALP) activity
measurement as described previously (18).
Oil Red O staining
Bone marrow cells were cultured as described above. Cells were
fixed in 10% formaldehyde for 10 min, then in 60% isopropanol for 1
min, stained with Oil Red O for 30 min, and rinsed briefly with 60%
isopropanol. All Oil Red O-positive cells in each well were
counted.
RT-PCR
Bone marrow cells were cultured as described above in 6-cm
plates. Total RNA was isolated using Isogen (Nippon Gene Co., Toyama,
Japan) according to the manufacturer’s instruction. The resultant RNA
samples were further purified by a round of lithium chloride (2
M) precipitation, followed by a round of ethanol
precipitation. The amount of RNA was calculated with the absorbance at
260 nm, and all the samples were diluted to the same concentration
(0.1–0.2 µg/µl). One microgram of the diluted RNA samples was
electrophoresed on a 1% agarose gel, stained with ethidium bromide
(0.5 µg/ml), and visualized by UV transilluminator. The integrity and
equality of RNA samples were verified by the band intensity of
ribosomal RNA 28S and 18S. RT-PCR was performed as described with
modification (19). One microgram of RNA was reverse transcribed by
incubation for 5 min at room temperature and then for 90 min at 42 C
with 100 U Moloney murine leukemia virus transcriptase (Life Technologies, Inc.); 5 mM random hexamer
(Roche Molecular Biochemicals, Indianapolis, IN); 2.5
µM oligo(deoxythymidine)16 (Roche Molecular Biochemicals); 1 U/µl ribonuclease inhibitor
(Promega Corp., Madison, WI); 1 mM each of
deoxy (d)-ATP, dCTP, dGTP, and dTTP; and 1 x Taq
reaction buffer (Promega Corp.) supplemented with 5
mM MgCl2 in a total volume of 20 µl. After
the reaction, the mixture was heated for 5 min at 95 C and diluted to
60 µl with 1 x Taq buffer. Three microliters of the
products were used for PCR amplification in a total volume of 20
µl containing 1 x Taq reaction buffer, 0.2
mM dNTPs, 1.5 mM MgCl2, 1
µM of each primer, and 0.5 U AmpliTaq Gold DNA polymerase
(Perkin-Elmer Corp., Norwalk, CT). The primers used were
5'-gtgaaccactgatattcagg-3' and 5'-ctgatgcactgcctatgagc-3' for PPAR1,
5'-gggtcagctcttgtgaatgg-3' and 5'-ctgatgcactgcctatgagc-3' for PPAR2,
5'-tcttgatttacacggaggtg-3' and 5'-tcttgtttgtttgtccagtg-3' for
lipoprotein lipase (LPL), 5'-atacgaggacaaacaagtgg-3' and
5'-gtaaccacaccttcgagtg-3' for adipsin, 5'-aaacacacgaactgcagcac-3'
and 5'-tcttcttcccaggcaggctc-3' for osteoprotegerin (OPG) or
osteoclastogenesis inhibitory factor, and5'-tcttcagctgatggtgtatg-3'
and 5'-gagtctcagtctatgtcctg-3' for osteoclast differentiation factor
(ODF) or OPG ligand. PCR cycles were performed in GeneAmp PCR system
2400 (Perkin-Elmer Corp.) with the following temperature
profile: denaturation at 95 C for 30 sec, and primer annealing and
primer extension at 60 C, each for 30 sec. After initial denaturation
(9 min), the cycle was repeated 20–45 times, followed by a final
extension step of 10 min at 60 C. Half of the PCR product was
electrophoresed on a 2.5% NuSieve 3:1 (FMC BioProducts, Rockland, ME)
agarose gel and stained with 0.5 µg/ml ethidium bromide, and bands
were visualized by UV transilluminator. The identity of PCR products
was confirmed by fluorescence-based dideoxy sequencing of each PCR
product using ABI PRISM 310 genetic analyzer (Perkin-Elmer Corp.).
Statistical analyses
All statistical analyses were performed using StatView software
(version 4.5, Abacus Concepts, Inc., Berkeley, CA). The results were
analyzed with one-way ANOVA followed by Bonferroni/Dunn’s test.
P < 0.05 was considered significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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activation in bone marrow stromal
cells suppress osteoblastic differentiation, we measured ALP activity
as an early osteoblastic differentiation marker. As shown in Table 1
, Tro had no significant effect on ALP
activity in bone marrow cultures. Neither Pio (up to 50
µM) nor BRL (up to 50 µM) affected ALP
activity (data not shown).
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). 1,25-(OH)2D3
(1 nM) alone did not induce Oil Red O-positive cells or
influence the effect of TZDs. As expected from previous reports (13),
adipocytic differentiation was associated with increased expression of
PPAR2 and adipsin, but was not associated with the changes in
PPAR1 or LPL expression (Fig. 1).
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). At 10
µM, BRL and Pio decreased TRAP(+)MNC formation to a
similar level as Tro (Fig. 2B). These TZDs also suppressed TRAP(+)MNC
formation induced by 10 nM PTH (data not shown).
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). Tro inhibited both
1,25-(OH)2D3- and PTH-induced pit formation at
10 µM (Fig. 3, B and C). To check whether the inhibitory
effect of Tro on pit formation is mediated by general cytotoxicity or
promotion of apoptosis, we assessed cell viability using the MTT assay.
As shown in Table 3, Tro up to 10
µM did not significantly change MTT values, suggesting
that inhibition of pit formation by Tro was not mediated by its
cytotoxicity. However, at 30 µM, Tro decreased MTT
values, which was less marked than its inhibition of pit formation.
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). However, rhIL-11 up to 10
nM failed to restore the decreased number of TRAP(+)MNC in
the presence of 10 µM Tro (Fig. 4B).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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in the differentiation of adipocytes has
been established (8). Because bone marrow stromal cells are the origin
for not only adipocytes but also osteoblasts and because they are
necessary for osteoclast generation, it is presumable that PPAR
activation affects differentiation of osteoblasts and/or osteoclasts.
In the present study we demonstrated that TZDs inhibit the formation of
osteoclast-like cells and bone resorption in vitro. To our
surprise, although TZDs markedly promoted adipocytic differentiation of
the marrow cells, this was not associated with a decrease in ALP
activity, a representative early marker of osteoblastic
differentiation. Effects of TZDs on osteoblastic cells have been studied by other investigators. Gimble and co-workers (13) used a mouse bone marrow-derived adipocytic cell line, BMS2, that can be differentiated into osteoblastic cells with the stimulation by bone morphogenetic protein-2 and primary bone marrow stromal cells that are similar to the cells we used in the present study. They reported that low dose TZDs promote adipocytic differentiation, but did not touch on the possible effects of these compounds on osteoblastic differentiation. Recently, Nuttall et al. (27) reported that primary human osteoblast-like cells could be transdifferentiated into adipocytes after stimulation by TZD; however, whether this was associated with the loss of osteoblastic phenotype was not mentioned. The increase in adipocyte number in the bone marrow associates with the decrease in osteoblast number as evident in the aged subjects (6, 28). The reasons why the effects of TZDs on osteoblastic differentiation have not been demonstrated in vitro may be due to the limitation of the presently available osteoblastic markers, such as ALP and osteocalcin, because cells in the adipocyte lineage may express them (29). As suggested by Nuttall et al. (27), in the bone marrow, adipocytes and osteoblasts are too closely related to be discriminated in the presence of TZDs.
In contrast to the lack of the effects on early osteoblastic
differentiation, TZDs clearly inhibited osteoclast-like cell formation
and bone resorption in vitro. Although the mechanisms by
which TZDs inhibit osteoclastic bone resorption are not clear from the
present study, we are able to speculate several possibilities and
estimate whether they are probable based on our observations. Firstly,
because both vitamin D receptor and PPAR heterodimerize with
retinoid X receptor to act as transcription factors (8), TZDs could
antagonize vitamin D action at this level and inhibit osteoclastic bone
resorption. However, this is unlikely, because TZDs inhibited
osteoclast-like cell formation and pit formation induced not only by
1,25-(OH)2D3 but also by PTH.
Secondly, adipogenic differentiation could cause a loss of the ability of stromal cells to support osteoclastogenesis. However, the fact that IL-11 failed to restore osteoclast-like cell formation at doses that significantly blocked adipogenic differentiation of bone marrow cells suggests otherwise. Moreover, Kelly et al. (30) reported that a bone marrow stromal cell line, BMS-2, supports osteoclastogenesis even after adipogenic differentiation. These results suggest that adipogenic differentiation of stromal cells per se does not affect their capability to support osteoclastogenesis.
Thirdly, TZDs could affect actions of cytokines that control
osteoclastogenesis. Osteoclastogenesis is a complex process that is
under the control of a variety of hormones and cytokines (5, 31). We
first examined the possible contribution of IL-11-dependent processes,
because IL-11 is secreted by bone marrow stromal cells, inhibits
adipogenesis, and stimulates osteoclastogenesis in the bone marrow (20, 21, 32). However, our findings suggest that the inhibitory effect of
TZDs on osteoclastogenesis is not mediated by this cytokine pathway.
The recently identified ODF/OPG system plays the most important role in
the regulation of osteoclastogenesis dependent on stromal/hemopoietic
stem cell interaction (22, 23, 24, 25). We also explored possible involvement
of this newly established pathway in TZD inhibition of
osteoclastogenesis. Although our data are limited, out results suggest
that the ODF/OPG pathway is unlikely to contribute to TZD effects on
osteoclastogenesis. IL-6 is also secreted by bone marrow stromal cells
and has similar effects as IL-11 (5, 31). IL-6 secretion by bone marrow
cells was not affected by Tro up to 10 µM and was only
slightly decreased at 30 µM (Takeuchi, Y., unpublished
observation). Among other candidate mediators are IL-1 and tumor
necrosis factor-, both are known to promote osteoclastogenesis
in vitro (31) and are reported to be decreased in human
peripheral blood mononuclear cells with TZDs (33). These cytokines
along with macrophage colony-stimulating factor are all candidates that
mediate TZD’s effects on bone resorption, and they are now studied in
our laboratories.
Finally, TZDs could act on osteoclast progenitors instead of stromal
cells. Whether osteoclast progenitors express PPAR is unknown;
however, it is expressed in several myeloid cell lines, spleen that is
rich in osteoclast precursors, and tissue macrophages, which originate
from the same progenitor as osteoclasts (34, 35). Interestingly,
PPAR activation in a monocytic leukemia cell line, HL-60, promotes,
but does not inhibit, differentiation of these cells into mature
macrophages (35). This suggests that TZDs may act on hemopoietic cells
not to stimulate their osteoclastic differentiation but to push them
toward macrophage differentiation. However, this possibility has to be
tested in a system totally devoid of stromal cells and is beyond the
scope of the present study.
Our results agree with those reported by Gimble and co-workers (13, 30, 36), in that TZDs promote adipocytic differentiation of marrow cells, but differ in several respects. First, we were unable to see any antagonistic effect of 1,25-(OH)2D3 on adipogenesis induced by TZDs. They reported that 1,25-(OH)2D3 was a potent inhibitor of adipogenesis induced by glucocorticoid and insulin and a weak inhibitor of that induced by TZDs (36). Second, they did not see any inhibitory effect of TZDs on osteoclast-like cell formation (30). Because their culture system is completely different from ours, especially in the latter case, this could be a reason for the apparent discrepancies.
In conclusion, we demonstrated that TZDs were potent inhibitors of bone resorption in vitro. Our preliminary data (37) indicate that Tro administration to type 2 diabetics causes decreases in bone resorption markers before significantly improving glucose metabolism. We have previously reported that poorly controlled type 2 diabetes patients exhibit high bone resorption, which is normalized with glycemic control by treatment modalities other than TZDs (15). Taken together, our results suggest that TZDs inhibit bone resorption by dual mechanisms; locally acting on bone marrow cells and systemically improving glucose metabolism. Whether TZD treatment prevents bone loss in noninsulin-dependent diabetes patients has to be determined in future studies.
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Acknowledgments |
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Footnotes |
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Received March 16, 1999.
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References |
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