REPORT Recessive and Dominant De Novo ITPR1 Mutations Cause Gillespie Syndrome Sylvie Gerber,1,22 Kamil J. Alzayady,2,22 Lydie Burglen,3,4,22 Dominique Bre´mond-Gignac,5 Valentina Marchesin,6 Olivier Roche,5 Marle`ne Rio,7 Benoit Funalot,7,8 Raphae¨l Calmon,9,10 Alexandra Durr,11 Vera Lucia Gil-da-Silva-Lopes,12 Maria Fernanda Ribeiro Bittar,12 Christophe Orssaud,5 Be´ne´dicte He´ron,13,14 Edward Ayoub,2 Patrick Berquin,15 Nadia Bahi-Buisson,16 Christine Bole,17 Ce´cile Masson,18 Arnold Munnich,7 Matias Simons,6 Marion Delous,19 Helene Dollfus,20 Nathalie Boddaert,9,10 Stanislas Lyonnet,7,16 Josseline Kaplan,1 Patrick Calvas,21,23 David I. Yule,2,23 Jean-Michel Rozet,1,23,* and Lucas Fares Taie1,23 Gillespie syndrome (GS) is a rare variant form of aniridia characterized by non-progressive cerebellar ataxia, intellectual disability, and iris hypoplasia. Unlike the more common dominant and sporadic forms of aniridia, there has been no significant association with PAX6 mutations in individuals with GS and the mode of inheritance of the disease had long been regarded as uncertain. Using a combination of triobased whole-exome sequencing and Sanger sequencing in five simplex GS-affected families, we found homozygous or compound heterozygous truncating mutations (c.4672C>T [p.Gln1558*], c.2182C>T [p.Arg728*], c.6366þ3A>T [p.Gly2102Valfs5*], and c.6664þ5G>T [p.Ala2221Valfs23*]) and de novo heterozygous mutations (c.7687_7689del [p.Lys2563del] and c.7659T>G [p.Phe2553Leu]) in the inositol 1,4,5-trisphosphate receptor type 1 gene (ITPR1). ITPR1 encodes one of the three members of the IP3-receptors family that form Ca2þ release channels localized predominantly in membranes of endoplasmic reticulum Ca2þ stores. The truncation mutants, which encompass the IP3-binding domain and varying lengths of the modulatory domain, did not form functional channels when produced in a heterologous cell system. Furthermore, ITPR1 p.Lys2563del mutant did not form IP3-induced Ca2þ channels but exerted a negative effect when co-produced with wild-type ITPR1 channel activity. In total, these results demonstrate biallelic and monoallelic ITPR1 mutations as the underlying genetic defects for Gillespie syndrome, further extending the spectrum of ITPR1-related diseases.

Aniridia (MIM: 106210) refers to a rare inborn error of eye development whose most prominent sign is the partial or complete absence of iris. Associated ocular malformations include macular and optic nerve hypoplasia, cataract, and corneal opacification. Most aniridia cases are accounted for by dominant mutations affecting PAX6 (MIM: 607108) at the AN locus (MIM: 106210).1 Wilms tumor, genitourinary abnormalities, and intellectual disability are not uncommon in individuals with aniridia. This complex form of the disease known as WAGR syndrome (MIM: 194072) has been ascribed to 11q13 deletions involving the PAX6 locus and the adjacent Wilms tumor 1 (WT1) locus (MIM: 607102).1

Occasionally, aniridia can be associated with cerebellar ataxia and intellectual disability, defining a clinical entity known as Gillespie syndrome (GS [MIM: 206700]).2 GS is usually diagnosed in the first year of life by the presence of fixed dilated pupils in a hypotonic infant. Ophthalmologic examination reveals partial aniridia with the pupil border of the iris containing festooned edge and iris strands extending onto the anterior lens surface at regular intervals.3 These eye findings are mandatory to make the diagnosis of GS because they demonstrate the developmental nature of the iris anomaly, thereby excluding neurogenic congenital mydriasis in an infant with neuropsychomotor symptoms.

1 Laboratory of Genetics in Ophthalmology (LGO), INSERM UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 2Department of Pharmacology and Physiology, University of Rochester, Rochester, NY 14526, USA; 3Reference Center for Cerebellar Malformations and Congenital Diseases, Department of Genetics, Trousseau Hospital, AP-HP, 75012 Paris; 4INSERM U1141, DHU PROTECT, Robert Debre´ Hospital, 75019 Paris, France; 5Department of Ophthalmology, IHU Necker-Enfants Malades, Paris Descartes University, 75015 Paris, France; 6Epithelial biology and disease – Liliane Bettencourt Chair of Developmental Biology, INSERM UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 7Department of Genetics, IHU Necker-Enfants Malades, University Paris Descartes, 75015 Paris, France; 8Department of Genetics, GHU Henri Mondor, 94010 Cre´teil, France; 9Department of Neuroradiology, IHU Necker-Enfants Malades, Paris Descartes University, 75015 Paris, France; 10 Image at Imagine, INSERM UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 11Maladies Neurode´ge´nˆ pital Pitie´-Salpeˆtrie`re, 75013 Paris, France; e´ratives, Institut du Cerveau et de la Moe¨lle Epinie`re, CHU Paris-GH La Pitie´ Salpeˆtrie`re-Charles Foix, Ho 12 Department of Medical Genetics, Faculty of Medical Sciences, University of Campinas, CEP 13083-887, Sao Paulo, Brasil; 13Department of Neuropediatˆ pital Jean Verdier, 93143 Bondy, France; 15Po ˆ le de pe´diatrie, Centre d’activite´ de ˆ pital Trousseau, 75012 Paris, France; 14Department of Pediatrics, Ho rics, Ho ˆ pital Nord, 80054 Amiens, Cedex 1, France; 16Embryology and Genetics of Human Malformation, INSERM neurologie pe´diatrique, CHU d’Amiens - Ho UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 17Genomics Platform, INSERM UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 18Bioinformatics Platform, INSERM UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 19Laboratory of Hereditary Kidney Diseases, INSERM UMR1163, Imagine – Institute of Genetic Diseases, Paris Descartes University, 75015 Paris, France; 20Laboratoire de Ge´ne´tique Me´dicale, Institut de Ge´ne´tique Me´dicale d’Alsace, INSERM U1112, Fe´de´ration de Me´decine Translationnelle de Strasbourg (FMTS), Universite´ de Strasbourg, 67085 Strasbourg, France; 21Service de Ge´ne´tique Clinique, ˆ pital Purpan, 31300 Toulouse, France Ho 22 These authors contributed equally to this work 23 These authors contributed equally to this work *Correspondence: [email protected] http://dx.doi.org/10.1016/j.ajhg.2016.03.004. Ó2016 by The American Society of Human Genetics. All rights reserved.

The American Journal of Human Genetics 98, 971–980, May 5, 2016 971

Since Gillespie’s initial description in 1965, 22 affected families have been reported.2–19 Current GS-affected families include three sibpairs describing an autosomal-recessive inheritance, two families with disease transmission from an affected mother to the progeny, and 18 simplex families in which recessive and dominant de novo mutations might underlie the disease. De novo PAX6 mutations have been identified in two simplex families but otherwise molecular studies have failed to identify the diseasecausing mutation, supporting genetic heterogeneity of GS.9,13,15 We collected five trios with index case subjects presenting with partial aniridia, cerebellar ataxia, and intellectual disability with or without additional systemic symptoms (Table 1 and Figures 1A and 2). Informed consents were obtained from all the research participants. The study received approval from the Comite´ de Protection des Personnes ‘‘Ile-de-France II.’’ Three index case subjects (F1:II1, F2:II1, F5:II1), one of which was reported previously (F2:II1),14 were born to consanguineous parents (Figure 1A). F1:II1, F3:II1, F4:II1, and F5:II1 had neither point mutations nor micro-rearrangements affecting PAX6 as determined by Sanger sequencing of the 50 regulatory elements,20 the exons and the intron-exon boundaries, nor deletions in, or around, PAX6 as determined by array comparative genomic hybridization and multiplex ligation-dependent probe amplification (MLPA; SALSA MLPA kit P219 PAX6, In vitrotek), respectively. To identify the molecular defect underlying GS, we subjected the genomic DNA (1 mg) from an index case subject (F1:II1) and her consanguineous parents of Tunisian origin (F1:I1 and F1:I2) to exome resequencing, using the SureSelect Human All Exon kit version 5 (Agilent). Each genomic DNA fragment was analyzed on a sequencer using the paired-end strategy and an average read length of 75 bases (Illumina HISEQ, Illumina). Image analysis and base calling were performed with Real Time Analysis (RTA) Pipeline v.1.9 with default parameters (Illumina). Sequences were aligned to the human genome reference sequence (hg19 assembly) and SNPs were called based on allele calls and read depth using the CASAVA pipeline (Consensus Assessment of Sequence and Variation 1.8, Illumina). Genetic variation annotation was performed by an in-house pipeline. We first focused our analysis on consensus splice-site changes, non-synonymous variants, and insertion and/or deletion in coding regions. Considering the rarity of the disease and the uncertain mode of inheritance, we assumed that the affected individual was either single heterozygous for a variant absent in parental DNA (dominant de novo model) and in the Exome Aggregation Consortium (ExAC), 1000 Genomes, dbSNP132, and Imagine de´ja`-vu databases, or homozygous for a variant found in the heterozygote state in parental DNA and absent or with a minor allele frequency < 0.001 in databases (recessive model). This filtering strategy pointed to one de novo and three homozygous variants (Tables S1 and S2). The de novo change occurred in KLK9

(c.88C>T [p.Arg30Cys]; GenBank: NM_012315.1), which encodes the kallikrein-related peptidase 9 (MIM: 605504). This variant was predicted to be deleterious both by PolyPhen-2 and SIFT softwares (Table S2), but its frequency in the ExAC database (7.5:10,000) was significantly higher than that of Gillespie syndrome, making unlikely its causality in the disease. Two out of the three homozygote variants were predicted to be benign according to PolyPhen-2 and SIFT and were filtered out (Table S2). The unique remaining candidate variant consisted of a nonsense mutation (c.4672C>T [p.Gln1558*]) in ITPR1 (GenBank: NM_001099952.2), which encodes the inositol 1,4,5-trisphosphate (IP3) receptor (IP3R), type 1 also known as IP3R1 (UniProt: Q14643; MIM: 147265) (Tables 2 and S2). Sanger sequencing confirmed homozygosity and single heterozygosity of the change in the affected child and her parents, respectively (Figure 1A). Three major ITPR1 mRNA isoforms are reported which in total represents 62 unique exons.21 Variant 1 (GenBank: NM_001099952.2) includes 59 exons, 57 of which are coding. Compared to variant 1, variant 2 (GenBank: NM_0022225) lacks exon 12 whereas isoform 3 (GenBank: NM_001168272.1) lacks exon 12, uses an alternate inframe splice site in intron 23, and includes three additional exons between variant 1 exons 39 and 40. Sanger sequencing of the 62 ITPR1 exons and intron-exon boundaries (Table S3) detected five additional unique ITPR1 mutations in the other four index case subjects (Table 2 and Figure 1A). F2:II1, born to consanguineous parents from Brazil,14 carried another homozygous nonsense mutation (c.2182C>T [p.Arg728*]) and F3:II1, born to non-consanguineous parents from mainland France, had two splicesite mutations (c.6366þ3A>T and c.6664þ5G>T). Segregation analysis confirmed the biparental transmission in the two families (Table 2 and Figure 1A). F4:II1, born to nonconsanguineous parents from mainland France, carried a 3-base pair (bp) deletion (c.7687_7689del [p.Lys2563del]) absent from parental DNA. (F4:II1 is the same individual as 1388_1388 who is described, with the same ITPR1 mutation, in the accompanying report by McEntagert et al.22) Likewise, F5:II1, born to consanguineous parents from the French Caribbean island La Guadeloupe, carried a missense change (c.7659T>G [p.Phe2553Leu]) absent in parental DNA. All variant numbers refer to ITPR1 isoform 1 (GenBank: NM_001099952.2). The six unique ITPR1 mutations were absent from all databases and were predicted to be deleterious according to the Alamut Mutation Interpretation Software, a decision support system for mutation interpretation based on Align DGVD, PolyPhen-2, SIFT, SpliceSiteFinder-like, MaxEntScan, NNSPLICE, and Human Splicing Finder (Table 2). ITPR1, ITPR2 (MIM: 600144), and ITPR3 (MIM: 147267) encode the three subtypes of the IP3-receptor family (ITPR1, ITPR2, and ITPR3 are also known as IP3R1, IP3R2, and IP3R3, respectively) that form large (>1 MDa), homoand heterotetrameric Ca2þ release channels localized

972 The American Journal of Human Genetics 98, 971–980, May 5, 2016

Table 1.

Clinical Features of the Individuals Affected with Gillespie Syndrome F1:II1

F2:II114

F3:II1

F4:II1

F5:II1

Age (years)

4.5

16

7.5

18

1.5

Gender

F

F

F

F

F

Family history

no

no

no

no

no

Consanguinity

yes

yes

no

no

yes

Prenatal history

persistent truncus arteriosus

no

no

no

no

Iris presentation

bilateral partial aniridia

bilateral partial aniridia

bilateral partial aniridia

bilateral partial aniridia

bilateral partial aniridia

Fundus aspect

unremarkable

unremarkable

unremarkable

unremarkable

unremarkable

OCT

unremarkable

unknown

unknown

unknown

unknown

Ocular Features

Neurologic Features Age at onset

birth

3 months

birth

birth

birth

Initial symptom

hypotonia with iris hypoplasia

hypotonia with iris hypoplasia

hypotonia with iris hypoplasia

hypotonia with iris hypoplasia

hypotonia with iris hypoplasia

Nystagmus

Yes

yes

yes

yes

yes

Ataxia

yes

yes

yes

yes

yes

Postural tremor

yes

unknown

yes

no

unknown

Slurred speech

not evaluable

yes

yes

yes

unknown

General hypotonia

yes

yes

yes

yes

yes

Pyramidal sign

no

unknown

no

no

no

Extrapyramidal sign

no

unknown

no

no

yes

Peripheral neuropathy

no

unknown

areflexia of lower limbs

no

no

Epileptic seizures

no

no

no

no

transitory myoclonus in the neonatal period

Intellectual disability

severe (uses two/ three words)

mild (phrases at 4 years)

moderate

no

unevaluable

Head control

delayed

12 months

18 months

18 months

10 months

Sit without support

3.5 years

3 years

18 months (with support)

3 years

no

Stand with support

4.5 years

yes

yes

3 years

no

Walk unassisted

no

no

no

16 years (a few steps only)

no

MRI findings

unremarkable at 1.5 months; marked cerebellar atrophy, thin corpus callosum, discrete ventricular dilatation at 4.5 years

reported unremarkable at ages 6 months and 8 years

moderate cerebellar atrophy at 2.5 years; marked cerebellar atrophy, at 8 years

unremarkable at 6 months; cerebellar atrophy at 4 years (vermis > hemispheres)

unremarkable at 3.5 months; marked cerebellar atrophy at 1.5 years

Other findings

hymeneal imperforation

facial dysmorphy



short 4th metatarsal

marked kyphosis, left pectoral agenesis

Cerebellar Signs

Motor Development

predominantly in membranes of endoplasmic reticulum (ER) Ca2þ stores.23,24 ITPR subtypes are ubiquitously expressed throughout tissues and all cell types express at least one subtype. Notably, the majority of cell types outside the

central nervous system express multiple isoforms.25–27 Although the expression of various ITPR subtypes is overlapping, the expression of certain subtypes is more enriched in a particular tissue to meet the developmental and

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Figure 1. Recessive and Dominant De Novo ITPR1 Mutations in Families with GS (A) Pedigree of the families and chromatograms showing the mutations at the genomic level. In families F1, F2, and F3, the affected individual is homozygous or compound heterozygous for parental mutations whereas in families F4 and F5, the affected individual is single heterozygous for a mutation absent in parental DNA, suggesting a de novo event. (B) Agarose gel separation of PCR products obtained from reverse-transcribed leucocyte mRNAs from individuals of family F3 and a control subject using the following primers: forward (exon 49), 50 -caatgcctcgaagttgctc-30 and reverse (exon 53) 50 -cgtgggttgtaacccgact-30 . A Formamide gel-loading buffer was used to avoid formation of heteroduplexes. The 605 bp wild-type fragment in the control is seen in all three members of the family. The presence of this band in II1 demonstrates that both the c.6366þ3A>T and c.6664þ5G>T allow the production of a wild-type transcript (chromatogram not shown). The mutant mRNA transcribed from the c.6366þ3A>T and c.6664þ5G>T alleles are seen as additional 543 bp and 615 bp fragments in I1 and II1 and I2 and II1, respectively. (C) Sanger sequencing of additional 543 and 615 bp fragments amplified from reverse-transcribed mRNAs from individuals of family F3 is consistent with the activation of cryptic splice sites in exon 50 and intron 52, respectively.

physiological needs of that tissue. For example, ITPR1 is specifically enriched in the nervous system whereas ITPR2 and ITPR3 are highly expressed in the digestive system. The structure of ITPR is generally divided into an N-terminal ligand-binding domain, a central transducing/modulatory domain, and a C-terminal channel domain23 (Figure 3A). IP3, generated after phospholipase C-mediated hydrolysis of phosphatidyl-inositol 4,5-bisphosphate, engages IP3 receptors and results in Ca2þ release from ER stores.28 Here we utilized biological samples obtained from GSaffected individuals and healthy control subjects to characterize ITPR1 mRNA levels and integrity. Further, we generated ITPR1 constructs corresponding to mutants identified in GS-affected individuals and assessed their activity in heterologous cell systems. The effect of the c.4672C>T nonsense mutation and the c.6366þ3A>T and c.6664þ5G>T splice-site mutations was assessed on mRNA from GS-affected individuals and control leucocytes or immortalized lymphocytes (biological samples from family 2 individuals were unavailable for analysis). Considering the c.4672C>T nonsense mutation, we measured the abundance of ITPR1 mRNAs by quantitative reverse-tran-

scription PCR (RT-qPCR) in immortalized lymphocytes of the homozygous index case subject of family F1 and two unrelated control subjects. Compared to control subjects, GS-affected individuals’ lymphoblasts had significantly reduced levels of ITPR1 mRNAs (mean relative expression levels of 0.74 5 0.05 and 0.21 5 0.01 in control and GSaffected individual cell lines, respectively; Figure S1), suggesting degradation through nonsense-mediated mRNA decay (NMD). According to the SpliceSiteFinder-like, NNSPLICE, and Human Splicing Finder prediction softwares, the c.6366þ 3A>T and c.6664þ5G>T mutations identified in family F3 reduced exons 50 and 52 splice-donor splicing scores below the scores of neighboring exonic and intronic cryptic splice-donor sites, respectively (Figure S2). To assess the activation of these cryptic splice-donor sites, reversetranscribed leucocyte mRNAs from F3:II1 and her parents F3:I1 and F3:I2 were amplified using primers designed in exons 49 and 53, respectively. Three PCR products were obtained from F3:II1 reverse-transcribed mRNA which Sanger sequencing revealed use of both the wild-type splice-donor sites and neighboring exonic and intronic

974 The American Journal of Human Genetics 98, 971–980, May 5, 2016

Figure 2. Brain MRI and Eye Findings in Individuals with Mutations in ITPR1 (A–J) MRIs were performed at 1.5 months and 4.5 years, 2.5 and 8 years, 3.5 months and 1.5 years, and 8 years of age in individuals F1:II1 (A–C), F3:II1 (D–F), F5:II1 (G–I), and F4:II1 (J) by means of the following modalities: sagittal T1 (A, B, D, E, G, H) or T2 (J) and axial T2 (C, F, I). In the first months of life, sagittal T1 images are unremarkable (A, G). The cerebellar atrophy appears with age as shown on sagittal T1 or T2 images by the moderate atrophy at age 1.5 years and 2.5 years (**) (D, H) and the marked cerebellar atrophy at 4.5 and 8 years (**) (B, D, J). In addition to cerebellar atrophy, individual F1:II1 presents with thin corpus callosum (*) (B) and discrete ventricular dilatation (***) (C). (K) Photograph of an eye of individual F5:II1 showing typical partial iris hypoplasia with iris strands extending onto the anterior lens surface.

cryptic splice-donor sites, respectively (Figures 1B and S2). Sequencing analysis showed that both the c.6366þ3A>T and c.6664þ5G>T mutant alleles encoded wild-type and mutant mRNAs. The mutant mRNA arising from the c.6366þ3A>T allele lacked the last 62 bp of exon 50 (r.[ ¼ , 6305_6366del; 6366þ3a>u]; Figure 1B) and was predicted to encode a truncated ITPR1 (p.Gly2102Valfs5*; Figure 3B). The mutant transcript arising from the c.6664þ5G>T allele retained the first 70 bp of intron 52 (r.[ ¼ , 6664_6665ins6664þ1_6664þ70; 6664þ5g>u]; Figure 1B) and was predicted to encode another truncated ITPR1 (p.Ala2221Valfs23*; Figure 3B). Consistent with parental genotypes, the shortened and enlarged mutant mRNA products were specifically detected in the father and mother leucocytes, respectively (Figure 1B). None of the aberrant splice products were detected in control individuals (Figure 1B). The truncation constructs encompass the ligand-binding domain and most of the central regulatory domain

but lack the channel domain. To evaluate the impact of these mutants on Ca2þ signaling, mammalian expression vectors co-expressing hcRed and human ITPR1 (ITPR1 WT), ITPR1 p.Gln1558*, ITPR1 p.Gly2102Valfs5*, or ITPR1 p.Ala2221Valfs23* were created. These truncation mutants (Figure 3B) were transiently transfected into human embryonic kidney cell line (HEK293). At 24 hr after transfection, immunoblot analysis was performed using antibodies that recognize an epitope located on the N-terminal region of ITPR1. The antibody detected a band corresponding to intact endogenous ITPR1 in all lysates (Figure S3A). The antibody also detected bands of appropriate molecular mobility in lysates from cells transfected with ITPR1 p.Gln1558*, ITPR1 p.Gly2102Valfs5*, and ITPR1 p.Ala2221Valfs23* but not in cells transfected with empty vector (Figure S3A). Furthermore, when expressed in DT40-3KO lymphoma cell line devoid of all three endogenous ITPRs,29,30 there was no Ca2þ release in response to IP3 generating stimuli (Figures S3B and S3C), consistent with the fact that these truncation mutants lack the channel domain and cannot mediate Ca2þ release. The truncation fragments represent the soluble part of the ITPR1 and contain the entire IP3-binding domain and most, if not all, of the modulatory domain that encompasses interaction sites for many regulatory factors. Accordingly, these mutants might impact endogenous intact ITPR activities by competing for or sequestering these factors. Theoretically, this scenario can occur in heterozygous states where intact ITPR1 (or even ITPR2 or ITPR3) and truncation mutants coexist in the same cells. To evaluate the impact of mutant constructs on Ca2þ signaling mediated by intact ITPRs, the truncation mutants were transiently transfected into a cell line designated as HEK-3KO cells. These cells were derived from HEK293 cells by disrupting all genes encoding IP3R subtypes using CRISPR/Cas9 editing system (Figure S4). Compared to DT40-3KO, these cells are easier to transfect and to manipulate during imaging. An empty vector (pDC315-hcRed) was used as a control. At 24 hr after transfection, cells were loaded with Ca2þ-sensitive dye Fura2AM and stimulated with trypsin to induce Ca2þ release from internal stores. Trypsin activates a protease-activated receptor-2 (PAR2), a G-protein-coupled receptor. Consistent with DT40-3KO data, cells expressing truncation mutants were refractory to stimulation, whereas cells expressing ITPR1 WT showed robust Ca2þ responses (Figures S4A and S4B). However, no significant difference between cells expressing ITPR1 WT alone and cells co-transfected with ITPR1 WT and various truncation mutants was observed (Figures S4A and S4B). Altogether, these data show that the truncation fragments do not affect—either positively or negatively—Ca2þ release mediated by endogenous IP3-receptors under these experimental conditions. Both p.Lys2563del and p.Phe2553Leu are conserved in all three ITPR subtypes and they are located in the distal portion of the sixth membrane-spanning helix or pore-lining helix that is predicted to project beyond the membrane

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Table 2.

ITPR1 Mutations Identified in the Individuals Affected with Gillespie Syndrome Allele 1 Pedigree

Parental Origin

Family 1

S, C

Family 2

S, C

Mutation (origin)

Predicted Effect

Mutation (origin)

Predicted Effect

Morocco

c.4672C>T (paternal)

p.Gln1558*

c.4672C>T (maternal)

p.Gln1558*

Brazil

c.2182C>T (paternal)

p.Arg728*

c.2182C>T (maternal)

p.Arg728*

c.6664þ5G>T (maternal)

p.Ala2221Valfs23*1

Family 3

S, NC

France

Family 4

S, NC

France

Family 5

S, NC

Allele 2

France

2

1

c.6366þ3A>T (paternal)

p.Gly2102Valfs5*

c.7687_7689del (de novo)

p.Lys2563del





c.7659T>G (de novo)

p.Phe2553Leu





Abbreviations are as follows: S, simplex family; C, consanguineous; NC, nonconsanguineous. 1 Predicted from mRNA analysis. Nucleotide positions according to the reference sequence GenBank: NM_001099952.2. The p.Phe2553Leu changes was predicted to be deleterious according to PolyPhen-2, SIFT, and Mutation Taster available through the Alamut Interpretation Software 2.0. 2 French Caribbean Island La Guadeloupe.

to the cytosol. This region of the receptor is believed to be critical for channel gating. To begin to assess the effect of these mutations on ITPR channel activity, ITPR1 WT was mutated by PCR to make ITPR1 p.Lys2563del and was transfected into DT40-3KO to generate stable cell lines. Figure S5A shows that anti-ITPR1 detected single bands in cells transfected with ITPR1 WT and ITPR1 p.Lys2563del constructs, corresponding to intact ITPR1, but no immunoreactivity was detected in DT40-3KO lysates. To measure intracellular Ca2þ release, cells were loaded with Fura-2AM and stimulated with 500 nM trypsin to activate the protease-activated receptor and induce IP3 formation. As shown previously, DT40-3KO cells do not respond to trypsin activation because they lack all three endogenous ITPRs.29 In contrast, stimulation of ITPR1 WT expressing cells induced a marked Ca2þ peak (Figures S5B and S5C). Cells expressing ITPR1 p.Lys2563del were not responsive, even to higher concentrations of trypsin (Figures S5B and S5C). In addition, the resting [Ca2þ] was not significantly different in DT40-3KO or in DT40-3KO expressing wild-type human ITPR1 or ITPR1 p.Lys2563del, indicating that these proteins do not perturb the integrity of intracellular stores (basal fluorescence 0.398 5 16.8 in ITPR1 WT and 0.379 5 4.34 in ITPR1 p.Lys2563del cells). In total, the above findings indicate that ITPR1 p.Lys2563del do not form functional calcium release channels when produced in isolation in DT40-3KO cells. To determine whether the assembly of ITPR1 p.Lys2563del into tetramers containing wild-type ITPR1 subunits affects the activity of channels formed, ITPR1 p.Lys2563del and ITPR1 WT were transfected either alone or in combination into HEK-3KO cells. When wildtype and mutant subunits are co-produced in the same cell, it would be predicted that they might associate or co-assemble into tetrameric channels. However, because the association or oligomerization is probably random, the number of wild-type and mutant subunits in the tetrameric channels formed in cells transfected with multiple cDNAs is difficult to predict. Initial experiments co-expressing ITPR1 p.Lys2563del and ITPR1 WT cDNA at 1:1 or 2:1 ratio cDNA indicated that mutant subunits did not

appear to markedly alter calcium release (Figures S6A and S6B). We reasoned that increasing the amount of mutant subunit would favor the formation of tetrameric channels incorporating mutant subunits. Therefore, to increase the likelihood that most assembled channels incorporate mutant subunits, an ITPR1 p.Lys2563del to ITPR1 WT ratio of 6:1 was used per transfection. Protein levels produced from the transfected constructs were verified via immunoblot analyses and Figure 4A shows that anti-ITPR1-detected single bands in HEK-3KO cells transfected with ITPR1 WT and ITPR1 p.Lys2563del corresponding to intact ITPR1, but no immunoreactive band was detected in HEK-3KO lysates transfected with empty vector. As expected, stimulation of HEK-3KO cells with 500 pM trypsin did not result in Ca2þ release, because they lack all three endogenous IP3Rs. In contrast, stimulation of HEK-3KO transfected with ITPR1 WT alone induced a marked Ca2þ peak (Figures 4B and 4C). Interestingly, co-expression of ITPR1 p.Lys2563del and ITPR1 WT cDNAs resulted in inhibition of both the amplitude of calcium signal (Figures 4B and 4C) and the percentage of responding cells (5.66% 5 1.46% in co-transfected cells compared to 20.71% 5 2.11% in cells transfected with ITPR1 WT only). The simplest interpretation of these data is that ITPR1 channels incorporating one or more mutant subunits are not functional. The fact that more mutant cDNA was required to see the inhibitory effect might suggest that numerous subunits are required to assemble into a tetrameric channel to render it dysfunctional. Alternatively, it is possible that the mutant subunit does not favor oligomerization or association with the wild-type subunits and thus a higher proportion of mutant subunits are necessary to increase the probability of this association. Here, we report that both recessive and dominant ITPR1 mutations cause Gillespie syndrome. ITPR1 is a predominant isoform in the brain among the three types of ITPRs and is strongly expressed in cerebellar Purkinje cells.31 Mice with complete homozygosity for Itpr1 ablation suffer from severe epilepsy and ataxia and die either in utero or before weaning.32 Consistently, ITPR1 mutations have been reported to cause cerebellar diseases including

976 The American Journal of Human Genetics 98, 971–980, May 5, 2016

Figure 3. Schematic Diagrams Depicting Full-Length ITPR1 and ITPR1 Mutation Associated with Human Diseases (A) The panel displays the functional domains of ITPR1. Shown are point mutations associated with spinocerebellar ataxia or ataxic cerebellar palsy (blue) or Gillespie syndrome not investigated in this study (red). (B) The lower four panels show the truncation mutants and p.Lys2563del mutant expressed in heterologous cell systems for functional analysis. The black boxes of the ITPR1 p.Gly2102Valfs5* and ITPR1 p.Ala2221Valfs23* mutants depict the VGSA and VGFQRISNNTERHSSGISQTWA amino acid sequences introduced by the c.6366þ3A>T and c.6664þ5G>T, respectively.

late-onset spinocerebellar ataxia type 15 (SCA15 [MIM: 606658]),33 congenital nonprogressive spinocerebellar ataxia and mild cognitive impairment (SCA29 [MIM: 117360]),34 infantile-onset cerebellar ataxia with mild cognitive deficit,35 and childhood-onset ataxic cerebellar palsy with moderate intellectual disability36 (see ITPR1 schematic diagram in Figure 3A). Affected individuals had similar iris anomalies and neonatal ataxia with progressive cerebellar atrophy (Figure 2). Moderate to severe intellectual disabilities were noted in the three individuals with recessive mutations (F1:II1, F2:II1, and F3:II1; Table 1). In contrast, the affected individual F4:II1 aged 18 years and harboring the de novo c.7687_7689del mutation was reported to have normal intelligence (Table 1). It will be interesting to follow the intellectual development of the second individual with a heterozygous de novo mutation to determine whether some genotype-phenotype correlations can exist (F5:II1, age 1.5 years; c.7659T>G). All individuals with adult-onset SCA15 but one show heterozygosity for partial 3 pter deletions encompassing ITPR1 (or at least the 50 region of the gene).33 The four ITPR1 mutations identified in GS-affected families F1–F3 are predicted to introduce a premature translation termination codon. Intriguingly, none of the heterozygous parents in families F1 and F3 (F2 unavailable) had neurologic symptoms, as assessed by clinical examination and 3 Tesla cerebral RMN imaging (Figure S7). It cannot be excluded that some or all these heterozygous individuals whose age ranged from 35 to 44 years might develop cerebellar symptoms later in life. However, it is also quite possible that they harbor mutations that are not as detrimental as gene ablation. The fact that homozygosity for null alleles is incompatible with a prolonged life in Itpr1/ mouse32 and the Italian Spinone with spinocerebellar ataxia37 but not in individuals with Gillepsie syndrome is consistent with this hypothesis. In addition, mRNA analysis in family F3 demonstrated that both parents carry an intronic mutation that allows for the production of the wild-type mRNA

from the mutant allele. The Purkinje cells of these individuals are likely to produce more ITPR1 than that of individuals with adult-onset cerebellar ataxia due to gene ablation, thereby bypassing the Ca2þ release defect. Along the same lines, premature termination codon (PTC) selfcorrecting mechanisms might have occurred in the Purkinje cells of individuals F1:I1, F1:I2, F2:I1, and F2:I2 harboring the c.4672C>T and c.2182C>T truncating mutations, respectively. These mechanisms include translational read-through that has recently been reported to be more abundant than expected in higher species, including human.38 Nonsense-associated altered splicing (NAS)39–42 and basal exon skipping (BES),43,44 which allow selective exclusion of an in-frame exon with a premature termination codon, are other correcting mechanisms that have recently been shown to modulate disease severity in human.43–45 Although experiments presented here did not show a clear impact of the p.Gln1558*, p.Gly2102Valfs5*, and p.Ala2221Valfs23* truncated ITPR1s on the Ca2þ release activity of channels assembled from wild-type ITPRs in HEK293 cells, it cannot be formally excluded that mutant proteins could functionally interact in vivo with wild-type ITPR channels and modulate their function. Unlike adults affected with SCA15 who consistently carry null ITPR1 alleles, children with early-onset cerebellar ataxia and moderate cognitive disability consistently carry missense ITPR1 mutations affecting the conserved IP3-binding domain or modulatory domain. Because ITPR channels can be homo- or heteromeric, it can be envisioned that ITPR channels are assembled from various combination of mutant ITPR1 (and wild-type in case of heterozygous state) as well as ITPR2 and ITPR3 subunits. How the incorporation of mutant ITPR1 affects channel activity probably depends on the severity or location of the mutation as well as on the stoichiometry of mutant subunits assembled within the tetrameric channel. It has been suggested that these mutations, most of which

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Figure 4. Production of the ITPR1 p.Lys2563del Mutant in HEK-3KO and Evaluation of Its Effect on the Formation of ITPR1 Channels (A) HEK-3KO cells were transfected as indicated. Lysates were prepared from transfected cells, quantified and equivalent amounts of proteins were separated on 4% SDS-PAGE and processed in immunoblots using a rabbit polyclonal antibody raised against the most carboxyl-terminal 19 amino acid residues of ITPR1 (generated on demand; Pocono Rabbit Farms and Laboratories). A representative experiment is shown. Lysates from WT-HEK, naive HEK293 cells, were used as a control. (B) HEK-3KO cells grown on coverslips were transfected with 2.4 mg empty vector or cDNA coding for ITPR1 p.Lys2563del either alone or with 0.4 mg ITPR1 WT as indicated. The total amount of DNA was kept constant by adding empty vector. At 24 hr after transfection, cells were loaded with Fura-2AM and stimulated with 500 pM trypsin to induce IP3 formation. Ca2þ release was determined as a change in the 340/380 fluorescence ratios. Fluorescent ratios were normalized to the ratios from the first 5 s after the start of run. Shown are representative traces. (C) Bar graphs depict the average maximum change over basal 340/380 fluorescence ratios resulting from trypsin stimulation of cells expressing the indicated constructs. Experiments were repeated at least three times. Data are presented as mean 5 SE.

occurred de novo, might exert a dominant-negative effect, possibly disrupting cellular calcium homeostasis.35 Interestingly, the two de novo mutations identified in individuals with GS affect the distal portion of the S6 pore-lining helix that is pivotal for channel gating and where no disease-associated mutation has been reported to our knowl-

edge. Co-expression of the ITPR1 p.Lys2563del mutant construct with the wild-type counterpart resulted in altered calcium release, strongly supporting the view that the mutation exerts a dominant-negative effect. Studies wherein the exact subunit composition of tetrameric channels is predetermined46 are required to define how many ITPR1 p.Lys2563del subunits per channel are necessary to exert the dominant-negative effect or disable the channel activity. To our knowledge, no abnormality in eye development has been described in heterozygous ITPR1-related individuals with SCA, the Itpr1 knock-out mouse,32,47 the naturally occurring Itpr1opt/opt mouse,48 or the Italian Spinone with spinocerebellar ataxia.37 Given that minor ocular symptoms could be overlooked, ophthalmological examinations including gonioscopy and funduscopy were undertaken in parents of individuals with GS (families 1 and 3) and in two brothers with SCA15 harboring a large gene deletion.49 None of the individuals had any ocular symptom, supporting the view that expression of one wild-type allele is sufficient to allow proper eye development in these individuals. There is accumulating evidence for the involvement of Ca2þ in developmental events and gene expression. Furthermore, ITPRs have proven important in early embryogenesis, in particular to determine the dorsoventral axis formation.50 The role of ITPR1 in eye development has not been documented thus far. In the Drosophila, however, it has been reported that the unique IP3R isoform (DmIP3R) is detected ubiquitously throughout development51–53 and is essential for embryonic and larval development. In Ip3r mutants, maternal Ip3r mRNA is sufficient for progression through the embryonic stages, but larval organs show asynchronous and defective cell divisions and imaginal discs arrest early and fail to differentiate.54 Adult mosaic Drosophila containing homozygous Insp3 mutant photoreceptors in an otherwise wild-type genetic background have been created, the analysis of which revealed disruptions of the onomatidal architecture and retinal degeneration, demonstrating a role in eye function.54 Hopefully, further studies in vertebrate animals will allow researchers to substantiate the role of ITPR1 in the development of the eye. In summary, we report on biallelism or single heterozygosity for ITPR1 mutations in 5/5 index case subjects with Gillespie syndrome. These findings demonstrate the long-suspected co-existence of autosomal-recessive and autosomal-dominant patterns of inheritance of Gillespie syndrome and further extend the spectrum of ITPR1 deficiencies in the eye.

Supplemental Data Supplemental Data include seven figures and three tables and can be found with this article online at http://dx.doi.org/10.1016/j. ajhg.2016.03.004.

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Acknowledgments We are grateful to the families for their participation in the study. This work was supported by grants from the Fe´de´ration des Aveugles de France (FAF) to L.F.T., Retina France to J.M.R., and Geniris to J.M.R. and P.C. K.J.A. and D.I.Y. were supported by NIH grants DE19245 and DE14756. Received: November 16, 2015 Accepted: March 7, 2016 Published: April 21, 2016

Web Resources 1000 Genomes, http://browser.1000genomes.org CONDOR, http://condor.nimr.mrc.ac.uk/ dbSNP, http://www.ncbi.nlm.nih.gov/projects/SNP/ ExAC Browser, http://exac.broadinstitute.org/ GenBank, http://www.ncbi.nlm.nih.gov/genbank/ NHLBI Exome Sequencing Project (ESP) Exome Variant Server, http://evs.gs.washington.edu/EVS/ OMIM, http://www.omim.org/ RefSeq, http://www.ncbi.nlm.nih.gov/RefSeq UCSC Genome Browser, http://genome.ucsc.edu UniProt, http://www.uniprot.org/help/uniprotkb

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Recessive and Dominant De Novo ITPR1 Mutations Cause Gillespie Syndrome.

Gillespie syndrome (GS) is a rare variant form of aniridia characterized by non-progressive cerebellar ataxia, intellectual disability, and iris hypop...
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