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V. A. Connecticut Healthcare System, Newington, Connecticut 06111; and The University of Connecticut Health Center, Farmington, Connecticut 06030
Address all correspondence and requests for reprints to: Dr. Sun-Kyeong Lee, Division of Endocrinology, Department of Medicine, University of Connecticut Health Center, 263 Farmington Avenue, Farmington, Connecticut 06030-1850.
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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B (RANK) messenger RNA
(mRNA) in cultured murine bone marrow, calvaria, and osteoblasts.
TRANCE, OPG, and RANK are recently identified regulators of osteoclast
formation. Bone marrow cells were cultured with or without PTH(1–34)
for 6 days. TRANCE, OPG, and RANK mRNA were measured by RT-PCR. In
6-day cultures, PTH stimulated the number of OCL/well in a
dose-dependent manner. A time course showed significant
(P < 0.01) increases in OCL/well after 24 h
of PTH (100 ng/ml). TRANCE mRNA expression, like OCL formation,
increased dose dependently and was maximal, with 10–100 ng/ml PTH. In
contrast, OPG mRNA expression was decreased by 0.1 ng/ml PTH (40%) and
completely abolished by 1 ng/ml. TRANCE mRNA expression was rapidly
stimulated by PTH (maximal response at 1 h, 8.1-fold over
control). Expression declined by 40% at 24 h but was still much
greater than control at 6 days (4.6-fold) in a time-course study. PTH
caused a transient stimulation of OPG mRNA at 1 h (2-fold), which
returned to basal levels by 2 h. After 6 h, PTH completely
inhibited OPG mRNA. There were only minor effects of PTH on RANK mRNA
expression. PTH had less potent effects on TRANCE and OPG mRNA
expression in calvaria organ cultures and osteoblasts. In mouse
calvaria cultures, TRANCE expression was detectable in controls and was
increased 2.9-fold by PTH at 24 h. PTH treatment of calvaria
decreased OPG expression by 30% at 6 h. MC3T3 E-1 osteoblastic
cells expressed minimal levels of TRANCE mRNA either before or after
PTH treatment. OPG mRNA was present in MC3T3 E-1 cells, but levels were
not modulated by PTH. In primary osteoblastic cells, PTH stimulated
TRANCE mRNA expression 4-fold at 2 h and inhibited OPG mRNA
expression by 46%.
These results demonstrate a tight correlation between the ability of
PTH to stimulate OCL formation in marrow culture and expression of
TRANCE (r = 0.87, P 
0.05) and OPG mRNA (r=
-0.88, P 0.05). Reciprocal regulation of TRANCE
and OPG mRNA by PTH preceded its effects on OCL formation by 18–23 h.
Hence, it is likely that PTH regulates bone resorption, at least in
part, via its effects on TRANCE and OPG expression.
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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TRANCE [tumor necrosis factor (TNF)-related activation-induced cytokine], which is also known as ODF (osteoclast differentiation factor), RANKL (receptor activator of NF-kappaB ligand), and OPGL (osteoprotegerin ligand), is a new member of the TNF ligand family that is induced upon T cell receptor binding (4, 5, 6). TRANCE was cloned during a search for apoptosis-regulatory genes using a somatic cell genetic approach in T cell hybridomas (4). TRANCE stimulates osteoclast differentiation and bone resorption (7, 8). It induces osteoclast-like cell (OCL) formation in spleen cells that are cultured without osteoblasts or stromal cells but with M-CSF (macrophage colony stimulating factor) (9). TRANCE acts directly on osteoclast progenitors (10) and induces OCL formation in human peripheral blood mononuclear cell cultures in vitro (11). It also induces bone resorption via activation of mature osteoclasts and inhibition of osteoclast-apoptosis (7, 8, 12).
OPG, which is also known as osteoclastogenesis inhibitory factor
(OCIF), is a soluble receptor for TRANCE (7, 13). OPG inhibits
osteoclastogenesis by interrupting cell-to-cell signaling between ST-2
cells and osteoclast progenitors (10), and recombinant OPG blocks
osteoclastogenesis in vitro by inhibiting the
differentiation of osteoclasts (14, 15). OPG is present as a
heparin-binding secretory glycoprotein that forms both 60-kDa monomer
and a membrane-bound 120-kDa homodimer (16, 17, 18). Human and mouse OPG
genes have been cloned and characterized. OPG is a single-copy gene
with 5 exons that spans 29 kb in humans (19) and mice (20). OPG is also
a receptor for TRAIL (TNF-related ligand), which blocks the
antiosteoclastogenic activity of OPG (21). OPG knockout mice develop
severe osteoporosis and have increased osteoclastogenesis, marked bone
loss, destruction of their growth plate, and lack trabecular bone in
their long bones (22, 23). Transgenic mice that overexpress OPG are
osteopetrotic and lack osteoclasts (14). OPG prevents bone loss in
ovariectomized rats and increases bone mineral density and bone volume
in normal rats (14). Expression of OPG is down-regulated by PG
E2 treatment in human bone marrow cells (24). In contrast,
OPG is up-regulated by interleukin (IL)-1
in the human osteosarcoma
cell line MG-63 and human osteoblast-like cells (25) and by TNF- and
-ß in MG63 cells (26).
RANK (receptor activator of NF B), also known as ODAR (osteoclast
differentiation and activation receptor) (27), is a new member of the
TNF receptor family that was first identified in dendritic cells. It is
a membrane-bound receptor for TRANCE/RANKL (6). RANK directly mediates
TRANCE-induced osteoclast differentiation and activation in osteoclast
precursor cells (27). IL-4 and TGF-ß induced surface expression of
RANK on either phytohemagglutinin (PHA)- or anti-CD-3-activated
peripheral blood T lymphocytes (6).
In this study, we investigated whether PTH regulated TRANCE, OPG, and RANK messenger RNA (mRNA) expression in mouse bone marrow cells, mouse calvarial cultures, the immortalized osteoblastic cell line MC3T3 E-1, and primary osteoblastic cells.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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-MEM, and cultured (1 x 106
cells/cm2) in -MEM containing 10% heat-inactivated FCS
(HIFCS). Cultures were fed every 3 days with fresh medium. Bovine PTH
(1, 2, 3, 4, 5, 6, 7, 8, 9, 10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31, 32, 33, 34) (bPTH) [Bachem California, Inc. (Torrance,
CA); 1 pg–100 ng/ml] was added to cultures, as indicated in each
experiment. Cells were fixed, on day 6 of culture, with 2.5%
glutaraldehyde in PBS for 30 min at room temperature before being
stained for TRAP. Enzyme histochemistry for TRAP was performed with a
commercial kit (Sigma Chemical Co., St. Louis, MO). In
some experiments, radiolabeled [125I]-salmon CT
(NEN Life Science Products, Boston, MA) was incubated with
or without excess cold salmon CT (10-7 M, 10
million-fold excess), washed, and developed by autoradiography to
demonstrate the presence of CTR on cells. Briefly, cells were plated on
slideflasks (1 million cells/cm2) and, at the end of
culture cells, were incubated with radiolabeled
[125I]-salmon CT (0.04 µCi, 100,000 cpm/ml) in the
absence or presence of cold sCT (10-7 M;
Bachem California, Inc.) at room temperature for 2 h.
Cells were then washed with PBS twice to remove nonspecific
radioactivity and were fixed with 2.5% glutaraldehyde in PBS. Slides
were dipped in LM-1 photographic emulsion (1:1 dilution with 1.7%
glycerol; Amersham, Arlington Heights, IL) for
autoradiography. Cells were then developed and stained with Giemsa. For PCR amplification, cells were plated in 10-cm culture dishes, at 1 x 106 cells/cm2, with 10 ml of medium/dish. Cultures were fed every third day, and total RNA was extracted from cells on day 6 or at the appropriate time point.
Mouse calvaria were removed from 2- to 3-day-old CD-1 mice and dissected free from adhering soft tissue, as previously described (34). Calvaria were precultured overnight in DMEM with 5% HIFCS and stimulated with or without PTH (100 ng/ml) for up to 48 h.
Primary osteoblasts from neonatal calvaria of CD-1 mice (Charles River Farms) were produced by sequential digestion, as previously described (35), and MC3T3 E-1 cells were cultured in DMEM containing 10% HIFCS with or without PTH (100 ng/ml) for up to 2 h.
PCR amplification
Total RNA was extracted from the bone marrow cultures,
osteoblastic cells, MC3T3 E-1, or calvaria organ cultures, by either
the acid guanidine isothiocyanate extraction and cesium chloride
ultracentrifugation method (36) or with Tri-reagent (Molecular Research Center, Inc., Cincinnati, OH). Total RNA was converted
to complementary DNA (cDNA, Gaithersburg, MD) by reverse transcriptase
(Superscript II, Life Technologies, Inc., Gaithersburg, MD) and random hexamer. Sonicated salmon sperm DNA was
used as a carrier. The first-strand cDNA was extracted with
phenol/chloroform, precipitated, resuspended in sterile water, and
amplified by PCR.
PCR amplification was done using gene-specific PCR primers and Taq polymerase (Amplitaq, Perkin-Elmer Corp., Norwalk, CT). The PCR mixture (without enzyme) was overlaid with mineral oil and heated to 94 C for 5 min. During the last minute, Amplitaq was added (hot start) and amplification was allowed to proceed in a thermal cycler (Perkin-Elmer Corp.). The temperature cycling was as follows: denaturation at 94 C for 1 min, primer annealing at 65 C for 2 min, and extension at 72 C for 3 min for 10 cycles. In subsequent cycles, the primer annealing temperature was decreased stepwise (step-down method), by 5 C, every 5 cycles. After the last cycle, the mixture was incubated at 72 C for 7 min. To verify that amplifications were in the linear range of each PCR analysis, we performed PCR amplification for up to 36 cycles with each amplimer set using samples prepared from either control or PTH-treated cultures. For bone marrow studies, TRANCE was amplified from control cultures and cultures treated with PTH for 1 h, which is the time when TRANCE mRNA levels seemed to be maximal. For OPG, we used control bone marrow, because PTH treatment had a predominantly inhibitory effect. For RANK, we used mRNA from bone marrow cultures that were treated with PTH for 6 days. Specific amplimer sets were designed from published cDNA sequences: murine TRANCE (6) (antisense: 5'GGGAATTACAAAGTGCACCAG3'; sense: 5'GGTCGGG-CAATT CTGAATT3'), murine OPG (14) (antisense: 5'TCAAGTGCTTGAGGGCATAC3'; sense: 5'TGGAGATCGAATTCTGCTTG3'), murine RANK (6) (antisense: 5'GTCTTCTGGAACCA TCTTCTCC3'; sense: 5'CACAGACAAATGCAAACCTTG3'), ß-actin (37) (antisense: 5'CT-CTTTGATGTCACGCACGAT-TTC3'; sense: 5'GTGGGCCGCTCTAGGCACCAA3'). The amplified samples were run in a 1.5–2.0% agarose gel (depending on the product size), stained with ethidium bromide, and photographed under UV illumination. Images were captured by a FOTO/Analyst Archiver Electronic Documentation system (Fotodyne, Inc., Hartland WI), and optical density was determined using a digital image processing and analysis program (Scion Image, Scion Corp., Frederick, MD).
The identities of the amplified PCR products were confirmed by direct sequencing using an automatic DNA sequencer (PE Applied Biosystems, Norwalk, CT).
Statistical analysis
Statistical analysis was performed by one-way ANOVA, and by the
Bonferroni post hoc test when ANOVA demonstrated significant
differences. All experiments were repeated at least twice, and
representative experiments are shown.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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). The
effect was significant for PTH at 1 to 100 ng/ml. As shown in Table 2, PTH (100 ng/ml) significantly
(P < 0.01) increased OCL formation after 24 h,
and this effect was maximal after 4 days. In these time-course
experiments, PTH was added during the last period of culture. At all
time points, unstimulated cultures contained less than 10 OCL/well.
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, A and B). Binding of
[125I]-sCT was specific because there were only
background levels of grains accumulated over MNCs and mononucleated
cells in the presence of excess unlabeled sCT (10-7
M) (Fig. 1, C and D). In both control and PTH-treated
cultures, at all times, greater than 98% of the MNC were positive for
[125I]-sCT binding.
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), and was highly correlated with
increases in OCL numbers in Table 1
(r = 0.87, P
0.05). In contrast, OPG mRNA expression was decreased 40% by 0.1
ng/ml PTH and essentially abolished by concentrations of 1 ng/ml or
higher. Expression of OPG mRNA was inversely correlated with increases
in OCL numbers in Table 1 (r = -0.88, P 0.05).
RANK mRNA expression was low in control cultures, and PTH had a slight
biphasic effect in the dose response study shown in Fig. 2, causing an
approximately 50% inhibition at 0.001–0.1 ng/ml and no
consistent effect at 100 ng/ml (Figs. 2 and 3). In time-course studies, PTH
stimulated TRANCE mRNA expression approximately 8.1-fold between 1 and
6 h (Fig. 3). Expression declined from peak levels at 24 h
but was still much greater than control at 6 days (4.6-fold). PTH also
had a biphasic effect on OPG mRNA expression. It caused a transient
stimulation of levels at 1 h (2-fold), which returned to basal
values by 2 h. After 6 h, it completely inhibited OPG mRNA.
There was little effect of PTH (100 ng/ml) on RANK mRNA expression at
any time point. To investigate the consistency of PTH (100 ng/ml)
effects on TRANCE, OPG, and RANK mRNA expression in 6-day cultures and
to perform statistical analysis, results from four independent
experiments were pooled (Fig. 4). This
analysis demonstrated that PTH stimulated TRANCE 4.4-fold, completely
inhibited OPG, and had no effect on RANK mRNA expression.
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, TRANCE, OPG, and RANK were
amplified by PCR for 30 cycles, whereas ß-actin was amplified for 27
cycles. To verify that we measured TRANCE, OPG, RANK, and ß-actin
mRNA in the linear range of each PCR analysis, we performed PCR
amplification for each primer set over a range of 18–36 cycles (Fig. 5). In these experiments, samples from
either control or PTH-treated bone marrow cultures were used for TRANCE
mRNA. Samples from control bone marrow cultures were used for OPG mRNA.
Samples from PTH-treated bone marrow cultures were used for RANK mRNA,
and a combination of control and PTH-treated bone marrow cultures was
used for ß-actin mRNA. For TRANCE, PCR amplification was linear
between cycle numbers 27 and 36 for control cultures and between 24 and
36 cycles for PTH-treated cultures. For OPG and RANK, PCR amplification
was linear between cycle numbers 27 and 36. For ß-actin, PCR
amplification was linear between 24 and 36 cycles. Hence, for all the
data shown in
Figs. 2–4, measurement of TRANCE, OPG, RANK, and
ß-actin mRNA was performed in the midlinear range of PCR
amplification.
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(data not shown). As shown in
Fig. 6, the level of TRANCE mRNA
expression was minimal in control cultures and increased 1.5-, 2.8-,
and 2.9-fold at 1 h, 6 h, and 24 h, respectively, by PTH
treatment. At 48 h, TRANCE levels had decreased 30%, compared
with cultures treated for 24 h, but were still 2-fold greater than
control. In contrast to the marrow cultures, PTH had only a small
inhibitory effect on OPG mRNA expression at any time point (maximum
inhibition of 30% at 6 h). Dose response studies (Fig. 7), performed at 6 h, demonstrated
TRANCE mRNA expression to increase 2-, 3.9-, and 4.6-fold with 1, 10,
and 100 ng/ml PTH, respectively. As with bone marrow cultures, there
was little effect of PTH treatment on RANK mRNA expression in calvaria
cultures.
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). MC3T3 E-1 cells expressed
minimal levels of TRANCE mRNA, either in control cultures or after PTH
treatment. In contrast, OPG mRNA expression was detected in control
cultures of these cells. However, PTH did not affect OPG mRNA
expression in MC3T3 E-1 cells. Primary osteoblastic cells expressed
TRANCE mRNA in control cultures and PTH stimulated expression 2.2-fold
and 4-fold at 1 and 2 h, respectively, in 25 cycle-PCR
amplification. In addition, PTH decreased OPG mRNA levels by 30% at
1 h and 46% at 2 h in primary osteoblasts. RANK mRNA
expression was not detectable in MC3T3 E-1 cells, and only minimal
levels were seen in primary osteoblastic cells, which likely represents
contamination of these cultures with osteoclast precursors or other
hematopoietic cells.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Further evidence for a link between the effects of PTH on OCL formation and its effects on TRANCE and OPG mRNA comes from time-course studies. PTH rapidly stimulated TRANCE mRNA (at 1 h) and inhibited OPG mRNA (at 6 h). In contrast, PTH increased OCL numbers in the marrow cultures only after 24 h of treatment. These findings suggest that regulation of TRANCE and OPG mRNA is an early step in the stimulation of OCL by PTH in bone marrow cultures. The effects of PTH on OCL formation in murine bone marrow cultures may also be regulated by changes in M-CSF expression, because this cytokine is necessary for osteoclast formation from precursor cells (38), and PTH stimulates M-CSF expression in osteoblastic cells (39).
It is unlikely that PTH stimulates OCL formation through effects on RANK mRNA expression, because we found only small inhibitory effects of PTH on RANK mRNA levels in murine marrow cultures and no effect of PTH on RANK mRNA in calvaria organ cultures or primary osteoblastic cells. However, it is also possible that an additional receptor on osteoclast precursor cells may bind TRANCE and mediate its effects on osteoclast formation.
Expression of TRANCE and OPG in the murine bone marrow cultures probably occurs in the stromal cell populations, because these cells are known to be critical for osteoclast formation, and they respond to stimulators of resorption (7, 14). Interestingly, we found less regulation of OPG and TRANCE mRNA expression by PTH in primary mouse osteoblasts and murine calvaria cultures. Similar lower levels of TRANCE and OPG regulation, compared with bone marrow cultures, were demonstrated in primary osteoblast, in a recent report (40). Our findings demonstrate that expression of TRANCE and OPG mRNA in bone marrow cultures is extremely sensitive to PTH. Hence, these results suggest that a major role of PTH in bone is to regulate expression of TRANCE and OPG in cells of the bone marrow cultures. Because we found different levels of TRANCE and OPG regulation by PTH in bone marrow cells, calvaria, and osteoblasts, it is likely that there are complex mechanisms influencing TRANCE and OPG mRNA expression in bone. It seems that the greatest regulation of these factors is in the less differentiated stromal elements of the bone marrow cells, because these are present in bone marrow populations to a much greater degree than in primary osteoblasts or organ cultures.
MC3T3 E-1 cells are reported to not support osteoclast differentiation from precursor cells (3). Our data would suggest that a major reason for this is because they produce little TRANCE but do produce OPG in both the basal state and after PTH stimulation. Hence, these cells express the mRNA of an inhibitor of osteoclast formation without expressing significant amounts of a major stimulator, TRANCE.
Regulation of TRANCE and OPG are likely critical pathways through which many, and possibly all, stimulators of resorption generate an osteoclastic response. Therefore, it will be interesting to determine which mechanisms are involved in these responses. It is likely that multiple pathways are used and that regulation is complex and dependent on a number of factors, including the differentiation state of the cell types that are studied and the time after stimulation when they are examined.
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Footnotes |
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Received November 23, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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,25-dihydroxyvitamin D3,
prostaglandin E2, or parathyroid hormone in the
microenvironment of bone. Biochem Biophys Res Commun 246:337–341[CrossRef][Medline]
in
the human osteosarcoma cell line MG-63 and in human osteoblast-like
cells. Biochem Biophys Res Commun 248:696–700[CrossRef][Medline]
and -ß up-regulate
the levels of osteoprotegerin mRNA in human osteosarcoma MG-63 cells.
Biochem Biophys Res Commun 248:454–457[CrossRef][Medline]
, 25dihydroxyvitamin D3 in osteoclast-like
multinucleated cells formed from mouse bone marrow cells. Endocrinology 123:1504–1510[Abstract]
and ß by newborn
mouse calvarial cultures. J Bone Miner Res 5:77–83[Medline]
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