| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
Women’s Heath Research Institute (P.V.N.B., Y.P.K., F.J.B., B.S.K.), Wyeth Research, Collegeville, Pennsylvania 19426; Aventis Pharmaceuticals (W.Z.), Bridgewater, New Jersey 08807; Pathology (A.-J.L.), Wyeth Research, Chazy, New York 12921; Investigational Pathology (M.B.G.), Wyeth Research, Andover, Massachusetts 01818; and Department of Cell Biology (T.G., G.S.S., J.B.L.), University of Massachusetts Medical School, Worcester, Massachusetts 01655
Address all correspondence and requests for reprints to: Dr. Peter V. N. Bodine, Women’s Health Research Institute, Wyeth Research, 500 Arcola Road, Collegeville, Pennsylvania 19426. E-mail: bodinep{at}wyeth.com.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
rnusse/wntwindow.html). These proteins bind to a membrane receptor complex composed of a Frizzled (FZD) G protein-coupled receptor and a low-density lipoprotein receptor-related protein (LRP) (2, 3). This binding activates one of several intracellular signaling pathways depending upon the Wnt, FZD receptor, and cell type involved (1, 2, 3). One of these is the canonical or Wnt/ß-catenin pathway that starts with activation of disheveled (Dsh) and leads to inhibition of glycogen synthase kinase-3ß and subsequent stabilization of ß-catenin. This protein then translocates to the nucleus, where it binds to and activates lymphoid-enhancer binding factor/T-cell transcription factors (1, 2, 3, 4). Additional noncanonical pathways are also activated by Wnts (2, 3). These include the G protein-mediated or Wnt/calcium pathway (5) and the disheveled (Dsh)-mediated c-Jun NH2-terminal kinase pathway (6). Previous studies of Wnt function have elucidated the importance of these proteins in skeletal development (7). However, more recent work has also identified a role for Wnts in postnatal bone formation (8, 9). Loss-of-function mutations in the Wnt coreceptor LRP5 have been shown to cause osteoporosis pseudoglioma syndrome in humans (10). This syndrome leads to low peak bone mass and premature fractures that result from a defect in bone accrual. Characterization of an LRP5–/– mouse model for osteoporosis pseudoglioma showed that osteoblast proliferation and activity was suppressed, and that bone formation and trabecular bone volume was diminished (11). On the other hand, a gain-of-function mutation in LRP5 has the opposite effect and causes a high bone mass (HBM) trait in humans (12, 13) and increases trabecular bone volume, cortical thickness, and bone formation in mice (14). Additional gain-of-function mutations of human LRP5 also lead to endosteal hyperostosis, Van Buchem disease, autosomal dominate osteosclerosis, and osteopetrosis type I (15). Although these studies have defined a role for LRP5 in bone formation, the function of additional Wnt pathway regulators in this process remains to be established.
Bone is formed and maintained by two cell types: osteoblasts that synthesize and mineralize the extracellular matrix, and osteoclasts that resorb the calcified tissue (9, 16, 17). Osteoblasts arise from multipotent mesenchymal stem cells that are located in bone marrow (9, 17, 18), whereas osteoclasts originate from hematopoietic bone marrow cells (9, 19). These cells work together in a process known as bone remodeling, which is the mechanism by which immature, damaged, or aged bone is replaced with new lamellar bone (20). Bone remodeling is initiated by recruitment and activation of osteoclasts that remove the mineralized matrix. The process ends approximately 6 months later with the filling-in of the resorption pit with newly formed osteoid by the osteoblasts. At the end of this last phase, the bone-forming cells have one of three fates (9, 17, 21). They can differentiate to osteocytes upon entrapment within the mineralized matrix; they can become quiescent lining cells; or they can undergo apoptosis (21). Although 50–70% of osteoblasts die at the end of the remodeling cycle, the mechanisms controlling this fate are not well understood.
Many extracelluar and intracelluar proteins control Wnt signaling (1, 2, 3). Among the extracelluar regulators are a variety of secreted proteins that include Wnt inhibitory factor, secreted FZD-related proteins (sFRPs), Cerberus, and dickkopfs (DKKs) (1, 2, 3, 22, 23). These proteins bind Wnts directly (Wnt inhibitory factor, sFRPs, Cerberus), bind to FZD receptors (sFRPs), or interact directly or indirectly with LRPs (DKKs). Inhibitors like sFRPs that bind Wnts or FZD receptors have the ability to blunt all Wnt-activated pathways (23), whereas DKKs only suppress the canonical pathway (22).
In preliminary studies, we identified sFRP-1 as an important regulator of osteoblast and osteocyte survival in vitro (24). Expression of sFRP-1 increased with advancing osteoblast differentiation and peaked in the preosteocytic stage. This increased expression correlated with inactivation of the canonical pathway and acceleration of apoptosis, which has also been observed with other cell types (23, 25, 26). These observations led us to postulate that at least one function of Wnts in bone is to regulate osteoblast and osteocyte survival. Like other members of the sFRP family of Wnt antagonists, sFRP-1 has a cysteine-rich domain (CRD) in its amino terminus that sequesters Wnts and prevents these proteins from binding to the FZD receptor/LRP complex (23). The protein also binds directly to the FZD receptor and may subsequently block intracellular signaling (27). The carboxy terminus contains a netrin domain that has also been implicated in sFRP-1 function (28, 29, 30), as well as a hyaluronan binding motif (31).
To determine whether sFRP-1 regulates osteoblast and osteocyte viability in vivo, we developed an sFRP-1–/– mouse line. Using these animals, we show that deletion of sFRP-1 not only reduces osteoblast and osteocyte apoptosis, but also potentiates osteoblast proliferation and differentiation, and increases trabecular bone formation. We find that loss of sFRP-1 prolongs and enhances trabecular bone accumulation and inhibits bone loss in adult mice while having little effect on cortical bone and most nonskeletal tissues. Thus, the predominant role for sFRP-1 appears to be the regulation of Wnt signaling in osteoblasts that reside in adult bone.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
). This exon was replaced with a LacZ/MC1-Neo selection cassette so that sFRP-1 promoter activity could be followed with ß-galactosidase expression in –/– mice. PCR analysis of DNA samples collected from weanlings demonstrated that sFRP-1+/+ mice carried the +/+ allele, whereas sFRP-1–/– animals lacked this allele and contained the LacZ gene (Fig. 1B). As expected, sFRP-1+/– mice had both alleles.
|
). Other tissues like heart and spleen expressed intermediate levels of mRNA, whereas gut (small intestine and colon), lung, bone (tibia), brain, and skeletal muscle contained lower message levels. On the other hand, liver did not express the gene. Of note is the observation that murine ovaries express high levels of sFRP-1, which was not reported previously (32). The human tissue distribution of sFRP-1 is similar to that observed in mice, with the exception that the heart appears to express the highest levels of the message, whereas ovary, testis, and kidney contain intermediate amounts of mRNA (25, 33).
To confirm that –/– mice no longer expressed sFRP-1, Northern blot analysis was performed with poly adenylate [Poly A(+)] RNA isolated from kidneys of +/+ and –/– animals at 16–18 wk of age (Fig. 1D). As expected, this tissue showed high levels of sFRP-1 mRNA expression in +/+ females and males, whereas –/– mice did not express the message. Because the Northern blot was performed with a probe that hybridized to the CRD domain, we also isolated total RNA from calvarial bones of 32-wk-old females and measured sFRP-1 message levels by real-time RT-PCR analysis using primer and probe sets for either the 5' region (CRD) or the 3' region (netrin domain) (Fig. 1E). This analysis confirmed that sFRP-1–/– mice did not express either portion of the gene. Microarray analysis of these RNA samples indicated that expression of sFRP-2 and –4 messages did not change after loss of sFRP-1, whereas other sFRP family members were not detected in either +/+ or –/– bones (data not shown).
Despite the broad tissue distribution of sFRP-1 mRNA in +/+ animals, –/– mice appeared normal without an overt phenotype. They were also fertile and viable up to 18 months of age or more (data not shown). We did not observe consistent differences between +/+ and –/– mice in body or most organ weights when examined between 20 and 40 wk of age. Moreover, gross and microscopic histology of 41 nonskeletal tissues from males and females at 18–20 and 39–40 wk of age did not reveal any clinically relevant pathologic changes in sFRP-1–/– animals (listed in Materials and Methods). However, the analysis did identify slight hematologic changes in the 40-wk-old –/– males with decreases in total white blood cell and red blood cell counts, as well as decreased values for both segmented neutrophils and lymphocytes. There was also a decrease in the neutrophil to lymphocyte ratio in SFRP-1–/– males at 20 and 40 wk of age. sFRP-1–/– females had increased heart weights, and both –/– sexes had decreased brain weights at 20 and 40 wk of age. However, these weight changes were not reflected by gross or microscopic tissue modifications.
The overall skeletal morphology of sFRP-1–/– mice was normal as determined by radiographic analysis (data not shown). Using peripheral quantitative computed tomography (pQCT) to measure vBMD [volumetric bone mineral density (BMD)], we did not observe consistent changes up to 52 wk of age in cortical bone parameters such as femoral diaphyseal BMD, thickness, or periosteal and endosteal circumference, and femur length was not affected (data not shown). Consistent with these results, total body BMD and bone mineral content was not altered in sFRP-1–/– mice as determined by dual-energy x-ray absorptiometry (DEXA). However, percent body fat was reduced by 22% in sFRP-1–/– males at 39–40 wk of age when compared with +/+ controls (P < 0.05, data not shown).
Serum markers of bone turnover such as osteocalcin and type I collagen C terminal telopeptides (CTX) were within the normal range in sFRP-1–/– mice, as were serum calcium and bone-alkaline phosphatase (ALP) levels (Fig. 2). Similarly, 18 additional serum measurements were unaffected by loss of sFRP-1, and urinary deoxypyridinoline cross-link levels as well as 14 other urine parameters were unchanged in –/– animals (data not shown, listed in Materials and Methods). Serum phosphorous was elevated by 29% in –/– males at 18–20 wk of age when compared with +/+ controls (Fig. 2E) but was not altered in females or in either sex at 39–40 wk of age. An increase in serum phosphorous in the 20-wk-old sFRP-1–/– males supports observations made by Berndt et al. (34) that sFRP-4 is a circulating phosphaturic factor that antagonizes renal Wnt-signaling and inhibits inorganic phosphate reabsorption.
|
Deletion of sFRP-1 Increases Trabecular Bone Formation in Adults
Histology of proximal femurs from 35-wk-old mice indicated that deletion of sFRP-1 increased trabecular bone area by 143% in –/– females and by 62% in –/– males when compared with the +/+ controls (Fig. 3, A and B). At this age, femurs of sFRP-1+/+ mice contained relatively little trabecular bone due to age-dependent loss of the tissue. Histomorphometry of proximal femurs from 20- to 35-wk-old animals indicated that trabecular bone area was similar between +/+ and –/– mice at 20 wk of age (Fig. 3, C and D). However, between 20 and 35 wk in females and 27 and 35 wk in males, +/+ mice lost bone, whereas –/– animals retained trabecular bone area, suggesting that deletion of sFRP-1 prevents age-dependent bone loss.
|
, A and B), although it was not significantly altered in males (data not shown). Because MAR is a measurement of the amount of bone matrix produced by individual osteoblast teams (38), these findings indicate that ablation of sFRP-1 enhances osteoblast activity at least in females. The reasons that MAR did not increase in male sFRP-1–/– mice may be related to the smaller gains in trabecular vBMD and bone volume seen in this sex, or to an interaction between the phenotype and sex steroids.
|
, A and B). Detailed analysis of these data demonstrated that removal of sFRP-1 not only increased trabecular bone volume/tissue volume, but also elevated connectivity density (29–47%), trabecular number (18–25%), and trabecular thickness (4–19%) when compared with sFRP-1+/+ controls (Fig. 5, C and D). Moreover, as expected from these results, trabecular separation was decreased by 18–22% in sFRP-1–/– animals. These data demonstrate that loss of sFRP-1 leads to an enhancement of trabecular bone quality.
|
, A and B). Beyond this time point, the –/– males lost some vBMD, whereas the –/– females retained femoral trabecular bone. Compared with +/+ controls, vBMD was 75–127% higher in –/– females and 39–51% higher in –/– males between the ages of 28–52 wk. Thus, in the distal femur, increased trabecular vBMD was more pronounced in –/– females than in males, and ablation of sFRP-1 prolonged bone gain and delayed its loss. In contrast, analysis of tibias indicated that male sFRP-1–/– mice exhibited a 24% increase in trabecular vBMD at 35 wk of age when compared with +/+ controls, whereas no difference was observed at earlier or later ages (Fig. 6D). Although tibial vBMD was consistently higher in female sFRP-1–/– mice between 20 and 47 wk, this difference did not reach significance when compared with the +/+ controls (Fig. 6C). Thus, in contrast to the femur, deletion of sFRP-1 appeared to have a greater impact on tibial trabecular vBMD of males than females.
|
, A and B). Trabecular bone volume and trabecular number were also increased by approximately 60% in female sFRP-1–/– mice, but the differences between +/+ and –/– did not achieve significance, perhaps due to the small sample size (n = 6/group). A similar pattern was also seen in males, although the differences between –/– mice and +/+ mice were less pronounced (Fig. 7, C and D).
|
Deletion of sFRP-1 Decreases Apoptosis and Enhances Proliferation and Differentiation of Osteoblast Lineage Cells
To determine whether bone cell apoptosis was altered in sFRP-1–/– mice, we performed TUNEL (terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick end-labeling) staining of calvaria from 32- to 33-wk-old females. Deletion of sFRP-1 led to a 48–56% decrease in the number of apoptotic osteoblasts and osteocytes (Fig. 8, A and B). This decrease in apoptosis resulted in an 18% increase in calvarial thickness and a 5% increase in osteocyte number, although both were not significantly different from sFRP-1+/+ animals (data not shown). When the percentage of apoptotic osteocytes per square millimeter was calculated (Fig. 8C), sFRP-1–/– females exhibited a 65% decrease in this parameter when compared with +/+ controls. Thus, although sFRP-1 is not the only regulator of apoptosis in murine osteoblasts and osteocytes, it contributes to about half of this process.
|
). However, this effect was enhanced 3- to 4-fold with bone marrow from sFRP-1–/– mice. In addition, ß-galactosidase staining of –/– cultures intensified after 12 and 16 d of incubation, indicating that sFRP-1 promoter activity also increased with advancing osteoblast differentiation (Fig. 9B). Furthermore, this elevation correlated with an up-regulation of ALP staining in sFRP-1–/– cultures, as well as a marked enhancement of von Kossa staining when compared with +/+ cultures.
|
). In sFRP-1+/+ bone marrow cultures, the percentage of apoptotic cells increased with increasing osteoblast differentiation, which correlated with elevated sFRP-1 promoter activity. However, apoptosis was reduced by 51–68% in cells from –/– mice at d 15 and 21 of culture. To measure proliferation, DNA synthesis was quantified in nondifferentiated bone marrow cells (Fig. 9D). During the first 5 d in culture, stromal cell attachment occurred and there was no difference in [3H]thymidine incorporation between +/+ and –/– cells. But after 7 d in culture (proliferative phase), cells from sFRP-1–/– mice exhibited a 27% increase in [3H]thymidine incorporation when compared with cells from +/+ controls. Thus, a reduction in osteoblast lineage cell apoptosis appears to be the major contributing factor for the increased osteoblastogenesis in –/– bone marrow cultures, although proliferation is also modestly enhanced by deletion of sFRP-1.
Deletion of sFRP-1 Enhances Osteoclastogenesis in Vitro
Hausler et al. (40, 41) have presented preliminary results showing that sFRP-1 binds to and blocks the effects of receptor activator of nuclear factor-
B ligand (RANKL) and TNF-
on in vitro osteoclastogenesis. To determine whether osteoclastogenesis was altered by deletion of sFRP-1, we performed an in vitro osteoclast differentiation assay with bone marrow from the mice. As expected, incubation of bone marrow from 25- to 26-wk-old sFRP-1+/+ females for 21 d with soluble human RANKL and murine monocyte-colony-stimulating factor (M-CSF) led to an increase in the number of tartrate-resistant acid phosphatase (TRAP)-positive, multinuclear cells (Fig. 10). However, this effect was enhanced 3- to 50-fold with bone marrow from sFRP-1–/– mice. Although deletion of sFRP-1 resulted in stimulation of osteoclastogenesis in vitro, as noted before, we did not observe any changes in bone resorption markers in sFRP-1–/– mice. Moreover, the phenotype of these animals is consistent with an increase in osteoblastic function and not an increase in osteoclastic activity that results in osteoporosis (42). Consequently, it appears that other regulators of osteoclast differentiation (e.g. osteoprotegerin) are able to compensate for loss of sFRP-1 in vivo (9, 19).
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
A second important finding from these studies is that, despite its broad tissue distribution, deletion of sFRP-1 in mice does not affect fertility, body weight, blood and urine chemistries, as well as most hematologic and nonskeletal organs. These observations indicate that other sFRPs compensate for loss of the gene in nonskeletal tissues, whereas sFRP-1 plays a dominant role in bone. More importantly, our findings show that loss of sFRP-1 has selective and beneficial effects on active osteoblasts in trabecular bone, which is the site of homeostatic remodeling. Furthermore, the advantageous consequences of sFRP-1 deletion occur in adult animals when bone is normally lost. The lack of differences in serum biochemical markers of bone metabolism and turnover further points to a predominant action for sFRP-1 in modulating bone cell proliferation and apoptosis. The increases in trabecular bone that result from elimination of sFRP-1 correlate with an elevation of the bone formation rate, a suppression of osteoblast and osteocyte apoptosis, and an enhancement of osteoblast lineage cell proliferation, differentiation, and mineralization. Hence, sFRP-1 not only regulates osteoblast and osteocyte longevity, but it also modulates the production of these cells and their function. Although we cannot exclude the possibility that loss of sFRP-1 expression in a nonskeletal tissue contributes to the bone phenotype, the results are nevertheless consistent with a direct effect on cells of the osteoblast lineage. Thus, we propose that activation of the Wnt pathway in osteoblasts by elimination of sFRP-1 has important implications for maintaining bone architecture in the aging skeleton.
Comparisons of the sFRP-1–/–, LRP5–/–, and HBM transgenic mice highlight functional differences for Wnt signaling pathways in the bone formation process. As anticipated, the skeletal phenotype of sFRP-1–/– mice is distinct from that of LRP5–/– animals (11). Deletion of LRP5, a Wnt coreceptor, reduces trabecular bone volume in young animals as early as 2 wk of age, whereas loss of sFRP-1 prolongs and enhances bone accrual and inhibits senile bone loss in adults. Mechanistically, ablation of LRP5 decreases trabecular bone MAR, whereas loss of sFRP-1 increases this parameter. Moreover, deletion of LRP5 reduces the number of ALP+ stromal cell progenitors and suppresses osteoblast activity, whereas loss of sFRP-1 increases osteoblast lineage cell number and differentiation and enhances osteoblast function. Additionally, osteoblasts in LRP5–/– mice have a reduced proliferative capacity, whereas the apoptotic index of these cells is unaltered. Conversely, deletion of sFRP-1 modestly increases osteoblast lineage cell proliferation but considerably reduces apoptosis of these cells, as well as of osteoblasts and osteocytes. These opposing phenotypes support an emerging concept that regulation of Wnt signaling is an important component, not only of skeletal development, but also of bone renewal throughout adult life.
Our findings from sFRP-1 loss-of-function, like the LRP5 gain-of-function phenotype (14), indicate that activation of the Wnt pathway in the aging skeleton has beneficial effects. However, the differences between these two models further emphasize the many options for controlling Wnt signaling in bone. The HBM transgenes exhibit increased trabecular bone accretion as early as 5 wk of age (14), whereas loss of sFRP-1 has no effect on trabebular bone before 13 wk of age. Like the sFRP-1–/– mice, the HBM animals do not lose bone as they age, and osteoblast and osteocyte apoptosis is reduced. However, deletion of sFRP-1 only affects trabecular bone, whereas expression of the HBM mutation of LRP5 affects both trabecular and cortical compartments. Absence of a cortical bone phenotype in sFRP-1–/– mice may reflect a predominate role for the gene on remodeling, which is more pronounced in trabecular than in cortical bone.
Differences between the LRP5 mutants and the sFRP-1–/– mice also underscore the roles for various Wnt signaling pathways in bone formation because LRP5 only modulates the canonical pathway (22), whereas sFRP-1 potentially regulates multiple pathways (23). In addition, the differential expression profile of LRP5 and sFRP-1 in bone cells may be another contributing factor to these variances. LRP5 expression occurs in proliferative stage, preosteoblasts, and peaks when ALP and collagen mRNA levels are high but osteocalcin expression is low (10). In contrast, expression of sFRP-1 increases with advancing osteoblast differentiation and peaks in the post-proliferative, preosteocytic stage when osteocalcin secretion is high but ALP levels are low; coincidentally, this expression profile correlates with an increase in the apoptotic index (17, 18, 24). Moreover, sFRP-1 is a secreted protein and can therefore act on neighboring and even distant cells in a paracrine manner, whereas LRP5 activity is restricted to the cells that express it. Finally, Wnt and FZD specificity may also play a role in these differences. Although we cannot exclude the possibility that sFRP-1 has actions beyond the regulation of Wnt pathways, the phenotype of the –/– mice is not consistent with increased RANKL and TNF- signaling.
Collectively, the results of mouse models and human genetic studies have established an essential role for Wnt signaling pathways in osteoblast biology and bone formation. Although deletion of LRP5 leads to additional nonskeletal aberrations such as altered eye development (10, 11) and cholesterol metabolism (43), the HBM mutation of LRP5 (12, 13, 14) and deletion of sFRP-1 appear to predominantly affect the skeleton. Although redundancy of the LRP and sFRP gene families may explain some of this apparent tissue selectivity, it is also likely that the complexity and intricacies of Wnt signaling contributes to these phenotypes. Thus, additional research into these pathways is needed to uncover the mechanisms of Wnt action in not only mammalian development but also in the adult.
|
MATERIALS AND METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
The sFRP-1 mice were produced by Lexicon Genetics, Inc. (The Woodlands, TX). The targeting vector was derived from the Lambda KOS system (44). Genomic clones were isolated from a Lambda KOS library by PCR using the exon 1-specific forward primer (5'-CTGAGGCTGTGCCACAACG-3') and the reverse primer (5'-CATGACCGGCTCGCACGAG-3'). A yeast selection cassette containing the URA3 marker was generated by PCR using a sense primer containing the gene-specific sequence (5'-GCAGCGGGACGCGCGCGTGAAGGCAGCGTG-3') and an antisense primer containing the gene-specific sequence (5'-GGGGTTCGCGGGCGTGGGAAGGCATACCCT-3'). This marker was introduced into the genomic clone by yeast recombination and resulted in the deletion of 1176 bp of exon 1 that was subsequently replaced with a LacZ/MC1-Neo selection cassette. The NotI linearized vector was electroporated into 129 Sv/Evbrd (LEX1) embryonic stem (ES) cells. G418/FIAU-resistant ES cell clones were isolated and analyzed for homologous recombination using Southern analysis. The 5' probe was a 510-bp PCR fragment derived from the sense primer (5'-ATGTGTATCTTGAGTTGGTATC-3') and the antisense primer (5'-CATAATACTTGCAAATTGATGC-3'). Use of this probe on EcoRI-digested genomic DNA produced a 12-kb wild-type and 10-kb mutant band. The 3' probe was a 542-bp PCR fragment derived from the sense primer (5'-CAACATAGCACTACATCTTCG-3') and the antisense primer (5'-GGCCAACGCTGAAGCCAG-3'). Use of this probe on BamHI-digested genomic DNA produced a 13-kb wild-type and 9.5-kb mutant band. Four targeted ES cell clones were identified and injected into C57BL/6(albino) blasotocysts. The resulting chimeras were mated to C57BL/6(albino) females to generate sFRP-1+/– animals. Subsequent intercrosses of sFRP-1+/– mice were performed to produce +/+, +/–, and –/– animals. Additional matings of these mice were equally successful and produced litters of similar size.
Animals were grouped by sex, age, and genotype with ad libitum access to filtered chlorinated water from the municipal water supply and a standard rodent diet containing 0.9% (wt/vol) calcium and 0.7% (wt/vol) phosphorous (Purina No. 5001 Rodent Diet, PMI, Richmond, IN). Environmental conditions were set to maintain 22 ± 3 C, 40 ± 15% humidity, and 12-h light, 12-h dark cycle. Animal health observations and environmental parameters were monitored daily.
Weanlings were ear or tail biopsied at 3 wk of age. DNA was extracted, and mice were initially genotyped using conventional PCR with the following primer sets: sFRP-1 (forward 5'-GGCAGCCCCGACGTCGCCGAGCAAC-3', reverse 5'-CCTTGGGGTTAGAGGCTTCCGTGG-3'; a 379-bp fragment); ß-galactosidase (forward 5'-ACGGCATGGTGCCAA-TGAATCGTCTG-3', reverse 5'-CAAATAATATCGGTGGCCGTGGTGTC-3'; a 212-bp fragment). Genotyping was latter performed using real-time PCR with an ABT PRISM 7700-sequence detection system (PE Applied Biosystems, Foster City, CA) as described by the manufacturer. The following Taqman primers and probe sets were used: sFRP-1 (forward 5'-GCCACAACG TGGGCTACAA-3', reverse 5'-ACCTCTGCCATGGTCTCGTG-3', probe 6-FAM-5'-AGATGGTGCTGCCCAACCTGCTG-TAMRA-3'); ß-galactosidase (forward 5'-CTGCTGATGAAGCAGAACAACTTT-3', reverse 5'-GCGT-GTACCACAGCGGATG-3', probe VIC-5'-CGCCGTGCGCTGTTCGCATTA-TAMRA-3').
RNA Analysis
Total RNA was isolated from tissues using Trizol reagent according to the manufacturer’s instructions (Invitrogen Life Technologies, Gaithersburg, MD). The Poly A(+) RNA fraction was obtained using an Oligotex mRNA Maxi kit according to the manufacturer’s instructions (QIAGEN, Valencia, CA), and northern hybridization was performed as previously described (45). A 379-bp sFRP-1 cDNA fragment from the CRD was generated by PCR using the following primers (forward 5'-CGGCCAGCGAGTACGACTACGTGAGC-3', reverse 5'-GCATCTCGG GCCAGTAGAAGCCGAAG-3'), whereas a full-length GAPDH cDNA was obtained by restriction enzyme digestion.
Quantification of sFRP-1 mRNA was performed with total RNA using real-time RT-PCR on an ABT PRISM 7700-sequence detection system as described by the manufacturer. The following Taqman primers and probe sets were used: sFRP-1 CRD (forward 5'-GCCACAACG TGGGCTACAA-3', reverse 5'-ACCTCTGCCATGGTCTCGTG-3', probe 6-FAM-5'-AGATGGTGCTGCCCAACCTGCTG-TAMRA-3'); sFRP-1 netrin domain (forward 5'-CGCTTGTGCTGTTCCTGAAG-3', reverse 5'-CGGCCCATGATGAGAAAGTT-3', probe 6-FAM-5'-TGCCGACTGTCCCTGCCACCA-TAMRA-3'). The sFRP-1 results were normalized to 18S ribosomal RNA levels using VIC probe reagents from PE Applied Biosystems (part no. 4308329), and mRNA levels were calculated using the Standard Curve Method as described by Applied Biosystems in User Bulletin No. 2.
Clinical Pathology
A complete gross necropsy examination was performed on the mice after isoflurane anesthesia and blood collection. The animals were weighed, exsanguinated, and tissues and organs were examined and collected in 10% neutral buffered formalin. All gross observations and organ weights were captured using the Xybion Path/Tox System (Xybion Medical Systems, Cedar Knolls, NJ). Organs weighed before fixation were the brain, liver, adrenal glands (paired), heart, lungs, spleen, testes (paired), ovaries (paired), thymus, and kidneys (paired). In addition, representative portions of the entire epididymides, peripheral nerve (sciatic), spinal cord, urinary bladder, pancreas, coagulating gland, mandibular and mesenteric lymph nodes, skeletal muscle (thigh), vertebrae, small and large intestine (duodenum, jejunum, ileum, cecum, colon, rectum), stomach, uterus, cervix, vagina, oviduct, esophagus, parathyroid gland, seminal vesicles, skin, pituitary gland, Harderian gland, femur, mammary gland, gall bladder, salivary glands, trachea, prostate gland, thyroid gland, eyes, bone marrow, and aorta were collected and examined.
The following hematology parameters were measured: white blood cells, differential white blood cells, red blood cells, hemoglobin, hematocrit, mean cell volume, mean cell hemoglobin, and platelets.
Bone Histomorphometry, Histology, and Densitometry
In vivo whole body areal BMD (g/cm2) of anesthetized mice was evaluated using peripheral DEXA (PIXImus, GE-Lunar Corp., Madison, WI) as previously described (14).
After peripheral DEXA, mice were placed in a prone position and radiographed using a Faxitron radiographic unit (Faxitron X-Ray Corp., Buffalo Grove, IL) set at 25 kVp and a 10-sec exposure with Kodak X-Omat TL film (Eastman Kodak Co., Rochester, NY).
For ex vivo analyses, mice were euthanized by exposure to carbon dioxide, and bones were removed, dissected free of soft tissue, and stored in 70% (vol/vol) ethanol at 4 C until evaluation. The vertebral column was sectioned at the level of the intervertebral disc L1–L2 and the lumbar vertebrae L2–L6 with the tail.
Volumetric BMD (mg/cm3) and cortical bone measurements of femurs and tibias were evaluated using an XCT Research pQCT densitometer (Stratec Medizintechnik, Pforzheim, Germany) as previously described (14, 46).
Static and dynamic histomorphometry measurements of the femur were made using the R&M Biometrics Inc. Bioquant Image Analysis System (Bioquant Image Analysis Corp., Nashville, TN) as previously described (14).
µCT20 or 40 (Scanco Medical AG, Basserdorf, Switzerland) was used to evaluate trabecular bone volume fraction and microarchitecture of the distal femur as previously described (14, 46).
Vertebral histomorphomertry was performed as follows. L4 to L6 vertebrae were trimmed free of skeletal muscle, the spinal cord was removed, and the vertebrae were cleaned. Using a diamond saw, a parasagittal cut parallel to the median line on the left side of the vertebrae was made through the transverse process and the vertebral body. Decalcified vertebrae were embedded in methylmethacrylate medium, and 5–7 µm sections of vertebral cancellous bone were prepared using a Leica Polycut microtome (Leica Microsystems, Nussloch, Germany). Sections of decalcified bone were stained with Goldner’s, von Kossa or kept unstained.
The TUNEL method (Roche Diagnostics, Indianapolis, IN) was used to detect cell apoptosis in paraffin-embedded parietal bone as previously described (14, 47).
Serum and Urine Measurements of Bone Turnover
Serum osteocalcin and CTX were determined by ELISA using Rat-Mid Osteocalcin and RatLaps kits manufactured by Osteometer BioTech A/S (Herlev, Denmark). Additional serum and urine analyses were performed by Anilytics Inc. (Gaithersburg, MD).
The following serum parameters were measured: alanine aminotransferase, aspartate aminotransferase, alkaline phosphatase (total), creatine kinase, alkaline phosphatase (bone isoform), glucose, alkaline phosphatase (liver isoform), albumin, 
glutamyltransferase, albumin/globulin ratio, blood urea nitrogen, bilirubin (direct), total protein potassium, globulin bicarbonate, bilirubin (total), phosphorus, sodium osteocalcin, chloride sorbitol dehydrogenase, calcium, and CTX.
The following urine parameters were measured: bilirubin, blood, color and appearance, glucose, ketones, pH, proteins, specific gravity, urobilinogen, microscopy of centrifuged deposit, urine output, urine creatinine, urine urea nitrogen, and deoxypyridinoline.
In Vitro Bone Cell Differentiation, Apoptosis, and Proliferation
Isolation and culture of bone marrow derived mesenchymal and hematopoietic stem cells was performed as previously described (48, 49). Cells were cultured in DMEM (high glucose) containing 1 mM GlutaMAX-1, 1% penicillin-streptomycin (all from Invitrogen Life Technologies) and 15% heat-inactivated charcoal-stripped fetal bovine serum (HyClone Laboratories, Inc., Logan, UT) at 37 C in a 5% CO2/95% humidified air incubator (Forma Scientific, Marietta, OH). Supplements for osteoblast differentiation included 10 mM ß-glycerol phosphate (Sigma-Aldrich, St. Louis, MO), 10–7 M dexamethasone, 50 µM ascorbate-2-phosphate (Wako, Richmond, VA) or 50 µg/ml ascorbic acid (Sigma-Aldrich) and were added to the medium with the first and all subsequent medium changes. ALP activity and mineralized nodules were quantified as previously described (48, 49, 50). To measure apoptosis, the cells were trypsinized, washed with PBS (pH 7.2), fixed with 70% ethanol at –20 C for 2 h, and analyzed by FACStar using the Consort 30 software (Becton Dickinson, Mountain View, CA). Cellular DNA synthesis was measured by [3H]thymidine incorporation as previously described (51). Staining for ß-galactosidase was performed as follows. Cell layers were fixed at 4 C for 10 min in PBS (pH 8.8), that contained 2% paraformaldehyde, 0.2% glutaraldehyde, 5 mM EGTA (pH 7.3), and 2 mM MgCl2 as previously described (52). ß-Galactosidase staining of the cell layers was determined after 4 h of incubation at 37 C using PBS (pH 7.2), that contained 5 mM potassium ferricyanide, 2 mM MgCl2, 0.2% Nonidet P-40 (USB, Cleveland, OH), 0.01% sodium deoxycholate and 1 mg/ml X-Gal (Gold Biosciences, St. Louis, MO). Supplements for osteoclast differentiation included 30 nM srhRANKL (Research Diagnostics Inc., Flanders, NJ) and 10 nM recombinant murine M-CSF (R&D Systems, Minneapolis, MN) and were added to the medium with the first and all subsequent medium changes. Preosteoclasts (mononuclear cells) and mature osteoclasts (multinuclear cells, >2 nuclei/cell) were stained with TRAP and quantified with a commercially available kit (Sigma-Aldrich) as previously described (53).
Statistical Analysis
Data were analyzed for statistical significance by one-way ANOVA using the Dunnett’s test with the JMP software (SAS Institute, Cary, NC).
|
ACKNOWLEDGMENTS |
|---|
|
FOOTNOTES |
|---|
B ligand; sFRP, secreted FZD-related protein; TRAP, tartrate-resistant acid phosphatase; TUNEL, terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate nick end-labeling; vBMD, volumetric BMD. Received for publication December 31, 2003. Accepted for publication February 11, 2004.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
or RANKL-dependent osteoclast formation. J Bone Miner Res 17:S349
This article has been cited by other articles:
 |
|
 |
|
  L. Wang, D. Gesty-Palmer, T. A. Fields, and R. F. Spurney Inhibition of WNT Signaling by G Protein-Coupled Receptor (GPCR) Kinase 2 (GRK2) Mol. Endocrinol., September 1, 2009; 23(9): 1455 - 1465. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. S. Friedman, S. M. Oyserman, and K. D. Hankenson Wnt11 Promotes Osteoblast Maturation and Mineralization through R-spondin 2 J. Biol. Chem., May 22, 2009; 284(21): 14117 - 14125. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
L. Xiao, P. Liu, X. Li, T. Doetschman, J. D. Coffin, H. Drissi, and M. M. Hurley Exported 18-kDa Isoform of Fibroblast Growth Factor-2 Is a Critical Determinant of Bone Mass in Mice J. Biol. Chem., January 30, 2009; 284(5): 3170 - 3182. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. Pederson, M. Ruan, J. J. Westendorf, S. Khosla, and M. J. Oursler Regulation of bone formation by osteoclasts involves Wnt/BMP signaling and the chemokine sphingosine-1-phosphate PNAS, December 30, 2008; 105(52): 20764 - 20769. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Yokota, K. Oritani, K. P. Garrett, T. Kouro, M. Nishida, I. Takahashi, M. Ichii, Y. Satoh, P. W. Kincade, and Y. Kanakura Soluble Frizzled-Related Protein 1 Is Estrogen Inducible in Bone Marrow Stromal Cells and Suppresses the Earliest Events in Lymphopoiesis J. Immunol., November 1, 2008; 181(9): 6061 - 6072. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Matsubara, K. Kida, A. Yamaguchi, K. Hata, F. Ichida, H. Meguro, H. Aburatani, R. Nishimura, and T. Yoneda BMP2 Regulates Osterix through Msx2 and Runx2 during Osteoblast Differentiation J. Biol. Chem., October 24, 2008; 283(43): 29119 - 29125. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. Bovolenta, P. Esteve, J. M. Ruiz, E. Cisneros, and J. Lopez-Rios Beyond Wnt inhibition: new functions of secreted Frizzled-related proteins in development and disease J. Cell Sci., March 15, 2008; 121(6): 737 - 746. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. G. Robling, P. J. Niziolek, L. A. Baldridge, K. W. Condon, M. R. Allen, I. Alam, S. M. Mantila, J. Gluhak-Heinrich, T. M. Bellido, S. E. Harris, et al. Mechanical Stimulation of Bone in Vivo Reduces Osteocyte Expression of Sost/Sclerostin J. Biol. Chem., February 29, 2008; 283(9): 5866 - 5875. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
H. Zhou, W. Mak, Y. Zheng, C. R. Dunstan, and M. J. Seibel Osteoblasts Directly Control Lineage Commitment of Mesenchymal Progenitor Cells through Wnt Signaling J. Biol. Chem., January 25, 2008; 283(4): 1936 - 1945. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. C. Manolagas and M. Almeida Gone with the Wnts: {beta}-Catenin, T-Cell Factor, Forkhead Box O, and Oxidative Stress in Age-Dependent Diseases of Bone, Lipid, and Glucose Metabolism Mol. Endocrinol., November 1, 2007; 21(11): 2605 - 2614. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C.H. Li and S. Amar Inhibition of SFRP1 Reduces Severity of Periodontitis Journal of Dental Research, September 1, 2007; 86(9): 873 - 877. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. C. Kudo, S. L. Karsten, J. Chen, P. Levitt, and D. H. Geschwind Genetic Analysis of Anterior Posterior Expression Gradients in the Developing Mammalian Forebrain Cereb Cortex, September 1, 2007; 17(9): 2108 - 2122. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J Rodriguez-Lopez, M Pombo-Suarez, M Liz, J J Gomez-Reino, and A Gonzalez Further evidence of the role of frizzled-related protein gene polymorphisms in osteoarthritis Ann Rheum Dis, August 1, 2007; 66(8): 1052 - 1055. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
X. Zhong, T. Desilva, L. Lin, P. Bodine, R. A. Bhat, E. Presman, J. Pocas, M. Stahl, and R. Kriz Regulation of Secreted Frizzled-related Protein-1 by Heparin J. Biol. Chem., July 13, 2007; 282(28): 20523 - 20533. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Baron and G. Rawadi Targeting the Wnt/{beta}-Catenin Pathway to Regulate Bone Formation in the Adult Skeleton Endocrinology, June 1, 2007; 148(6): 2635 - 2643. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Kang, C. N. Bennett, I. Gerin, L. A. Rapp, K. D. Hankenson, and O. A. MacDougald Wnt Signaling Stimulates Osteoblastogenesis of Mesenchymal Precursors by Suppressing CCAAT/Enhancer-binding Protein {alpha} and Peroxisome Proliferator-activated Receptor {gamma} J. Biol. Chem., May 11, 2007; 282(19): 14515 - 14524. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. V. Bodine Wnt Signaling in Bone IBMS BoneKEy, March 1, 2007; 4(3): 108 - 123. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. A. Clines, K. S. Mohammad, Y. Bao, O. W. Stephens, L. J. Suva, J. D. Shaughnessy Jr., J. W. Fox, J. M. Chirgwin, and T. A. Guise Dickkopf Homolog 1 Mediates Endothelin-1-Stimulated New Bone Formation Mol. Endocrinol., February 1, 2007; 21(2): 486 - 498. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Liu, R. A. Bhat, L. M. Seestaller-Wehr, S. Fukayama, A. Mangine, R. A. Moran, B. S. Komm, P. V. N. Bodine, and J. Billiard The Orphan Receptor Tyrosine Kinase Ror2 Promotes Osteoblast Differentiation and Enhances ex Vivo Bone Formation Mol. Endocrinol., February 1, 2007; 21(2): 376 - 387. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 W. S. Wright, K. A. Longo, V. W. Dolinsky, I. Gerin, S. Kang, C. N. Bennett, S.-H. Chiang, T. C. Prestwich, C. Gress, C. F. Burant, et al. Wnt10b Inhibits Obesity in ob/ob and Agouti Mice Diabetes, February 1, 2007; 56(2): 295 - 303. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 L. Sun, Y. Peng, N. Zaidi, L.-L. Zhu, J. Iqbal, K. Yamoah, X. Wang, P. Liu, E. Abe, B. S. Moonga, et al. Evidence that calcineurin is required for the genesis of bone-resorbing osteoclasts Am J Physiol Renal Physiol, January 1, 2007; 292(1): F285 - F291. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. N. Pearse Wnt antagonism in multiple myeloma: a potential cause of uncoupled bone remodeling. Clin. Cancer Res., October 15, 2006; 12(20): 6274s - 6278s. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C.H. Li and S. Amar Role of Secreted Frizzled-related Protein 1 (SFRP1) in Wound Healing Journal of Dental Research, April 1, 2006; 85(4): 374 - 378. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. J. Spencer, J. C. Utting, S. L. Etheridge, T. R. Arnett, and P. G. Genever Wnt signalling in osteoblasts regulates expression of the receptor activator of NF{kappa}B ligand and inhibits osteoclastogenesis in vitro J. Cell Sci., April 1, 2006; 119(7): 1283 - 1296. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 W. Satoh, T. Gotoh, Y. Tsunematsu, S. Aizawa, and A. Shimono Sfrp1 and Sfrp2 regulate anteroposterior axis elongation and somite segmentation during mouse embryogenesis Development, March 15, 2006; 133(6): 989 - 999. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. R Mundy Nutritional modulators of bone remodeling during aging Am. J. Clinical Nutrition, February 1, 2006; 83(2): 427S - 430S. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. J. Lories, S. Boonen, J. Peeters, K. de Vlam, and F. P. Luyten Evidence for a differential association of the Arg200Trp single-nucleotide polymorphism in FRZB with hip osteoarthritis and osteoporosis Rheumatology, January 1, 2006; 45(1): 113 - 114. [Full Text] [PDF]
|
|
|
|
|
|
M. Almeida, L. Han, T. Bellido, S. C. Manolagas, and S. Kousteni Wnt Proteins Prevent Apoptosis of Both Uncommitted Osteoblast Progenitors and Differentiated Osteoblasts by {beta}-Catenin-dependent and -independent Signaling Cascades Involving Src/ERK and Phosphatidylinositol 3-Kinase/AKT J. Biol. Chem., December 16, 2005; 280(50): 41342 - 41351. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P. Clement-Lacroix, M. Ai, F. Morvan, S. Roman-Roman, B. Vayssiere, C. Belleville, K. Estrera, M. L. Warman, R. Baron, and G. Rawadi Lrp5-independent activation of Wnt signaling by lithium chloride increases bone formation and bone mass in mice PNAS, November 29, 2005; 102(48): 17406 - 17411. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Oshima, M. Abe, J. Asano, T. Hara, K. Kitazoe, E. Sekimoto, Y. Tanaka, H. Shibata, T. Hashimoto, S. Ozaki, et al. Myeloma cells suppress bone formation by secreting a soluble Wnt inhibitor, sFRP-2 Blood, November 1, 2005; 106(9): 3160 - 3165. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. J. Martin New Aspects of Wnt Signalling Revealed by Mouse and Human Genetics IBMS BoneKEy, October 1, 2005; 2(10): 7 - 11. [Full Text] [PDF]
|
|
|
|
|
|
T. Gaur, C. J. Lengner, H. Hovhannisyan, R. A. Bhat, P. V. N. Bodine, B. S. Komm, A. Javed, A. J. van Wijnen, J. L. Stein, G. S. Stein, et al. Canonical WNT Signaling Promotes Osteogenesis by Directly Stimulating Runx2 Gene Expression J. Biol. Chem., September 30, 2005; 280(39): 33132 - 33140. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. Hamerman Osteoporosis and atherosclerosis: biological linkages and the emergence of dual-purpose therapies QJM, July 1, 2005; 98(7): 467 - 484. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. Morgan, G. J. Atkins, M. K. Trivett, S. A. Johnson, M. Kansara, S. L. Schlicht, J. L. Slavin, P. Simmons, I. Dickinson, G. Powell, et al. Molecular Profiling of Giant Cell Tumor of Bone and the Osteoclastic Localization of Ligand for Receptor Activator of Nuclear Factor {kappa}B Am. J. Pathol., July 1, 2005; 167(1): 117 - 128. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Ai, S. L. Holmen, W. Van Hul, B. O. Williams, and M. L. Warman Reduced Affinity to and Inhibition by DKK1 Form a Common Mechanism by Which High Bone Mass-Associated Missense Mutations in LRP5 Affect Canonical Wnt Signaling Mol. Cell. Biol., June 15, 2005; 25(12): 4946 - 4955. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
X. Li, Y. Zhang, H. Kang, W. Liu, P. Liu, J. Zhang, S. E. Harris, and D. Wu Sclerostin Binds to LRP5/6 and Antagonizes Canonical Wnt Signaling J. Biol. Chem., May 20, 2005; 280(20): 19883 - 19887. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. N. Bennett, K. A. Longo, W. S. Wright, L. J. Suva, T. F. Lane, K. D. Hankenson, and O. A. MacDougald Regulation of osteoblastogenesis and bone mass by Wnt10b PNAS, March 1, 2005; 102(9): 3324 - 3329. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. Gazzerro, R. C. Pereira, V. Jorgetti, S. Olson, A. N. Economides, and E. Canalis Skeletal Overexpression of Gremlin Impairs Bone Formation and Causes Osteopenia Endocrinology, February 1, 2005; 146(2): 655 - 665. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Billiard, D. S. Way, L. M. Seestaller-Wehr, R. A. Moran, A. Mangine, and P. V. N. Bodine The Orphan Receptor Tyrosine Kinase Ror2 Modulates Canonical Wnt Signaling in Osteoblastic Cells Mol. Endocrinol., January 1, 2005; 19(1): 90 - 101. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
