Abstract

Hirschsprung disease (HD) has been described in association with microcephaly, mental retardation and characteristic facial features, delineating a syndrome possibly caused by mutations localized at chromosome 2q22–q23. We have analyzed a de novo translocation breakpoint at 2q22 in one patient presenting with this syndrome, and identified a gene, SIP1, which is disrupted by this chromosomal rearrangement. SIP1 encodes Smad interacting protein 1, a new member of the δEF1/Zfh-1 family of two-handed zinc finger/homeodomain transcription factors. We determined the genomic structure and expression of the human SIP1 gene. Further analysis of four independent patients showed that SIP1 is altered by heterozygous frameshift mutations causing early truncation of the protein. SIP1, among other functions, seems to play crucial roles in normal embryonic development of neural structures and neural crest. Its deficiency, in altering function of the TGFβ/BMP/Smad-mediated signalling cascade, is consistent with some of the dysmorphic features observed in this syndrome, in particular the enteric nervous system defect that underlies HD.

Received April 11, 2001; Revised and Accepted May 18, 2001.

INTRODUCTION

Hirschsprung disease (HD), or aganglionic megacolon, is a congenital disorder characterized by intestinal obstruction due to an absence of enteric neurones along a variable length of the intestine. This relatively common disease affects approximately 1 in 5000 infants, with a 4:1 male predominance. HD results from the failure of neural crest cells to develop normally. The genetic aetiology of this neurocristopathy is complex. Simple Mendelian inheritance is rarely observed, and the disease is presumed to be multifactorial, with contributions from several genes and possibly environmental factors. Epidemiological studies have shown differences in the recurrence risks and mode of inheritance, based on the length of the aganglionic bowel. Short-segment HD (involving the rectum and sigmoid only), which accounts for ∼80% of cases, is compatible with an autosomal recessive or multifactorial mode of inheritance. In contrast, segregation analysis of families with long-segment HD suggested a dominant mode of inheritance with incomplete penetrance (1).

HD is associated with a wide variety of other congenital disorders, some of which share the features of a neurocristopathy. It is a frequent finding in trisomy 21 (Down’s syndrome) and it has been associated with other chromosomal abnormalities, including partial deletion of chromosome 13q and 22q11.2 deletions. Other congenital anomalies of the central nervous system, the heart, and the gastrointestinal and genitourinary tracts have been reported with increased frequency in HD. Other syndromes with Mendelian patterns of inheritance in which HD occurs include Bardet–Biedl syndrome, cartilage hair hypoplasia, primary central hypoventilation syndrome (Ondine’s curse), familial dysautonomia, Goldberg–Shprintzen syndrome, neurofibromatosis I, MEN 2A and 2B, X-linked hydrocephalus, the Smith–Lemli–Opitz syndrome type II and Waardenburg syndrome (2).

Linkage studies of familial cases of HD, and the analysis of several mouse or rat models of HD, contributed to the identification of at least six genes mutated in patients with this disease. The RET receptor tyrosine kinase gene, which has been implicated in at least four different disorders, was the first to be identified as a major HD susceptibility gene (3,4). Heterozygous loss-of-function mutations of RET are responsible for approximately half of the familial and 15–20% of the sporadic HD cases following a reduced-penetrance dominant pattern of inheritance (5). Mutations in the RET ligands, glial cell line-derived neurotrophic factor (GDNF) and neurturin (NRTN), have been shown to contribute to the phenotype in a small minority of cases of HD, in combination with mutations in other genes (68). In mice, homozygous mutations in either the endothelin-B receptor gene (EdnrB) or endothelin-3 (Edn3), a ligand of EDNRB in neural crest-derived enteric neural precursors and cutaneous melanoblasts, lead to intestinal aganglionosis and piebaldism (9,10). In humans, a few cases of isolated HD are caused by mutations in EDNRB or in EDN3 genes (2,11). WS4 or Shah–Waardenburg syndrome, a syndrome that associates HD, pigmentation defects of skin and hair and sensorineural hearing loss, has been described in rare individuals with homozygous or heterozygous mutations in either EDNRB or EDN3 (2,12). Dominant mutations of SOX10, a SRY-related HMG box-containing transcriptional regulator that is mutated in Dominant megacolon (Dom), a spontaneous dominant mouse model of HD, have also been shown to cause WS4 (13). Finally, syndromic HD has also been associated with a mutation in the endothelin converting enzyme-1 gene (ECE-1), a gene involved in EDN3 biosynthesis (14).

Despite significant advances in understanding the genetic background in HD, the molecular basis of some syndromic HD remains elusive. Here, we have investigated patients in whom HD is associated with mental retardation (MR), microcephaly, a distinctive facial phenotype and other anomalies. We have previously reported patients having this form of syndromic HD, one presenting with a de novo chromosome del(2)(q22–q23) interstitial deletion, the other with a de novo apparently balanced translocation t(2;11)(q22.2;q21), suggesting the existence of a HD gene at a chromosome 2q22 locus (15,16). Taking advantage of the t(2;11) rearrangement, we found that SIP1, a gene that encodes Smad interacting protein 1, a member of the transforming growth factor β (TGF-β) signalling pathway, is disrupted by the breakpoint. Furthermore, we showed that mutations in SIP1 cause the condition in four other cases, defining a new HD gene. These results provide clues as to the complex aetiology of HD and open new avenues of investigation into the mechanisms behind the various phenotypic features of this neurocristopathy.

RESULTS

Determination of the candidate interval

To fine-map the chromosome 2 breakpoint, we performed fluorescence in situ hybridization (FISH) analysis on metaphase chromosomes from patient 1, using Alu-PCR products from a series of yeast artificial chromosome (YAC) clones as probes. This led us to identify a YAC clone (954e8) which covers the breakpoint. Using STS markers mapping, we assembled a set of different bacterial artificial chromosome (BAC) clones encompassing the region (Fig. 1A). One of the BAC clones (RP11-107E5) was shown to span the translocation breakpoint.

Isolation and structure determination of SIP1

Computational analysis of the candidate region revealed that a gene product, KIAA0569, was contained within two overlapping BAC clones, one of which is disrupted by the translocation breakpoint. Alignment of the KIAA0569 sequence against that of the two BAC clones (RP11-95O9 and RP11-107E5) allowed us to determine the genomic structure of a gene presenting sequence identity with a recently described new gene in the mouse, sip1 (GenBank accession no. Mus musculus sip1 AF033116) (17). Human SIP1 is composed of 10 exons that span ≥120 kb (GenBank accession no. SIP1 AY029472) (Fig. 1B). The initiation codon is located in exon 2 and the gene encodes a putative protein of 1215 amino acids. Comparison of the predicted amino acid sequence of the human SIP1 with mouse or Xenopus (GenBank accession no. XSIP1 AF237679) orthologues showed 96.6 and 85.5% identity, respectively. The Smad-binding domain (SBD) was assigned to amino acids 437–488 by sequence homology. Furthermore, strong similarity conservation exists among different functional domains (homeodomain, zinc finger domains) between SIP1 and δ-crystallin enhancer binding factor (δEF1) (Fig. 2).

Implication of SIP1 gene in the t(2;11) translocation

To assess whether SIP1 is disrupted by the chromosome 2 breakpoint, we performed FISH analysis using a pool of long-range PCR products covering the whole genomic sequence. Metaphase spreads from patient 1 showed FISH signals on both derivative chromosomes (Fig. 3). To precisely map the chromosome 2 breakpoint within the gene, we used several individual long-range PCR products covering intron 2. One of these PCR products showed a hybridization signal on both derivative chromosomes, demonstrating that the translocation breakpoint mapped within the second intron of the SIP1 gene (data not shown).

SIP1 mutation analysis

To further demonstrate the implication of SIP1 in patients with syndromic HD, we searched for mutations in four independent patients with a similar phenotype. Direct sequencing of PCR products allowed identification of four different mutations in the heterozygous state (Table 1). In all four cases, SIP1 is altered by frameshift mutations (nucleotide deletion or insertion) causing early truncation of the protein, deleting both the SBD and the homeodomain. Three of these mutations are located in exon 8 and one in exon 6. Analysis of DNA from both parents of patients 2 and 3, available for study, showed the absence of SIP1 mutations, confirming the de novo occurrence of the defect in their children.

Expression pattern of SIP1

Hybridizations of multiple-tissue northern blots with the partial SIP1 probe A detected a specific transcript of ∼5.5 kb, a size compatible with the expected length of the mRNA (Fig. 4A). A similar result was obtained with probe B (data not shown).

SIP1 mRNA was detected with high levels in almost all adult human tissues tested (heart, brain, placenta, lung, liver, squeletal muscle), as well as in mouse heart and brain. Sip1 expression in the mouse seems to begin in early stages of development, since transcripts were already detected in embryo at 7 days. In some of the tissues tested, hybridization also detected a faint higher molecular weight signal suggestive of the existence of related transcripts, the nature of which has yet to be investigated.

Hybridization of a human multiple-tissue expression array with probe A detected SIP1 expression in different brain structures (whole brain, cerebral cortex, cerebellum, corpus callosum and thalamus) except in the pituitary gland. This tissue array also detected consequent expression in transverse and descending colon, in different areas of the heart (aorta, atrium, ventricles and inter-ventricular septum) and different regions of the digestive system (oesophagus, stomach, duodenum and jejunum). In addition, no signal was visible in prostate, testis, ovary, pancreas, thyroid, salivary and mammary glands. SIP1 expression was also detected in several fetal human tissues like brain, heart, kidney, liver or spleen (data not shown).

The detection of a strong sip1 signal in whole adult mouse brain prompted us to look in more detail at sip1 gene expression in various brain areas by RT–PCR. Sip1 was found to be expressed in each mouse brain region analysed (hypothalamus, cerebellum, cortex, rhombencephale, hippocampus, mid brain and spinal cord) (Fig. 4B).

DISCUSSION

We report here that heterozygous mutations of SIP1, a recently described member of the δEF1/Zfh-1 family of two-handed zinc finger/homeodomain proteins, cause a form of HD which manifests in association with MR, microcephaly and distinctive dysmorphic facial features. The fact that two of the patients previously reported with this syndrome had an interstitial deletion of chromosome 2 suggested that some isolated cases could have a contiguous gene syndrome or a dominant single gene disorder involving a locus for HD at 2q22–q23 (15,18). The observation in one of our patients of a de novo translocation at 2q22 prompted us to analyse the breakpoint region. This led us to identify SIP1 as the gene disrupted in intron 2 by this chromosomal rearrangement. In our four other patients, all isolated cases, we identified a point mutation in SIP1. All mutations found are frameshift defects that result in early truncation of the encoded protein, removing most of the functional domains (SBD, homeodomain and C-terminal zinc finger). Our observation closely follows the report of Wakamatsu et al. (19), who independently identified loss-of-function mutations of the same gene in patients with identical phenotypic features. It is striking to see that most of the SIP1 mutations described to date (6,7) in this syndrome lie in exon 8 (19 and this work). Further studies of other cases may indicate whether this gene region is a hotspot for mutations.

In all these patients with a severe phenotype, the presence of heterozygous frameshift mutationsis likely to result from haploinsufficiency caused by null mutation of one allele, rather than from a dominant-negative effect of the mutant product. This is supported by the finding of the same phenotype in the patients with cytogenetic 2q22 deletions, including SIP1.

Nevertheless, the resulting change in gene dosage may not always be sufficient to cause malfunction. Indeed, it is possible that some SIP1 mutations could result in a milder or incomplete phenotype. For instance, SIP1 mutations could produce the same distinctive dysmorphism and MR without HD. Some types of defects, such as missense mutations, might be compatible with the maintenance of a clinically normal phenotype in the heterozygote state. This question is of importance as there is clearly genetic heterogeneity within the group of patients presenting with HD, MR and microcephaly. It will be particularly interesting to see whether patients with Goldberg–Shprintzen syndrome, a similar clinical entity in which a recessive mode of inheritance seems likely (2022), have a separate disorder or also have SIP1 missense mutations.

Like several other transcription modulators, SIP1 is likely to have multiple essential functions during development, at different stages and in various locations and tissues. It was recently reported that XSIP1, a Xenopus homologue of SIP1, is activated very early in embryogenesis and expressed throughout development and in the adult. Interestingly, XSIP1 is strongly expressed early in prospective neuroectoderm, then in the neural plate, and later in the neural tube and neural crest (23,24).

SIP1, a transcriptional repressor which was cloned as a Smad interacting protein 1, also interacts with Smad2, Smad3, and Smad5 through an SBD (17). Smad proteins are signal transducers involved in the TGF-β family signalling cascade. TGF-β family members have critical roles during embryogenesis and in maintaining tissue homeostasis during adult life (2528). Upon binding to the serine/threonine kinase receptor complex, each member of this family of secreted polypeptide growth factors [i.e. TGF-β, activins and bone morphogenetic proteins (BMPs)] activates a subset of Smad proteins. Activated Smads form hetero-oligomeric complexes with common partner Smads that translocate to the nucleus, where they control the expression of target genes in a cell-specific manner.

The identification of SIP1 as a gene involved in syndromic HD opens new avenues of investigation into the mechanisms behind the various phenotypic features, and could help understand the underlying developmental processes. One possible clue to understand how SIP1 deficiency results in a developmental defect of the enteric nervous system (ENS) could lie in its possible involvement in the RET activation pathway. Indeed, this receptor tyrosine kinase and its functional ligands, GDNF and NRTN, have been identified to control, either directly or indirectly, morphogenesis and differentiation of the ENS (reviewed in ref. 29).

Among the TGF-β family members that are involved in nervous system patterning, BMPs play an important role. BMP2/4 were demonstrated to induce in vitro determination of neural crest cells into catecholaminergic derivatives that can generate either sympathic, adrenal or enteric neurones (30,31). In this process, MASH-1 and Phox2 genes are important players, since in sympathetic neurones they are downstream effectors of BMPs. Furthermore, regulatory relationships between MASH-1, Phox2a and RET have been demonstrated in sympathetic neurones, in which MASH-1 activates RET indirectly via induction of Phox2a. In addition, Phox2b is likely to activate expression of RET as well (32,33). It is not known whether BMPs promote neurogenesis in the ENS as it does in sympathetic neurones. Nevertheless, the observation that SIP1 deficiency results in a defect in ENS development invites investigation of a possible functional link between SIP1 and RET. Furthermore, it should prompt testing of the involvement of SIP1 in other neurocristopathies.

MATERIALS AND METHODS

Patients

Lymphoblastoid cell lines from patients 1 and 2 were obtained from the Department of Medical Genetics, the Family Federation of Finland (Helsinki, Finland), and clinically described as case report nos 1 and 3, respectively, by Kääriäinen et al. (16). Patient 1 carried a t(2;11)(q22.2;q21) de novo reciprocal translocation.

DNA from patients 3, 4 and 5 was obtained from the Department of Clinical Genetics, The Children’s Hospital Westmead (Sydney, Australia), and clinically described as case report nos 1, 2 and 3, respectively, by Mowat et al. (15).

All five patients have HD, microcephaly, strikingly similar dysmorphic facial features and severe MR.

FISH probes

YAC clones were obtained from the CEPH library and were Alu-PCR amplified using standard procedures (34). BAC clones were provided by the CHORI BAC-PAC Resources (Oakland, CA). DNA was extracted from YAC and BAC clones according to standard techniques.

Long-range PCR probes were obtained from human normal genomic DNA using the Expand Long Template PCR System kit (Roche Diagnostics, Mannheim, Germany) with various sets of primers designed based on the human SIP1 gene sequences. PCR products were gel-purified using a gel extraction kit (Macherey-Nalgen, Düren, Germany) prior to labelling.

Physical mapping of the 2q22 region

CEPH YAC clones belonging to the 2q22 region were selected according to the Unified Database for Human Genome Mapping information (http://bioinformatics.weizmann.ac.il/udb) and GDB database (http://www.gdb.org) about their location. The YAC contig was assembled by STS content, using several markers (sequences available at the GDB database). BAC clones were determined by their STS content by using BlastN search from the GenBank/htgs database (http://www.ncbi.nlm.nih.gov).

Determination of the genomic structure of SIP1

The genomic structure of SIP1 was determined by direct comparison of its full-length cDNA (KIAAO569, GenBank accession no. AB011141) with the sequence of two human chromosome 2 BAC clones, RP11-107E5 and RP11-95O9 (GenBank accession nos AC009951 and AC010130) retrieved by BlastN search.

FISH analysis

FISH analysis was carried out essentially as described elsewhere (35,36). Metaphase chromosomes of the cell line with the translocation were prepared by standard techniques. All probes were labelled by nick-translation with Bio11-dUTP (Sigma-Aldrich, St Louis, MO). Twenty nanograms of labelled BAC clones or long-range PCR fragments and 200 ng of labelled Alu-PCR amplified YAC clones were used. The probes were blocked with Cot-1 DNA (Life Technologies, Gaithersburg, MD). Fluorescent signals were enhanced by avidin fluorescein-isothiocyanate. Chromosomes were counterstained with DAPI and the signal was analysed using a Leica fluorescence microscope equipped with the Cytogen Fluoquant program (Imstar, Paris, France).

Mutation analysis

In each individual, the nine coding exons and flanking sequences were amplified from genomic DNA using specific primers and 0.5 U Taq polymerase (Life Technologies). The primers and cycling conditions used were as follows:

Exon 2 (253 bp fragment), 2A (5′-TTTCAATGGGCGCGCGATGC-3′) and 2B (5′-TCCTTCTCCCTGGGTCTCGA-3′), annealing temperature 65°C, 2.5 mM MgCl2.

Exon 3 (601 bp fragment), 3A (5′-ACCTGAAGGATATTAATTATATTTTCCTGA-3′) and 3B (5′-GATGTAACTGCCGCAATGTGATA-3′), annealing temperature 55°C, 2.5 mM MgCl2.

Exon 4 (230 bp fragment), 4A (5′-CATGCTTAGTATAGTAAGCCT-3′) and 4B (5′-TGGAGATACAAATGGATGTGTC-3′), annealing temperature 55°C, 1.5 mM MgCl2.

For exons 5, 6, 7, 8, 9 and 10, we performed long-range PCR reactions (according to the manufacturer’s instructions) to amplify two different products: the first for exons 5–7 (8705 bp fragment) with primers 5A (5′-TTTCTTGGCTCACGGAATTTAAG-3′) and 7B (5′-ATGTTCACAGTGGAAGGTAAGGGCTATCTA-3′); and the second for exons 8–10 (12536 bp fragment) with primers 8A (5′-TAGATAGCCCTTACCTTCCACTGTGAACAT-3′) and 10B (5′-AGTTTGGCTACATTTTTATTCGAGCATGGTC-3′).

All PCR products were gel-purified using a gel extraction kit (Macherey-Nalgen) and sequenced with the ABI PRISM BigDye Terminator cycle sequencing kit (Perkin Elmer, Foster City, CA) using an Applied Biosystems model 373A.

Northern blot analysis

Two different RT–PCR fragments were used as α[32P]dCTP-radiolabelled probes for northern blot analysis: probe A, a 975 bp fragment covering the N-terminal region of the cDNA (forward primer, 5′-AAGATGAAATAAGGGAGGGTGGA-3′; reverse primer, 5′-CCGTGTGTAGCCATAAGAAC-3′; annealing temperature 58°C, 1.5 mM MgCl2), and probe B, a 1886 bp fragment corresponding to the 3′ end of SIP1 cDNA (forward primer, 5′-CAACGAAAAGTCTACCAGTA-3′; reverse primer, 5′-GTGATACATAAGTAGAGTGC-3′, annealing temperature 58°C, 1.5 mM MgCl2).

Three human or mouse multiple-tissue northern blot membranes (Clontech, Palo Alto, CA) were hybridized with these two SIP1 radiolabelled probes, following the manufacturer’s instructions. A human multiple-tissue expression array (Clontech) containing polyA+ RNA from 76 different tissues was also hybridized with probe A.

RT–PCR

Approximately 1 µg of total RNA was reverse-transcribed using the Superscript II kit (Life Technologies). This reaction mix (1 µl) was subjected to amplification with SIP1-specific primers (forward primer, 5′-GTCCATGCGAACTGCCATCTGATCCGCTCT-3′; reverse primer, 5′-GGCTTGCAGAATCTCGCCAC-3′; annealing temperature 58°C, 1.5 mM MgCl2) and β-actin.

ACKNOWLEDGEMENTS

We thank S. Amselem for helpful comments on the manuscript and M.F. Belin for the gift of mouse brain mRNAs. This work was supported by grants from INSERM.

+

To whom correspondence should be addressed. Tel: +33 1 49 81 28 61; Fax: +33 1 48 99 33 45; Email: michel.goossens@im3.inserm.frThe authors wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors

Figure 1. Physical map of the chromosome 2 region involving the t(2;11) translocation breakpoint. (A) The relative position of 15 STS in the region, BAC and YAC clones located between the markers D2S1290 and D2S1738 are indicated. (B) Genomic organization of human SIP1 gene: coding exons are shown as closed black boxes and non-coding regions are shown as open boxes. Sizes of exons and introns are indicated in the table.

Figure 1. Physical map of the chromosome 2 region involving the t(2;11) translocation breakpoint. (A) The relative position of 15 STS in the region, BAC and YAC clones located between the markers D2S1290 and D2S1738 are indicated. (B) Genomic organization of human SIP1 gene: coding exons are shown as closed black boxes and non-coding regions are shown as open boxes. Sizes of exons and introns are indicated in the table.

Figure 2. Predicted amino acid sequence of SIP1 and protein homology. Amino acids identical in at least two species are shown with black background, whereas similar residues are shaded. Gaps that were introduced to optimize the alignment are indicated by dashes. Open bars indicate the eight zinc fingers and the solid and shaded bars show the homeodomain and the SBD, respectively.

Figure 2. Predicted amino acid sequence of SIP1 and protein homology. Amino acids identical in at least two species are shown with black background, whereas similar residues are shaded. Gaps that were introduced to optimize the alignment are indicated by dashes. Open bars indicate the eight zinc fingers and the solid and shaded bars show the homeodomain and the SBD, respectively.

Figure 3. Metaphase spread from patient 1 carrying the t(2;11)(q22.2;q21) hybridized with a pool of PCR products covering the entire SIP1 gene. Breakpoint spanning the SIP1 genomic probe in green gives signals on the normal chromosome 2 and both derivative chromosomes 2 and 11.

Figure 3. Metaphase spread from patient 1 carrying the t(2;11)(q22.2;q21) hybridized with a pool of PCR products covering the entire SIP1 gene. Breakpoint spanning the SIP1 genomic probe in green gives signals on the normal chromosome 2 and both derivative chromosomes 2 and 11.

Figure 4. Tissue distribution of SIP1 mRNA. (A) Northern blot analysis performed with human and mouse multiple-tissue membranes hybridized with probe A (a 975 bp fragment covering the N-terminal region of the SIP1 cDNA). A major transcript of 5.5 kb was detected both in adult human and mouse tissues and in mouse embryo. (B) RT–PCR analysis of SIP1 expression in different structures of mouse brain (top). β-actin RT–PCR was used as an internal control for the amount of RNA in each sample (bottom).

Figure 4. Tissue distribution of SIP1 mRNA. (A) Northern blot analysis performed with human and mouse multiple-tissue membranes hybridized with probe A (a 975 bp fragment covering the N-terminal region of the SIP1 cDNA). A major transcript of 5.5 kb was detected both in adult human and mouse tissues and in mouse embryo. (B) RT–PCR analysis of SIP1 expression in different structures of mouse brain (top). β-actin RT–PCR was used as an internal control for the amount of RNA in each sample (bottom).

Table 1.

SIP1 mutations detected

Patients Exon Mutations Effect on protein sequence Functional domain    
    NZFa SBDb HDc CZFa 
1216 delAC Frameshift→stop at 409 – – – 
920 delA Frameshift→stop at 332 Partial – – – 
594 delC  Frameshift→stop at 211 – – – – 
1421 insA  Frameshift→stop at 481 – – – 
Patients Exon Mutations Effect on protein sequence Functional domain    
    NZFa SBDb HDc CZFa 
1216 delAC Frameshift→stop at 409 – – – 
920 delA Frameshift→stop at 332 Partial – – – 
594 delC  Frameshift→stop at 211 – – – – 
1421 insA  Frameshift→stop at 481 – – – 

aNZF and CZF, N- and C-terminal clusters of zinc fingers, respectively.

bSBD, Smad-binding domain.

cHD, homeodomain.

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