Patients and methods

BWS PATIENTS

A diagnosis of BWS was made according to standard diagnostic criteria, either (1) three major features (anterior abdominal wall defects, macroglossia and pre-/postnatal growth >90th centile) or (2) two major plus three or more minor features (characteristic ear signs (ear lobe creases or posterior helical ear pits), facial naevus flammeus, hypoglycaemia, nephromegaly, and hemihypertrophy).1 Seventy patients with classical BWS were analysed. Results of uniparental disomy analysis and H19 methylation have been reported previously.13 14 21 In addition, six patients with Wilms tumour and overgrowth (>90th centile) were also investigated.

MOLECULAR GENETIC ANALYSIS

Genomic DNA was isolated from whole blood as described previously.13 14 The following five sets of primers were used to amplify the entire coding region of the CDKN1C gene and its exon-intron boundaries in overlapping fragments (fig 1). Set A forward primer, 5′-CGTTCCACAGGCCAAGTGCG-3′ and reverse primer 5′-GCTGGTGCGCACTAGT ACTG-3′; set B forward primer, 5′-CGTCCC TCCGCAGCACATCC-3′ and reverse primer 5′-CCTGCACCGTCTCGCGGTAG-3′; set C forward primer, 5′- TGGACCGAAGTGGA CAGCGA-3′ and reverse primer 5′- GGGGCCAGGACCGCGACC-3′; set F forward primer, 5′-CGGAATTCCGGAGC AGCTGCCTAGTGTC-3′ and reverse primer 5′- CTTTAATGCCACGGGAGGAGG-3′; set H 5′-CGGCGACGTAAACAAAGCTG-3′ and reverse primer 5′-GGTTGCTGCTACATGA CCGG-3′.

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Figure 1

Structure of the CDKN1C (p57^KIP2) gene, including coding regions and protein domains. The relative positions of primer pairs used for template preparation in the sequencing reactions are also shown.

The reaction mixtures used for fragments A, B, C, and H were 10 mmol/l Tris-HC1, pH 8.8, 50 mmol/l KC1, 1.0 mmol/l MgC1₂, 0.25 mmol/l dATP, dTTP, dCTP, dGTP, 10% dimethyl sulfoxide, and 0.75 unitsTaq polymerase (Amplitaq) in a final volume of 30 μl. For fragment F the reaction mixtures were 1 × LA PCR Buffer II (Takara), 0.2 mmol/l dATP, 7-deaza dGTP, dCTP, dTTP, 5% dimethyl sulfoxide, and 2.5 units Taqpolymerase in a final volume of 50 μl. A Perkin-Elmer Thermocycler was used for PCR analysis. Temperature conditions comprised: fragment A, a single denaturing step at 95°C for four minutes followed by 35 cycles of 94°C for 30 seconds, 55°C for 30 seconds, and 72°C for one minute, no primer extension step was used; fragment B, a single denaturing step at 95°C for four minutes followed by 35 cycles of 94°C for 30 seconds, 64°C for 30 seconds, and 72°C for one minute, no primer extension step was used; fragment C, a single denaturing step at 95°C for four minutes followed by 35 cycles of 94°C for 30 seconds, 59°C for 30 seconds, and 72°C for one minute, no primer extension step was used; fragment F, a single denaturing step at 95°C for four minutes followed by 40 cycles of 95°C for one minute, 65°C for one minute, and 72°C for 1.5 minutes, and a primer extension step for 72°C for 10 minutes was used; fragment H, a single denaturing step at 95°C for four minutes followed by 35 cycles of 94°C for 45 seconds, 53°C for 30 seconds, and 72°C for one minute, no primer extension step was used.

PCR fragments were purified before sequencing using Amicon columns according to the manufacturer’s instructions. Sequencing was performed on an ABI PRISM 377 semiautomated sequencer. Both forward and reverse strands were sequenced in every case using the di-rhodamine dye-terminator chemistry sequencing kit and protocol (Applied Biosystems). Reactions were set up as follows: 4 μl di-rhodamine kit reaction mix, 16 pmol sequencing primer, PCR template (1-5 μl depending upon the product concentration), and sterile distilled water to 10 μl (final volume) and overlayed with mineral oil (20 μl), before thermal cycling on a Omn-E thermal cycler (Hybaid) as follows: 96°C for three minutes followed by 30 cycles of 98°C for 10 seconds, 55°C for 20 seconds, 60°C for four minutes. Samples were then precipitated, washed, resuspended, denatured, and loaded onto the ABI 377 sequencer according to the manufacturer’s instructions. CDKN1C genomic sequence numbering is as given in Genbank Accession no D64137. The CDKN1C coding sequence starts at nucleotide 1261.

GENOTYPE-PHENOTYPE ANALYSIS

The clinical features of BWS patients with confirmed germline CDKN1C mutations identified in this study and those by Hatadaet al 31 32 were compared to the clinical phenotypes of patients we have identified previously with uniparental disomy or an IGF2/H19 ICD.13 14 21

Results

CDKN1C MUTATION ANALYSIS

Seventy patients with classical BWS were examined for germline mutations in the CDKN1C gene by direct sequencing of the coding region and intron-exon boundaries. Fifty four were unrelated patients with sporadic BWS and no evidence of uniparental disomy for chromosome 11p15.5 markers.13 14 Five of the 54 non-familial patients had H19 hypermethylation suggesting an ICD14 21and 49 had a normal biparental H19 methylation pattern. Sixteen of the 70 patients examined were familial cases from seven kindreds. In addition, six patients with Wilms tumour and overgrowth (>90th centile) were analysed.

Germline CDKN1C sequence variants were identified in seven BWS probands. In four cases, the nucleotide change occurred within the coding sequence and was predicted to cause a change in protein sequence (two missense and two protein truncating) (table 1, fig 2). The two protein truncating mutations were from familial cases (F1 and F2), and in both instances the mutation was maternally inherited. The human CDKN1C protein consists of three structurally distinct domains (fig 1): (1) the 5′-amino-terminal, which is significantly similar to regions of p21^CIP1 and p27^KIP1 and has been shown to be necessary for Cdk inhibition, (2) a highly polymorphic hexanucleotide repeat encoding a proline-alanine series of repeats, and (3) a highly conserved 3′-QT box containing glutamine and threonine residues that is conserved between CDKN1C (p57^KIP2) and p27^KIP1. Both familial protein truncating mutations occurred within exon 2. Two non-familial patients (S1 and S2) had previously unreported nucleotide substitutions within the coding region producing putative missense mutations. S1 showed a de novo non-conservative proline to leucine substitution within the conserved Cdk inhibitory domain. The parental transmission of the non-conservative arginine to tryptophan substitution in S2 could not be established with certainty. This mutation arises in the C-terminal amino acid and was not present in the mother of S2, but the father’s DNA was unavailable. In a further familial case (F3), a C to G substitution at position -3 of the exon 3 splice acceptor was characterised in the proband, an affected sib, and their mother. This change was not identified in 50 control chromosomes and in a survey of human splice sites,36 no wild type splice acceptor sites with a G at position -3 were reported. Further supportive evidence that this transversion was likely to be pathogenic was provided when the wild type and mutant sequences were analysed by the Splice Site Prediction Neural Network Program (Reeseet al),http://www-hgc.lbl.gov/project/splice.html). The program correctly identified all the splice sites within the wild type sequence, but there was a complete failure to recognise the mutant 3′ splice acceptor site.

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Figure 2

(A) Sequence analysis of patient F3 showing the C→G splice site mutation (see table 1 for details). (B) Sequence analysis for S2 illustrating the C→T transition causing an arginine→tryptophan missense mutation in the QT domain (see table 1for details).

Overall, germline CDKN1C mutations were identified in 43% (three of seven) familial cases analysed, and 4% (two of 55) non-disomic sporadic cases. In addition, two previously unreported nucleotide substitutions in the 5′ untranslated region were identified in two sporadic patients (S3 and S4). In both cases the sequence variant was maternally inherited and may represent either a regulatory mutation or a rare polymorphism. None of the putative regulatory mutations in S3 or S4 or the missense mutation in S2 have been detected in 100 control chromosomes (unpublished observations), which, together with the pattern of inheritance in the familial cases, favours the interpretation that these are disease associated mutations. No germline mutations (or sequence variants) were identified in six children with Wilms tumour and overgrowth.

PHENOTYPIC EXPRESSION OF CDKN1C MUTATIONS IN BWS

BWS is genetically heterogeneous and we therefore investigated the relationship between phenotypic expression of BWS and molecular pathology. We compared the phenotype of BWS patients with the CDKN1C coding sequence/splice site mutations identified in this study (n=8) and those reported previously by Hatada et al 31 ³²¹ (n=5) to that of BWS patients with other types of defined molecular pathology (table 2). A striking difference was a very high incidence of exomphalos (11/13) among patients with CDKN1C mutations. The incidence of exomphalos in CDKN1C mutation cases was significantly higher than in patients with an ICD (0/5) (p<0.005) or those cases with UPD (0/9) (p<0.0005). However, germline CDKN1C mutations do not account for all BWS patients with exomphalos and 12 BWS cases with exomphalos did not have a detectable CDKN1C mutation. There were no other significant phenotypic differences between the three molecular subgroups (data not shown). None of the patients with CDKN1C mutations developed Wilms tumour.

Discussion

We have confirmed that germline CDKN1C mutations account for a subgroup of patients with BWS. Our results suggest that the molecular pathology of familial and sporadic BWS differs, such that mutations in CDKN1C account for only a minority of sporadic classical BWS cases but a substantial proportion of familial cases. Unfortunately, it is not possible to elicit the relationship between family history and CDKN1C mutation status in studies by other groups. However, in Japanese BWS patients, Hatada et al 31 32found germline CDKN1C mutations in four (one familial, three sporadic) of 24 cases. Lee et al 33 found CDKN1C mutations in two (one familial and one unknown) of 40 BWS patients. Although O’Keefe et al 34 identified a germline CDKN1C mutation in the only familial case studied (and in none of four sporadic cases), a full breakdown of family history status is not available for the patients studied by Hatada et al 31 32 and Lee et al.33 Finally, Okamoto et al,35 using SSCP analysis, did not detect germline CDKN1C mutations in 40 BWS patients (five of whom were familial). The mutation detection techniques used by us and others would not have identified large CDKN1C deletions or epigenetic silencing of CDKN1C. Thus the frequency of CDKN1C involvement in BWS may have been underestimated. Our observation that germline CDKN1C mutations are significantly more frequent in familial than in sporadic BWS cases is consistent with our previous findings that the majority of sporadic BWS cases have LOI of IGF2.17 Nevertheless, although CDKN1C mutations are uncommon in sporadic cases, the identification of a germline CDKN1C mutation has major implications for genetic counselling and is therefore of considerable clinical importance.

To date no recurrent CDKN1C mutations have been reported. Familial BWS is characterised by a higher penetrance and more severe phenotypic expression when the disease is maternally transmitted, features that are compatible with the preferential maternal expression of CDKN1C. However, Lee et al 33 reported partial features of BWS in a mother of a classical BWS patient who had inherited a CDKN1C mutation from her father.

In our study, all patients with a germline CDKN1C mutation had a normal H19/IGF2 methylation pattern indicating that the CDKN1C gene does not represent the H19/IGF2 regional imprinting control centre inactivated in patients with an ICD. Furthermore, CDKN1C mutations were not identified in the five ICD patients analysed and in 11 sporadic BWS patients with loss of imprinting of IGF2 and normal H19/IGF2 methylation status.17 Direct analysis of IGF2 imprinting status has not been possible in patients with CDKN1C mutations because of absence of suitable material or uninformativeness of transcribed polymorphisms.

BWS is clinically and molecularly heterogeneous. The variability of the BWS phenotype may be determined by the differential involvement of several imprinted genes according to the precise molecular pathology. This hypothesis is compatible with our findings of genotype-phenotype correlations in BWS. Overall, exomphalos occurs in ∼45% of cases1; however, we have shown that CDKN1C inactivation is associated with a high risk of exomphalos (87%), while patients with UPD and IGF2/H19 hypermethylation (ICD) have a low frequency of exomphalos. The absence of CDKN1C mutations in non-BWS overgrown patients with Wilms tumour is compatible with these findings. As we might expect UPD to result in reduced CDKN1C expression, these findings are at first sight contradictory. Nevertheless, as UPD cases are mosaic, the presence of a proportion of normal cells may markedly reduce the severity of the phenotype. This may be further offset by the relatively relaxed imprinting of the paternal allele, which expresses detectable levels of CDKN1C mRNA.37 Consistent with our predictions, BWS patients with a chromosome 11p15.5 duplication do not have exomphalos.38 It is possible that some patients with exomphalos and no detectable CDKN1C mutation have epigenetic silencing of CDKN1C mRNA expression.

Our genotype-phenotype findings are reinforced by the results of two recent mouse models of BWS. Thus a cdkn1cknockout mouse displayed a high frequency of anterior abdominal wall abnormalities including exomphalos, but lacked some other features of BWS (including overgrowth).39 Furthermore, mice with IGF2 protein overexpression mimic many features of BWS but do not show exomphalos.15 Exomphalos in BWS has generally been attributed to result from organomegaly and a concomitant increased intra-abdominal pressure. However, the preferential association of exomphalos with CDKN1C mutations suggests that this explanation is insufficient and that additional factors (for example, disturbance of normal cellular migration or adhesion patterns) are likely to be indicated. Genitourinary development abnormalities are common in BWS, but the association with prune belly syndrome (seen in case F3a) is rare. It is not known if this association will be specific to CDKN1C mutation cases or if it is coincidental (although the incidence of isolated prune belly syndrome is ∼1 in 3000). Further studies of the molecular pathology of BWS including the imprinting status of IGF2, H19, CDKN1C, and KCNQ1 in patients with different subgroups of molecular lesions, followed by further analysis of genotype/phenotype relationships should provide insight into the pathogenesis of BWS.

The mechanism by which mutations in CDKN1C may cause BWS is unclear. Overexpression of CDKN1C causes cell arrest at G1 and so inactivating mutations might lead directly to abnormal cellular proliferation. In the mouse, IGF2 protein overexpression but not CDKN1C inactivation is associated with overgrowth.14 39 As we did not find evidence that CDKN1C mutations directly affect IGF2 imprinting (see above), the most likely explanation for the phenotypic overlap of CDKN1C mutations and IGF2 overexpression is that the two proteins are implicated in a common growth control pathway. If so, this throws a new light on the observation that imprinted genes tend to be clustered in the genome.2 Within the 11p15.5 imprinted gene cluster, IGF2 and H19 have been functionally linked by the enhancer competition model and further studies of IGF2, CDKN1C, and other 11p15.5 imprinted genes may elucidate functional relationships that provide an explanation for their genomic clustering.