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Human Molecular Genetics Pages 1021-1028

Elastin point mutations cause an obstructive vascular disease, supravalvular aortic stenosis
Introduction
Results
   Genomic structure of ELN
   SSCP analyses reveal ELN point mutations
Discussion
Materials And Methods
   Phenotypic evaluation
   DNA sequence and exon-intron determination
   SSCP analysis
Acknowledgements
References

Elastin point mutations cause an obstructive vascular disease, supravalvular aortic stenosis

Elastin point mutations cause an obstructive vascular disease, supravalvular aortic stenosis Dean Y. Li1,3,4, Amanda E. Toland2,3, Beth B. Boak3,4, Donald L. Atkinson2,3,4, Gregory J. Ensing5, Colleen A. Morris6 and Mark T. Keating1-4,*

1Cardiology Division, University of Utah Health Sciences Center, 2Department of Human Genetics, 3Eccles Institute of Human Genetics, 4Howard Hughes Medical Institute, Salt Lake City, Utah 84112, USA, 5Department of Pediatrics, Division of Pediatric Cardiology, Indiana University School of Medicine, Indianapolis, IN 46202, USA and 6Department of Pediatrics, University of Nevada, Las Vegas, NE 89102, USA

Received December 12, 1996; Revised and Accepted April 8, 1997

Supravalvular aortic stenosis (SVAS) is an inherited obstructive vascular disease that affects the aorta, carotid, coronary and pulmonary arteries. Previous molecular genetic data have led to the hypothesis that SVAS results from mutations in the elastin gene, ELN. In these studies, the disease phenotype was linked to gross DNA rearrangements (35 and 85 kb deletions and a translocation) in three SVAS families. However, gross rearrangements of ELN have not been identified in most cases of autosomal dominant SVAS. To define the spectrum of ELN mutations responsible for this disorder, we refined the genomic structure of human ELN and used this information in mutational analyses. ELN point mutations co-segregate with the disease in four familial cases and are associated with SVAS in three sporadic cases. Two of the mutations are nonsense, one is a single base pair deletion and four are splice site mutations. In one sporadic case, the mutation arose de novo. These data demonstrate that point mutations of ELN cause autosomal dominant SVAS.

INTRODUCTION

With the exception of hypercholesterolemia and hypertension, little is known about the genetic factors that contribute to vascular disease. Supravalvular aortic stenosis (SVAS) is an inherited obstructive vascular disorder that causes hemodynamically significant narrowing of large arteries (1 ). Although the aorta is most frequently diseased, any artery can be affected, including the pulmonary, carotid and coronary arteries. The onset and severity of vascular disease in SVAS is variable. If untreated, this disorder can lead to heart failure, myocardial infarction and death. Currently, the only treatment option is surgery.

SVAS can be inherited as an autosomal dominant trait (1 -3 ) or as one feature of Williams syndrome (Williams syndrome) (2 -4 ). In addition to vascular disease, Williams syndrome manifestations include mental retardation, characteristic personality, specific cognitive profile, infantile hypercalcemia, dysmorphic facial features and other connective tissue abnormalities. The incidence of familial and syndromic SVAS is estimated at 1 in 20 000 live births (5 ).

Four lines of evidence support a role for ELN in SVAS. First, ELN is genetically linked to the disease phenotype in autosomal dominant SVAS (5 -7 ). Second, large DNA rearrangements that disrupt ELN are associated with the disease in three SVAS families (8 -10 ). Third, hemizygosity of ELN is observed in individuals with Williams syndrome (11 ). Fourth, the structural and functional importance of elastin in large elastic arteries is consistent with a role for ELN in this disease. However, all molecular genetic data implicating ELN in SVAS involve large DNA rearrangements (>550 kb deletions in Williams syndrome; 35 and 85 kb deletions and a translocation in familial SVAS). Most cases of non-syndromic autosomal dominant SVAS are not associated with large DNA rearrangements.

In this study, we cloned human ELN, refined its genomic structure and used this information to develop oligonucleotide primers suitable for mutational analysis. Single strand conformation polymorphism analysis (SSCP) identified ELN point mutations in four familial and three sporadic cases of SVAS. Two of the mutations were nonsense, one was a frameshift and four were splice site mutations. In one sporadic case, the mutation arose de novo. These data establish that point mutations in ELN can cause autosomal dominant SVAS.

RESULTS

Genomic structure of ELN

Previous work on the genomic structure of human ELN did not define intron-exon boundaries or intronic sequences (12 ,13 ). To facilitate identification of point mutations in human ELN, we constructed a contiguous physical map comprised of cosmid and phage clones that span 55 kb of chromosome 7q11.23 (Fig. 1 ). The numbering of 34 exons in human ELN was based on homology with bovine ELN, which has 36 exons (12 ,13 ). Exons 34 and 35 of bovine ELN have no counterparts in human ELN. The intron-exon boundaries are shown in Table 1 and are available in GenBank. Analysis of this structure indicated that the last nucleotide of each ELN exon is spliced with the first two nucleotides of the following exon to form a codon. Thus, removal of any internal ELN exon would not result in a frameshift but in the loss of amino acids encoded by the spliced exon. We designed oligonucleotide primers for mutational analyses from intronic sequences (Table 2 ).

Table 1 Intron-exon boundaries of human ELN
  Exon Intron Exon
Exon 1 CGG CCT GGA G gtaaggacc....tcctttcag GG GTC CCT GGG Exon 2
  R P G   G V P G
Exon 2 TTT TAT CCA G gtaacgtac....cccccacag GG GCT GGT CTC Exon 3
  F Y P   G A G L
Exon 3 GGA GGA GGA G gtgagctca....cctctgcag CG CTG GGG CCT Exon 4
  G G G   A L G P
Exon 4 CTT AAG CCA G gtaagaccc....tatccacag TT CCC GGA GGG Exon 5
  L K P   V P G G
Exon 5 CTT GGG GCA G gtgagtgct....cctccgcag GG CTC GGC GCC Exon 6
  L G A   G L G A
Exon 6 GCT AAG GCT G gtgagtggt....gctctgcag GC GCT GGG CTT Exon 7
  A K A   G A G L
Exon 7 GTG TCT GCA G gtacgatgg....ctgtgccag GT GCG GTG GTT Exon 8
  V S A   G A V V
Exon 8 AAA GTG CCG G gtcagtgcg....tggccacag GT GTG GGG CTG Exon 9
  K V P   G V G L
Exon 9 GTG CTC CCA G gtgagagca....ccccggcag GA GCT CGG TTC Exon 10
  V L P   G A R F
Exon 10 AAG GCT CCA G gtatgcagc....tcattacag GT GTA GGT GGA Exon 11
  K A P   G V G G
Exon 11 GGA ATC CCA G gttgaggca....ttcccacag GA GTT GGA CCC Exon 12
  G I P   G V G P
Exon 12 AAG CTG CCT G gtaagtcag....tctctccag GT GGC TAT GGA Exon 13
  K L P   G G Y G
Exon 13 CTG CCC TAT G gtgagtgag....gttttgcag GC TAT GGG CCC Exon 14
  L P Y   G Y G P
Exon 14 ACA GGG ACA G gtaaggaaa....ttcccccag GG GTT GGC CCC Exon 15
  T G T   G V G P
Exon 15 GCA AAG TTC G gtgagtgcc....tctcaacag GT GCT GGA GCA Exon 16
  A K F   G A G A
Exon 16 GGC ATC GCA G gtaacatct....tgttttcag GC GTT GGG ACT Exon 17
  G I A   G V G T
Exon 17 GCC AAG TAT G gtgagtgcc....cttccacag GA GCT GCT GCA Exon 18
  A K Y   G A A A
Exon 18 GCG GTT CCA G gtgagctgg....tctctccag GG GTT GTG TCA Exon 19
  A V P   G V V S
Exon 19 GCC AAA TAC G gtgagtgct....cctctgcag GG GCC AGG CCC Exon 20
  A K Y   G A R P
Exon 20 GGC ATT TCC C gtgagcctt....tccttgtag CC GAA GCT CAG Exon 21
  G I S   P E A Q
Exon 21 GCC AAG TAC G gtaagtgcc....ctccccgag GT GCT GCA GGA Exon 22
  A K Y   G A A G
Exon 22 GGA GTG CCA G gtgagctgt....cacctccag GA GTG GGG ACC Exon 23
  G V P   G V G T
Exon 23 GCC CAG TTT G gtaagtccc....aatgatcag CT CTT CTC AAT Exon 24
  A Q F   A L L N
Exon 24 GGA GTT GCA G gtgagtttc....ctccctcag CT GCA GCA AAA Exon 25
  G V A   A A A K
Exon 25 GCC CAG CTC C gtgagtgcc....tgtctgcag GA GCT GCA GCT Exon 26
  A Q L   R A A A
Exon 26a TTC GGG GCA G gtgcagatg....  
  F G A    
Table 1. Continued
Exon 26b TCA CCC AGG G gtgcatagt....tctccccag TA CCT GGA GCC Exon 27
  S P R   V P G A
Exon 27 GCC AAA TAT G gtgagtgca....tccccgcag GA GCA GCA GTG Exon 28
  A K Y   G A A V
Exon 28 GGT GTG GTG G gtgagttga....tgttcccag GA GCC GGA CCC Exon 29
  G V V   G A G P
Exon 29 GCC CAG TTT G gtgagcact....tctccacag GC CTA GTG GGA Exon 30
  A Q F   G L V G
Exon 30 GGC CTT GGA G gtgagagtt....tctccccag GT ATA CCT CCA Exon 31
  G L G   G I P P
Exon 31 GCT AAA TAC G gtgagttcc....tcttctcag GT GCT GCT GGC Exon 32
  A K Y   G A A G
Exon 32 CCA CTT GGA G gtaggggtg....ctcctgcag GA GTG GCA GCA Exon 33
  P L G   G V A A
Exon 33 ATT TTC CCA G gtatgccag....cccgtccag GT GGG GCC TGC Exon 36
  I F P   G G A C
Numbering of the 34 exons in human ELN is based on homology with the bovine elastin gene, which has 36 exons. Exons 34 and 35 of bovine elastin have no counterparts in human ELN. DNA sequence for intron-exon boundaries are shown using the consensus sequence defined by Shapiro and Senapathy (32). Intron sequences are shown in lower case, exon sequences in uppercase. The corresponding amino acids are listed below the exon sequences. The last nucleotide of each exon forms a codon with the first two nucleotides of the following exon. Exon 26 of human ELN has two alternative donor splice sites, resulting in an exon of either 126 or 225 bp. The donor site for the 126 bp product is listed as exon 26a while the donor site for the 225 product is listed as exon 26b.

SSCP analyses reveal ELN point mutations

To determine if ELN point mutations cause SVAS, we performed SSCP analyses on DNA samples of 20 unrelated individuals with familial and sporadic SVAS. No ELN rearrangements were identified in these DNA samples by fluorescence in situ hybridization, pulsed-field gel electrophoresis or Southern analyses (data not shown).

Previous work demonstrated that SVAS was linked to a polymorphic marker near ELN in kindred 1779 (K1779; 5). SSCP analyses using primers amplifying exon 26 identified an anomalous band that co-segregated with the disease in this family (Fig. 2 ). The aberrant conformer was not observed in DNA samples of >375 unaffected and unrelated control individuals. The conformer was cloned and sequenced, leading to the identification of a single nucleotide deletion at position 1821 in exon 26, 1821delC. This deletion caused a frameshift, resulting in a premature stop codon in exon 28. A second change, 1824C -> T, identified in the aberrant conformer is silent.


Figure 1. Physical map of the human ELN. The genomic organization of human ELN is shown (A). Two lambda and two cosmid clones spanning the entire gene are indicated (B) followed by a BamHI restriction map (C).


Figure 2. Frameshift mutation in ELN exon 26 associated with familial SVAS. The pedigree structure for K1779 is shown. Individuals with SVAS are indicated by filled circles (females) or squares (males). Phenotypically unaffected individuals are denoted by open symbols. Individuals with an unknown phenotype are indicated by gray symbols. The results of SSCP analyses using primers that amplify ELN exon 26 are shown below the pedigree. Anomalous bands, indicated by arrows, co-segregate with the SVAS phenotype in this kindred. DNA sequence analyses of aberrant conformers revealed deletion of a C residue at nucleotide 1821,1821delC, resulting in a premature stop codon in exon 28 (nucleotide 2020). A second change, 1824C -> T, identified in the aberrant conformers is silent. The deleted nucleotide is enclosed by a box. The corresponding amino acids are indicated below the DNA sequence.


Figure 3. Nonsense mutations in ELN exons 21 and 25 associated with sporadic SVAS. SSCP analyses using primers to exons 21 and 25 in two sporadic cases of SVAS, K2178 (A) and K2016 (B), are shown. Sequence analysis of DNA obtained from the affected members of K2178 revealed a C to T transition at nucleotide 1324, 1324C -> T, resulting in a premature stop codon in exon 21. In K2016, analysis defined a C to T transition at nucleotide 1708, resulting in a premature stop codon in exon 25, 1708C -> T. The nucleotides involved are indicated by boxes.

Mutational analyses using primers for exons 21 and 25 defined aberrant conformers in two sporadic cases of SVAS, K2178 (Fig. 3 A) and K2016 (Fig. 3 B). These anomalies were not seen in >375 control DNA samples. In K2178, a C to T transition was identified at nucleotide 1324, 1324C -> T, resulting in a nonsense mutation in exon 21. In K2016, a C to T transition was identified at nucleotide 1708, 1708C -> T, resulting in a nonsense mutation in exon 25. DNA samples could not be obtained from the parents of either proband.

SSCP analyses using primers for exon 16 demonstrated an identical anomalous conformer that co-segregated with the disease in two unrelated kindreds, K2260 and K2205 (Fig. 4 ). Analyses of a third kindred, K2044, revealed a second anomalous band. These conformers were not seen in >375 control DNA samples. DNA sequence analyses revealed an A to G transition, IVS15-2A -> G, (K2260 and K2205) and a C to G transversion, IVS15-3C -> G, (K2044) in the splice acceptor site preceding exon 16.

Mutational analyses using primers flanking exon 3 revealed an anomalous conformer in a sporadic case of SVAS (K2017; Fig. 5 ). This conformer was not observed in the proband's parents, indicating that it arose de novo. Paternity and maternity were confirmed by genotypic analysis using seven short tandem repeat polymorphisms (data not shown). The anomaly was not observed in analyses of >375 control DNA samples. DNA sequence analyses revealed a G to A transition in the 5' splice donor site following exon 3, IVS3+1G -> A.

To determine if the point mutations described here result in the synthesis of abnormal or reduced mRNA ELN transcript, we used reverse transcriptase-polymerase chain reaction (RT-PCR) to isolate ELN transcripts from lymphoblastoid cells. However, we were unable to detect ELN transcripts from experimental and control lymphoblastoid cells. Skin fibroblasts were not available for study.

To determine if a specific ELN mutation could be used to predict the severity of vascular disease, we re-examined the phenotypic data in this genotypically defined population. We found that intrafamilial variability of phenotypic expression was as great as interfamilial variability. For example in K1779, the severity of the phenotype ranged from an asymptomatic 65-year-old male who had mild SVAS by echocardiography to a severely affected 16-year-old female who required surgical correction. In K2205, phenotypic variation ranged from an asymptomatic 50-year-old female who had minimal SVAS by cardiac angiography to a 4-year-old male with severe disease requiring surgical palliation. Of the 17 individuals with ELN point mutations, 16 have documented SVAS by echocardiography and/or angiography. For one individual with an ELN point mutation, the mother of the affected individual in K2260, we could not obtain phenotypic data. Thus, the penetrance in this study was 100%.

DISCUSSION

We conclude that point mutations in ELN cause an inherited vascular disorder, SVAS. In this study, we cloned human ELN, refined its genomic structure and used this information to synthesize oligonucleotide primers for mutational analyses. Using SSCP analyses, we identified ELN point mutations associated with disease in four SVAS families and three sporadic cases. These mutations included nonsense, frameshift and splice site mutations. In one case, we demonstrated thatthe ELN mutation arises de novo. Combined with existing molecular and histologic data, these findings establish that point mutations in ELN cause SVAS.

Table 2 PCR primers for human elastin exons
Exon Forward Reverse Product size
1 CTCTTTCCCTCACAGCCGAC GTCTAGTCACCTGCCAAAGGG 229 bp
2 CAGCATTATTCTTGTTTCCA GACTTCTGGTAGGCGTGTCA 296 bp
3 CAAGGTCACGTAGTTAGGCA GCCTCCTCCCACCGCCTGGA 284 bp
4 GTGATTCCACACTGCCCACA GTTGCCTTGTCCTGTTCCTA 287 bp
5 GATCGACCCTGAGCATCACA GTCCCCGTCAGTGCCCAGGA 305 bp
6 GTCTGGCAGAGAGCGGAAGA GGAGGGCCCCACGGAAGCCA 314 bp
7 GCTAGCTGTGCCCAGTCTGA CTCCGGCTCCAGGCTGAGGA 298 bp
8 GCAAGGCTTGGTGGAGCCAA CGCAGGCACCAGCCTGGCTA 288 bp
9 CTGCCCCTGTCGGGCACAGA GCCTCAGTCTCCCAAAGCAA 322 bp
10 GAGGGGAGGGGTTCCCAGCA GGATCCCACCCCATTTCACA 225 bp
11 CCTTGGTGCTGTCTGGCCCA GGGTGCCACTTTGTGCTACA 292 bp
12 GAAGCAGGATGGTTTCTGGA GAACCACCACACCCGGCCGA 285 bp
13 CTTTAGCACCTGTGGGGGTA CGAGGTGCTCTCCTGGCCCA 278 bp
14 GGTGATGTCTGCACAGATGA CTGGAGCCTGGAAGATCAGT 160 bp
15 GAAGCTCCCATGTATACCCA CTTTGGAGGCAGGCTGTCAA 258 bp
16 CCCAGACAGGCCCATCTGGA GTGGCCATCTCAAGGACACA 279 bp
17 GTGTCCTTGAGATGGCCACA CCCGAGCCGTGGACTTACCA 170 bp
18 GAAATGTCAACCCACCTGCA GTCTCAATATTTCTCTCTAA 360 bp
18B   CACACACACAGCCCAGCTCA 293 bp
19 CCAACACACAGATGGGTAGA GGCAAGTAAAGGCCAGGACA 238 bp
20 CTCACTGAGCTTCTTTTCTA CTTCTCAACCCATGTCCCCA 287 bp
21 AGGTCGTTCCATGCCTTACA TGGGAACCAGTTTCCCTGAG 224 bp
22 CCATTCTCTTCTTCCTCCGA CTCCAGGGACCCTTCCATGA 278 bp
23 GGGAAGACTGAGCCTAGAGA CTGGGCGGTCAGACCCCAGA 245 bp
24 CTTTGATCAGGTCTTGGTTA GAGGGGGATTAGAGATGGAG 285 bp
25 CCCGCCTCCATCTCTAATCC GGCAGTGGTGGCGACAGCAG 285 bp
26 CTGCCTGCTGTCGCCACCAC GGAGAACCTAACAAGGATTT 288 bp
27 GAAGGTGGAGGCACTGTTCA GTGGTGGCAAGGGAGCCACA 250 bp
28 CGTTCAGAAATGGAACACTC CTGGGAACACAGACCGAGTT 196 bp
29 CCTCCCTGAACTCGGTCTGT GTGGAGGGCGGAGGTGTCCT 272 bp
30 TTCTGACCAGCGGAGTCTAA CCACCGCACCTGGCCAAGAG 245 bp
31 GCCGTCATGTGCCTCATCTC ATATCCCTGCTGGGTTCTGG 244 bp
32 ATCAGGGCCTCTTCCCGATG ATCCTAAAGTGCCCTCACCA 190 bp
33 CACCAACCTGAAATCTCTCC CGTGTGACCTGAGGCTGTGG 202 bp
36 GAAACTGAGAGGGGCCGGAC TGCCCTGTGGATCTGCAAGC 246 bp
All primers are shown 5' to 3'. The size of the amplified product is listed on the right. Exon 18 has two sets of primers. Because of repetitive sequences near the splice donor site following exon 18, two 3' or reverse primers were needed to amplify exon 18. The product from the first set of primers (18 forward and 18 reverse) is used as template in a second PCR reaction using 18 forward and 18B reverse.

The molecular consequences of the ELN point mutations described here are not yet known. It is of note that no missense mutations were identified in this study. The mutations defined here may lead to the synthesis of aberrant transcripts or may result in functional hemizygosity. Hemizygosity of ELN is established as the mechanism of SVAS in Williams syndrome (11 ). If mutant transcripts are stable, some would result in the production of elastin protein lacking the carboxy-terminus (Fig. 6 ). The carboxyl end of elastin contains two cysteine residues that are thought to be critical for interaction with microfibrillar-associated glycoprotein during elastogenesis (14 ,15 ). The loss of the carboxyl end of elastin protein could result in aberrant elastic fiber formation during vasculogenesis (16 ).

We identified three splice site mutations in individuals with SVAS. These mutations affected exons 3 and 16. Previous studies have demonstrated that mutations in the consensus sequence of either the splice acceptor site preceding an exon, or a splice donor site following an exon, can result in removal of that exon from the mature transcript (17 ,18 ). If no cryptic splice sites are present, the splice mutations of exons 3 and 16 would delete these exons (Fig. 6 ). Because of ELN's genomic organization, deletion of an exon results in the loss of the amino acids encoded by the exon, not in a frameshift. Exon 3 encodes a hydrophobic domain that may be important for the tertiary structure of elastin. Exon 16 encodes a relatively large hydrophobic domain that separates two alanine-rich cross-linking sites near the center of the molecule. Deletion of exon 16 would result in the juxtaposition of two cross-linking domains and may have a deleterious effect on the spatial symmetry required to form a functional interaction between cross-linking sites. In addition, exon 16 contains a hexapeptide sequence (PGAIPG) that interacts with elastin-binding proteins on the surface of cells (19 ,20 ). Thus, deletion of exon 16 could disrupt both the structural integrity and the cellular interaction of elastin.

We performed mutational analysis in 20 unrelated cases of SVAS and discovered mutations in seven. This relative insensitivity could be due to the technique (SSCP), but the gels were run under two conditions to increase sensitivity. In addition, we did not examine the 5'- or 3'-untranslated region of ELN. Finally, it is possible that SVAS is genetically heterogeneous. These issues must be addressed if genetic testing for SVAS is to become routine.


Figure 4. Splice site mutations affecting exon 16 of ELN in familial SVAS.Pedigree structures for (A) K2260, (B) K2205 and (C) K2044 are shown. SSCP analyses using primers for exons 16 demonstrate anomalous conformers that co-segregate with the disease in each kindred. DNA sequence analyses of the normal and abnormal conformers for K2260 and K2205 reveals an identical A to G transition within the consensus splice acceptor site preceding exon 16, IVS15-2A -> G. Sequence analysis of K2044 identifies a C to G transversion within the consensus splice acceptor site of exon 16, IVS15-3C -> G. Exons are denoted by uppercase letters and introns are denoted by lowercase letters. The nucleotides involved are highlighted by arrows.


Figure 5. De novo ELN point mutation in a sporadic case of SVAS. Results of SSCP analysis for K2017 using primers for exon 3 are shown below the pedigree. An aberrant SSCP conformer is amplified from DNA of an affected individual but not from DNA of his unaffected parents. DNA sequence analysis of the conformer revealed a G to A transition in the consensus splice donor site following exon 3, IVS3+1G -> A.


Figure 6. Schematic representation of elastin protein, showing the location of mutations. The elastin protein is divided into 34 domains, each encoded by a discrete exon. The first exon encodes the signal peptide and is indicated by the cross-hatched bar. Domains that participate in intermolecular cross-linking are denoted by gray bars. Hydrophobic domains are indicated by white bars. The domain encoded by exon 36 is a conserved, hydrophilic carboxy-terminus important for binding of elastin to the microfibrillar scaffold during elastogenesis and is indicated by hatched bars. Domains thought to be important for cell binding are denoted with asterisks. Positions and types of mutations are indicated.

In this study, we found that intrafamilial variability of phenotypic expression in SVAS is high, a finding consistent with previous studies (9 ,10 ,21 ,22 ). In one study, Olson and co-workers described a family with an intragenic deletion of ELN. In a second report, Curran et al. characterized an SVAS family harboring a balanced translocation that disrupted the 3' end of ELN. Phenotypes in both families ranged from severe generalized obstructive disease requiring surgery at an early age to modest asymptomatic focal vascular abnormalities documented late in life through clinical testing. This phenotypic variability will limit the prognostic utility of genetic testing.

Epidemiological and biochemical studies have led to the hypothesis that the pathogenesis of obstructive vascular disease is complex, resulting from the interplay of multiple heritable and environmental factors. Several major risk factors for vascular disease have been identified, including hypercholesterolemia, hypertension and diabetes, but these factors are thought to account for only 50% of heritable risk (23 -25 ). The abundance of elastin in the vasculature, the importance of elasticity to the modulation of hemodynamic stress, and our demonstration that ELN mutations cause SVAS suggest that elastin abnormalities could contribute to common vascular disease. The synthesis of oligonucleotide primers useful for mutational analyses will enable future studies on the role of ELN as a genetic risk factor for common vascular disease.

MATERIALS AND METHODS

Phenotypic evaluation

Informed consent was obtained from all study participants in accordance with standards established by local institutional review boards. To determine if family members and spouses had SVAS or Williams syndrome, physical examinations and echocardiograms were performed by a medical geneticist and a cardiologist, as previously described (26 ). In this study, individuals with an unknown phenotype were given that designation due to our inability to obtain an echocardiogram.

DNA sequence and exon-intron determination

Genomic clones were isolated from phage and cosmid libraries by probing with ELN cDNA. Four clones contained elastin exons ([lambda]-4, [lambda]-5, cELN-11d and cELN-6). These clones were mapped by restriction analyses and spanned exons 1-36 of ELN (Fig. 1 ).

Phage clones [lambda]-4 and [lambda]-5 and cosmid clones cELN-6 and cELN-11d were subcloned into pBluescript II SK+ from Stratagene (La Jolla). Subclones were sequenced in both orientations with dye-labeled primers (M13-21 and M13RP) on an ABI 373 automated sequencer. Sequenced fragments were assembled using the genome assembly program (27 ,28 ). Continuous coverage of this region was obtained by generation of nested deletions using Deletion Factory System Version 2.0 from Gibco-BRL (Gaithersberg, MD).

The complete sequence spanning exon 5 to 10 kb 3' of exon 36, and 70% of the region between exons 2 and 5 were sequenced. The intron-exon boundaries for exons 2-36 were defined. The intron-exon boundary for exon 1 was described previously (12 ). Restriction maps predicted by the sequence data were identical to maps generated with EcoRI, HindIII and BamHI. These data were submitted to GenBank (Accession nos U93034, U93035, U93036 and U93037).

SSCP analysis

Oligonucleotide primers amplifying each exon of ELN were designed and used for mutational analyses. Genomic DNA samples were amplified by PCR and used in SSCP analyses as described (29 ,30 ). The annealing temperature was 58oC for all PCR reactions. Reactions (10 [mu]l) were diluted with 40 [mu]l of 0.1% SDS/1 mM EDTA and 30 [mu]l of 95% formamide dye. Diluted products were denatured by heating at 100oC for 5 min, and 3 [mu]l of each sample was electrophoresed on 7.5% non-denaturing acrylamide gels (49:1 polyacrylamide:bis-acrylamide) at 4oC. Electrophoresis was carried out at 40 W for 2-5 h. Gels were dried and exposed to X-ray film at -80oC for 12-36 h.

Normal and aberrant SSCP conformers were cut directly from dried gels and eluted in 75 [mu]l of distilled water at 37oC for 30 min. Ten [mu]l of the eluted DNA was used as template for a second PCR reaction using the original primer pair. Products were fractionated in 2% low melting temperature agarose gels, and bands cloned directly into pBluescript II SK+ using the T-vector method as described (31 ). Plasmid DNA samples were purified and sequenced.

ACKNOWLEDGEMENTS

We are indebted to the family members for their participation. We are grateful for the advice of Drs R. Mecham and K. Ward. We thank Drs L. Urness and S. Odelberg for editorial comments. This work was supported by National Heart, Lung, and Blood Institute grants RO1HL4807 and 1K08HL3490-01, National Institute of Neurological Disorders and Stroke grant NS35102, a grant from the American Heart Association, a grant from the March of Dimes Foundation and an award from Bristol-Myers Squibb.

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16 Hinek, A. and Rabinovitch, M. (1993) The ductus arteriosus migratory smooth muscle phenotype processes tropoelastin to a 52-kDa product associated with impaired assembly of elastic laminae. J. Biol. Chem., 268, 1405-1413. MEDLINE Abstract

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18 Bonne, G., Carrier, L., Bercovici, J., Cruaud, C., Richard, P., Hainque, B., Gautel, M., Labeit, S., James, M., Beckmann, J., Weissenbach, J., Vosberg, H.-P., Fiszman, M., Komajda, M. and Schwartz, K. (1995) Cardiac myosin binding protein-C gene splice acceptor site mutation is associated with familial hypertrophic cardiomyopathy. Nature Genet., 11, 438-440. MEDLINE Abstract

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20 Grosso, L.E. and Scott, M. (1993) PGAIPG, a repeated hexapeptide of bovine tropoelastin, is a ligand for the 67-kDa bovine elastin receptor. Matrix, 13, 157-164. MEDLINE Abstract

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*To whom correspondence should be addressed at the Eccles Institute of Human Genetics. Tel: +1 801 581 8904; Fax: +1 801 585 7423; Email: mark.keating{at}genetics.utah.edu

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