Extending the spectrum of CLRN1- and ABCA4-associated inherited retinal dystrophies caused by novel and recurrent variants using exome sequencing.

BACKGROUND: Inherited retinal dystrophies (IRDs) are characterized by extreme genetic and clinical heterogeneity. There are many genes that are known to cause IRD which makes the identification of the underlying genetic causes quite challenging. And in view of the emergence of therapeutic options, it is essential to combine molecular and clinical data to correctly diagnose IRD patients. In this study, we aimed to identify the disease-causing variants (DCVs) in four consanguineous Jordanian families with IRDs and describe genotype-phenotype correlations.
METHODS: Exome sequencing (ES) was employed on the proband patients of each family, followed by segregation analysis of candidate variants in affected and unaffected family members by Sanger sequencing. Simulation analysis was done on one novel CLRN1 variant to characterize its effect on mRNA processing. Clinical evaluation included history, slit-lamp biomicroscopy, and indirect ophthalmoscopy.
RESULTS: We identified two novel variants in CLRN1 [(c.433+1G>A) and (c.323T>C, p.Leu108Pro)], and two recurrent variants in ABCA4 [(c.1648G>A, p.Gly550Arg) and (c.5460+1G>A)]. Two families with the same DCV were found to have different phenotypes and another family was shown to have sector RP. Moreover, simulation analysis for the CLRN1 splice donor variant (c.433+1G>A) showed that the variant might affect mRNA processing resulting in the formation of an abnormal receptor. Also, a family that was previously diagnosed with nonsyndromic RP was found to have Usher syndrome based on their genetic assessment and audiometry.
CONCLUSION: Our findings extend the spectrum of CLRN1- and ABCA4-associated IRDs and describe new phenotypes for these genes. We also highlighted the importance of combining molecular and clinical data to correctly diagnose IRDs and the utility of simulation analysis to predict the effect of splice donor variants on protein formation and function.

INTRODUCTION

Inherited retinal dystrophies (IRDs) are a group of clinically and genetically heterogeneous inherited retinal disorders with an estimated worldwide prevalence of about 1:3,000 (Hartong, Berson, & Dryja, 2006; Wang et al., 2018). IRDs can be either syndromic or nonsyndromic. Of the nonsyndromic IRDs, retinitis pigmentosa (RP) is the most common (Chizzolini et al., 2011; Werdich, Place, & Pierce, 2014; Wright, Chakarova, Abd El‐Aziz, & Bhattacharya, 2010), whereas Usher syndrome is the most common syndromic IRD that is characterized by hearing loss in addition to RP features (Cosgrove & Zallocchi, 2014; Mathur & Yang, 2015).

To date, over 300 genes were shown to be associated with IRDs (https://sph.uth.edu/Retnet/), and all the genetic inheritance patterns are involved (Astuti et al., 2018; Kajiwara, Berson, & Dryja, 1994; Wang et al., 2018). Not only variants in the same gene but also the same genetic variants have been shown to cause variable phenotypes in inter‐ and intrafamilial cases (Fahim, Daiger, & Weleber, 1993).

This broad genetic and clinical complexity warrants highly efficient and high‐throughput genetic testing approach to uncover the underlying genetic causes and reach to the accurate diagnosis of IRDs.

CLRN1 which is located on chromosome 3q25.1 and codes for clarin‐1 protein that has functions in the retina and the cochlea (Adato et al., 2002; Geng et al., 2009). Therefore, different DCVs in this gene were shown to cause RP and Usher syndrome type III in previous studies (Fields et al., 2002; Khan et al., 2011). ABCA4 is expressed in retinal photoreceptors and was implicated in different IRDs like cone‐rod dystrophy and Stargardt's disease (Jiang et al., 2016; Molday, Zhong, & Quazi, 2009).

Exome sequencing (ES) is an affordable technology that covers the coding regions and splice‐site variants and is powerful tool for identification of disease‐causing variants (DCVs) for Mendelian diseases (Dixon‐Salazar et al., 2012; Goldfeder, Wall, Khoury, Ioannidis, & Ashley, 2017; Petersen, Fredrich, Hoeppner, Ellinghaus, & Franke, 2017). Although the coding region represents only 1%–2% of the human genome, it carries 85% of the disease‐related variants (Gilissen, Hoischen, Brunner, & Veltman, 2012).

Therefore, in this study, we aimed to identify the DCVs in four Jordanian pedigrees with autosomal recessive IRDs using ES. Moreover, full ophthalmic examination was performed for genotype–phenotype correlations.

MATERIALS AND METHODS

Study subjects and clinical examination

This study was performed in accordance with the tenets of the Declaration of Helsinki and approved by the Institutional Review Board of the Cell Therapy Center, Amman, Jordan. Written informed consent was obtained from all participants prior to their inclusion in the study.

Thirteen patients from four unrelated Jordanian families with inherited visual problems were recruited into the study (Figure 1). Only family members labeled with a number in the pedigrees were included.

Figure 1

Pedigrees of the four Jordanian families participating in the study. Squares, circles, and dashes represent males, females, and deceased individuals, respectively. Arrows indicate probands. Solid symbols refer to affected individuals and open symbols to normal or carrier individuals. The two dashed squares in family F1 pedigree divide it into F2‐A and F2‐B. Double horizontal lines indicate consanguinity. M/M, homozygous mutated; M/W, heterozygous mutated; W/W, homozygous wild type. Zygosity for individuals III‐2 and III‐3 in family F1‐A is written in two lines, the above one for CLRN1 (c.433+1G>A), and the lower one for ABCA4 (c.5460+1G>A)

A detailed history was taken from all patients including their age at onset, first symptom, progression, and other co‐morbidities. All affected individuals underwent standard ophthalmic examination, including Snellen chart uncorrected visual acuity, best‐corrected visual acuity (BCVA), slit‐lamp biomicroscopy (Haag‐Streit BM 900, Switzerland), fundus photography (Optos 200Tx, Scotland), Optical Coherence Tomography (OCT), Keratometry (Oculus Pentacam Typ70700, Germany), and full‐field Electroretinography (ffERG) (Roland consult color Ganzfeld Q450 C, Germany). ERG recordings were performed according to the 2015 update of the International Society for Clinical Electrophysiology of Vision Standards (ISCEV) (McCulloch et al., 2015). Where necessary, pure‐tone audiometry was done to evaluate hearing status for suspected Usher syndrome cases.

Exome sequencing and bioinformatics analysis

Genomic DNA was isolated from peripheral blood samples of the patients and participating family members using QIAprep Spin Miniprep Kit. ES was performed on the proband patients of each family (Figure 1) by Partners HealthCare Personalized Medicine (PPM), Harvard, USA. Briefly, extracted DNA was sheared to 150–200 bp fragments. Exome enrichment was performed using Agilent SureSelect Human All Exon V5 kit. DNA was sequenced using 100 bp paired‐end reads on Hiseq 2500 platform (Illumina Inc.). Raw data were processed by Illumina base‐calling software 1.7 using default parameters. The sequencing reads were aligned to the NCBI reference sequence (GRCh37), using the Burrows‐Wheeler Aligner (BWA), and variant calls were made using the Genomic Analysis Tool Kit (GATK). The detected variants were annotated and filtered against four databases (i.e., NCBI CCDS [http://www.ncbi.nlm.nih.gov/CCDS/CcdsBrowse.cgi], RefSeq [http://www.ncbi.nlm.nih.-gov/RefSeq/], Ensembl [http://www.ensembl.org], and Encode [http://genome.ucsc.edu/ENCODE]).

Variants filtration and prioritization

To detect the candidate DCVs, multistep filtration approach was applied using Illumina basespace variant interpreter tool (https://variantinterpreter.informatics.illumina.com/) (Table S1). Variants were filtered based on (a) quality (total read depth ≥10), (b) genomic position (within exons and flanking intronic regions), (c) genes of interest (list of 186 genes) (Azab et al. 2019), (d) allele frequencies; we only included variants with MAF <0.01 in EXAC, gnomAD, 1000 Genomes project, or ESP, and (e) mode of inheritance; our four pedigrees follow an autosomal recessive mode of inheritance, therefore only homozygous or compound heterozygous variants were included (Table S1). Ensembl was also used as a reference dataset of population‐based allele frequencies.

To assess the pathogenicity of the identified variants, we used a combination of computational tools. The possible damaging impacts of missense variants on protein structure and/or function were predicted using SIFT (http://sift.jcvi.org), PolyPhen‐2 (http://genetics.bwh.harvard.edu/pph2), Provean (http://provean.jcvi.org/index.php), CONDEL (http://bg.upf.edu/fannsdb/) and MutationTaster2 (Schwarz, Cooper, Schuelke, & Seelow, 2014). Evolutionary conservations across species for missense variants were also assessed using Alamut Visual Prediction Software. For the impact of splice site variants, the following prediction tools were used: Human Splicer Finder (http://www.umd.be/HSF/), NNSPLICE (http://www.fruitfly.org/seq_tools/splice.html), SSF, and MaxEnt.

Variants validation and segregation analysis

Sanger sequencing was performed to verify the candidate DCVs identified by ES and for segregation analysis in all family members available for testing. Primers flanking the candidate loci were designed using Primer3 software and synthesized by IDT (S) (Table S2). The extracted DNA was PCR‐amplified by Platinum™ PCR SuperMix (Invitrogen), and purified by GeneJET PCR purification kit (Invitrogen). Sequencing was performed with the BigDye Terminator V3.1 Cycle Sequencing kit (Applied Biosystems) on an ABI 3500 genetic analyzer. The sequences data were analyzed by SeqA software (Applied Biosystems) and Chromas Pro software (Technolysium LTD). The variants were then classified as benign, likely benign, variant of unknown significance, likely pathogenic, and pathogenic according to the American College of Medical Genetics and Genomics (ACMG) standards and guidelines for the interpretation of sequence variants (Richards et al., 2015). Variants were submitted to ClinVar database (https://www.ncbi.nlm.nih.gov/clinvar/).

Simulation of CLRN1 (c.433+1G>A) splice donor variant

Simulation analysis including docking and molecular dynamic simulation was used to assess the effect of CLRN1 (c.433+1G>A) variant on mRNA splicing. The detailed explanation of the experiment methods is available in the supporting information.

RESULTS

Genetic findings

Aiming to identify the DCVs, ES was performed on five probands with autosomal recessive IRDs. Around six Gb of data was obtained, on average 80% of the target region was at least 10X covered, and about 70k variants were identified in each proband. To identify the DCV in each family, we did a multistep filtration approach as described in the methods section. In total, four pathogenic variants in CLRN1 and ABCA4 were identified in the four families (Table 1), in which two were novel CLRN1 variants and two were recurrent ABCA4 variants.

Table 1

Candidate variants identified in four inherited retinal dystrophy pedigrees by exome sequencing

Family Number	Gene	Variant Coordinate (hg19)	Transcript (Ensembl)	dbSNP ID	Variation				Total MAF (Gnomad)	ClinVar	Classification of variantsa	Associated phenotype
					HGVS cDNA	HGVS aa	Type	Zygosity
1‐A & 4	ABCA4	chr1:94480098	ENST00000370225.3	rs61753030	c.5460+1G>A	NA	Splice donor	Hom	0.00003185	Pathogenic	Pathogenic	RP in F1‐A and CRD in F4
1‐B	CLRN1	chr3:150659368	ENST00000327047.5	rs201205811	c.433+1G>A	NA	Splice donor	Hom	0.000007970	NA	Pathogenic	Usher syndrome type III
2	CLRN1	chr3:150659479	ENST00000327047.5	NA	c.323T>C	p.Leu108Pro	Missense	Hom	NA	NA	Uncertain Significance	Usher syndrome type III
3	ABCA4	chr1:94528780	ENST00000370225.3	rs61748558	c.1648G>A	p.Gly550Arg	Missense	Hom	0.000003977	Likely Pathogenic	LikelyPathogenic	CRD

Novel variants are highlighted in bold. Abbreviations: CRD, cone‐rod dystrophy; Hom, homozygous; MAF, minor allele frequency; NA, not available; RP, retinitis pigmentosa.

Classification is based on the ACMG guidelines for interpretation of sequence variants (See the text).

Family F1 was a large extended family that has two probands, one from each side of this pedigree (F1‐A and F1‐B) (Figure 1). ES revealed ABCA4 splice site donor variant (c.5460+1G>A) in F1‐A. Interestingly, this variant (c.5460+1G>A) was also found in family F4. On the other hand, for F1‐B novel splice site donor variant in CLRN1 (c.433+1G>A) was detected. The second novel CLRN1 variant was a missense variant (c.323T>C, p.Leu108Pro) identified in family F2. In F3, a missense variant in ABCA4 (c.1648G>A, p.Gly550Arg) was identified (Table 1; Figure S1).

ABCA4 variants

The splice site donor variant (c.5460+1G>A) in families F1‐A and F4 is predicted by the different in silico tools to affect splicing causing exon skipping resulting in out‐of‐frame transcripts (Table 2). This variant has been reported previously in the literature to be pathogenic (Rivera et al., 2000; Ścieżyńska et al., 2016; Xiong et al., 2015). The missense ABCA4 (c.1648G>A) variant introduces a substitution from glycine to arginine at amino acid residue 550 (p.Gly550Arg). It is located in a conserved genomic region and in silico analyses tools predicted it as pathogenic (Table 2). Moreover, this variant is already reported in ClinVar as likely pathogenic.

Table 2

In silico prediction of the identified candidate pathogenic variants

Gene	Variant (HGVS cDNA)	Variant type	Missense prediction					Splice site prediction				Overall prediction of pathogenicity
			SIFT	Polyphen	CONDEL	PROVEAN	MutationTaster2	SSF	MaxEnt	NNSPLICE	HSF
CLRN1	c.433+1G>A	Splice donor	−	−	−	−	−	+	+	+	+	Pathogenic
ABCA4	c.5460+1G>A	Splice donor	−	−	−	−	−	+	+	+	+	Pathogenic
ABCA4	c.1648G>A	Missense	D	PD	D	DE	Disease causing	−	−	−	−	Pathogenic
CLRN1	c.323T>C	Missense	D	PD	D	NA	Disease causing	−	−	−	−	Pathogenic

Abbreviations: D, damaging; DE, deleterious; NA, not available; PD, probably damaging.

CLRN1 variants

All splicing‐related in silico platforms we utilized unanimously predicted the novel CLRN1 splice site variant (c.433+1G>A) to break the splice donor site (Table 2). To further assess the effect of this novel variant, Sanger sequencing revealed that the four identified candidate variants completely segregated with disease phenotypes following an autosomal recessive inheritance pattern (i.e., DCVs were homozygous in affected individuals, heterozygous in their parents, and normal/heterozygous in unaffected relatives) (Figure 1). According to the ACMG guidelines, we classified the two novel CLRN1 ((c.433+1G>A) and (c.323T>C)) as pathogenic and uncertain significance, respectively. We also confirmed the previous ClinVar classification of ABCA4 (c.5460+1G>A) and ABCA4 (c.1648G>A, p.Gly550Arg) as pathogenic and likely pathogenic, respectively.

Simulation analysis

To characterize the effect of the splice‐donor variant (c.433+1G>A) in CLRN1 on splicing and the function of the transmembrane protein clarin‐1, simulation analysis was performed. Docking simulation was used to assess the interaction between U1 snRNP and RNA splice site in both the wild‐type and mutated forms. Figure 2a shows the best oriented pose of the wild‐type RNA and U1 snRNP. Molecular dynamics simulation showed that mutating G21 to A in the best‐docked pose between wild‐type RNA and U1 snRNP would result in loss of noncanonical interactions between certain atoms in wild‐type RNA and U1 snRNP. Figure 2b shows the interacting atoms between G21 (from the wild‐type RNA strand) with C8 (from U1 snRNP) as annotated spheres. These noncanonical interactions include hydrogen bonds, and interactions among C–H and O/N groups. These interacting atoms have not been found when mutating G21 to A, indicating the importance of G21 for such binding interactions (Figure 2c). Further details of the simulation analysis results are illustrated in the supporting information.

Figure 2

Simulation analysis of the splice donor variant results CLRN1 (c.433+1G>A). (a) Best oriented docked pose of the wild‐type RNA with U1 snRNP. The wild‐type RNA and the U1 snRNP are shown in green and red colors, respectively. The wild‐type polymorphism (Guanine) is shown as G21, it is binding with Cytosine 8 of U1 snRNP, (b) The main noncanonical interacting atoms between Guanine (G21) from wild‐type RNA with Cytosine (C8) from U1 snRNP. (c) Noncanonical interactions became less frequent when G21 is mutated to A. Noncanonical interacting atoms are shown as annotated spheres

Clinical results

Thirteen patients from four consanguineous Jordanian families with autosomal recessive visual problems were recruited to this study for further evaluation and molecular diagnosis. The detailed clinical information of all patients is summarized in Table 3.

Table 3

Clinical data of all participated inherited retinal dystrophy patients in the study

Family	Affected members	Age		First symptom	BCVA		ffERG		Fundus examination		Slit‐lamp biomicroscopy	OCT		Keratoconus	Hearing status (audiometry)	Inherited retinopathy phenotype
		At exam	Onset		OD	OS	Scotopic response	Photopic response	OD	OS		OD	OS
1A	IV‐7	49	10	Low VA	CF at 1.25 m	0.05	Severely reduced	Severely reduced	Typical RP	Typical RP	No cataract	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	Topographic keratoconus in both eyes	Normal	RP
	IV‐8	37	10	Low VA	HM	HM	Severely reduced	Severely reduced	Typical RP	Typical RP	No cataract Horizontal nystagmus	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	No keratoconus	Normal	RP
	V‐6	11	9	Low VA	0.1	0.1	Slightly reduced	Normal	Atypical RP	Atypical RP	No cataract	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	No keratoconus	Normal	RP
	V‐7	12	6	Low VA	0.1	0.1	Slightly reduced	Normal	Atypical RP	Atypical RP	No cataract	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	No keratoconus	Normal	RP
1B	IV‐16	48	5	Nyctalopia	HM	CF closely	Severely reduced	Severely reduced	Typical RP	Typical RP	PSCC in both eyes	Blurred OCT due to cataract	Blurred OCT due to cataract	No keratoconus	Bilateral moderate‐to‐severe hearing loss	USH III
	IV‐17a	54	7	Nyctalopia	No LP	0.4	Severely reduced	Severely reduced	Typical RP	Typical RP	Had cataract surgery in the left eye	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	OD: cannot comment due to trauma OS: no keratoconus	Bilateral moderate‐to‐severe hearing loss	USH III
	IV‐18	44	6	Nyctalopia	CF at 1 m	CF at 1.5 m	Severely reduced	Severely reduced	Typical RP	Typical RP	Posterior polar cataract with nuclear sclerosis	Mild central atrophic maculopathy with no CME	Mild central atrophic maculopathy with no CME	No keratoconus	Bilateral mild‐to‐moderate hearing loss	USH III
	IV‐19	48	7	Nyctalopia	CF at 2 m	CF at 2 m	Severely reduced	Severely reduced	Typical RP	Typical RP	PSCC in both eyes	Mild central atrophic maculopathy with no CME	Poor quality due to cataract	Keratometry not available	Not available	USH III
2	II‐3	41	36	Nyctalopia	0.3	CF at 2 m	Severely reduced	Severely reduced	Typical RP	Typical RP	PSCC in both eyes	Mild central atrophic maculopathy with no CME	Mild central atrophic maculopathy CME with epiretinal membrane	No keratoconus	Bilateral mild hearing loss	USH III
3	IV‐4	NA	11	Low VA	CF at 1 m	0.1	OD: severely reduced OS: moderately reduced	Severely reduced	Atypical RP	Atypical RP	Congenital cataract in the left eye	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	No keratoconus	Normal	CRD
4	IV‐4	19	8	Low VA	CF at 2 m	CF at 2 m	Slightly reduced	Moderately reduced	Atypical RP	Atypical RP	Had cataract surgery	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	No keratoconus	Normal	CRD
	IV‐6	24	8	Low VA	CF at 2 m	CF at 2 m	Moderately reduced	Severely reduced	Atypical RP	Atypical RP	No cataract	Central atrophic maculopathy with no CME	Central atrophic maculopathy with no CME	No keratoconus	Normal	CRD

Typical RP denotes the presence of the classical triad (attenuated arterioles, disc pallor, and bone spicules) in fundus photography. Atypical RP means that a part of the triad is missing.

Abbreviations: BCVA, best‐corrected visual acuity; CF, counting fingers; CME, cystoid macular edema; CRD, cone‐rod dystrophy; ffERG, full‐field electroretinography; HM, hand motion; LP, light perception; NA, not available; OCT, optical coherence tomography; OD, right rye; OS, left eye; PSCC, posterior subcapsular cataract; RP, retinitis pigmentosa; USH III, Usher syndrome III.

This patient had cataract surgery in the left eye, and his right eye was lost due to trauma.

Clinical consequences of the identified CLRN1 variants

Two families (F1B and F2) were found to have their DCVs in CLRN1 gene (Table 1) (Figure 1). The patients' ages in F1B and F2 ranged between 35 and 54 years old at the time of examination. Almost all of them were legally blind at time of examination (best‐corrected visual acuity <3/60 (0.05) in the better‐seeing eye, according to WHO) (World Health Organization, 2016). Patients in F1‐B started to have night blindness during early childhood, followed by progressive constriction of peripheral vision and central vision later. Fundus examination showed typical RP features (Figure 3; Figure S2). Clinically, F1‐B patients had no associated symptoms with RP and were thus considered to have nonsyndromic RP. However, ES results showed CLRN1 as disease causing and this gene has been classically associated with Usher syndrome. This warranted further audiometric evaluation to rule in/out hearing problems.

Figure 3

Optical coherence tomography (OCT) (a) and (b) fundus photography of probands of all families (OD and OS). White arrows show central atrophic maculopathy with no cystoid macular edema, blue arrow indicates waxy pallor of optic disc, and red arrow indicates attenuation of retinal arterioles. Fundus images also show bone spicules

Audiometry showed different degrees of bilateral sensorineural deafness in all patients, confirming the diagnosis of Usher syndrome (Figure S3). Patients' hearing was normal at birth, their speech is normal, and they do not have balance problems, all suggesting type III Usher syndrome (Mathur & Yang, 2015). Being at late stages, ffERG in all patients showed severe reduction in both scotopic and photopic responses (Figure 4; Figure S5). OCT revealed variable degrees of central macular atrophy (Figure 3; Figure S4). Besides, they have cataract in both eyes (Table 3).

Figure 4

Electroretinograms (ERGs) of both eyes in all probands including the standard six recordings denoted beside each row

On the other hand, patient F2: II‐3 had a late‐onset RP (at the age of 36), unlike patients in F1‐B family. Although her ffERG showed severe reduction in both photopic and scotopic responses in both eyes, her visual acuity is 0.3 in the better‐seeing eye (Figure 4; Figure S5). Audiometry was also done for this patient revealing mild bilateral sensorineural deafness. Her affected sister (F2: II‐3) declined to participate in the clinical assessments.

Clinical consequences of the identified ABCA4 variants

The molecular assessment for families; F1‐A, F3, and F4 revealed pathogenic variants in ABCA4 (Figure 1). Patients in F1‐A and F4 share the same DCV (ABCA4, c.5460+1G>A) (Table 1). F1‐A has four affected members; two older individuals (IV‐7 and IV‐8) whose both night vision and visual acuity are affected, and two younger individuals (V‐6 and V‐7) who are still at an early stage of the disease (Figure 1; Table 3). The presenting symptom for all of them was decreased visual acuity in the first decade of life. ffERG showed severe reduction in both photopic and scotopic responses for the patients (F1‐A: IV‐7 and IV‐8) (Figure 4; Figure S5). Whereas, those of the younger siblings (F1‐A: V‐6 and V‐7) showed slight impairment of scotopic responses only. Hence, the disease in this family (F1‐A) started in the rods followed by the cones (rod‐cone dystrophy (RCD).

Family F4 patients (IV‐4 and IV‐6) who harbor the same DCV as family F1: A also reported decreased visual acuity as the first symptom. However, their ffERGs showed more severe impairment of photopic responses than scotopic responses (Figure 4). Additionally, fundus photography showed atypical RP features; RP sine pigmentoin addition to the presence of bone spicules that were restricted to the central part of the retina (Figure 3; Figure S2). OCT of all patients showed central macular atrophy (Figure 3; Figure S4). Therefore, this family was diagnosed with cone‐rod dystrophy (CRD).

In the third family (F3) with ABCA4 DCV (Figure 1), IV‐5 reported decreased visual acuity as the first symptom, which was followed by gradual constriction of peripheral vision. Funduscopic examination showed bone spicules only in the central macula and in the inferior part of the retina, suggesting sector RP features (Figure 3). OCT revealed central atrophic maculopathy in both retinae which was more severe in the left eye (Figure 3). FfERG showed severe reduction in both scotopic and photopic responses, except for scotopic responses in the left eye which were moderately reduced (Figure 4; Table 3). Both OCT and ERG results were consistent with visual acuity being better in the left eye than the right (0.1 and CF at 2 m, respectively). This family was diagnosed with CRD.

Associated ocular diseases

Among patients with available clinical data, seven of 12 patients either have cataract or underwent cataract surgery in the past. Three patients (25%) had posterior subcapsular cataract in both eyes, one has posterior polar cataract with nuclear sclerosis, and one with congenital cataract (Table 1). Keratometry showed topographic keratoconus only in patient IV‐7 in family F1‐A (Figure S6).

DISCUSSION

In this study, we combined clinical and molecular data to characterize the phenotype of patients with various forms of IRDs. We identified two novel DCVs in CLRN1; one splice donor variant (c.433+1G>A) and another missense variant (c.323T>C, p.Leu108Pro). Moreover, we added thorough clinical descriptions to two recurrent ABCA4 variants ((c.5460+1G>A), (c.1648G>A, p.Gly550Arg)).

CLRN1 codes for a four‐transmembrane domain protein called clarin‐1 that localizes to the plasma membrane of auditory hair cells and spiral ganglion cells in the retina (Adato et al., 2002). Clarin‐1 has multiple functions in the retina and auditory hair cells including organization of actin cytoskeleton, and sensory perception of visual and auditory stimuli (Aller et al., 2004; Tian et al., 2009). CLRN1 variants have been classically associated with Usher syndrome (Aller et al., 2004; Khan et al., 2011; Pennings, Kremer, Deutman, Kimberling, & Cremers, 2002; Roux, 2005). A study published in 2011 described two missense CLRN1 variants causing nonsyndromic IRD (Khan et al., 2011). This study suggested that missense CLRN1 variants located in the boundaries of transmembrane domains, and resulting in no change in the polarity of the amino acid are more likely to cause nonsyndromic IRD (Khan et al., 2011). In other studies, Usher syndrome was mostly caused by CLRN1 variants located in the extra‐ or intracellular domain of clarin‐1 (Garcia‐Garcia et al., 2012; Isosomppi et al., 2009). The CLRN1 (c.323T>C, p.Leu108Pro) missense variant identified in family F2 is located in the middle of the second transmembrane domain of clarin‐1 and caused Usher syndrome. This is similar to two missense variants (c.449T>C, p.Leu150Pro) and (c.313T>C, p.S105P) identified before and were causative of Usher syndrome (Fields et al., 2002; Sadeghi, Cohn, Kimberling, Tranebjaerg, & Moller, 2005). Therefore, our study and those of others suggest that the location of the substituted amino acid within clarin‐1 domains as well as the change in its polarity appear to be reliable predictors of the resultant phenotype. This variant was associated with late‐onset Usher syndrome (late onset of both RP and hearing loss). On the other hand, family F1‐B patients who harbor the splice donor CLRN1 DCV had a much earlier onset and more severe disease as evident by their lower BCVA. Noteworthy, cataract also contributes to the lower VA of some patients of family F1‐B compared to F2.

The story of patients in these two families highlights the importance of integrating ES into clinical diagnosis of IRDs. Prior to molecular assessment, all our patients with CLRN1 variants had no hearing or balance complaints and were thus diagnosed with nonsyndromic RP. Only after molecular testing showed CLRN1 as disease causing, audiometry was done and revealed sensorineural hearing loss in all patients. Consequently, all patients in families F1‐B and F2 were diagnosed with Usher syndrome III. This illustrates the utility of audiometry to diagnose asymptomatic hearing impairment as well as the importance of genetic testing to accurately diagnose IRD caused by CLRN1 variants and differentiate between syndromic and nonsyndromic phenotypes.

Simulation analysis including docking and molecular dynamics simulation was performed to the novel CLRN1 splice donor variant (c.433+1G>A) to study the effects of a splice donor variant in CLRN1 on mRNA processing. Docking simulations showed that mutating guanine to adenine at the splice site affects binding with U1 snRNP tremendously. Molecular dynamic simulations clarified that even when the mutated RNA binds U1 snRNP, there would be lessening of the main noncanonical contacting interactions between RNA and U1 snRNP. Taken together, these results indicate that this variant is predicted to affect splicing and thus produce an abnormal disease‐causing protein.

The other families (F1‐A, F3, and F4) have DCVs in ABCA4. ABCA4 is exclusively expressed in the retina, especially in its photoreceptors and functions to translocate diverse intermediates of visual cycle out of them (Allikmets et al., 1997). Reduced activity of ABCA4 protein leads to accumulation of its substrates in the photoreceptors and RPE which consequently leads to apoptosis (Cideciyan et al., 2009; Sun, Molday, & Nathans, 1999). Death of RPE cells occurs within and near the macula where cones and most of the rods are located, respectively (Martinez‐Mir et al., 1998; Tsybovsky, Molday, & Palczewski, 2010). This might explain why ABCA4 variants may cause both RCDs and CRDs. ABCA4 variants have been linked to Stargardt's disease, CRD, and RP (Lois, Holder, Bunce, Fitzke, & Bird, 2001). Of interest, we identified the same rare variant (ABCA4 (c.5460 G>A)), whose MAF is 0.00003185, as disease causing in two unrelated families (F1‐A and F4). More surprisingly, they had different clinical phenotypes. F1‐A patients have RCD, while F4 patients have CRD based on their ERGs (Table 3). This illustrates the extensive heterogeneity of IRDs as they do not only exhibit genetic heterogeneity and variable expressivity but also "variant pleiotropy". F3 has ABCA4 (c.1648G>A, p.Gly550Arg) as the DCV. This variant has been previously reported to cause CRD, Stargardt's disease, and generalized choriocapillaris dystrophy (Shroyer, Lewis, Yatsenko, Wensel, & Lupski, 2001; Thiadens et al., 2012). Here, we report this variant with CRD and sector RP; a form of atypical RP. To the best of our knowledge, this variant has never been reported to cause sector RP. Heterozygous ABCA4 variants were also associated with age‐related macular degeneration (ARMD) (Kjellström, 2015; Schulz et al., 2017). Hence, both affected and carrier individuals of families F1‐A, F3, and F4 might be at increased risk of ARMD in the future. In conclusion, our findings emphasize the importance of combining ophthalmic and genetic tests to correctly diagnose rare diseases like IRDs and provide proper counseling. It also characterizes detailed explanation of the effect of a splice donor variant in CLRN1 clinically and the level of mRNA processing. Furthermore, we emphasized that ABCA4 variants can cause a spectrum of retinal diseases rather than a single entity, even when the same DCV exists in two unrelated families. This might be an indication of the presence of other environmental or epigenetic factors that govern genotype–phenotype correlations.

CONFLICT OF INTEREST

All authors declare that they do not have any competing interests.

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