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* Bone and Mineral Research Program, Garvan Institute of Medical Research, Darlinghurst, Sydney, New South Wales, Australia;
Department of Pediatrics, Medical University of South Carolina, Charleston, South Carolina, USA; and
Department of Orthopaedics, Prince of Wales Hospital, Sydney, New South Wales
1Correspondence: Garvan Institute of Medical Research, 384 Victoria St., Darlinghurst, Sydney, New South Wales 2010, Australia. E-mail e.gardiner{at}garvan.unsw.edu.au
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: osteoclast • osteocyte • periosteum • uncoupling • turnover
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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and primarily affects cortical bone (2)
. In both
situations, there is gradual deterioration of the bone
microarchitecture as well as the overall loss of bone mass.
Effective osteoporosis therapies minimize bone loss, essentially by
reducing resorption (3
, 4)
. Therapeutic use of the active
1
-hydroxylated forms of vitamin D (i.e., 1,25-dihydroxyvitamin
D2 and D3, referred to
collectively as 1,25-dihydroxyvitamin D) has yielded somewhat disparate
clinical results (5
6
7
8)
, presumably because of diverse
actions on the osteoblastic and osteoclastic cell lineages as well as
on other tissues such as intestine, parathyroid, and kidney.
1,25-Dihydroxyvitamin D directly inhibits osteoblastic differentiation
(9
, 10)
, but stimulates mineralized matrix formation by
mature osteoblasts (11)
; however, it also indirectly
stimulates osteoclastic recruitment and differentiation, acting through
immature cells of the osteoblastic lineage via the vitamin D receptor
(VDR) (12)
. Regulation of osteoclastic bone resorption by
cells of the bone-forming lineage provides a mechanism by which the
opposing actions of these two cell lineages are coordinated in healthy
bone, maintaining overall bone mass and calcium homeostasis
(13)
. This model, however, is inherently incomplete as it
does not explain how bone resorption is restrained once homeostatic
requirements have been met. An inhibitory pathway that locally limits
bone resorption could complete the model of coupled bone turnover,
ensuring the maintenance of bone microarchitectural integrity.
The present study tested the hypothesis that if mature osteoblastic cells were made more responsive to 1,25-dihydroxyvitamin D relative to immature osteoblasts as well as to the osteoclastic lineage and other responsive tissues, a net positive bone effect would ensue. This strategy has been shown to be effective in a transgenic model system, with an increase in cortical bone formation. However, presence of the transgene was also associated with a decrease in trabecular bone resorption, indicating for the first time that a counter-regulatory pathway acts via mature osteoblastic cells to restrain osteoclastic activity.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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into the
human osteocalcin-based pGOSCAS vector (15)
, followed by
SV40 small t antigen splice and polyadenylation signals (Promega
Corporation, Madison, Wis.) immediately downstream. The human
osteocalcin gene is expressed solely in mature cells of the
osteoblastic lineage, and this cell type specificity was duplicated by
a pGOSCAS reporter transgene in previous work (15)
.
Transgenic mice were generated by pronuclear injection of FVB/N
embryos. The OSV9 and OSV3 lines were independently derived, each
carrying a single insertion of 5 to 10 copies of the transgene.
Hemizygous experimental animals, bred by mating homozygous males to
FVB/N females, were studied.
Age-matched female nontransgenic and transgenic mice were mixed and
group housed at weaning. This study was conducted twice. The first
study included FVB/N and OSV9; a repetition including these lines plus
OSV3 confirmed initial observations. Data shown are from the second
study. A third line, OSV8 line, showed inconsistent growth
characteristics unrelated to transgene status and was excluded from
final analyses as the level of VDR protein in its bones was not
significantly different from the FVB/N level (P=0.35). Four
weeks prior to collection, the mice were randomly assigned to two
groups and changed from standard laboratory chow (0.9% calcium) to
semi-synthetic diets (16)
with moderate (0.5%) or low
(0.1%) calcium content. Vitamin D was supplied (1000 IU/kg) in all
diets. Mice were injected with the fluorescent tetracycline compounds
calcein and demeclocycline (Sigma Chemical Company, St. Louis, Mo.),
each at 15 mg/kg, 10 and 3 days prior to collection. Tibiae were
collected and stored for mechanical testing at -20°C in
phosphate-buffered saline. Femora and vertebrae were collected, fixed
in 4% paraformaldehyde and prepared for histomorphometry. Calvaria,
radius, kidney, liver, brain, muscle, heart, lung, and spleen were
collected for molecular analyses.
Analysis of transgene expression
Total RNA was prepared from tissues of 8 wk old mice and
analyzed by Northern blot as described previously (15)
.
Filters were probed with a random primed
-32P-dCTP- (Amersham, Buckinghamshire,
England) labeled mouse VDR cDNA fragment that was cloned after reverse
transcription-polymerase chain reaction (RT-PCR) from nontransgenic
mouse kidney RNA, using primers derived from the human cDNA sequence
(forward primer 5'-CGGAATTCTCATTCTGACAGATGAGGAAGTGC-3' and reverse
primer 5'-AACTGCAGTCCTGGTATCATCTTAGCAAAGCC-3'). The filter was stripped
and reprobed for osteocalcin using a radiolabeled rat osteocalcin cDNA
insert from pOC918 (a kind gift from Dr. S. E. Harris) and for
GAPDH using a radiolabeled PCR product. Relative signals were
quantitated by PhosphorImager (Molecular Dynamics 445SI, Sunnyvale,
Calif.). Total VDR protein was measured by ELISA (see method below)
from long bones of six to eight 9 month old mice for each line, with
equal numbers of mice from the low and moderate calcium diet groups.
Biochemistry
Serum 1,25-dihydroxyvitamin D was measured by radioimmunoassay
(RIA) (17)
. PTH was also measured by RIA (Immutopics, San
Clemente, Calif.), as was serum osteocalcin, using the method of
Gundberg (18)
except 50 µl sample sizes were assayed.
Primary antibody and osteocalcin standards were generously provided by
Dr. C. Gundberg. Iodinated osteocalcin was purchased from Biomedical
Technologies, Inc. (Stoughton, Mass.) and donkey anti-goat IgG
secondary antibody from Sigma.
Total VDR protein (sample collection described above) was measured by
ELISA (19)
using antibodies generously provided by Dr. H.
DeLuca, commercially supplied biotin-conjugated alkaline phosphatase
(Bio-Rad, Hercules, Calif.), and purified VDR protein standards (Pan
Vera, Madison, Wis.). Nuclear protein extracts for VDR assay were
prepared using a protocol adapted from Pierce (Rockford, Ill.)
(20)
. Whole bones were homogenized initially using a
Polytron homogenizer and subsequently by Dounce homogenizer. Total
protein levels were determined by Bradford colorimetric assay
(Bio-Rad). Values are means ± SE.
Histology
For histology, 8 wk old animals were treated with a single
intraperitoneal injection of 1,25-dihydroxyvitamin
D3 (Tetrionics Inc., Madison, Wis.) at a dose of
2 µg per kg body weight. Femora were collected 6 h later, distal
segments were paraffin-embedded, and sagittal sections were analyzed as
previously (15)
. In situ hybridization used
antisense and sense human VDR cDNA riboprobes generated from linearized
pGhVcEBx, which contains 400 bp of 5' sequence from the human VDR cDNA.
The antisense riboprobe detects both mouse and transgenic human
transcripts. Hybridized probe was detected using the alkaline
phosphatase-coupled anti-digoxigenin antibody method (Boehringer
Mannheim, Mannheim, Germany) with the addition of 1.2 mg/ml levamisole
(Sigma) in the final staining solution. Immunohistochemistry on
sections of the same specimens used a human-specific anti-VDR antibody
(21)
, kindly provided by Dr. P. Tuohimaa. Biotinylated
goat anti-rabbit IgG was used as secondary antibody, and
avidin-biotinylated peroxidase complex and diamino-benzidine staining
(Vector Laboratories, Burlingame, Calif.) were used for detection.
Sections were not counterstained.
Histomorphometry
The fourth caudal vertebra and the distal half of the right
femur from 4- and 9-month-old animals were fixed and embedded
undecalcified in K-Plast resin (Medim-Medizinische Diagnostik, Giessen,
Germany) and 5 µm sagittal sections were analyzed (Bioquant, R&M
Biometrics Inc., Nashville, Tenn.). Femoral width was measured using
bright field microscopy, and periosteal mineral appositional rate and
vertebral bone formation rate (BFR=double-labeled surface x MAR)
by fluorescence microscopy (Leica, Heerbrugg, Switzerland). Sections
were stained for mineralized bone (22)
; trabecular bone
volume (BV/TV), thickness (Tb.Th), and number (Tb.N) were quantitated
(23)
. For measurements of osteoclast surface (Oc.S) and
number (Oc.N), sections were stained for tartrate-resistant acid
phosphatase activity as described previously (15)
.
Mechanical testing and morphometry
The biomechanical and physiological consequences of osteoblastic
and osteocytic VDR elevation were evaluated in 4 and 9 month old female
mice. Tibiae were dissected free of remaining soft tissue prior to
three-point bending tests using an MTS 858 Bionix Testing Machine (MTS
Systems Corporation, Minneapolis, Minn.) at 2 mm/min until failure.
Samples were tested immersed in normal saline at room temperature with
a support span of 10 mm. After mechanical testing, tibiae were imaged
at the fracture site at 40x magnification using a Leica stereo
microscope. Measurements and analyses were completed using the Bioquant
System (R&M Biometrics). Cortical height (diameter) and width were
measured in the sagittal plane and cortical moments of inertia were
calculated (24)
.
Statistics
Statistical analyses were performed by one-way analysis of
variance (ANOVA) within age groups with linear contrasts selected
a priori to compare results from each of the transgenic
lines with those from the FVB/N control line, *P < 0.05
(SPSS for Macintosh v. 4.02; SPSS Inc., Chicago, Ill.). The effects of
transgenic line and dietary calcium group were examined by two-way
ANOVA; n=17–20 mice per line per age group unless otherwise
indicated.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). VDR expression was undetectable in transgenic or
nontransgenic bones by immunohistochemistry or in situ
hybridization. Injection of mice with 1,25-dihydroxyvitamin
D3, however, elevated VDR expression in cuboidal
and flattened osteoblasts (Fig. 1C
, D
) and in hypertrophic
chondrocytes of transgenic but not nontransgenic bones, as expected
based on previous studies of the transgenic human osteocalcin promoter
(15)
. Signal was also detected in a minority of osteocytes
(10–20%), primarily those immediately adjacent to positively stained
osteoblastic surfaces. Immunohistochemistry with an antibody specific
for human but not mouse VDR detected VDR protein in osteoblasts and
osteocytes of bones from treated transgenic but not nontransgenic mice
(Fig. 1E
, F
). Elevation of VDR protein in untreated
transgenic mice was also apparent by ELISA measurement (Fig. 1B
), with a threefold greater level in OSV3 bones relative
to FVB/N (7.0±0.6 vs. 1.8±0.5 fmol/mg protein) and intermediate
elevation in OSV9 bones (5.7±0.7).
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Transgenic animals were healthy and phenotypically normal. As there
were no interactions between transgenic line and dietary calcium
content, diet groups were combined for all analyses. There was no
consistent transgene-associated effect on body weight, bone length
(tibia), or serum levels of calcium or calcium homeostatic hormones
(Table 1
). The transient elevation of serum PTH at 4 months was not present in
older animals. Similarly, serum osteocalcin was lower in transgenic
mice at 9 months but not at 4 months (Table 1)
or in older animals
(data not shown). Northern blot analysis revealed a reduction in
osteocalcin gene expression in 8 wk old transgenic animals (Fig. 1A
).
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Cortical bone
Tibiae from transgenic females were significantly stronger in a
three-point bending test than nontransgenic bones (mean peak load for
OSV9 and OSV3, 16% higher than FVB/N at 4 months and up to 25% higher
at 9 months; Fig. 2A
). Transgenic bones were also stiffer than their wild-type
counterparts (up to 24%, Fig. 2B
). Similar differences in
peak load and stiffness were also observed for males at the same ages,
with mean peak loads up to 27% higher than FVB/N males (16.3±0.3,
OSV3 vs. 12.8±0.3, FVB/N at 9 months, P=0.001) and
stiffness values elevated by up to 30% (78.0±2.2, OSV3 vs. 62.7±1.7,
FVB/N at 9 months, P=0.001).
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Cortical area moments of inertia (a measure of bone geometry and
determinant of bone strength) (24)
of the female tibiae
were greater in OSV9 and OSV3 than in FVB/N mice by 8 and 11%,
respectively, at 4 months, and by 23 and 34% at 9 months (Fig. 2C
). There were similar differences in tibial cortical areas
at both ages, with OSV3 up to 18% larger than FVB/N (Fig. 2D
). The greater cross-sectional areas of the transgenic
tibiae were positively correlated with their greater strengths in the
three-point bending test (Fig. 2E
). The increases in moments
of inertia and cross-sectional areas were associated with increased
cortical diameter (Fig. 2F
), consistent with an
increase in periosteal bone formation. This parameter was therefore
investigated in femora from these mice.
By histomorphometry, the OSV3 femora were also wider than those of
FVB/N femora, with diameter 7% greater at 4 and 9 months (Fig. 3A
). Femoral periosteal mineral appositional rate (MAR),
indicated by greater separation of tetracycline labels (pictured at 4
months, Fig. 3B
) was elevated in both transgenic lines at 4
months (130% increase in OSV3, 66% in OSV9), and showed a similar
pattern at 9 months (P=0.07; Fig. 3C
). These MAR
increases are consistent with the greater cortical dimensions of the
OSV3 bones. Endocortical mineral apposition rate was not altered in the
transgenic femora (data not shown).
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Trabecular bone
Trabecular bone volume measured in the fourth caudal vertebral
body (Fig. 4A
) was at least 17% greater in OSV3 and OSV9 mice than in
FVB/N at 4 months and 25% greater at 9 months (Fig. 4B
).
This difference was associated with thicker trabeculae (OSV3, 17%
greater than FVB/N at both ages) without a change in trabecular number
(Fig. 4C
, D
). In contrast to the pattern observed in
cortical bone, however, this difference in trabecular thickness was not
attributable to greater bone formation in the transgenic mice, as
neither trabecular mineral apposition rate nor double-labeled surface
(data not shown), nor bone formation rate (Fig. 4E
) was
affected by transgene status. The observed increase in trabecular bone
volume was instead associated with a reduction in bone resorption, with
osteoclast surface on vertebral trabeculae of both transgenic lines
reduced by 32% at 4 months and by greater than 40% at 9 months (Fig. 4F
). An apparent transgene-related reduction in
osteoclast number was not significant (Fig. 4G
). A similar
trend to increased trabecular bone volume and reduction in osteoclast
surface was also apparent at 4 months in the femoral metaphysis but not
the epiphysis (Fig. 4H
, I
, J
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Increased mineral apposition rate reflects an increase in the anabolic
ability of individual osteoblasts rather than a change in osteoblastic
proliferation or survival (1)
. The increased osteoblastic
bone formation on the OSV periosteal surface may most simply be
explained by a transgene-enhanced ability of 1,25-dihydroxyvitamin D to
regulate expression of bone structural or regulatory genes
(25
26
27
28)
. Although the effect of 1,25-dihydroxyvitamin D
on expression of individual genes may be either positive or negative,
the net consequence of VDR elevation in mature osteoblasts in these
mice is anabolic. As body weight and tibial length were not altered by
the transgene, it appears that periosteal osteoblasts in the transgenic
mice have a greater osteogenic response for a given range of body
weights. This is of particular interest in light of the suggestion that
age-related deficiencies in the anabolic potential of osteoblasts may
be a major contributor to age-related bone loss in humans
(29)
. The change in osteoblast activity was associated
with wider tibiae and femora in OSVDR mice. As a consequence,
transgenic tibiae were stronger and stiffer than nontransgenic bones.
The modest increases in periosteal bone deposition result in
substantial gains in long bone strength because area moment of inertia
increases with the fourth power of the radius of the bone
(24)
.
The transgene effect on bone formation was envelope specific, being evident only on the periosteal surface of the long bones but not on endosteal or trabecular surfaces, suggesting that the transgene affects osteoblasts of periosteal but not bone marrow origin. In contrast, the transgene effect on bone resorption presumably was mediated by osteoblastic cells originating from the bone marrow, as trabecular osteoclast surface was reduced in the fourth caudal vertebrae and the femoral metaphyseal region. No transgene effect on resorption or formation was detected in the femoral epiphysis, indicating site specificity. These variations in cellular responses to transgene expression suggest that the osteoblastic and osteoclastic phenotypes may result from transgene interaction with local factors in the bone microenvironment, as discussed below.
In caudal vertebrae, thickening of trabeculae and reduction of bone
resorption occurred despite the maintenance of normal or slightly
elevated levels of serum 1,25-dihydroxyvitamin D and PTH in the
transgenic mice. This surprising result was not predicted by the large
number of previous studies showing that 1,25-dihydroxyvitamin D acts
through immature osteoblasts and stromal cells to stimulate
osteoclastic recruitment and activity by direct and indirect mechanisms
(12)
. Rather, the inhibition of bone resorption evidently
results from the elevated sensitivity of mature osteoblasts to normal
endogenous levels of 1,25-dihydroxyvitamin D. Such a coupling of
pathways may provide a system for local control of bone turnover and
maintenance of microarchitectural integrity. The present approach
allowed the specific responses of mature osteoblasts to be assessed
without altering the sensitivity of immature osteoblastic and stromal
cells, in contrast to earlier studies in which intact animals, mixed
cell populations in culture, or tissues were treated with active
vitamin D compounds. Given the strength of the proresorptive response
of immature osteoblastic and stromal cells to 1,25-dihydroxyvitamin D,
it is unlikely that this counter-regulatory pathway would be detected
in vivo without an experimental enhancement of the mature
osteoblast response.
A previous study that used a similar mouse osteocalcin gene-based
transgene expression system to transiently or chronically ablate
differentiated but still proliferating osteoblastic cells indicated
that there is no obligatory cross-regulation between bone formation and
resorption (30)
. Although our data do not address the
issue of necessity, they do indicate that differentiated osteoblastic
cells are able to decrease resorption. There was no indication in the
previous study that ablation of differentiated osteoblastic cells
caused an increase in osteoclastic resorption, as would be expected
from the OSVDR findings. This may be because only proliferating cells
were ablated, leaving lining cells and osteocytes intact and
functional. If so, the implication is that the negative regulation of
osteoclastic resorption may be mediated by these post-proliferative
cells.
Possible mechanisms
Levels of the circulatory factors 1,25-dihydroxyvitamin D and PTH
were not consistently changed and certainly were not reduced in the
OSVDR mice, indicating that cortical and trabecular transgene effects
are paracrine rather than endocrine. Paracrine pathways may act via
soluble mediators such as growth factors or cytokines or via cell–cell
or cell–matrix interactions. The mechanisms underlying the transgene
effects on bone formation and resorption may involve multiple vitamin
D-responsive pathways, some of which may affect both processes.
One system that could mediate the resorption effect is the network of
tumor necrosis factor family members, which has recently been shown to
regulate osteoclastogenesis. This family includes the NF
-B receptor
activator RANK on osteoclast precursors, its ligand RANKL on immature
osteoblastic cells, and the soluble decoy receptor osteoprotegerin
(OPG) (31)
. As RANKL and OPG are normally expressed by
osteoblastic cells and regulated by 1,25-dihydroxyvitamin D
(32)
, a transgene-associated decrease in the local
RANKL/OPG ratio could reduce osteoclastic recruitment/activation. To
date, however, there is limited information about the regulation of
these genes in mature osteoblastic cells. Another cytokine that may
participate in similar regulatory systems is OCIL, a recently described
cytokine that inhibits osteoclastogenesis, is expressed by mature
osteoblastic cells and is up regulated by 1,25-dihydroxyvitamin D
(33
34
35)
. Identification of the factor(s) involved in the
transgenic reduction in bone resorption is the subject of ongoing
investigation.
In addition to the structural changes already described, material
properties of the transgenic bones may also contribute to increased
OSVDR bone strength and/or stiffness. Moreover, extracellular matrix
composition can alter bone cell biology and gene expression
(36)
and thus may also contribute to the OSVDR bone
phenotype, as 1,25-dihydroxyvitamin D is a common regulator of bone
matrix protein genes. For example, a reduction in osteocalcin protein,
as suggested by Northern blots, could contribute to the increase in
periosteal mineral apposition rate, as it has been suggested that
osteocalcin is an inhibitor of bone formation (37
, 38)
.
Studies of the osteocalcin-deficient mouse indicate, however, that a
significant decrease in osteocalcin protein would cause an increase in
osteoclast surface on trabecular bone rather than the decrease observed
in the OSVDR mice (38)
.
Given that the mechano-transducing osteocytes (39)
and
osteoblastic lining cells are sites of elevated VDR in the transgenic
mice, response to weight bearing may be altered in the OSVDR bones. As
noted above, however, periosteal bone formation was enhanced but
endosteal and trabecular formation were unaffected by the transgene,
indicating that any mechano-sensory contributions to the transgenic
osteoblastic phenotype may be envelope specific. The transgene effect
on trabecular osteoclast surface is also site specific, as it was
evident in the fourth caudal vertebrae and the femoral metaphyseal
region but not apparent in the more heavily loaded (40)
femoral epiphysis. These local differences in cortical and trabecular
bone responses to OSVDR could relate to differential expression of
regulatory genes that are expressed by osteocytes, regulated by
1,25-dihydroxyvitamin D, and change levels in response to mechanical
loading such as c-fos or osteopontin (28
, 41
42
43)
.
This project was undertaken to model a potential therapeutic approach based on the hypothesis that net bone formation would result if mature osteoblastic cells could be made more responsive to vitamin D. As the VDR is normally expressed in this cell type, this genetic alteration represents a subtle enhancement in the context of otherwise normal calcium homeostasis and bone physiology. The reduced trabecular bone resorption in the OSVDR mice highlights a specific osteoclastic inhibitory mechanism. It acts via mature osteoblasts, where the transgene is expressed, and counterbalances osteoclastogenic signals from immature osteoblasts and osteoblastic stromal cells. This novel negative regulatory activity may also be controlled by other calcium homeostatic regulators such as PTH, interleukins, and prostanoids. A therapeutic approach that specifically enhances mature osteoblastic responses to endogenous or exogenous agents could constitute an ideal strategy for osteoporosis treatment, decreasing trabecular bone resorption and increasing cortical bone formation, thereby reversing the typical patterns of osteoporotic and age-related bone loss.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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3 Current address: Department of Endocrinology, Prince of Wales Hospital, Sydney, New South Wales 2031 Australia.
4 Current address: Genomics Research Centre, Griffith University Gold Coast Campus, Queensland 9726, Australia.
Received for publication December 24, 1999.
Revision received March 29, 2000.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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