1 Endocrine Unit, Massachusetts General Hospital, Boston, MA 02114, USA
2 Department of Human and Molecular Genetics, Baylor College of Medicine, Houston, TX 77030, USA
Present address: Department of Biochemistry, School of Dentistry, Showa University, Japan
Present address: Department of Pathology Angell Memorial Animal Hospital, Boston, MA, USA
Present address: Department of Oral Biology, Forsyth Institute, Boston, MA, USA
*Author for correspondence (e-mail: kronenberg.henry{at}mgh.harvard.edu)
Accepted 20 March 2002
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SUMMARY |
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Key words: PTHrP, PTH/PTHrP receptor, Indian hedgehog, Growth plate, Chondrocyte differentiation, Cre-loxP, Mouse
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INTRODUCTION |
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Previous studies using chimeric growth plates comprising wild-type and PPR-null cells have demonstrated that loss of PPR signaling in proliferating chondrocytes caused premature hypertrophic differentiation (Chung et al., 1998). However, Ppr/ cells in the periarticular region of the chimeric growth plate are not morphologically distinct from wild-type cells. The possible influence of PPR signaling on the differentiation of periarticular chondrocytes to columnar chondrocytes is, therefore, unclear.
To understand better how PTHrP and Ihh signaling regulate chondrocyte differentiation, we have developed mice with PPR ablation in chondrocytes using the Cre-loxP system (Pluck, 1996). During generation of floxed mice, we also established a mouse line with reduced PPR expression. Through the analysis of these novel mouse models with abnormal PPR signaling, we show that loss or impairment of PPR signaling is associated with chondrocyte differentiation not only at the terminal step but also at an earlier step. In another model with mosaic ablation of the PPR in the growth plate, we show that the differentiation of early chondrocytes is correlated with Ihh action but is not directly regulated by PTHrP. Based on these findings, we propose a model in which PTHrP and Ihh control chondrocyte differentiation at multiple steps.
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MATERIALS AND METHODS |
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Generation of Cre transgenic mice
Col2-Cre transgenic mice where Cre recombinase (OGorman et al., 1997) is expressed under the rat collagen type II promoter (Yamada, 1990
) were generated as described elsewhere (Schipani et al., 2001
). Ost-Cre transgenic mice bear a fusion gene composed of a 1.3 kb fragment of the mouse OG2 promoter (Ferendo et al., 1998
) fused to Cre recombinase and polyA signal excised from a pCBM-9 vector (Saur and Henderson, 1989
). After removing vector sequence, DNA was subjected to pronuclear injection. Cre activity was assessed using Rosa26-R (R26R) reporter mice (Soriano, 1999
): Cre transgenic mice were crossed with R26R mice. Embryos were collected, fixed and stained with X-gal, as described elsewhere (Chung et al., 1998
).
Genotyping of mice
The floxed PPR allele and the unanticipated d allele were analyzed by PCR using primers P1 (5'-TGGACGCAGACGATGTCTTTACCA-3') and P2 (5'-ACATGGCCATGCCTGGGTCTGAGA-3'), which recognize the sequences spanning the 3' loxP site. Wild-type and mutant alleles give 450 bp and 490 bp PCR products, respectively. The PPR null locus was detected using PCR primers, P3 (5'-CCACCAATGTGAGTTCCTACAGAAA-3') for an intronic sequence between exons E2 and E3, and P4 (5'-TCCAGACTGCCTTGGGAAAAGCGC-3') for the PGK promoter used for the neomycin resistant marker. The mutant allele with a retained PGK promoter sequence gives a 500 bp band. Cre sequences were detected by PCR using primers recognizing internal sequence of the transgene (P5, 5'-CGCGGTCTGGCAGTAAAAACTATC-3'; P6, 5'-CCCACCGTCAGTACGTGAGATATC-3'). All PCR reactions used the following program: 94°C for 10 minutes, followed by 35 cycles of 95°C for 30 seconds, 68°C for 30 seconds, and 72°C for 1 minute.
Southern and northern blotting
DNA was prepared from the livers of mice homozygous for the wild-type PPR, the floxed PPR and the mutated PPR described in the text. After overnight enzyme digestion, DNA was separated in 0.7% agarose gels, denatured and transferred onto nylon or nitrocellulose membranes.
Total RNA was extracted from the kidneys of 3-week-old mice. RNA was separated in a 1% agarose gel and transferred onto a nitrocellulose membrane. The probe for the mouse PPR was generated by a random priming method (Megaprime, Amersham), using the DNA template amplified from mouse kidneys by RT-PCR using primers P7 (5'-ACCAACTACTACTGGATTCTGGTGG-3') and P8 (5'-CGGCTCCAAGACTTCCTAATCTCTG-3'). The probes used for Southern analysis are indicated in Fig. 1A. RNA and DNA hybridization was performed using the QuickHyb Kit (Stratagene) according to the manufacturers instruction. Signals were visualized on X-ray films or by the Cyclone storage phosphor system (Packard). PPR and GAPDH mRNA were quantified using the Cyclone storage phosphor system. PPR mRNA levels normalized for GAPDH mRNA were compared. The band intensity of the Southern analysis was similarly quantified and normalized with the Gapdh gene. The normalized band intensities obtained from at least three independent blots for each set were used for the comparisons.
PPR cDNA sequencing
Total RNA was prepared from the kidney of homozygous d/d mutant mice. Reverse transcription was performed using Superscript reverse transcriptase (Gibco/BRL) followed by PCR using primers P9 (5'-CCGAGGGACGCGGCCCTAG-3') and P10 (5'-AGTCCTGAATAGACAGCCAGCCAAA-3') to amplify the entire coding sequence of the PPR. DNA sequence was determined by direct sequencing bi-directionally using the following sequencing primers: forward primers, P9, P11 (5'-GCTGCTCAAGGAAGTTCTGCACACA-3'), P12 (5'-GATCTACACCGTGGGATATTCCATG-3'), P13 (5'-ACCAACTACTACTGGATTCTGGTGG-3'), P14 (5'-CACTGTGGCAGATCCAGATGCACTA-3'); backward primers, P10, P14 (5'-CGGCTCCAAGACTTCCTAATCTCTG-3'), P15 (5'-AGTGTTGGCCAAGGTTGCTCTGACA-3'), P16 (5'-GCAGCATAAACGACAGGAACATGTG-3'), P17 (5'-CAGCAAACGATGTTGTCCCACTCTG-3').
In situ hybridization
Tissues were fixed in 4% paraformaldehyde/PBS overnight at 4°C, processed, embedded in paraffin wax and cut. Sections were stained with H&E or nuclear Fast Red (Vector laboratories). In situ hybridization was performed as described previously (Lee et al., 1995) by using complimentary 35S-labeled riboprobes. The probes for mouse type X collagen, mouse patched 1, mouse Ihh and rat PTHrP were obtained from Dr Bjorn Olsen (Harvard Medical School), Dr Ron Johnson (Stanford University, Stanford) Dr Benoit St-Jacques (Harvard University) and Dr Andrew C. Karaplis (McGill University), respectively. Rat full-length PPR cDNA probe, R15B has been described previously (Calvi et al., 2001
). An exon E1 specific PPR probe was generated by PCR using primers: F, 5'-GTGGACGCAGACGATGTCTTTACC-3'; and R, 5'-CTGCTGTGTGCAGAACTTCCTTGA-3'. PCR product was subcloned into pGEMT Easy vector (Promega).
BrdU labeling
Pregnant mice received intra-peritoneal injections of 50 µg BrdU/g of body weight and were sacrificed 1 or 24 hours later. Limbs were dissected and fixed in 4% paraformaldehyde overnight at 4°C. Tissues were processed, embedded and sectioned using standard procedures. BrdU was detected using a BrdU Staining Kit (Zymed Laboratories). The BrdU-positive and -negative nuclei were counted in the periarticular region and the columnar region separately. The border between the periarticular and columnar regions was defined as the line separating these two morphologically distinct groups of chondrocytes.
For counting of BrdU-positive cells in the periarticular region, using sections with smaller periarticular regions, we first determined an area for BrdU counting by drawing a closed line that made the area as large as possible, while avoiding the border containing ambiguous cells. Then, we applied the same area on the control sections as it was placed in the center of the corresponding region. We confirmed that the area only included cells with the typical morphological appearance of periarticular chondrocytes. For counting of BrdU-positive cells in the columnar region, as Pprd/ growth plates lack sharp transition of between columnar and hypertrophic regions, we first chose top one third of the columnar region of Pprd/ mice not to include any hypertrophic cells. We set a rectangular area excluding periarticular and perichondrial cells for BrdU counting. The same rectangle was applied onto control sections. Exclusion of other types of cells was similarly confirmed.
Mutants and controls used in this study were littermates. Nine sections from at least three independent mice per group were counted. Statistical analysis was done by ANOVA.
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RESULTS |
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Mice with chondrocyte-specific PPR ablation have a growth plate phenotype similar to that of Ppr-null mice
Homozygous floxed, Pprfl/fl mice and compound heterozygous Pprfl/ mice have no abnormality in growth plate cartilage (data not shown). Chondrocyte-specific Ppr gene ablation was carried out by mating these floxed mice with Cre transgenic mice expressing Cre under the control of the rat type II collagen promoter (Col2-Cre). Cre activity was determined using R26R reporter mice, and was present in growth plate chondrocytes but limited to the chondrocytes and a part of the perichondrium, intra-joint tissues, ligaments and tendons (Fig. 2A). PPR is predominantly expressed in the prehypertrophic region and weakly in columnar chondrocytes (Fig. 2J). PPR expression determined by an exon E1-specific probe was lost in the growth plate of double mutant mice, Col2-Cre:Pprfl/fl (Fig. 2K). Col2-Cre:Pprfl/fl mice develop chondrodysplasia that resembles that of Ppr/ (Fig. 2B,C): the tibial growth plate is shortened and lacks most of the columnar chondrocytes. Reduction on the proliferating chondrocytes with preservation of a fairly normal hypertrophic layer was confirmed by the expression patterns of type II and type X collagens (Fig. 2D-I). The periarticular region is flattened (Fig. 2C). At E16.5, the mutant sternum is mostly occupied by hypertrophic cells, whereas there are few hypertrophic cells in the control (Fig. 2L,M). However, unlike Ppr/ mice in some genetic backgrounds, mutant mice survive until birth and they are not as small as Ppr/ mice (data not shown). From these observations, we conclude that the Ppr/ growth plate phenotype is primarily caused by loss of PPR signaling in the chondrocytes themselves.
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To support the hypothesis further that reduced PPR signaling in chondrocytes is responsible for the cartilage phenotype of Pprd/ mice, we crossed Pprd/ mice with transgenic mice that express a constitutively active Ppr mutant gene in chondrocytes using rat type II collagen promoter (caPpr). This gene has previously been shown to rescue the growth plate abnormality of Pthrp/ mice (Schipani et al., 1997). The proximal tibias of E17.5 caPpr mice, Pprd/ mice and caPpr:Pprd/ mice were compared (Fig. 3D). The tibia of the caPpr mouse is characterized by an expansion of fully differentiated hypertrophic chondrocytes that express osteopontin and a decreased amount of type X collagen mRNA. Although both the caPpr and Pprd/ mice show expansion of the hypertrophic region, the hypertrophic region of Pprd/ mice expresses high levels of type X collagen mRNA and little osteopontin mRNA. This characteristic abnormality of the Pprd/ growth plate disappears when the caPpr gene is introduced, and the growth plate of the double mutant mice, caPpr:Pprd/ is indistinguishable from that of caPpr.
Impairment or loss of PPR signaling causes acceleration of early chondrocyte differentiation
The number of cells in each chondrocyte layer represents the balance between the number of cells entering and leaving the layer as well as proliferation within the layer. To analyze these steps, first, we performed BrdU labeling (Fig. 4A): when E17.5 mice were sacrificed 1 hour after BrdU administration, no BrdU-positive cells were found in the hypertrophic region either in Pprd/ or in wild-type controls. The fraction of cells labeled with BrdU in the columnar region was unchanged in Pprd/ mice. It was, however, significantly increased in the periarticular region of Pprd/ mice, even though the region was smaller. This was also confirmed in E16.5 and E18.5 mice (data not shown). This suggests that periarticular chondrocytes differentiate into columnar chondrocytes and therefore leave the periarticular region at a greater rate than that of controls. Then, we performed BrdU pulse-chase assay to see whether hypertrophic chondrocytes are also generated at a greater rate in Pprd/ mice. Proliferating chondrocytes of E17.5 embryos were pulse labeled with BrdU. Mice were sacrificed 24 hours after BrdU labeling. BrdU-positive hypertrophic chondrocytes are generated during the period. We found that those BrdU-positive hypertrophic chondrocyte did not reach chondro-osseous junction at this condition; therefore, we did not lose BrdU-positive hypertrophic chondrocytes by the replacement of cartilage by bone cells. As hypertrophic chondrocytes do not incorporate BrdU (Fig. 4A), these BrdU-positive hypertrophic chondrocytes are generated by differentiation of columnar chondrocytes labeled with BrdU 24 hours before sacrifice. The number of BrdU-positive hypertrophic chondrocytes is determined by the number and proliferation of BrdU-labeled proliferating chondrocytes and the rate of their hypertrophic differentiation. The number of BrdU-positive hypertrophic cells produced in Pprd/ mice during this time was greater than that of controls, and the area encompassed by BrdU-positive hypertrophic cells was greater (Fig. 4B). As the number and the proliferation rate of columnar chondrocytes of Pprd/ mice were not greater than the control (Fig. 4A), we concluded that the increased number of BrdU-positive hypertrophic chondrocytes was due to an increase in the rate of hypertrophic differentiation. The observations that osteopontin is only expressed in the terminal end of the hypertrophic region of Pprd/ mice with no expansion in number of osteopontin-positive cells suggest that this acceleration does not continue in the step of further hypertrophic maturation. Thus, acceleration of the differentiation rate of periarticular into columnar chondrocytes as well as of columnar to hypertrophic chondrocytes appear to have caused an accumulation of early hypertrophic chondrocytes in Pprd/ mice. The acceleration of periarticular to columnar differentiation was also observed in mice with PPR ablation in chondrocytes of E17.5 mice (Fig. 4C). Despite the decreased size of the periarticular region, chondrocyte proliferation was increased.
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DISCUSSION |
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The observation that Col2-Cre:Pprfl/fl mice showed a growth plate abnormality similar to that of the Ppr/ mice is consistent with the previous finding that loss of PPR signaling in columnar chondrocytes per se accelerates their terminal differentiation (Chung et al., 1998). However, chondrocyte-specific Ppr-null mice differ from Ppr/ mice in body size and embryonic lethality. Further, in the Ppr/ growth plate, the initial hypertrophic differentiation is delayed (Lanske et al., 1999
), whereas it is accelerated in chondrocyte-specific Ppr-null mice (e.g. Fig. 2J,K). Thus, the loss of the PPR in the cartilage as well as in the other tissues contribute to the Ppr/ growth plate phenotype. Nevertheless, the strong similarities of the cartilage phenotypes seen among Ppr/, Pthrp/ and Col2-Cre:Pprfl/fl mice suggests that these phenotypes are primarily caused by the loss of PPR signaling in chondrocytes.
Although universal gene ablation is a powerful method for analyzing the roles of genes in vivo, the extreme nature of the phenotypes can limit conclusions, especially when gene ablation causes early embryonic lethality. Partial knockout (knock-down) of gene expression can, therefore, be revealing. To date, several different methods have been reported to generate mice that have phenotypes due to reduced gene expression or function: transgenic animals that express antisense RNA (Nemir et al., 2000) or dominant negative proteins (Go et al., 2000
), introduction of hypomorphic mutation into proteins (Tang et al., 1997
) and insertion of foreign sequences into intronic sequences to disturb efficient RNA splicing (Nagy et al., 1998
; Meyers et al., 1998
; Mohn et al., 1999
). The mice with reduced PPR expression presented here were unintentionally obtained during an attempt to generate floxed PPR mice. Although the genetic structure of the d allele is not completely understood, the following findings indicate that the phenotypes of Pprd/ mice are caused by reduction of normal PPR mRNA expression: (1) PPR mRNA expression was reduced, as determined by RNA blot and in situ hybridization analysis; (2) the RT-PCR product had the normal coding sequence of PPR and the mRNA was of normal size; (3) homozygous Pprd/d mice have milder phenotypes than do Pprd/ mice (data not shown), a result expected if the d allele generates a smaller amount of a normal mRNA; (4) heterozygous Pprd/+ mice are virtually normal (data not shown); and (5) overactivity of the PPR caused by a constitutively active PPR mutation completely reverses the Pprd/ mutant phenotype. Based on northern blot analysis (Fig. 1B,C and data not shown) and the fact that the Pprd/d mice have slightly abnormal growth plates while Ppr+/ mice do not have apparent morphologic abnormalities, PPR expression level in Pprd/ mice is estimated to be less than 50% of that of heterozygous Ppr+/ mice. The growth plate phenotype of Pprd/ mice is superficially very different from that of Ppr/ mice. Close observations, however, revealed relative reductions of the periarticular and the columnar regions in Pprd/ mice, which are also seen in Ppr/ and Col2-Cre:Pprfl/fl mice in extreme forms. The diminished extent of the periarticular region, despite increased proliferation in this region in both Pprd/ and Col2-Cre:Pprfl/fl mice, suggests that loss or impairment of PPR signaling accelerates the differentiation of periarticular cells to columnar cells. The BrdU study also excluded the possibility that an increase in proliferation of columnar cells might have caused the early hypertrophic expansion in the Pprd/ growth plate. Thus, the expansion of the hypertrophic layer in the Pprd/ mice is probably caused both by acceleration of differentiation of periarticular to columnar cells and acceleration of differentiation of columnar cells into hypertrophic cells. The former process supplies cells to the pool with the highest proliferation rate; therefore, modest acceleration of this step may cause a substantial difference in the production of hypertrophic cells, further abetted by early conversion of proliferating cells to hypertrophic cells. This combination of cell transformation causes the Pprd/ mouse to have a tibia longer than normal at E17.5.
PTHrP expression in the periarticular region is dependent on Ihh expressed predominantly in the prehypertrophic chondrocytes. PTHrP, in turn, blocks premature hypertrophic differentiation of columnar chondrocytes (Kronenberg and Chung, 2001). However, Ihh clearly has roles in cartilage development independent of PTHrP, as Ihh/ mice show a growth plate phenotype distinct from that of Ppr/ mice, with marked reduction of chondrocyte proliferation. Expression of constitutively active PPR in the cartilage is able to reverse only acceleration of terminal differentiation of Ihh-null chondrocytes and is not able to rescue the reduced proliferation (Karp et al., 2000
). The positive association between cellular proliferation and Ihh activity is also observed in periarticular chondrocytes of the different models presented here. The increased proliferation accompanies increased rates of differentiation of periarticular cells to columnar/hypertrophic cells.
Based on the observations above, we propose that alteration of PPR signaling in chondrocytes changes the rates of differentiation of periarticular to the columnar chondrocytes (arrow 1) as well as the generation of hypertrophic chondrocytes (arrow 2) (Fig. 7A). This model can explain the diversified phenotypes in various PPR mutant growth plates (Fig. 7B). However, the data from mosaic PPR ablation demonstrate that PPR signaling itself does not directly regulate the first step. Conversely, Ihh activity is positively correlated with the acceleration of the first step, along with an increase in proliferation, suggesting that Ihh action may positively control this step (Fig. 7C). There remains a possibility that another factor secreted from the ectopic hypertrophic chondrocytes might be responsible for this step. This possibility, however, appears unlikely because a previous study of chimeric growth plates composed of wild-type cells and Ppr/;Ihh/ doubly mutant cells showed that Ihh in the ectopic hypertrophic chondrocytes was responsible for the characteristic elongation of the columnar layer of the growth plate (Chung et al., 2001).
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ACKNOWLEDGMENTS |
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