Intragenic microdeletion of RUNX2 is a novel mechanism for cleidocranial dysplasia
© Springer Science+Business Media B.V. 2008
Received: 24 June 2008
Accepted: 22 July 2008
Published: 12 August 2008
Cleidocranial dysplasia (CCD; MIM 119600) is a rare autosomal dominant disorder characterized by facial, dental, and skeletal malformations. To date, rearrangement and mutations involving RUNX2, which encodes a transcription factor required for osteoblast differentiation on 6p21, has been the only known molecular etiology for CCD. However, only 70% patients were found to have point mutations, 13% large/contiguous deletion but the rest of 17% remains unknown. We ascertained a family consisted of eight affected individuals with CCD phenotypes. Direct sequencing analysis revealed no mutations in the RUNX2. Real time quantitative PCR were performed which revealed an exon 2 to exon 6 intragenic deletion in RUNX2. Our patients not only demonstrated a unique gene change as a novel mechanism for CCD, but also highlight the importance of considering “deletion” and “duplication” in suspected familial cases before extensive effort of gene hunting be carried.
KeywordsCleidocranial dysplasia Intragenic deletion Mechanism RUNX2
Cleidocranial dysplasia (CCD; MIM 119600) is a rare autosomal dominant human skeletal disorder. The clinical features of CCD include facial and dental malformations characterized by delayed closure frontanelles, frontal bossing, absent clavicles, short stature, late eruption, and supernumerary permanent teeth and other skeletal anomalies (Mundlos 1999). There is considerable phenotypic variation for CCD, even within families (Chitayat et al. 1992). Mutations in the runt-related transcription factor 2 gene (RUNX2, also known as CBFA1, PEBP2αA, and AML3) located on chromosome 6p21 (Mundlos et al. 1997) have been identified as the cause of CCD. RUNX2 is one of the three mammalian homologs of the Drosophila runt gene, which encodes a transcription factor required for osteoblast differentiation. RUNX2 spans a region over 220 kb in 6p21 and is composed of eight exons and several splice variants have been described (Geoffroy et al. 1998). It has also been reported that RUNX2 is transcribed from two promoters (the distal promoter P1 and the proximal promoter P2) (Stewart et al. 1997). Numerous mutations in RUNX2 have been identified in patients with CCD (Otto et al. 2002; Yoshida et al. 2002; Zhou et al. 1999). Most of the missense mutations were located in the runt region (Baumert et al. 2005; Otto et al. 2002; Yoshida et al. 2002) involving heterodimerization and DNA binding with CBFβ. This discrepancy in distribution could be explained by that the runt domain is highly conserved and is less resistant to single nucleotide changes. Nonsense, splicing mutation, and insertion/deletions were also found and they were scattered throughout the entire RUNX2 gene. Deletion of the entire RUNX2 gene or larger has been described (Mundlos et al. 1995, 1997; Otto et al. 2002; Quack et al. 1999) and in one case the deletion spanning both RUNX2 and its upstream VEGF gene with the patient exhibiting both CCD and cardiovascular defects (Izumi et al. 2006).
Numerous CCD patients without any detectable mutations in RUNX2 by sequencing or FISH have been identified (Kim et al. 2006; Otto et al. 2002; Quack et al. 1999; Yoshida et al. 2002). This would indicate a genetic heterogeneity such as mutation in RUNX2 gene’s interacting proteins or regulatory elements or due to other mechanism that was not yet reported. One recent study identified a case with CBFβ mutation which encodes an interacting molecule of RUNX2. This individual did not have classical CCD phenotypes but exhibited delayed skull ossification and cleft palate (Khan et al. 2006).
In this study, we ascertained an extended family with many have classic yet severe CCD phenotypes. However, sequencing analysis did not reveal any mutations in RUNX2 and the results of FISH study were not confirmative. Further analysis using real time PCR, Southern blot, and reverse PCR revealed a novel microdeletion of about 125.6 kb and defined the breakage points in one allele of the gene. While intra-gene deletion involving multiple exons has been reported in many other genes, it has not been reported in CCD. The molecular mechanism for such deletion and the characteristic phenotype in this family are also discussed.
Materials and methods
Real time quantitative PCR (qPCR) for copy number analysis
RUNX2 copy number was determined by real time quantitative PCR reactions performed using Power SYBR GREEN PCR Master kit (Applied Biosystems, Foster City, CA, USA). Three independent experiments were performed to determine the variation in copy number between CCD patients and normal individuals with duplicate samples for each experiment. The RT-qPCR primers were designed according to manufacturer’s instruction. Primers were designed to detect copy number of the promoter, exons, and 3′ UTR regions of RUNX2. The qPCR reactions were performed using the ABI Prism® 7900HT Sequence Detection system and the fluorescent signal intensity was recorded on ABI Prism 7900HT Sequence Detection system and analyzed by Sequence Detector v2.3 software. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as control. The formula for calculating copy number was: copy number = 2 * 2−(ΔCtp − ΔCtn) where Ct was the threshold cycle defined as the mean cycle at which the fluorescence curve reached an arbitrary threshold; ΔCt was calculated as Ct of RUNX2 − Ct of GAPDH, ΔCtp was the ΔCt of patients, and ΔCtn was the ΔCt of normal individuals. Two normal individuals were used in the experiments.
Inverse PCR for deletion mapping
Inverse PCR is a method for the rapid in vitro amplification of unknown DNA sequences that is flanked by a region of known sequences (Ochman et al. 1988). Restriction digestion was first carried out as described above and the enzymes were inactivated at 65°C for 20 min. The digested DNA fragments were allowed for self-ligation to generate circular DNA. Amplification was then performed with outward facing primers 5′-GTTCCTGCAAAGAATGGTCC-3′ and 5′-TAGAGCAGGGAAACCCACAG-3′. Sequencing of the unknown region can then be performed on the amplified DNA with the above primers.
Molecular analysis of the RUNX2 gene
Initial sequencing of all exons, intro-exon junctions, and 2 kb upstream of transcription start site of the RUNX2 failed to identify any mutations in the gene. FISH analysis was subsequently performed on one patient (I:8) and one normal control in this extended family to determine the presence of deletions in RUNX2. The signals on one of the chromosome 6 homologs in the patient’s cells appeared weakened (data not shown). These findings suggested that one of the chromosome 6 of the CCD patient could have deletion involving a portion of the RUNX2 gene.
Southern blot analysis was next performed to narrow down the region that harbored the 3′ break point and mapped the 3′ breakage point to within 1 kb between exon 6 and exon 6.1 (data not shown). This also confirmed the qPCR data that the deletion was indeed present.
The work presented here described the first large intragenic microdeletion: exons 2–6 were deleted in the three generation CCD family. This deletion created a truncated protein without most of the N-terminal domain. Without the DNA binding runt domain, this protein was unable to modulate transcription of RUNX2 downstream genes. Consequently, CCD phenotype arose as a result of haploinsufficiency of RUNX2 (Mundlos et al. 1997; Otto et al. 1997).
This family was significantly shorter than the reported cases (137 vs 156) and (150 vs 165) (Cooper et al. 2001). The average height of the affected adults was 148 cm. While we cannot exclude the contribution of ethnic background for the shorter stature, the short distal phalangeal hyperplasia and small hands were very significant in this family and could be explained by this intragenic deletion. Cesarean section has been reported to be unusually high up to 69% (Cooper et al. 2001); however, it is not reported in this family.
Using Southern blot and inverse PCR, we have determined precisely the breakage points of the RUNX2 deletion in this three generation CCD family. It is interesting to note that both ends contain the same three nucleotides ATC. However, the sequence homology is probably too short for homologous recombination to occur. Thus, this particular intragenic deletion is most likely generated through non-homologous end joining.
FISH is a useful tool to detect microdeletion; however, its sensitivity depends on the size of the microdeletion and the location and size of the probe. The BAC clone used, RP11-1019C24 (191 kb), located within the RUNX2 gene (220 kb) should be the best probe for the detection of RUNX2 deletions. However, even with this probe the FISH study was inconclusive due to the partial RUNX2 deletion.
The work described here can also be a good example for other studies. While genetic heterogeneity and pathway molecules can be the alternative mechanism when a mutation is not found, we suggest detailed study for a known gene before ever-ending effort in linkage be put forward as a general rule. When sequence variants are not detected by direct sequencing, real time PCR assays similar to the one used in this study or MLPA would allow detection of gene deletion or duplication efficiently.
In summary, we have identified the first intragenic microdeletion in RUNX2 in a CCD family. Current clinical testing by sequence-based study only detects 60–70% of individuals with a clinical diagnosis of CCD. Microdeletion with contiguous deletion has been suggested to account for another 13% (Mendoza-Londono and Lee 2008). In our cohort, 28% is due to deletion (unpublished data). Our patients demonstrated a rare and novel deletion for CCD. We therefore suggest that in patients whose mutation is not found by traditional sequencing, the deletion/duplication assay, either RT-qPCR/MLPA, needs to be done particularly in a disease haploid insufficiency is thought to be the main cause. The deletion/duplication assay can improve the molecular diagnosis of CCD and likely change the statistics of molecular mechanism of this disease.
We would like to thank Yi-Wen Lin for her assistance in the Southern blot analysis. This research project was supported by grants from the National Research Program for Genomic Medicine, National Science Council, Taiwan (National Clinical Core, NSC95-3112-B-001-010 and National Genotyping Center, NSC95-3112-B -001-011). The authors declare no conflict of interests.
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