Department of Medicine, Emory University School of Medicine and Veterans Affairs Medical Center, Atlanta, Georgia 30033
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ABSTRACT |
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Macrophage
colony-stimulating factor (MCSF) and osteoprotegerin ligand (OPGL),
both produced by osteoblasts/stromal cells, are essential factors for
osteoclastogenesis. Whether local MCSF levels regulate the amount of
osteoclast formation is unclear. Two culture systems, ST-2 and Chinese
hamster ovary-membrane-bound MCSF (CHO-mMCSF)-Tet-OFF cells, were used
to study the role of mMCSF in osteoclast formation. Cells from bone
marrow (BMM) or spleen were cultured with soluble OPGL on
glutaraldehyde-fixed cell layers; osteoclasts formed after 7 days.
Osteoclast number was proportional to the amount of soluble OPGL added.
In contrast, varying mMCSF levels in the ST-2 or CHO-mMCSF-Tet-OFF cell
layers, respectively by variable plating or by addition of doxycycline, did not affect BMM osteoclastogenesis: 20-450 U of mMCSF per well generated similar osteoclast numbers. In contrast, spleen cells were
resistant to mMCSF: osteoclastogenesis required 250 U per well and
further increased as mMCSF rose higher. Our results demonstrate that
osteoclast formation in the local bone environment is dominated by
OPGL. Increasing mMCSF above basal levels does not further enhance
osteoclast formation from BMMs, indicating that mMCSF does not play a
dominant regulatory role in the bone marrow.
tetracycline regulation; osteoclast differentiation factor; TRANCE; colony stimulating factor-1; bone marrow
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INTRODUCTION |
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OSTEOBLASTS PLAY AN IMPORTANT ROLE in the regulation of osteoclastogenesis. Macrophage colony-stimulating factor (MCSF or CSF-1), released by osteoblasts and stromal cells in the bone microenvironment, appears to be essential for the proliferation of the early osteoclast progenitor as well as for the survival of the mature osteoclast (4). Posttranscriptional modifications of MCSF mRNA determine whether the final protein is destined for secretion (secreted MCSF, or sMCSF) or insertion into the cell membrane (membrane-bound MCSF, or mMCSF). Although <5% of the total MCSF made by cultured osteoblasts and stromal cells is membrane bound (7, 24), mMCSF can support macrophage proliferation (7) as well as formation of osteoclast-like cells (33). Because mMCSF is anchored and displayed by cells within bone, this protein isoform might represent a local regulatory signal by which osteoclast development is controlled. However, differentiating the effects of the membrane-bound from the secreted form of MCSF has posed a problem: sMCSF is rapidly removed from the microenvironment by MCSF receptor-positive cells, whose numbers are directly proportional to the amount of MCSF available. Thus the role of mMCSF in regulating osteoclast formation has been difficult to ascertain in the presence of significantly larger amounts of the secreted form.
Recently another factor critical to osteoclast maturation has been identified: osteoprotegerin ligand (OPGL/ODF/TRANCE), a member of the membrane-associated tumor necrosis factor (TNF) ligand family (15, 34). OPGL is displayed by osteoblast and/or stromal cells in bone and is a critical factor allowing entry of progenitors into the osteoclast differentiation pathway. When spleen cells (34) or nonadherent bone marrow cells (15) are cultured in the presence of MCSF and OPGL in the absence of osteoblasts or stromal cells, osteoclasts are formed. Thus MCSF and OPGL, both existing in forms expressed on cell membranes, are necessary for the process of osteoclastogenesis.
These findings beg the question of which factor, MCSF or OPGL, is dominant, sufficient, or regulatory. Several studies have suggested that MCSF levels might regulate the rate of osteoclastogenesis. Because we and others have shown that high levels of sMCSF are inhibitory to osteoclast formation because of increased entry of precursors into nonosteoclast lineages (6, 20), the possibility that the membrane-bound form of MCSF functions as a critical regulatory signal in bone is raised. In fact, Lea et al. (17) found large changes in bone marrow mMCSF after ovariectomy, a condition that is associated with increased osteoclast differentiation and subsequent bone resorption. In this study we explore the effects of mMCSF during osteoclast generation. Our findings provide the first definitive evidence that mMCSF serves a permissive role during bone marrow cell selection of osteoclast lineage. Whereas osteoclastogenesis is dependent on and sensitive to the amount of OPGL available, increasing mMCSF 20-fold, i.e., to levels surpassing in vivo levels of expression, it does not increase osteoclast formation.
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MATERIALS AND METHODS |
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Materials. 1,25-Hydroxyvitamin D3, or 1,25(OH)2D3, was obtained from Biomol Research Laboratories (Plymouth Meeting, PA). C57BL/6 male mice were purchased from the Frederick Cancer Center (Frederick, MD). Fetal bovine serum (FBS) was obtained from Atlanta Biological (Atlanta, GA). Mouse OPGL was obtained from Amgen (Thousand Oaks, CA). Other chemicals and media were obtained from Sigma (St. Louis, MO) except as noted.
Fixed ST-2 cell culture systems.
ST-2 cells (Riken Cell Bank, Tsukuba Science City, Japan), original
clones from murine bone marrow stromal cells, were plated in 12-well
plates overnight and treated with 10 nM
1,25(OH)2D3 (1,25D) and 100 nM dexamethasone
(Dex) for 2 days. ST-2 cell layers were fixed in 2.5% glutaraldehyde
for 1 min, followed by the addition of 1.5% glycine. The cell layers
were washed with PBS three times and covered with -MEM containing
10% FBS (33). Nonadherent bone marrow cells (BMMs) were
isolated from 3- to 4-wk-old C57BL/6 male mice. Briefly, BMMs were
collected from mouse tibia and forma by flushing bone marrow cavities.
The cells were incubated in
-MEM containing 10% FBS at 37°C
overnight. On the next day, nonadherent BMMs were collected and
overlaid on prefixed ST-2 cell layers at a density of 5.5-6
million/well. After 7 days of culture, the cultures were fixed with
ethanol-acetone and stained for tartrate-resistant acid phosphatase
(TRAP; Sigma, St. Louis, MO). A TRAP-positive cell containing three or
more nuclei was counted as a TRAP-positive multinuclear cell
(TRAP+MNC).
mMCSF extraction.
Live cell layers were washed with PBS and then treated for 10 min at
37°C with trypsin solution (1 mg/ml) to release mMCSF. Adding FBS for
a final concentration of 15% stopped the reaction. The cell
suspensions were centrifuged at 1,500 g for 5 min, and supernatants were stored at 30°C until bioassay (7, 24, 30).
MCSF bioassay. MCSF activity was determined by measuring the proliferation of MCSF-dependent M-NFS-60 cell line (American Type Tissue Collection, Rockville MD); 10,000 cells/well were cultured with test samples in a final volume of 100 µl as previously described (24). M-NFS-60 cell proliferation was assessed using MTT, a colorimetric assay to detect mitochondrial dehydrogenase levels. Recombinant human MCSF (6.94 × 107 CFU/mg; Cetus, Emeryville, CA) was used as the standard.
Blocking mMCSF. ST-2 cells were plated in 24-well plates at a density of 15,000/well and cultured with 10 nM 1,25D and 100 nM Dex for 2 days. ST-2 cell layers were treated at 50% confluence with 10 µg/ml of goat anti-mouse MCSF neutralizing polyclonal antibody (pAb; R&D System, Minneapolis, MN) or 10 µg/ml goat IgG at 37°C for 2 h before glutaraldehyde fixation. To determine the dose of blocking pAb, 0.1, 1, 10, or 100 µg/ml of pAb or IgG were added to ST-2 cells before fixation. Spleen cells were plated over fixed layers, and [3H]thymidine incorporation was determined. pAb at a dose of 1 µg/ml blocked one-half of the mMCSF-dependent proliferation, and complete blocking was seen by 10 µg/ml (no further decrease by 100 µg/ml).
Preparation of spleen cells.
Spleen cells were isolated from 3- to 4-wk-old C57BL/6 male mice.
Briefly, spleens were washed, minced, and suspended in PBS. Contaminating erythrocytes were eliminated from the cell pellet by
adding 0.83% NH4Cl in 10 mM Tris buffer (pH 7.4). The
cells were washed three times with PBS, suspended in -MEM containing 10% FBS, and cultured in 75-cm2 flasks overnight. The
nonadherent spleen cells were collected and overlaid on prefixed ST-2
or CHO-MCSF Tet-OFF cell layers.
TRAP assay.
After culture for 7 days, the cell layers were washed twice with PBS
and lysed with 0.05% Triton X-100. The cell lysates were centrifuged
at 10,000 g for 15 min. The supernatants were stored at
30°C. For assay, 20 µl of cell lysate and 180 µl of substrate buffer [0.48 M acetate buffer pH 5, 2 mM methylumbelliferyl phosphate (MUP), 83 mM tartaric acid] were added to each well in Microtiter 96-well white plates (Dynex Technologies, Chantilly, VA). After incubation at 37°C for 30 min in the dark, 100 µl of 0.5 M glycine solution containing 50 mM EDTA (pH 10.4) were added to stop the reaction. Fluorescence was measured on an LB 50 Plate Reader (Perkin Elmer, Buckinghamshire, England) at excitation wavelength 366 nm and
emission wavelength 456 nm. A serial dilution of methyl-umbelliferone (0-800 µM) was used to generate a standard curve. The enzyme
activity was represented as micromoles of MUP hydrolyzed per milligram of protein per min.
Cell proliferation. [3H]thymidine (2 µCi/well) was added to cultures on the specific day. After 24 h, cells were lysed in 20 mM NaOH containing 1% SDS, transferred to scintillation vials, and counted by a PACKARD 2500 TR Liquid Scintillation Analyzer (Downers Grove, IL).
RT-PCR.
Total RNA was prepared in TRIzol (Life Technologies, Gaithersburg, MD).
For measurement of cathepsin K, 0.5 µg of total RNA was added to an
RT reaction containing 1 mM dNTP, 0.5 µM forward primer, 100 U of
MMLV reverse transcriptase, and 20 U of RNAsin. The RT reaction was
incubated for 30 min at 37°C. For quantitation of PCR products, 1 µCi of -[32P]dCTP was used in each standard PCR
reaction (cathepsin K PCR forward primer 5'-CCAGTGAA-GAAGTGGTTCAG,
reverse primer 5'-TATCCTTTCTTTCGATAGTCG). PCR products were
chromatographed and phosphorimages captured on a Molecular Dynamics
instrument. The cathepsin K density was normalized by
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) density obtained from
the same sample.
Tet-OFF system.
The mMCSF cDNA (1.6 kb) plasmid was the generous gift of Dr. Martine
Roussel (St. Jude's Children Research Hospital, Memphis, TN). The
region coding for mMCSF (i.e., exon 6) was obtained by PCR with
adapters added to mutate two base pairs on each PCR primer (forward
primer 5' CCC GGC CGC GGC CCA GCT GCC CGT ATG ACC G 3' and reverse
primer 5' GAG AGA TCT TAG AAT TCC CTC TAC ACT GGC 3') to create
Sac II and Bgl II sites, respectively. PCR was
performed by adding 50 ng of 1.6 kb mMCSF cDNA plasmid with 100 ng each
of primers into 50 µl of 200 µM dNTP, 2.2 µM Mg2+,
and 10 units of Pfu DNA polymerase (Stratagene, San Diego, CA). Twenty-five cycles of PCR were used at the following parameters: denature at 94°C for 30 s, anneal at 60°C for 30 s, and
extend at 72°C for 2 min. The PCR product was digested with Sac
II and Bgl II, and the appropriate band was purified
from 1% agarose using a gel extraction kit (Qiagen, Valencia, CA). The
pTRE vector obtained from Tet-OFF gene expression system (Clontech,
Palo Alto, CA) was opened with Sac II and BamH I
and then extracted and purified. The TRE-MCSF construct was generated
by ligating the above insert and vector, because BamH I
and Bgl II have compatible ends. Sequencing confirmed that
the entire coding region of mMCSF was inserted into the TRE response
vector. CHO cells purchased from Clontech had appropriate
tetracycline-regulatory protein (Tet-R) expression. CHO cells cultured
in
-MEM containing 10% Tet-free FBS were cotransfected with
pTRE-MCSF and a vector carrying resistance to hygromycin (pTK-Hyg) at a
10:1 ratio. Stable cell lines were selected under 400 µg/ml G418 and
200 µg/ml hygromycin (Sigma). Thirty-six clones were isolated and
tested by MCSF bioassay in the presence and absence of doxycycline
(Dox). Clone 4 produced the highest mMCSF expression in the
absence of Dox vs. low expression with 1 µg/ml Dox. The doubly stable
CHO-mMCSF cell line was maintained under 200 µg/ml G418, 100 µg/ml
hygromycin, and 1 µg/ml Dox constraints.
Statistical analysis. Data were analyzed by Tukey ANOVA by use of Prism software.
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RESULTS |
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mMCSF stimulates bone marrow macrophage and spleen
cell proliferation.
ST-2 cells were plated at densities from 7,500 to 120,000 per well.
After culture with 1,25D and Dex for 2 days, cell number and mMCSF were
assayed. The ST-2 cell doubling time of 28 h was highly
reproducible, as was the final mMCSF level achieved. Figure 1A shows that mMCSF levels as
measured by bioassay before fixation increased from 3.5 to 138 U/well
as final cell numbers increased from 56,000 to 266,000/well.
Glutaraldehyde treatment at this time can thus ensure a controlled
amount of mMCSF for study.
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OPGL regulation of osteoclast formation is dose dependent. ST-2 cells were treated for 2 days with 1,25D (10 nM) and Dex (100 nM), which induces maximal expression of both mMCSF (22) and OPGL (10, 34). At this time, cells were fixed with 2.5% glutaraldehyde. To test whether the fixed ST-2 cells were able to support osteoclastogenesis, nonadherent BMMs were plated over the fixed ST-2 layers in the presence or absence of murine sOPGL (0-10 ng/ml). After 7 days of culture, no TRAP+MNCs appeared when BMMs were cultured over ST-2 cells alone, but they were induced with the addition of exogenous OPGL. Because ST-2 cells are well known to display OPGL and support osteoclastogenesis under the culture conditions used here (34), the glutaraldehyde fixation procedure blocked or destroyed the OPGL molecule on the cell surface. In the presence of exogenous OPGL, however, glutaraldehyde-fixed layers were able to support osteoclast formation. This result suggested that the process of glutaraldehyde fixation removed the contribution of endogenous OPGL and allowed us to study the effects of mMCSF availability on osteoclast formation.
Osteoclast formation increased in an sOPGL dose-dependent fashion as seen both visually (Fig. 4A) and by counting TRAP+MNCs (Fig. 4B). As a corroborative measure of increase in the osteoclast phenotype, TRAP activity was measured using a sensitive methylumbilliferone assay. In Fig. 4C, TRAP activity dose dependently increased in proportion to the added sOPGL. ST-2 cells express mMCSF in the absence of 1,25D or Dex, as we have previously shown (11). To assure that a lower level of mMCSF expression also supported osteoclastogenesis, sOPGL was added to cocultures of ST-2 cells and BMMs in the absence of 1,25D or Dex. sOPGL caused a dose-dependent increase in osteoclast numbers as follows: sOPGL at 0 ng/ml = 6 ± 2 osteoclasts, at 2.5 ng/ml = 74 ± 7 osteoclasts, at 5 ng/ml = 353 ± 16 osteoclasts, and at 10 ng/ml = 783 ± 43 osteoclasts. Although these data cannot be compared directly with the results obtained when ST2 cells are incubated in the presence of 1,25D and Dex, they serve to show that basal levels of mMCSF are adequate to support osteoclastogenesis in the presence of sOPGL.
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Role of mMCSF in regulation of osteoclast formation from bone marrow progenitors. To evaluate whether mMCSF might play a dominant role in regulating osteoclast precursor differentiation, we varied concentrations of the presented growth factor before fixation. In the first system, the ST-2 cell-plating density regulated mMCSF availability. In the second, mMCSF expression was regulated in the tetracycline-sensitive CHO-mMCSF clone with Dox.
The range of mMCSF in fixed ST-2 cell layers was between 35 and 120 U/well. Soluble OPGL was added to stimulate osteoclastogenesis: 5 ng/ml sOPGL represented the ED50 for sOPGL-induced osteoclast formation in the fixed ST-2 culture system (see Fig. 4). After 7 days of culture with BMMs, cultures were fixed and stained for TRAP. As shown in Fig. 5A, TRAP+MNCs did not increase when mMCSF availability rose from 34 to 120 U/well. The sensitive TRAP activity assay also revealed no changes as mMCSF units increased, as shown in Fig. 5 as TRAP activity. There was even a trend toward decreased osteoclast formation at the higher mMCSF availability. In addition, experiments were performed with 2.5 ng/ml sOPGL, which confirmed that increasing mMCSF did not increase TRAP activity (data not shown).
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Requirements for MCSF and OPGL during
spleen progenitor to osteoclast differentiation.
Spleen cells can proliferate with added MCSF and differentiate into the
osteoclast lineage when sOPGL is added to the culture medium in the
absence of accessory stromal cells (34). We were not able
to generate osteoclasts from spleen cells over fixed ST-2 cell layers;
however, the CHO-mMCSF-Tet-OFF system, which displays higher amounts of
mMCSF, was successful. TRAP+MNCs were present at the
highest doses of fixed mMCSF (632 and 320 U/well), disappearing when
Dox decreased mMCSF expression below 100 U/well (Fig.
6). Figure 6, which compiles two separate experiments, did show a statistical trend toward an increase in osteoclast numbers when mMCSF rose above 300 U/well, suggesting that
spleen cells respond differently than do BMMs.
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DISCUSSION |
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In the work presented here, we wished to clarify whether mMCSF levels could regulate the level of osteoclast formation. During the past decade, MCSF elaborated by stromal cells has been recognized to be an essential factor for osteoclast precursor proliferation and differentiation and osteoclast survival (27). Stromal cells from op/op mice that are deficient in MCSF do not support osteoclastogenesis in the absence of exogenous MCSF (32) Antibodies that completely block the MCSF receptor inhibit the development of osteoclasts in vitro (9). Furthermore, even with the addition of substantial amounts of sOPGL, monocytic precursors do not appear to generate osteoclasts in the absence of MCSF (3, 15, 34).
Osteoclastogenesis is known to be modulated by systemic hormones and
local stimulatory factors, such as 1,25D, parathyroid hormone (PTH),
interleukins, TNF-, and TGF-
, among others (8, 19, 25,
27). Many of these factors stimulate the expression of MCSF,
leading to the hypothesis that the level of MCSF present within the
local bone environment might dose dependently direct osteoclast
formation. For instance, 1,25D, TNF-
, and PTH, all of which potently
induce osteoclast formation, cause increased secretion of sMCSF
(11, 24, 31). Additionally, several investigators have
suggested that estrogen deficiency might promote osteoclastogenesis by
increasing the production of sMCSF by bone stromal cells (12, 26). However, very high levels of added sMCSF inhibit
recruitment of progenitors into the osteoclast lineage despite
increased proliferation of the predominant clone (6, 20, 29,
34). Further confounding these issues is a lack of data
demonstrating significant transcriptional control within the murine
MCSF promoter by osteoclastogenic factors known to affect sMCSF
expression (11, 23). We and others have suggested that
sMCSF secretion might be controlled posttranscriptionally (5) or posttranslationally via the trafficking of the
secreted protein within the stromal cell (22). In sum, the
role of sMCSF in regulating osteoclastogenesis, directly or indirectly,
has never been firmly established.
Alternatively, mMCSF, which is directed to the cell membrane after posttranscriptional excision of exon 6 (16), might represent a more finely tuned and significant regulatory molecule as expressed by cells within bone. Glucocorticoids, which promote bone resorption, increase the expression of mMCSF in bone stromal cells while decreasing secretion of the soluble form (22). As well, estrogen deficiency may specifically increase the expression of mMCSF, as opposed to sMCSF, within the bone marrow (17). Recent studies have shown that mMCSF is sufficient to support osteoclast formation in vitro (33), suggesting a role for this isoform in regulating osteoclast lineage selection.
To investigate effects of mMCSF on regulation of osteoclastogenesis, we used both fixed ST-2 cells and CHO-MCSF Tet-OFF cells overlaid with osteoclast progenitors. In both systems, proliferation of the osteoclast progenitors was proportional to mMCSF availability, as expected (6). Neither cell system, however, supported osteoclastogenesis in the absence of exogenous OPGL. This was unexpected in the case of the ST-2 cells, which express OPGL under the control of hormones (13), indicating that OPGL was destroyed by our fixation procedure or was fixed in an inactive conformation. In contrast, Kong et al. (14) reported that fixed activated T cells expressing OPGL could trigger osteoclastogenesis; this difference may be due to differences in fixation or in aspects of lymphocyte presentation of the molecule. Nonetheless, our system allowed investigation of the role of mMCSF in directing osteoclast formation.
Our experiments showed that BMMs cultured on fixed ST-2 cell layers displaying varying amounts of mMCSF (34-120 U) showed no increase in osteoclast formation as mMCSF availability increased. The concentration of sOPGL, on the other hand, was intrinsically related to the amount of osteoclastogenesis. In confirmation of this finding, experiments using fixed CHO-mMCSF cell layers showed that BMM osteoclastogenesis was regulatable by sOPGL and that osteoclast formation was not changed as mMCSF was increased 20-fold. These data indicate that increasing mMCSF expression stimulates precursor proliferation but does not enhance osteoclast formation. Thus, whereas mMCSF undoubtedly has an important role in the recruitment of monocytes and may have effects on both osteoclast activity and fusion (2, 18), its regulatory function during osteoclast recruitment from marrow cells appears to be minor. MCSF's ability to enhance the expression of RANK, the receptor for OPGL (3), suggests a competence role for MCSF, rather than one that predicates final selection of the osteoclast phenotype.
Both marrow and spleen cells represent sources of osteoclast
precursors. In our experiments, these two cell sources were not interchangeable. Whereas BMMs formed osteoclasts in the presence of
minimal amounts of mMCSF (34 U/well) and sOPGL (2 ng/ml), spleen cells
required nearly ten times as much mMCSF and 10 ng/ml sOPGL to induce
osteoclast formation. This could be due to the differentiation state of
the spleen cells, which may harbor fewer cells capable of proliferating
and responding to sOPGL, perhaps suggesting that fewer spleen cells
express either RANK or MCSF receptors. In addition, spleen cells may
not provide other stimulatory factors that enhance osteoclast
development; for instance, Sells Galvin et al. (25) reported that TGF- had a direct stimulatory effect on
osteoclastogenesis in hematopoietic cells treated with sOPGL/ODF and
MCSF. Furthermore, our data did show that increasing mMCSF far above
levels that would generally be expressed in bone would further enhance
osteoclast formation from spleen progenitors. The source of osteoclast
progenitors thus represents a significant variable in studies that
evaluate the effects of stimulators and inhibitors of osteoclast
development. Because marrow cells, and not spleen or peripheral blood
cells, are precursors for bone osteoclasts in most organisms, lower
levels of mMCSF are likely adequate to support osteoclastogenesis.
Thus the major limiting factor for osteoclast induction in marrow
culture is OPGL. Previous results showing that very high levels of
sMCSF were associated with decreased entry of proliferating cells into
the osteoclast lineage can now be reinterpreted as being limited by the
fixed OPGL potential presented by stromal cells (6, 20).
Furthermore, it is unlikely that small changes in mMCSF induced by
resorptive factors such as TNF- (33) and 1,25D
(24) or physical factors such as hydrostatic pressure (21) have significant effects on osteoclast recruitment.
Alternatively, BMMs cultured with increasing amounts of sOPGL on fixed
ST-2 cell layers providing equivalent mMCSF showed a clear sOPGL dose
dependence in terms of osteoclast formation. In summary, our results
suggest that the osteoclastogenic potential of the marrow environment is dominantly regulated by OPGL, whereas mMCSF serves as a competence factor. It is likely that agents regulating osteoclastogenesis do so
directly or indirectly through OPGL.
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ACKNOWLEDGEMENTS |
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This work was supported by the Veterans Administration (Merit and Research Enhancement Award Program) and National Institute of Arthritis and Musculoskeletal and Skin Diseases R01-AR-42360.
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FOOTNOTES |
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Address for reprint requests and other correspondence: X. Fan, Veterans Affairs Medical Center - 151, Emory Univ. Medical School, 1670 Clairmont Rd., Decatur, GA 30033 (E-mail: xfan{at}emory.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 25 February 2000; accepted in final form 26 September 2000.
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