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Department of Medicine (T.Y., H.M., Y.T., S.C.M., E.A.), Division of Endocrinology and Metabolism, the UAMS Center for Osteoporosis and Metabolic Bone Diseases, and the McClellan VA Medical Center GRECC; Department of Microbiology and Immunology (J.U.I.), University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205
Address all correspondence and requests for reprints to: Etsuko Abe, Ph.D., Division of Endocrinology and Metabolism, University of Arkansas for Medical Sciences, 4301 West Markham, Mail Slot 587, Little Rock, Arkansas 72205. E-mail: eabe{at}acrc.uams.edu
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Abstract |
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Introduction |
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Osteopontin, a noncollagenous phosphoprotein, contains an RGD tripeptide identical to that in fibronectin, vitronectin, osteonectin, and fibrinogen (11, 12). Based on this, it is thought that like the rest of these proteins, osteopontin may be involved in cell-to-cell and cell-matrix interactions (13, 14) during cell proliferation and cell migration. Osteopontin was originally isolated from bone matrix (15). Subsequent studies demonstrated expression of this protein in various cells and tissues, including osteoblasts, osteoclasts, and macrophages (11). During early stages of embryonic life, osteopontin expression is very low, but it increases progressively during development (16, 17), reaching a peak in calvaria and tibiae at 4–14 days postpartum (16, 18). This progressive increase parallels the increase of several other proteins, including alkaline phosphatase (AP) and osteocalcin.
In the adult rat and human skeletons, osteopontin is localized in sites of bone formation as well as in sites of bone resorption (19, 20, 21, 22). Specifically, the protein has been demonstrated in cement lines, mineralized bone matrix during endochondral and intramembranous ossification, cartilage matrix, and osteoclastic resorption lacunae. Based on this, we have hypothesized that osteopontin plays a role in osteoclast and perhaps osteoblast development in the bone marrow; therefore its expression must be up-regulated in states where these processes are up-regulated, such as estrogen deficiency (23). Because osteopontin is a secreted protein, to identify its cellular origin in heterogenous bone marrow cell cultures, we examined osteopontin messenger RNA (mRNA) expression by in situ hybridization. We present evidence that the osteopontin mRNA is expressed during the differentiation of at least three distinct populations of bone marrow cells: stromal/osteoblast-like cells, osteoclast-like cells, and macrophage-like cells. Moreover, osteopontin seems to be critical for osteoclastogenesis, as an antiosteopontin antibody or an RGD-containing peptide inhibit this process.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-32P]
triphosphate (3000 Ci/mmol) was purchased from Amersham (Arlington
Heights, IL). Peptides (GRGDS and GRGES) were
purchased from American Peptide Company (Sunnyvale, CA).
Cell preparations and culture conditions
Swiss Webster mice (6-week-old male or 60-day-old female),
maintained in accordance with the NIH Guidelines for Care and Use of
Laboratory Animals, were killed by cervical dislocation, and femurs and
tibiae were aseptically removed and dissected free of adherent tissue.
Both ends of the bone were cut, and the bone cavity was flushed out
with culture medium slowly injected at the end of the bone using a
sterile 27-gauge needle. Bone marrow cells were subsequently
resuspended (106 cells/ml) in phenol red free -MEM
containing 10% heat-inactivated FCS (Sigma) and were maintained with
or without 10-8 M
1,25-(OH)2D3 for 7 days. Eighty percent of the
medium was replaced with fresh medium every 3 days.
In the experiments examining the effect of the antiosteopontin antibody on osteoclast formation, a rabbit antimurine osteopontin antiserum (antiosteopontin Ab) or nonimmune rabbit serum (NIRS) were used at 1/100 or 1/250 dilutions. Similarly, in the experiments examining RGD-containing peptide, GRGDS or GRGES (negative control) were used at 10-4 to 10-7 M. At the end of the culture (7 days), cells were fixed with 10% neutralized formalin and then stained for either TRAP, NSE, or AP. For the remaining experiments, 60-day-old female mice were either sham-operated or ovariectomized (OVX), and bone marrow cells were obtained 2 weeks after the operation. Bone marrow cells were then seeded in 75 cm2 flasks for Northern blot analysis or in four-chamber slide glass for in situ hybridization (Nunc Inc., Nashville, TN).
Northern blot analysis
Total RNA was extracted from freshly isolated bone marrow cells
or from bone marrow cells cultured for 7 days using Ultraspec RNA
(Biotecx Lab. Inc., Houston, TX). Total RNA (20 µg) was
electrophoresed on a 1% agarose gel, transferred to a nylon membrane
(Hybond N+, Amersham), and the blot was hybridized with
either a 32P-labeled full-length osteopontin cDNA probe or
the housekeeping control gene glyceraldehyde-3-phosphate dehydrogenase
(GAPDH) cDNA. The cDNA probe for murine GAPDH (987 bp), used as a
control in these experiments, was amplified by PCR from cultured murine
bone marrow cells using 5'-TGAAGGTCGGTGTGAACGGATTTGGC and
5'-CATGTAGGCCATGAGGTCCACCAC as the forward and reverse primers,
respectively. Both cDNAs were labeled using the Random Primed DNA
labeling kit (Boehringer-Mannheim, Mannheim, Germany) according to the
manufacturer’s protocol. The amount of osteopontin mRNA and GAPDH mRNA
was quantified using a PhosphorImager (Molecular Dynamics, Sunnyvale,
CA).
Combination of histostaining and in situ hybridization
Cell fixation and histostaining. After 3, 5, or 7 days of
culture, bone marrow cells from either sham-operated or OVX mice were
washed with calcium-free PBS [PBS(-), treated with 0.1% diethyl
pyrocarbonate (DEPC) (Sigma)]. Subsequently, the cells were fixed with
4% paraformaldehyde in 0.1 M DEPC-treated phosphate
buffer, pH 7.4, for 60 min at 4 C. Fixed cells were washed with PBS(-)
three times, and subsequently stained for TRAP or NSE using a Sigma
diagnostic kit, or for AP using an AP substrate kit I (Vector
Laboratories, Burlingame, CA). Stained cells were washed with PBS(-),
then soaked in 0.2 N HCl for 20 min at room temperature.
After this step, cells were subjected to three washes: first, with
DEPC-treated water; second, with 2x SSC; and finally, with PBS(-). At
this stage, the cells were digested in 2 µg/ml proteinase K in 10
mM Tris buffer (pH 8.0) for 10 min at 37 C, fixed again in
4% paraformaldehyde for 30 min at 4 C, washed 3 times with PBS(-),
and dehydrated sequentially with 70%, 80%, 90%, and 100%
ethanol.
In situ hybridization. Digoxigenin RNA probes were prepared from a plasmid containing the entire murine osteopontin cDNA (1.3 kb). The plasmid was either linearized with BamHI and transcribed with SP6 RNA polymerase to generate an antisense probe or linearized with NheI and transcribed with T7 RNA polymerase to generate a sense probe. The linearized cDNA (1.3 kb) was subsequently purified by phenol/chloroform. Digoxigenin-labeled RNA was synthesized using an RNA labeling kit (Boehringer-Mannheim). For hybridization, 3 µg/ml of digoxigenin-labeled RNA probe was dissolved in hybridization buffer containing 50% formamide, 1 mg/ml transfer RNA, 20 mM Tris (pH 8.0), 2.5 mM EDTA (pH 8.0), 1x Denhardt’s solution, 0.3 M NaCl, and 10% dextran sulfate. The mixture was heated at 90 C for 10 min and then cooled on ice to linearize RNA. The hybridization buffer (50 µl) was then applied to each chamber of a 4-chamber glass slide. The slides were covered with sterile silicon-coated glass cover slips (Sigmacoat, Sigma) to prevent evaporation and kept at 50 C overnight in a humidifier. After the overnight incubation, the slides were dipped into warmed (50 C) 5x SSC to gently remove the cover slips. The slides were then washed with 2x SSC for 30 min containing 50% formamide. This step was followed by a second 10-min wash at 37 C using a TNE buffer (10 mM Tris, 1 mM EDTA and 500 mM NaCl, pH 7.6). The slides were then treated with 5 µg/ml RNase A in TNE buffer for 30 min to remove nonspecific hybridizing probe. After another wash in TNE buffer for 10 min at 37 C, more stringent washes in 2x SSC at 50 C for 20 min and 0.2x SSC at 50 C for 20 min were performed to remove any residual nonspecific binding. Messenger RNA hybridized to digoxigenin-labeled antisense probe was detected with AP-conjugated antidigoxigenin antibody and visualized with NBT and BCIP (DIG nucleic acid detection kit, Boehringer-Mannheim) after soaking in 1% blocking reagent for 30 min at room temperature. Cells expressing osteopontin mRNA stained dark purple. No staining was detected when a 1.3-kb sense probe (negative control) was used. Cells expressing osteopontin and/or staining positive for either AP, TRAP (only those containing over 3 nuclei), or NSE were counted under a light microscope using 200x magnification. A total of five fields containing 200–300 cells per field were randomly selected and analyzed by two different observers, neither of whom knew the identity of the sample.
Statistics
Data are expressed as the mean ± SD from four
replicate cultures per experiment, with a minimum of three experiments
for each experimental condition. Differences between groups were
analyzed by one-way ANOVA. Student’s t test or the
Bonferroni test were used to estimate the levels of significances
between means.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). In culture maintained in the presence of
1,25-(OH)2D3, a potent prodifferentiation
agent, the abundance of this mRNA increased. Moreover, the abundance of
osteopontin mRNA in four separate experiments was consistently greater
in bone marrow cell cultures from OVX mice compared with sham-operated
controls. The mean increase, however, was small (1.2-fold) and
statistically insignificant (P > 0.05). In view of
these results, we suspected that the small but consistent increases in
osteopontin mRNA in bone marrow from estrogen deficient mice could be
due to the fact that only a subset of cells in these heterogenous cell
preparations expressed osteopontin. Therefore, for further analysis of
osteopontin mRNA expression in bone marrow cell cultures, we employed
in situ hybridization alone or in combination with
histostaining.
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).
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). In the
absence of 1,25-(OH)2D3, the percentage of
cells expressing osteopontin mRNA was significantly lower (14% sham
and 22% OVX) compared with the percentage of cells expressing the
osteopontin mRNA in the presence of
1,25-(OH)2D3 (20% sham, 30% OVX), most likely
due to the decrease of total cell numbers described in the results of
Fig. 2
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). The presence of 1,25-(OH)2D3
decreased the number of AP-positive cells but increased the number of
TRAP-positive and NSE-positive cells, independent of whether the
animals were sham-operated or ovariectomized. However, there was a
larger increase in TRAP-positive cells in OVX compared with
sham-operated animals. This was not the case with AP-positive or
NSE-positive cells. In experiments not shown here, we added 17-ß
estradiol (10-10 to 10-6 M) to
bone marrow cultures from ovariectomized mice and found no change in
the number of TRAP-positive cells formed in culture (control, 50
± 5/well; 10-6 M 17-ß estradiol, 48 ±
11/well).
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and 6. Osteopontin mRNA expression, together with AP, TRAP,
or NSE staining, was seen in the same cells. A representative
microphotograph of cells expressing osteopontin mRNA along with either
one of the three markers is depicted in Fig. 5. Interestingly, only a
fraction (30–50%) of all AP-positive, TRAP-positive, or NSE-positive
cells expressed osteopontin mRNA. The percentage of mononucleated
TRAP-positive expressing osteopontin was very similar to the percentage
of the multinucleated TRAP-positive cells expressing osteopontin. In
view of the evidence that the AP and TRAP appear later in these
cultures (after day 5) than osteopontin mRNA (day 3), these results
suggest that osteopontin mRNA expression is an early occurrence during
osteoclastic and osteoblastic cell differentiation. As in the case of
histostaining alone, staining of cells for osteopontin mRNA by in
situ hybridization in combination with histostaining for the
phenotypic markers revealed that the presence of
1,25-(OH)2D3 decreased the number of
osteopontin-positive/AP-positive cells, whereas it increased the number
of osteopontin-positive/TRAP-positive or
osteopontin-positive/NSE-positive cells. However, the combination of
the in situ hybridization and the histostaining techniques
revealed that ovariectomy caused an increase in the numbers of all
three subsets of cells, i.e. AP-positive cells expressing
osteopontin mRNA in the absence of 1,25-(OH)2D3
and TRAP-positive and NSE-positive cells expressing osteopontin mRNA in
the presence of 1,25-(OH)2D3.
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and 8). As seen in Fig. 7A, in
contrast to cultures receiving nonimmune rabbit serum (NIRS), cultures
treated with the antiosteopontin antibody (1/250 and 1/100 dilutions)
exhibited decreased formation of osteoclast-like cells, identified as
multinucleated cells staining positive for TRAP. Whereas the number of
osteoclast-like cells was indistinguishable between cultures treated
with vehicle or with NIRS, cultures treated with 1/250 dilution of the
antiosteopontin antibody exhibited more than 50% reduction in the
number of osteoclastic cells formed. This effect appeared to be
dilution dependent, as lower dilution of the antiosteopontin antibody
(1/100) decreased the number of osteoclast-like cells even further. At
the lower dilution (1/100), NIRS had an inhibiting effect on osteoclast
formation, suggesting the presence of an inhibitory factor for
osteoclast development in rabbit serum. However, the specificity of the
antiosteopontin antibody was indicated by its effectiveness at a
dilution at which the NIRS was ineffective. As shown in Fig. 7B, an
inhibitory effect of the antiosteopontin antibody was also found in
macrophage-like cells (identified by their ability to express NSE).
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). When the antibody was
present during the entire period of culture (days 0–7), there was
inhibition of the numbers of TRAP-positive multinucleated cells. The
extent of such inhibition was similar in cultures exposed to the
antibody during days 3–5. Significant inhibition was also seen when
the antibody was present from days 0–3; however, it was smaller than
that seen in cultures treated with the antibody during days 3–5. The
greater effectiveness of the antiosteopontin antibody during the early
stages of the culture is consistent with the time course of osteopontin
mRNA expression.
Finally, in view of the possibility that osteopontin affects
osteoclastogenesis because of its ability to influence cell to cell
contact through RGD sequences, we examined the effect of RGD (or
control) peptide addition in our bone marrow cultures for independent
confirmation of the observations with the osteopontin antibody. As
shown in Fig. 9A, at 10-4 M the
RGD peptide significantly inhibited osteoclast formation, whereas the
RGE peptide (negative control) did not. RGD or RGE peptides at
concentrations as high as 10-4 M had no effect
on cell viability as evidenced by trypan blue exclusion examination.
The effective concentration (10-4 M) of the
RGD peptide was not different from that required to prevent the
attachment of mature osteoclasts to precoated matrix proteins
(osteopontin, bone sialoprotein, fibronectin) on culture dishes (24, 25). The inhibition of osteoclast formation by RGD peptide was more
pronounced when the peptide was present during the entire period (days
0–7), compared with when the peptide was added at the early stages
(days 0–3) or intermediate and late stages (days 3–7), of the culture
(Fig. 9B).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Using a combination of in situ hybridization and histochemistry, we were able to localize osteopontin expression within particular cell subtypes present in murine bone marrow cell cultures. Specifically, we determined that osteopontin is expressed by at least three different cell subtypes: TRAP-positive, AP-positive, and NSE-positive cells. TRAP is a specific and fairly reliable phenotypic markers of preosteoclasts and osteoclasts, especially when combined with multinucleation. As AP is the only available marker for cells of the stromal/osteoblastic lineage, we have used this enzyme as a putative marker for these cells. Nonetheless, we are quick to point out that several other cells of the bone marrow, such as leukocytes and preadipocytes, also express AP. Hence, the identity of the AP-positive cells in these experiments is assumed. Be that as it may, based on AP expression in combination with the morphologic features of the AP-positive cells, we are led to believe that the majority of the AP-positive cells represent bone marrow stromal/osteoblastic cells. Based on the fact that NSE-positive cells express C3 receptors staining with Mac-1 and Mac-2 antibodies, it is safe to suggest that NSE-positive cells are most likely macrophage precursors (26). NSE-positive cells appeared at the third day of culture, and their percentage reached 40–50% of the total cell number at the end of 7 days. In view of evidence that both macrophages and osteoclast-like TRAP-positive cells are derived from common precursors that stained positive for NSE (26), the appearance of NSE-positive cells early in our culture is consistent with the notion that osteoclast differentiation may be already in progress at 3 days. Among the AP-positive, TRAP-positive, or NSE-positive cell populations, only 30%–50% expressed osteopontin mRNA. However, osteopontin mRNA was expressed earlier than AP or TRAP-positive cells.
The absolute number of osteopontin-expressing cells was not affected by
1,25-(OH)2D3, even though the percentage of
osteopontin-positive cells was increased in the presence of
1,25-(OH)2D3 as a result of a decrease in the
total number of adherent cells (Figs. 2 and 3). Hence, it seems that,
unlike the case in other cell types (27, 28),
1,25-(OH)2D3 may not regulate the expression of
osteopontin in bone marrow cell progenitors.
Increases in resorption, as well as formation, in bone remodeling are well established features of estrogen deficiency in rodents and humans (2). The former, of course, exceeds the latter, thus causing loss of bone. We have reported elsewhere that ovariectomy in mice causes not only up-regulation of osteoclastogenesis but also an increase in the number of osteoblast progenitors in the bone marrow, and that the latter change is temporally associated with increased bone formation, as measured by changes in serum osteocalcin (29). In the present studies, we found that ovariectomy up-regulated the number of cells expressing TRAP alone or TRAP in combination with osteopontin. In addition, ovariectomy caused an increase in the number of AP-positive cells that coexpressed osteopontin. Furthermore, NSE-positive cells coexpressing osteopontin were more numerous in bone marrow cultures from OVX animals compared with sham-operated animals. The results of our earlier report (29) are in full agreement with the above-mentioned observations and add strength to the contention that OVX up-regulates both osteoclastogenesis and osteoblastogenesis. The findings of the present studies are also in agreement with in vivo studies indicating that osteopontin mRNA expression is increased in osteoblasts, osteoclasts, and osteocytes in metaphysical trabecular bone after ovariectomy (30). The present results, taken together with the rest of the available evidence, strongly suggest that loss of ovarian function promotes differentiation of bone marrow cells, resulting, among other changes, in increased expression of osteopontin.
The demonstration of increased expression of a gene after loss of estrogen could be either a direct consequence of the removal of an inhibitory effect of the hormone on this particular gene or a secondary event caused by an increase in the number of cells that express this gene. In line with the former scenario, we and others have demonstrated that sex steroids directly suppress the expression of the IL-6 and the gp80 and gp130 subunits of the IL-6 receptor and that ovariectomy causes an increase in the expression of all of these three genes, as determined on an individual cell basis using in situ RT-PCR (31). Because of lack of evidence that estrogen regulates the osteopontin gene directly, the present observation that ovariectomy increases the number of osteopontin-expressing cells is consistent with the latter scenario. Further, the finding that the increased number of TRAP-positive cells in bone marrow cultures after ovariectomy was not reversed by adding estrogen back into the in vitro cultures, suggests that estrogen deficiency must have caused irreversible commitment of progenitors to the osteoclast differentiation pathway.
The exact nature of the contact mediated by osteopontin in the bone
marrow is only a matter of conjecture at this stage. Nonetheless, it is
likely that osteopontin facilitates the attachment of cells to matrix
substances, as well as to other cells, and that its ability to
facilitate interactions between stromal/osteoblastic cells and
hematopoietic osteoclast precursors is responsible for the observation
that neutralization of osteopontin decreases osteoclast and macrophage
development. Osteopontin contains RGD sequences required for
interaction with the vß3 integrin family of receptors (such as
vitronectin receptors) that are expressed in osteoclasts (32, 33, 34).
Osteoclast attachment to glass surfaces precoated with osteopontin,
bone sialoprotein, or fibronectin is inhibited by RGD-containing
proteins, such as vitronectin or echistatin. The evidence that an
RGD-containing peptide inhibited osteoclast formation suggests that
osteopontin plays a role in the development of osteoclasts. Of course,
our findings do not rule out the possibility that several other
adhesion proteins (e.g. bone sialoprotein or fibronectin,
which also contain the RGD sequence) play also a role in the process of
osteoclast development.
How osteopontin can influence osteoclast development is only a matter
of conjecture. Nonetheless, it is possible that osteopontin by
mediating cell-to-cell contact between stromal/osteoblastic cells and
osteoclast progenitors, facilitates osteoclast differentiation by
enhancing the support provided by stromal/osteoblastic cells.
Alternatively, osteopontin may act to potentiate the action of
paracrine cytokines produced by stromal/osteoblastic cells that in turn
serve to promote proliferation or differentiation of the hematopoietic
precursors. The first possibility is supported by the evidence that the
formation of stromal/osteoblastic cells was decreased in the presence
of the antiosteopontin antibody. Specifically, the antiosteopontin
antibody (at 1/250 dilution) significantly decreased the formation of
large size colonies of stromal/osteoblast-like cells in the presence of
1,25-(OH)2D3 (data not shown), raising the
possibility that osteopontin may affect their proliferation. The second
possibility (i.e. osteopontin enhances the action of
autocrine cytokines), is supported by evidence that cell-to-cell
contact is essential for the action of paracrine factors anchored to
matrices on the surface of cells, such as M-CSF and GM-CSF (35, 36), as
both these cytokines exert their actions while anchored to the cell
membrane, as opposed to being released from the cells in a soluble
form. Finally, the idea that osteopontin-mediated cell-to-cell contact
influences the function of other molecules is supported by recent
evidence that osteopontin recognizes the CD44 molecule expressed on the
surface of lymphocytes (37) and osteoclasts (38), thereby inducing
CD44-dependent chemotactic activity, which does not involve the actions
of integrins. CD44 is expressed not only in osteoclasts but also in
stromal/osteoblastic cells in bone marrow cultures, and an anti-CD44
antibody inhibits generation of TRAP-positive osteoclast-like cells
(39). Thus, the interaction of osteopontin with CD44 may prove as
important for osteoclast development as the vß3 integrin mediated
cell-to-cell interactions.
In conclusion, the evidence presented in this paper provides direct demonstration that an adhesion molecule, osteopontin, is involved in the process of osteoclast and, perhaps, osteoblast progenitor differentiation in the bone marrow: and that it may be in fact required for the former process. In addition, the results of these studies with bone marrow cultures from ovariectomized mice add strength to the contention that loss of ovarian function up-regulates the differentiation of early osteoclast as well as osteoblast progenitors in the bone marrow.
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Acknowledgments |
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Footnotes |
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Received January 13, 1997.
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References |
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, 25-dihydroxyvitamin
D3 in bone marrow derived stromal cells (ST2) and primary
osteoblastic cells. Endocrinology 129:2774–2779[Abstract]
, 25-Dihydroxyvitamin
D3 regulates in vivo production of the third
component of complement (C3) in bone. Endocrinology 131:2468–2475[Abstract]
2HS-glycoprotein) proteins in rat bone. J Bone Miner
Res 8:485–496[Medline]
mRNA in the bone
marrow following estrogen loss. J Bone Miner Res 11:S105
vß3 potentiate bone
resorption. J Biol Chem 268:9901–9907vß3 by osteopontin
stimulates phosphatidylinosytol 3-hydroxy kinase activity.
Endocrinology 136:2984–2992[Abstract]
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