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Human Molecular Genetics Pages 625-637  

Pleiotropic skeletal and ocular phenotypes of the mouse mutation congenital hydrocephalus (ch/Mf1) arise from a winged helix/forkhead transcription factor gene
Introduction
Results
   Evaluation of the ch/ch hydrocephalic phenotype
   Identification of markers closely linked to ch in the CHMU/Le inbred strain
   Mapping ch in interspecific crosses
   Physical map of the ch-Mf1 interval
   Evaluation of the ch/+ ocular phenotype
   Mapping of mouse Hfh1
Discussion
Materials And Methods
   Animals
   Skeletal and slit lamp analysis
   Genotyping
   Construction of YAC and BAC contigs
   IRS-PCR probe generation
   PCR analysis of Mf1 and Hfh1
   Radiation hybrid data analysis
Acknowledgements
References

Pleiotropic skeletal and ocular phenotypes of the mouse mutation congenital hydrocephalus (ch/Mf1) arise from a winged helix/forkhead transcription factor gene

Pleiotropic skeletal and ocular phenotypes of the mouse mutation congenital hydrocephalus (ch/Mf1) arise from a winged helix/forkhead transcription factor gene

Hee-Kyung Hong, Jonathan H. Lass1 and Aravinda Chakravarti*

Department of Genetics BRB 721 and Center for Human Genetics and 1Department of Ophthalmology,Case Western Reserve University School of Medicine and University Hospitals of Cleveland, 10900 Euclid Avenue, Cleveland, OH 44106-4955, USA

Received November 11, 1998; Revised and Accepted January 20, 1999

Congenital hydrocephalus is an etiologically diverse, poorly understood, but relatively common birth defect. Most human cases are sporadic with familial forms showing considerable phenotypic and etiologic hetero-geneity. We have studied the autosomal recessive mouse mutation congenital hydrocephalus (ch) to identify candidate human hydrocephalus genes and their modifiers. ch mice have a congenital, lethal hydrocephalus in association with multiple developmental defects, notably skeletal defects, in tissues derived from the cephalic neural crest. We utilized positional cloning methods to map ch in the vicinity of D13Mit294 and confirm that the ch phenotype is caused by homozygosity for a nonsense mutation in a gene encoding a winged helix/forkhead transcription factor (Mf1). Based on linked genetic markers, we performed detailed phenotypic characterization of mutant homozygotes and heterozygotes to demonstrate the pleiotropic effects of the mutant gene. Surprisingly, ch heterozygotes have the glaucoma-related distinct phenotype of multiple anterior segment defects resembling Axenfeld-Rieger anomaly. We also localized a second member of this gene family (Hfh1), a candidate for other developmental defects, ~470 kb proximal to Mf1.

INTRODUCTION

Congenital hydrocephalus, a common human birth defect with an estimated incidence of 2/1000 livebirths (1), is morphologically and clinically a heterogeneous disorder. The phenotype is defined as the abnormal accumulation of cerebrospinal fluid (CSF) in the cranial cavity resulting in an enlargement of the ventricular system; the accumulation arises from obstruction in CSF circulation via stenosis or occlusion of the pathways between ventricles, interference with CSF absorption or oversecretion. Genetically, the origins of hydrocephalus are diverse and both genetic and environmental forms are known. One rare familial form, the X-linked hydrocephalus due to congenital stenosis of the aqueduct of Sylvius (MIM 307000), is caused by mutations in the L1 cell adhesion molecule gene (2). Family studies imply the existence of autosomal genetic factors, some being compatible with Mendelian recessive or dominant inheritance; however, the majority of cases are probably multifactorial (3-8). The considerable heterogeneity makes genetic mapping and positional cloning of the relevant autosomal human genes difficult and, thus, progress in understanding its developmental and biological basis is a formidable task. A number of excellent Mendelian mouse mutations for hydrocephalus offer an alternative route to identifying the relevant genes and pathways.

In the mouse, several autosomal recessive hydrocephalus mutations are known and their phenotypes are quite distinct from one another (9). One such mutation is congenital hydrocephalus (ch), originally identified by Grüneberg (10,11; see also refs 12-17). The mutation resides in the proximal region of chromosome 13 (12,18). Mice of the ch strain have a congenital, lethal hydrocephalus in association with multiple developmental defects affecting the central nervous, urogenital and skeletal systems (10-17). ch homozygotes die at birth or shortly thereafter, as a result of asphyxia related to a defective tracheal cartilage. All hydrocephalic animals show a steeply bulging forehead, corres-ponding to the cerebral hemispheres, with grossly hemorrhagic CSF in the massively dilated ventricular system. Interestingly, these mutants show multiple skeletal abnormalities, the most distinctive of which are the absence of skull bones, lack of ossification and multiple malformations of the thorax.

Grüneberg’s (10,11) pioneering histological and developmental studies of ch suggested that the morphological defects arose from a defect in mesenchymal differentiation required for skeletal organo-genesis, specifically at the preskeletal condensation stage. Although much is known about the structural components of cartilage and bone (19), little is known about the molecular mechanisms and genes that regulate chondrogenesis and osteogenesis during development. Thus, identification of the ch gene can illuminate not only normal cranial development but also the developmental processes of cartilage and bone formation. To identify the ch gene by positional cloning we first defined the ch phenotype extensively, in particular bone abnormalities, and showed phenotypic differences between subspecies. We next refined the target ch locus to an ~360 kb interval. This proximal region of mouse chromosome 13 harbors three known winged helix/forkhead domain transcription factors (20,21), namely Hfh1 (HNF-3 forkhead homolog 1), Hfh10 (HNF-3 forkhead homolog 10) and Mf1 (mesoderm/mesenchyme forkhead 1), of which two (Hfh1 and Mf1) are candidate genes for ch. While this study was in progress, Kume et al. (22) described the targeted disruption of the mouse Mf1 gene and subsequent breeding studies, with a mutant phenotype identical to ch. These authors identified a nonsense mutation in the Mf1 forkhead domain as the ch mutation in the CHMU/Le strain (22). We show, by independent genetic studies, that Hfh1 is very closely linked to Mf1 but recombines with ch and that Mf1 is the gene for ch, correcting its previous more distal placement by Labosky et al. (21). In addition, we conducted a detailed skeletal analysis of ch newborns in several different genetic backgrounds. Although similar severe skeletal abnormalities were noted, we show subtle modification of the mutant phenotype depending on the genetic background.

The proximal region of MMU 13 is syntenic to human chromosome 6p21-p25 (http://www.informatics.jax.org/ ), the predicted location for the human homolog of ch. Indeed, a forkhead transcription factor FKHL7 (also known as freac3) has been localized to 6p25 (23,24) and shows 90% amino acid sequence identity to ch/Mf1. This map location is consistent with the observed phenotypes of several human patients who harbor deletions of 6p22-p25 and have multiple developmental anomalies, including hydrocephalus, heart and kidney defects, craniofacial abnormalities such as clefts of the lip and/or palate and mental retardation (25,26). Nishimura et al. (27) have shown that the gene disrupted in two patients with rare chromosomal rearrangements involving 6p25 and with primary congenital glaucoma is FKHL7. In addition, mutations in FKHL7 have been identified in several patients with dominant disorders causing glaucoma, associated with abnormalities in the anterior segment of the eye, namely Rieger anomaly, Axenfeld anomaly and iridogoniodysgenesis anomaly (27). In all patients, except those with chromosome aberrations, only one copy of the mutation was associated with the disease phenotype and none of the human patients exhibited extra-ocular manifestations. However, eye abnormalities in ch heterozygous mice have not been described. We present evidence, based on detailed ocular examination of ch heterozygous mice, that haploinsufficiency of Mf1 leads to congenital developmental anomalies in the anterior segment of the eye and that these ocular changes are progressive, as seen in human patients. Thus, ch mice are an outstanding model for Axenfeld-Rieger anomaly.

RESULTS

Evaluation of the ch/ch hydrocephalic phenotype

The appearance of ch hydrocephalic mice was grossly similar in all three genetic backgrounds we examined, CHMU/Le, Mus mus castaneus (CAST) and Mus mus molossinus (MOLF) as first described by Grüneberg (10,11). All affected mice showed a steeply bulging forehead corresponding to the two cerebral hemispheres; Figure 1a-c shows the whole-animal phenotype corresponding to the three genotypes at ch. The twin bulging sacs, indicative of a massively dilated ventricular system, were also hemorrhagic and of a dark purplish-blue color. The mutants showed distinctive short snouts and open eyelids at birth. Most affected mice survived to term, made a few gasps to breathe at birth, but invariably died within a few minutes after parturition. The ch newborns were of normal size, but unlike their normal littermates, exhibit a bloated appearance with extensive edema, subsequently leading to an appearance of excess skin folds after death.


Figure 1. Newborn phenotypes of CHMU/Le inbred strain. (a) ch wild-type homozygote (mu+/mu+), (b) heterozygote (mu+/+ch) and (c) mutant homozygote (+ch/+ch) are shown from left to right. Skeletal defects observed on staining with alcian blue for cartilage and alizarin red for bone are shown for (d) heterozygote and (e) ch mutant newborn skeletons. The cranial vault is missing (arrow) in the mutant; note the lack of ossification of the thorax in the mutant.

Histological analysis of ch mutants, from embryonic stage 12 days post coitum (d.p.c.) to the newborn stage, was first described by Grüneberg (11) in the original genetic background. By gross examination, ch mutants are easily distinguishable from their unaffected littermates by frontal bossing of the head at embryonic day (E) 14.5. At E13.5, the enlargement of the cranium was perceptible but mutants were clearly not as discernible at E12.5. We re-evaluated the phenotype because the mutation has since been bred into several different genetic backgrounds. We conducted detailed studies of the skeletal abnormalities in the CHMU/Le, CAST and MOLF backgrounds. All mutants exhibited craniofacial defects, malformation of the cervical skeletal elements, abnormalities in the thorax and fusion and lack of ossification of the ribs. The most striking abnormality was the absence of bony calvarium in the hydrocephalics (Figs 1d-e and 2a-c); no traces of frontal, parietal and interparietal bones were observed except for a small portion at their lateral basal levels. The basioccipital, exoccipital and hyoid bones were smaller and supraoccipital bones were severely malformed. At the base of the skull, the basisphenoid bone was usually horseshoe-shaped with an anterior opening and presphenoid bones were absent (Fig. 2d and e). In contrast, the squamosal and zygomatic bones were abnormally shaped and massively ossified in the mutants compared with wild-type mice (Fig. 2a-c). In affecteds, the mandible exhibited more curvature and was also massively ossified in mutants compared with wild-type mice. Furthermore, the nasal septum was shorter and reduced giving the mutant the short-snout appearance. In the cervical region, the atlas and axis were grossly defective; however, no obvious malformation was observed in the thoracic or lumbar vertebrae and in limbs, except that the ossification centers of the limbs were smaller than in wild-type littermates. The rib cages of ch mutant mice were very fragile due to the lack of a sternum and the left and right costal cartilages were barely attached (Fig. 2f-h). Usually, there were no ossification centers in the sternebrae and in the upper region of the xiphoid process, whereas unaffected littermates had six calcification centers. The xiphoid process in mutants was fragmented and, in some instances, was barely present.


Figure 2. Skeletal defects of ch mutant mice. Skull and thoracic defects in CHMU/Le inbred strain. Lateral views in (a-c) are of wild-type (mu+/mu+), heterozygote (mu+/+ch) and homozygote mutant (+ch/+ch) newborn skulls, respectively. Dorsal views in (d) and (e) are of heterozygote and ch mutant skulls, respectively. The cranial vault of the heterozygote has been removed to view the skull base which is missing in the mutant. Ventral views in (f-h) are of wild-type, heterozygote and mutant newborn rib cages, respectively, showing the lack of ossification. Skeletal defects of ch mutant mice in the CAST and MOLF genetic backgrounds. (i) Lateral view of newborn mutant skulls from a mixed CAST and MOLF background. Note the small ossification centers on the dome-shaped membrane of the head; similar ossification centers were observed in each of the CAST and MOLF backgrounds. (j and k) Ventral view of mutant newborn rib cages in the CAST genetic background. Most mutants showed no ossification centers (j), but few do demonstrate some progression of ossification (k, arrow). at, atlas; ax, axis; bo, basioccipital bone; bs, basisphenoid bone; e, exoccipital bone; f, frontal bone; i, interparietal bone; ns, nasal septum; p, parietal bone; pa, palatine bone; ps, presphenoid bone; so, supraoccipital bone; x, xiphoid process; z, zygomatic bone.

Although similar severe skeletal abnormalities were noted in the CAST, MOLF and CAST-MOLF mixed genetic backgrounds, subtle yet consistent phenotypic differences were identified. Invariably, mutants propagated in the CAST and/or MOLF backgrounds showed further progression in ossification than seen in the CHMU/Le strain; small reddish spots of alizarin red were observed in the sternum and on the twin bulging membranes of the future calvarium in some mutants (Fig. 2i-k). Such variation was also observed for the ocular phenotype in mutant heterozygotes (see below). These small phenotypic differences are likely influenced by modifying genes in the subspecies but this hypothesis needs to be rigorously investigated by genetic mapping.

Mutant heterozygotes, which have a normal lifespan, have been reported to have a low frequency of abnormalities of the urogenital system and skeleton, but much less severe, similar to those found in homozygotes (12). We observed that ossification centers of the sternum were slightly smaller in heterozygotes as compared with wild-type homozygotes and that there is some delay in ossification.

Identification of markers closely linked to ch in the CHMU/Le inbred strain

ch has been propagated in several different genetic backgrounds; it is currently maintained in the inbred strain CHMU/Le which also carries the recessive coat color mutation muted (mu). Muted is localized to MMU 13 in close proximity (3 cM) to ch (28). Within CHMU/Le, ch and mu are maintained in repulsion so that eye and coat color markers can be used to distinguish all three ch genotypes (Fig. 1): homozygous muted mice (mu+/mu+) have light eyes at birth and develop fur of a light brown shade; heterozygotes (mu+/+ch) exhibit dark eyes at birth and develop an agouti coat color; mutant homozygotes (+ch/+ch) develop congenital hydrocephalus.

The CHMU/Le inbred strain has been maintained for >50 generations by brother-sister mating of heterozygotes at the Jackson Laboratory (Bar Harbor, ME). Theoretically, after 20 generations of free inbreeding, 98.7% of the genome in each animal is expected to be homozygous (29). However, this strain is a ‘segregating inbred strain’ maintained by purposeful selection of mice heterozygous for both ch and mu. Consequently, markers that are very closely linked to ch or mu are also expected to be heterozygous and we sought to identify these regions directly. Under forced heterozygosis by brother-sister mating across g generations for an autosomal lethal trait, the mean genetic length of chromosome heterozygous on each side of the principal locus is ~1/3g, when g is large (30). Since g = 50 for the CHMU/Le strain, we expect ~0.6 cM or 1.3 Mb around ch and mu to remain heterozygous. We searched for heterozygous markers in carriers (mu+/+ch), by screening >110 microsatellite markers that map to the proximal one-third of MMU 13 and identified four informative loci: D13Mit79, D13Mit275, D13Mit117 and D13Mit88. We used these markers to genotype 45 progeny (10 muted, 22 agouti and 13 mutants) from the intercross CHMU/Le mu+/+ch × CHMU/Le mu+/+ch: markers D13Mit275 and D13Mit117 were linked to ch and marker D13Mit88 was linked to mu (data not shown); marker D13Mit79 was not linked toeither locus. This analysis identified two recombinants amongst 47 mutants for D13Mit275, D13Mit117 and D13Mit88,which localized ch proximal to D13Mit275 and suggested thefollowing gene order: (centromere)-D13Mit79-ch-D13Mit275-D13Mit117-mu-D13Mit88-(telomere). O’Brien et al. (28) have genetically mapped mu in a 528 animal backcross panel and found no recombinants between mu, D13Mit88 and D13Mit87, estimating that these non-recombinant loci lie within 0.6 cM of one another. Our mapping of the mu locus to the same region is consistent with this study.

Mapping ch in interspecific crosses

We constructed interspecific crosses between CHMU/Le mu+/+ch carrier females and normal CAST and MOLF males for fine structure mapping. These crosses produced two types of offspring each with half their genome from CAST or MOLF and who are normal carriers of ch or normal carriers of mu (i.e. non-carriers of ch). These genotypes are not distinguishable but, since we have previously identified the CHMU/Le marker haplotype associated with ch and mu, we could infer ch from mu carriers by genotype analysis. We performed brother-sister matings, each progeny yielding two informative meioses; hydrocephalic offspring were genotyped with chromosome 13 simple sequence length polymorphisms (SSLPs). We generated 303 homozygous hydrocephalic mutants: 266 from CAST, 23 from MOLF and 14 from a CAST-MOLF mixed background (Fig. 3a and b). We detected no recombinants with D13Mit294 among all hydrocephalic mutants (556 meioses) from the CAST interspecific cross suggesting that ch lies in a maximum genetic interval of 0.18 cM (~360 kb) surrounding D13Mit294. The marker D13Mit294 was not informative in the MOLF cross. Haplotype analysis further showed that D13Mit206 and D13Mit307, informative in both the CAST and MOLF crosses, are the closest proximal and distal flanking loci, respectively. A total of 18 recombinants were detected with D13Mit206 (606 meioses) suggesting a map position ~3 cM proximal to ch. Only one recombinant was detected with D13Mit307 (604 meioses) localizing the marker ~0.17 cM, or only 330 kb, distal to ch.


Figure 3. (a) Haplotypes for 303 intercross ch progeny (606 meioses) for 13 microsatellite markers within the linked segment. Black boxes represent CHMU/Le allele homozygotes; white boxes represent CAST or MOLF allele homozygotes and heterozygotes; untyped markers are shown as grey boxes. Markers with an asterisk are also informative in CHMU/Le. Marker D13Mit294, showing no recombination with ch, is underlined. The number of progeny inheriting each haplotype is listed at the bottom. (b) A genetic map of the ch region on MMU 13. The numbers of recombinants and total meioses, with the ch/Mf1 gene, are shown in parentheses. The location of Hfh1 (*) was obtained by physical mapping (see text).

Our marker map, for placement of D13Mit206, exhibited discrepancy when compared with the published MMU 13 linkage maps (31,32) which place D13Mit206 further proximal and in the centromeric region. Numerous SSLP markers have been localized between D13Mit206 and D13Mit307, 19 of which were tested in our recombinant panel: D13Mit115, D13Mit55, D13Mit81, D13Mit83, D13Mit154, D13Mit84, D13Mit273, D13Mit220, D13Mit197, D13Mit163, D13Mit176, D13Mit58, D13Mit14, D13Mit17, D13Mit132, D13Mit16, D13Mit134, D13Mit301 and D13Mit304. Our analysis suggested that D13Mit206 is localized distal to these 19 markers consistent with our preliminary observations (Fig. 3a; unpublished data). Similar discrepancies in marker order at the proximal region of MMU 13 have also been observed during mapping of the beige-satin region (33). Our genetic analysis showed that D13Mit294 is localized proximal to D13Mit307, rather than further distal and near D13Mit275 as published (31). These differences are expected considering the small number of mice that were previously examined to generate the genome-wide linkage map (31).

The Bmp6 (bone morphogenetic protein 6) gene is a candidate gene for ch and was localized on our map. We synthesized oligonucleotide primers targeted to the 3[prime]-UTR based on the published amino acid sequence from EMBL/GenBank (34; GenBank accession nos J04566 and X80992) and identified variants between the CHMU/Le, CAST and MOLF strains. We genotyped the polymorphism in our interspecific crosses (n = 90) by single-strand conformational polymorphism (SSCP) analysis; haplotype analysis placed Bmp6 between D13Mit117 and D13Mit88 (data not shown). All genotype results from our interspecific crosses taken together suggested the following marker order: centromere-D13Mit206-ch, D13Mit294-D13Mit307, D13Mit265-D13Mit275-D13Mit117-mu, Bmp6, D13Mit87-D13Mit88 (Fig. 3b).

Physical map of the ch-Mf1 interval

We had predicted that ch was likely to lie in an interval of 0.18 cM (~360 kb) surrounding D13Mit294. Furthermore, progeny tested as obligate carriers of ch were found to be either heterozygous at D13Mit294 (i.e. both CHMU and CAST alleles were present) or homozygous for the CAST allele only. Therefore, the interval containing ch is even less than 360 kb. We pursued building a yeast artificial chromosome (YAC) and bacterial artificial chromosome (BAC) contig seeded at D13Mit294 and by ‘walking out’ ~400 kb distal and proximal to D13Mit294 (Fig. 4). We initially identified four YACs with D13Mit294 (Y63, Y88, Y97 and Y237) and one YAC with D13Mit307 (Y147) from the YAC library (35). An additional three YACs (Y139, Y289 and Y293) for D13Mit307 were identified by database screening (36; http://www.genome.wi.mit.edu/cgi-bin/mouse/index ); data for D13Mit294 were unavailable. We identified an additional five YACs (Y127, Y75, Y55, Y10 and Y123) using the sequence tagged sites (STSs) generated from the contig. A total of 13 YACs covered the region from the marker locus D13Mit307 and encompassing D13Mit294. The largest YAC (Y293), covering most of this interval, has an estimated insert size of 1 Mb; the smallest insert size (Y289) is ~200 kb. Each of the YACs contained only one band; however, four were unstable. For example, an SSCP marker isolated from one end of the YAC Y23 did not map to the contig and haplotype analysis showed that it was not ch linked (data not shown). Similarly, YAC Y75 was also shown to contain a chimeric end (P. Denny, personal communication). Three YACs (Y88, Y147 and Y75) showed internal deletions for some markers. We also isolated and mapped 14 BACs in the contig: seven encompassing D13Mit294, six surrounding D13Mit307 and one from a BAC which contains the mouse-specific Hfh1 (see below). STSs were generated from YAC/BAC ends and interspersed repetitive sequence-polymerase chain reaction (IRS-PCR) fragments from YACs and their order was determined by STS content mapping. We generated and ordered a total of 30 novel anonymous DNA markers in the contig: three SSLPs and eight SSCPs, which exhibited variants between CHMU/Le, CAST and MOLF strains, and 19 STSs, (Fig. 4 and Table 1).

Table 1. Oligonucleotide primer sequences for MMU 13 DNA markers
Marker Marker type Primer sequence (5[prime]->3[prime]) Temperature (°C) Product size (bp)aC HMU/CAST/MOLF
YAC ends
cwr6 STS GGTCCATCTGTTTAAGGACAGC
ATGTCTTGCTGTTATGGACGG		 55 214
cwr11 STS TTCCCACCCTCCTGTAAGC
CTTTCTGAGAAGGCCCGAG		 51 109
cwr21 STS AGGAGGGAATGTGTCGGAG
CTTTGCCCTGAATGTTCCAT		 55 251
cwr23 SSCP TTCTACCGTGGACGATACTGG
ACATCTGCTGCCCTTTGTCT		 59 309
cwr25 SSCP AATTGAAAGACACCTCCAAAGG
GGGCTGGTGTCAGTAGAGGT		 55 152
cwr32 STS TTTAGCACGAATTGGCTGC
ACTGGGGACTTGAACCCAG		 55 225
BAC ends
cwr1 STS AGCACTCGAGGTGGCTTTTA
CAGTTGGGATACTGCCAGGT		 55 234
cwr2 STS ATTCTCCCTGAGCCTGGG
GAAATTCAGGTGGCCAAGAA		 59 144
cwr3 STS CAGTGGGAAGGCTAAAGCAG
GGGAGTATCTTGCCAGGACA		 55 184
cwr4 SSCP GACCTAAGCACATGGGCATC
GGGCATGTAATCTGTGCATG		 52 220
cwr5 STS TGATGAAAAAGATGTGCCCA
CAACCAAGATGCTTGTGCC		 51 242
cwr7 SSLP/(ca)n GACACATATCTGCATGCTGTCA
TAAACAAGTTAGATAAGCCAC		 59 185/183/199
cwr8 STS CACACCTTCCTTGGTATTCCA
ATAAAAGCCCTCAAGGAGATCA		 59 100
cwr9 SSCP TCCATCCTGGACTTAACAGTTG
AAGTGGAATTGCCTGCAGAC		 52 193
cwr10 STS AAGTGTCCCATTGATGCCTC
TCATTTGCCAACAGACCAAA		 51 154
cwr12 STS GGCTGTGAAATGCCCTAAAA
TCACGTATCAGAGATGGAAAGG		 55 177
cwr15 STS AATGCACAAGGATACCTTCCC
GGATTCAGTTTAACTGGCAAGC		 55 193
cwr16 SSLP/(ca)n CTGCACCTTGGGATCAGTTT
GCATCTAAACATAAGGTTGGGC		 55 150/157/153
cwr17 STS AATGGCAATCTGTCTGGAGG
TCAATACCCCTGCTCCTTACC		 52 153
cwr18 STS TCGAGTTGTCCCCTCTGC
GGGAGTTTTCAGGACAAAAGG		 55 253
cwr19 STS CCCGCAGTTAATAGGCACAT
GTCCCTTTTTCCAGTAAAACG		 55 128
cwr20 SSCP CAGCAGAACACTGCTAACAACC
GGCAGACAGTAGACGGCTTC		 55 218
cwr22 SSLP/(ca)n CCCATCACAGGCTGGTAGAT
CTGCACTCTGATCTCAGATTCC		 59 136/146/166
cwr24 STS ATTTCAGAGCGGAAATTGTAGG
GTTCATGGGCCAACTCAGAT		 55 223
cwr26 SSCP TGGTGTTCAGTTACCTTGATGG
GCACCTGGGAGACTTAGAAAA		 55 185
cwr27 STS CCTCCAGAAGCCACAGAAAC
GTTTGTTTCTGCACCCATCA		 55 123
cwr30 SSCP TTTAAGCCCAGGTCCTCATG
GAGAATGACGGGACCTCAAA		 55 108
cwr31 STS GAGAGACGCTGGTTGTCACA
AGAACCTGGCTAGGACTAGCG		 55 199
Other markers
cwr13 STS AGCTCAAGCAGCTTTTATTTGG
CTTGGGCTAAAAAAAAGGGG		 55 311
cwr14 SSCP TGTGCTTTTGTCAGATTGAGTT
ATGCAGATAGTGATGCATACGG		 59 295
Genes
Bmp6b SSCP TAGTACAGGCCTGGAAACTGC
TCCACAGAGCCTGCTGATGG		 58 188
Mf1c 3[prime]-untranslated   AGATTTTTTTTCCCCCTACAGC
TCCGGGTACATTTGCTTCT		 55 222
Hfh1d KF/KR primers   TTCGGAAAAGCGTCTCTCTCGG
GATGAGTGCGATGTAGGAGTAGGG		 55 528
Hfh1d F3/R3 primers   ACCTCTCGCTCAACGACTGT
AAGGTGTATTCGCTGTTGGG		 55 103
STS, sequence tagged site; SSCP, single-strand conformational polymorphism; SSLP, simple sequence length polymorphism.
aProduct sizes for CHMU/Le, CAST/Ei and MOLF/Ei alleles were determined by PCR SSLP analysis.
bPrimers for Bmp6 were designed from the published sequence (34; GenBank accession nos J04566 and X80992).
cPrimers were designed from genomic sequence provided by B.L.M. Hogan (personal communication).
dPrimer sets for Hfh1 were designed from the published rat Hfh1 cDNA sequence (39; GenBank accession no. L13201).

From analysis of ch recombinants we were able to identify cwr30 and cwr14 as closer proximal and distal flanking markers, respectively, further narrowing the ch critical interval for constructing a transcript map. While this work was in progress, Kume et al. (22) reported that targeted disruption of the mouse gene Mf1 revealed a homozygous phenotype identical and allelic to ch. This was puzzling given that this group (21) had recently mapped Mf1 near the Bmp6 gene and hence further distal to D13Mit294 and ch. Consequently, we refined the location of Mf1 on our fine structure genetic map. We generated primers targeted to the Mf1 genomic sequence and assayed whether Mf1 was contained in our contig. Indeed, four YACs (Y289, Y75, Y88 and Y23) and three BACs, which are also positive for D13Mit294, harbor Mf1 and this confirms the report by Kume et al. (22). We sequenced the Mf1 gene in ch mutants and their normal littermates to confirm the nonsense mutation at bp 367 (C367T and Q123X) within the forkhead DNA binding domain.


Figure 4. A contig map of overlapping YACs and BACs of the ch/Mf1 region. The contig spans ~1 Mb but precise intermarker distances of the 30 new DNA markers have not been determined (Table 1). Open circles denote a chimeric YAC end; clones with an asterisk have not been tested for chimerism. YACs with open boxes indicate deletions of the corresponding markers.

Evaluation of the ch/+ ocular phenotype

As a part of the phenotypic examination of ch mice we examined the eyes of wild-type homozygotes (mu+/mu+) and ch heterozygotes (mu+/+ch) from the CHMU/Le inbred strain at various ages of the newborn and adult. We examined ch heterozygotes from the following age groups by slit lamp examination: 3-4 weeks old (n = 14), 1-2 months old (n = 15) and 2-9 months old (n = 20). For the age group of 2-9 months, we examined three mice of age 2.5 months, two of age 3 months, seven of age 4.5 months, four of age 5 months and four of age 9 months. In addition, we examined six mice of age 1 year or older. Thus, a total of 55 ch heterozygotes were examined at various ages. In all age groups, we observed numerous anterior segment abnormalities in all ch heterozygotes whereas the eyes of age-matched wild-type homozygotes were normal (Fig. 5). Moreover, ch heterozygotes showed ocular abnormalities with marked variable expressivity. The most common anterior segment defect observed in all ch/+ was corectopia (displaced pupil), present mostly as bilateral (n = 41/55, 75%) but sometimes as unilateral (25%) defects. Mild corectopia and irregularly shaped pupils were already present in ch heterozygote mice by weaning age. In addition, some mice from the age group 3-4 weeks old showed prominent Schwalbe’s line (posterior embryotoxon) and mild corneal opacity. By the age of 2 months, the corectopia had progressed and iridocorneal adhesion (anterior synechia) was more evident. Anterior synechia was observed ranging in size from broad sheets to thread-like strands in some mice. The thinning of the iris stroma undergoes progressive changes with age and, later, multiple iris holes (polycoria or pseudocoria) develop in opposite quadrants of the displaced pupil. In older ch heterozygote mice, a high incidence of corneal opacification, with neo-vascularization and cataracts, was also observed by 9 months. Similar anterior segment defects were noted in ch heterozygotes in the CAST genetic background as well; however, the ocular abnormalities were not fully penetrant. We examined seven ch heterozygotes from the CAST genetic background which included four mice of age 2 months old, one of age 5 months old and two of age 10 months old. Only one mouse (5 months old) showed unilateral corectopia (n = 1/7, 14%) and all others were normal. Although we examined few mice, the data indicate that the genetic background influences the penetrance of ocular defects in ch heterozygotes. Studies with additional mice at various ages are in process to examine the genetic background influences in greater detail.


Figure 5. Anterior segment phenotype of wild-type homozygote and ch heterozygote. (a) Eye of a wild-type homozygote from the CHMU/Le inbred strain (mu+/mu+). The eye pigment is reddish and lighter in color due to the effects of mu. (b) For comparison, the eye of a normal wild-type homozygote (2.5 months old) from an agouti CAST genetic background is shown. (c-e) Eyes of heterozygotes (mu+/+ch) at various ages from the CHMU/Le inbred strain showing variable and progressive expressivity of anterior segment abnormalities. (c) A 6-week-old heterozygote with corectopia of the right eye and an elongated pupil of the left eye. Both eyes exhibit prominent Schwalbe’s lines (also known as posterior embryotoxon) (arrow). (d) A 2.5-month-old heterozygote. The right eye shows an elongated pupil and posterior embryotoxon. The left eye shows a broad anterior synechia with corneal opacification. (e) Eyes of a >1-year-old heterozygote. Both eyes exhibit extensive corneal opacification with neo-vascularization. The right eye shows an enlarged pupil with extensive anterior synechia and cataract. The left eye shows multiple holes in the iris. R, right; L, left.

Mapping of mouse Hfh1

The vast majority of winged helix/forkhead genes are distributed across the genome, yet clustered arrangements have also been described (37,38). On MMU 13, another member of the winged helix/forkhead transcription factor Hfh1 has been localized to the proximal region, ~2.2 ± 1.5 cM proximal to Bmp6 (21). Based on our mapping data (Figs 3 and 4) Hfh1 and Mf1 may be closely linked. We designed two PCR primer sets from the published rat Hfh1 cDNA sequence to amplify the mouse-specific Hfh1 gene (Table 1): the mouse Hfh1 sequence has not been published; however, the amino acid sequences from rat and mouse are reported to be identical (39,40). By PCR analysis we examined whether the mouse Hfh1 maps within the ch/Mf1 region: indeed, two YACs (Y10 and Y123) contain Hfh1, placing it proximal to Mf1 (Fig. 4).

We also localized Hfh1 in relation to other SSLP markers and Mf1 by radiation hybrid mapping using the T31 radiation hybrid (RH) panel (41). Hfh1 was analyzed against the following markers/loci which we genotyped in the RH panel: centromere-D13Mit81-D13Mit206-Mf1, (ch)-D13Mit294-D13Mit307-D13Mit265-D13Mit116. The RH data gave an order consistent with our previous genetic and physical map (Fig. 3). As expected, the retention patterns of Mf1 and D13Mit294 were identical. Hfh1 exhibited almost identical retention patterns as Mf1 ([chi]2 = 84.23, df = 1, P = 4 × 10-20) and the DNA marker D13Mit294([chi]2 = 85.17, df = 1, P = 3 × 10-20): only three hybrids were discordant between Hfh1 and Mf1 (96.91 ± 1.76%, n = 97 hybrids) and between Hfh1 and D13Mit294 (99.97 ± 1.76%,n = 98 hybrids). However, we observed greater discordancy between Hfh1 and the rest of the DNA markers tested. We further analyzed our radiation hybrid mapping data using the MultiMap (42) and the RADMAP (T.C. Matise and A. Chakravarti, unpublished program) programs which suggested that the most likely location of Hfh1, at 1000:1 or better odds, was proximal to Mf1 and D13Mit294. Our analysis estimated the Hfh1 and Mf1 distance as 7 cR. In addition, the distance between the markers D13Mit294 and D13Mit307, which are 800 kb apart, is estimated to be 12 cR. This argues for a conversion rate of 67 kb/cR3000 placing the mouse Hfh1 gene ~470 kb proximal to Mf1. We also identified a BAC which contains Hfh1 to generate additional markers in this region by BAC end sequencing. We designed two novel markers, cwr31 and cwr30, which map proximal and distal to Hfh1, respectively. As mentioned previously, genotype analysis of ch recombinants revealed one recombinant with cwr30, suggesting that its map location is ~400 kb proximal to Mf1.

DISCUSSION

Grüneberg (11) postulated that ‘the site of basic disturbance’ in ch mutants is the delayed differentiation of the preskeletal mesenchyme resulting in striking anomalous shapes and absence of various skeletal elements. Our evaluations of skeletal systems of ch mutants from CHMU/Le are concurrent with Grüneberg’s original hypothesis. We have also shown similar skeletal malformations of ch in the CAST and MOLF genetic backgrounds, although with some delayed ossification. Green (12) was the first to show that the subarachnoid drainage system fails to form in embryos, resulting in expansion of the cerebral hemispheres. The overstretching of the osteogenic membrane may hamper aggregation of mesenchymal cells to reach a critical density, so that skeletogenesis is compromised. This is further corroborated by observations of meningeal defects in Mf1 knockout mutants; however, there is some ability of the dorsal head mesenchyme of 11.5 d.p.c. mutant embryos to give rise to cartilage and bone in micromass culture (22). It is interesting to note ossification spots on the expanded cerebral hemispheres in some of our ch mutants from the interspecific cross; thus, mesenchymal cells from these mutants do have the ability to differentiate into osseous bones. ch mutants from our interspecific cross also showed similar effects on the sternum (Fig. 2k); however, the sternal primordium of the Mf1 null mutants failed to differentiate in micromass culture, even in the presence of added chondrogenic factors such as BMP2 and TGF[beta]1 (22).

The molecular mechanisms and factors that regulate chondro-genesis and osteogenesis are poorly understood. Skeletal development is a multistep process which includes aggregation of undifferentiated mesenchyme into primordial condensation, growth and commitment of precursor cells to chondrogenic and osteogenic lineages and subsequent terminal differentiation to endochondral and membranous bones. Recent studies of loss-of-function mutants have identified several nuclear factors affecting bone formation and thus have begun to dissect molecular pathways of skeletal organogenesis. Two independent targeted disruptions of Cbfa1 (core-binding factor) result in homozygous mutant mice with a complete lack of bone formation owing to maturational arrest of osteoblasts and osteogenesis (43,44). The expression of Cbfa1 reaches a peak at 12.5 d.p.c., at which time cartilage formation commences: Cbfa1 expression is restricted to cells of the mesenchymal condensation of the developing future skeletal structures (45). It has since been shown that Cbfa1 is deleted in ccd (cleidocranial dysplasia) mutant mice (44) and that patients displaying cleidocranial dysplasia have mutations in the human CBFA1 gene (46). Another regulator of bone development is the paired-class homeobox-containing gene Cart1 (cartilage homeo-protein 1) that maps to MMU 10 (47). Cart1 null mice are born with severe craniofacial and neural tube closure defects which give rise to exencephaly. Interestingly, these mutants also show craniofacial skeletal defects very much like those in ch, in particular the lack of a cranial vault (48). Additional nuclear factors that affect skeletogenesis seem to regulate the level of mesenchymal differentiation during early organogenesis. Targeted disruption of homeodomain-containing transcription factors in the mouse, such as Mhox (49), goosecoid (50) or the msh-like gene msx-1 (51), have revealed absence and malformation of specific skeletal elements each resulting in distinctive craniofacial defects and abnormalities of the limb and vertebral skeletal structures. These studies implicate specific genes in the formation, proliferation or differentiation of chondrogenic and osteogenic precursors of affected skeletal elements. A winged helix trans-cription factor, Mfh1, which has restricted and temporal expression in the developing mesenchyme (52), has also been shown to regulate bone development. A null allele, obtained by homologous recombination, exhibits multiple craniofacial and vertebral defects that result from deletion or malformation of skeletal elements derived mainly from the head mesenchyme (53).

Grüneberg (11) argued that ch phenotypes could also arise from a defective condensation environment rather than a direct effect of the mutation on condensation. Interestingly, it has been reported that the defective sternocostal cartilages in ch embryos have a deficient extracellular matrix, with as much as 50% less glycosaminoglycans than the cartilage in wild-type embryos (13). Furthermore, extracellular matrix (ECM) disorganization in the sternum has been demonstrated by a scarcity of proteoglycans and collagen fibrils in the intercellular matrix by ultrastructural analysis (15-17). The extracellular collagen fibrils identified in ch were either present in abnormal aggregates or enlarged due to a deficiency of polyanion (proteoglycan) matrix substances. Similar findings of ECM disorganization were also found in the ch renal system, and all ch kidneys demonstrate poor development or immaturity (12,15-17). The development of cartilage and bone is initiated by the aggregation of mesenchyme, which is dependent on reaching a critical cell density and maintenance of cell proliferation, processes which are also regulated by epithelial-mesenchyme tissue interactions (54). Moreover, a complex ECM provides important cues to embryonic cell populations for migration and differentiation; a variety of microenvironmental factors are important in determining the fate of cells from the neural crest as well as in defining their migration pathways and differentiation (54-57). Proteoglycans represent a vast family of ECM components and have been implicated in key developmental events. Recent studies suggest that proteoglycans are crucially involved in the process of neural crest cell migration; a time-related and region-specific organization of diverse types of proteoglycans occurs during neural crest cell migration and gangliogenesis and the onset of vertebral chondrogenesis (56,57). It is important to note that the mature arachnoid is made up of a trabecular meshwork composed of collagen fibers. Perhaps the ubiquitous nature of the ECM defects also plays a role in systemic developmental phenotypes in ch mutants. Whether or not Mf1, which is a transcription factor, regulates the organization of ECM components, directly or indirectly, is not known. It is possible to postulate that proteoglycans and other ECM components may be one of the downstream targets of Mf1. Thus, ch is an attractive model system to identify additional key players of skeletogenesis. The next challenge is to identify these target genes which may reveal the developmental pathway of Mf1 action and help in understanding the ECM defects and the pleiotropic phenotype observed in ch mutants.

Much attention has been placed in investigating the homozygous ch hydrocephalic mouse as a genetic animal model for congenital hydrocephalus and skeletal organogenesis in the past 40 years. Although the urogenital system and the sternum have been investigated (12,13), the ch heterozygous mouse has been overlooked by investigators, partly because of their healthy appearance. Here, ocular defects in ch heterozygotes are described for the first time and we show that eye abnormalities are likely to be caused by malformations of the anterior chamber of the eye. In addition, ocular defects displayed in ch heterozygotes have characteristic features observed in human patients diagnosed with Axenfeld-Rieger anomaly and iridogoniodysgenesis anomaly. In humans, Axenfeld-Rieger/iridogoniodysgenesis eye malformations are a group of dominant disorders characterized by iris hypoplasia, anterior segment mesenchymal dysgenesis and glaucoma resulting from a neurocristopathy leading to a developmental arrest of the anterior chamber angle structures (58). From linkage analysis and the recent cloning of the relevant genes, several transcription factors have been identified as causing Axenfeld-Rieger/iridogoniodysgenesis eye malformations (27,59,60). Mutation analysis has shown that FKHL7 is one gene involved in human patients (27). Thus, haploinsufficiency of Mf1 in mice, as for FKHL7 in humans, leads to the disease phenotype, suggesting that eye development is sensitive to levels of functional Mf1 protein.

The anatomic components of the anterior chamber angle includes the Schwalbe’s line, the trabecular meshwork and the scleral spur, all lying at the junction of the peripheral cornea and at the root of the iris. Shields et al. (58) proposed that, as a consequence of a developmental arrest, abnormal retention and contraction of the embryonic endothelial layer on portions of the iris and anterior chamber angle leads to the iridic changes, the tissue strands and the abnormalities of Schwalbe’s line associated with Axenfeld-Rieger anomaly. Ocular defects in ch heterozygotes are progressive and, as in the human, feature displacement of the pupil with occasional hole formation in the iris, the abnormalities of Schwalbe’s line, anterior synechia and, invariably, corneal scarring in older mice. In addition, ch heterozygote mice, even in the CHMU/Le inbred genetic background, display widely variable ocular defects. Clearly, the observed variability is not due to a heterogeneous genetic background in this study but stems from developmental defects of the eye. Furthermore, a wide spectrum of ocular anomalies are often seen within the same animal. Concurrent with our observation, gene-targeted Mf1 homozygotes, at 16.5 d.p.c., reportedly had an incompletely developed anterior chamber, disorganized cornea and hypoplasia of the iris stromal mesenchyme (22).

Glaucoma occurs in ~50% of human patients with Axenfeld-Rieger anomaly due to the secondary effect of the incomplete maturation of the anterior chamber structures (58). Aqueous humor produced by the ciliary body enters the posterior chamber and passes through the pupil into the anterior chamber (61). Drainage occurs mainly at the canal of Schlemm and contraction of the ciliary muscle though its insertion into the trabecular meshwork determines the rate of aqueous drainage. Ultrastructural analyses of the anterior chamber angle from human patients have shown failure of the intertrabecular spaces and Schlemm’s canal to develop, thus leading to obstruction to aqueous flow and glaucoma (58). Like the arachnoid layer of the brain meninges, the ocular trabecular meshwork is composed of collagen and elastic tissue, thus similar developmental defects may account for the inadequate drainage found for both the CNS and ocular system in ch homozygotes and heterozygotes. Whether ch heterozygotes develop glaucoma remains to be studied. However, ch heterozygotes are an excellent model system for Axenfeld-Rieger/iridogoniodysgenesis eye malformations.

It is interesting to note the high incidence of cataracts in ch heterozygotes; Mf1 is reported to be expressed in the periocular mesenchyme, in some cells of the cornea and in endothelial cells of the hyaloid vessels, but not expressed in the developing lens (22). Thus, cataract formation in ch heterozygotes may not be the direct result of Mf1 haploinsufficiency, but rather a secondary effect. Numerous mouse mutations associated with cataract formation frequently demonstrate abnormalities in the formation of other ocular structures (62,63). Therefore, ch heterozygotes are of potential value in deciphering the molecular mechanisms involved in secondary cataract formation. It is also interesting to note the expression of Mf1 in the developing hyaloid system, which mainly forms the ECM-containing vitreous. However, the developmental role of Mf1 in the hyaloid system is undefined at this time. Certainly further detailed analysis will elucidate the embryology and progression of ocular defects, as well as the events that lead to cataracts in ch heterozygotes.

The forkhead or winged helix transcription factors are a growing gene family identified in species ranging from yeast to human, all recognized by a conserved 110 residue DNA-binding domain. These genes participate in diverse biological functions, such as in tumorigenesis, cellular differentiation and a wide range of developmental processes (64). The Drosophila forkhead gene (fkh) and a family of hepatocyte nuclear factor 3 protein genes (HNF3), the products of which share a strikingly similar DNA-binding motif, are considered to be the founders of this gene family (65-67). Other members of this gene family have been isolated by low-stringency hybridization screening of genomic libraries using the HNF3/fkh motif as a probe. In this study, we present a fine localization of another member, Hfh1. By radiation hybrid mapping and physical mapping of the ch/Mf1 critical region, we have shown that Hfh1 is localized ~400 kb from Mf1. Whether this clustered arrangement has any implications regarding a common usage of regulatory elements governing their expression is unknown. The existing, limited expression studies show that Hfh1 is diffusely expressed throughout the lung tissue of the adult rat/mouse using in situ hybridization (40); Hfh1 is also moderately expressed in adult kidney tissue (39). Embryonic expression of Hfh1 has not been addressed and, in addition, the genomic structure and sequence of Hfh1 are undetermined. Finally, it would be useful to investigate whether the human ortholog of the mouse Hfh1 gene is also localized to the 6p terminal region and whether it is involved in the 6p deletion syndrome.

MATERIALS AND METHODS

Animals

Inbred strains of mice carrying ch (CHMU/Le mu+/+ch), M.m.castaneus (CAST/Ei) and M.m.molossinus (MOLF/Ei) were purchased from the Jackson Laboratory (Bar Harbor, ME) and maintained in accordance with animal usage guidelines. We conducted three types of matings: (i) brother-sister matings of ch carriers [CHMU/Le mu+/+ch × CHMU/Le mu+/+ch]; (ii) intersubspecific crosses between a ch carrier from CHMU/Le and M.m.castaneus (CAST) [CHMU/Le mu+/+ch × CAST/Ei] and M.m.molossinus (MOLF) [CHMU/Le mu+/+ch × MOLF/Ei]; and (iii) intercrosses of F1 ch carriers from the intersubspecific crosses to obtain recessive offspring in the CAST or MOLF genetic background.

Skeletal and slit lamp analysis

Skeletal preparations were made by the alcian blue/alizarin red clearance method (68), with some modifications. Eyes of anesthetized mice were examined using a Zeiss slit lamp and photographed with a vertical photo slit lamp (Marco Technology, Fl) (69).

Genotyping

Primer sets of SSLP markers were purchased from Research Genetics (Huntsville, AL). PCR was performed on a PTC-200 thermal cycler (MJ Research, Watertown, MA) in 20 µl volumes containing 100 ng of genomic DNA, 6.6 pmol of unlabeled primer, 1.3 pmol of labeled primer, 200 µM each dNTP and 0.5 U Taq polymerase with the recommended assay buffer (Boehringer Mannheim, Indianapolis, IN). Each primer was 5[prime]-end-labeled with [[gamma]-33P]ATP (Andotek Life Science, Irvine, CA) using 0.4 µCi/1.3 pmol primer. Amplified products were separated on 6% denaturing polyacrylamide gels followed by drying and autoradiography. Sizes of alleles were determined by comparison with M13 DNA sequencing ladders. For SSCP analysis, PCR was performed under standard reaction conditions without radioactive label and analyzed using the LKB PHAST system (Pharmacia, Piscataway, NJ). Samples of a 2:1 mixture of PCR product and stop solution were resolved on 20% homogeneous gels at 15°C for the recommended V/h depending on product size. Gels were stained and fixed as suggested by the manufacturer.

Construction of YAC and BAC contigs

Mouse YAC libraries from Research Genetics were screened by PCR with primer sets for appropriate microsatellite markers. The ends of YACs were isolated by the bubble-vector PCR method (70) or by homologous recombination/plasmid-end rescue (71). The ends of each YAC were sequenced using the ABI PRISM Dye Terminator Cycle Sequencing Ready Reaction kit (Perkin Elmer, Foster City, CA), as specified by the manufacturer. For estimation of YAC insert size, clones were grown and agarose plugs prepared in low melt agarose (72). Plugs were run in 1% agarose gels in the Bio-Rad Pulse Field Chef-DR II system (Bio-Rad, Hercules, CA), following the manufacturer’s protocols, and immobilized on a nylon filter (MSI, Westborough, MA) by the Southern method (73). pBR322 was digested to completion with BamHI and PvuII and the resulting 2.7 and 1.7 kb fragments were used as hybridization probes for the left and right ends of the YAC vector.

Mouse BAC clones arrayed on high density membranes (Research Genetics) were screened with appropriate STSs or oligonucleotides by hybridization. STS and oligonucleotide probes were labeled using the Redi-Prime random primer kit (Amersham, Arlington Heights, IL) and [[alpha]-32P]dCTP (NEN-Dupont, Boston, MA) and purified using Quick Spin G50 or G25 columns (Boehringer Mannheim), respectively. BAC DNAs were purified by the alkaline lysis method (73) using the Nucleobond AX 500 Kit (Clontech, Palo Alto, CA). BAC ends were directly sequenced with the ABI PRISM BigDye Terminator Cycle Sequencing Ready Reaction kit (Perkin Elmer), as specified by the manufacturer with some modifications. The sequencing reactions were run on an automated ABI model 377 sequencer and gels analyzed by the Sequence Analysis v.3.0 program (Perkin Elmer).

IRS-PCR probe generation

To generate fragments the following mouse-specific IRS primers were used: B1R, 5[prime]-CCCTGGCTGTCCTGGAACTCACTCTG-3[prime] (74); L1, 5[prime]-GGTATGGGGGACTTTTGGGAT-3[prime] (75). YACs were amplified under recommended conditions (74,75) and PCR products were analyzed on 2% agarose gels. IRS-PCR products were cloned into pCR2.1 plasmid using the Original TA cloning kit (Invitrogen, Carlsbad, CA) and sequenced.

PCR analysis of Mf1 and Hfh1

Genomic sequence for the mouse Mf1 gene was obtained from B.L.M. Hogan (22; GenBank accession no. AF045017) and primers which target the 3[prime]-UTR were used for mapping (Table 1). Two primer sets (Table 1, KF and KR and F3 and R3) were designed for the analysis of the mouse-specific Hfh1 locus from published rat Hfh1 cDNA sequence (39; GenBank accession no. L13201). PCR analyses for both Mf1 and Hfh1 were carried out under standard reaction conditions and analyzed on 2% agarose gels. The cycling conditions used for both Mf1 and Hfh1 were: an initial denaturation of 4 min at 94°C; 35 cycles of 30 s at 94°C, 30 s at 55°C, 30 s at 72°C; followed by 10 min elongation at 72°C.

Radiation hybrid data analysis

A panel of 100 whole-genome radiation hybrids for the mouse (T31) was purchased from Research Genetics. PCR reactions were carried under standard conditions as described above. All markers were genotyped in duplicate and consensus readings were analyzed using the computer programs MultiMap (42) and RADMAP (T.C. Matise and A. Chakravarti, unpublished program).

ACKNOWLEDGEMENTS

We wish to thank Drs Shukti Chakravarti, Ron Conlon and Terry Magnuson for their many helpful discussions, Drs Cindy Faust, Armin Schumacher and David Threadgill for extensive and persistent technical advice, Carl Kashuk for developing the mouse database, Kim Bentley and Nydia Bringht-Twumasi for technical assistance, Dr Shukti Chakravarti and Eugenia Diaconu for help in slit lamp analysis and Drs Brigid Hogan and Tsutomu Kume for providing us with Mf1 genomic sequence and primers. This work was supported by NIH grant HD 34857 to A.C., a grant from the Willson Foundation to the Center for Human Genetics and Research to Prevent Blindness Foundation, Ohio Lions Eye Research Foundation and EY11373-01A1 (J.H.L.).

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*To whom correspondence should be addressed. Tel: +1 216 368 5847; Fax: +1 216 368 5857; Email: axc39@po.cwru.edu

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