A Novel c.3636-4 A>G Mutation in the CCDC88C Plays a Causative Role in Familial Spinocerebellar Ataxia

Spinocerebellar ataxia (SCA) is an autosomal dominant disorder and the most common type of hereditary ataxia. It is characterized by high clinical and genetic heterogeneity, with 48 pathogenic genes having been identified. The main clinical symptoms of SCA are ataxia, some of which are accompanied by seizures, dementia, nystagmus, muscle tremor, and intellectual decline [1‒3]. Its pathogenesis involves abnormal aggregation of the polyglutamic acid complex; moreover, trinucleotide repeat amplification is the most common mutation subtype in patients with SCA and they typically exhibit genetic prematurity [4]. SCA is characterized by familial genetic history, cerebellar ataxia, and pathological damage to the cerebellum and spinal cord; however, its etiology and pathogenesis remain unclear. It is mainly caused by mutations in the identified pathogenic genes. Additionally, the microenvironment and local biochemical metabolic abnormalities are crucially involved in the occurrence and development of different SCA phenotypes. Moreover, stress, emotional fluctuations, and hormone imbalances may induce or aggravate SCA symptoms. Given the high disability and mortality rate, as well as the ineffectiveness of currently available treatments, SCA exerts a significant emotional burden on patients, adversely affects their quality of life, and exerts a heavy economic burden on society and families.

Gene mutations generally cause structural and functional changes in the coded proteins. Moreover, mutations can cause abnormalities in other components, including molecular chaperones, protease ubiquitination systems, and caspases. This can result in the formation of extracellular, cytoplasmic, or nucleic inclusion bodies and activation of various metabolic pathways and cytokines. Trinucleotide repeat amplification causes multicopy polyglutamine (polyQ) and other toxic fragments, which accumulate in the nucleus of neurons to form inclusion bodies, resulting in cytotoxicity [5], protein misfolding, dysregulation of the protease ubiquitination system, imbalanced Ca²⁺ homeostasis, and loss of molecular chaperones. This causes selective damage and functional loss in the nervous system [6]. Moreover, this toxicity resulting from trinucleotide repeat amplification at the RNA level is an important causative factor for polyglutamine diseases [7].

According to different pathogenic genes, SCA can be classified into different subtypes. With the development of gene sequencing, more than 40 different SCA subtypes have been identified [8]. Tsoi et al. [9] first reported the type spinocerebellar ataxia type 40 (SCA40) caused by the coiled-coil domain containing 88c (CCDC88C) gene mutation. The gene is located in 14q32.11-q32.12 and is highly expressed in the cerebellum. The first clinical features of this type are cerebellar ataxia, with or without vertical gaze disorder, spastic paraplegia, and cerebellar atrophy. It is characterized by concealed onset and gradually aggravation. In this study, we report a rare case of SCA40 in a four-generation Chinese kindred comprising eight affected individuals. Through gene sequencing and minigene assays, we identified a de novo insertion mutation in CCDC88C in this family, promoting further application of mutation screening in genetic counseling.

The pedigree of the family with eight affected individuals was from Central China and is presented in Figure 1a. There were no consanguineous relationships in the four-generation family (Fig. 1a). This study was approved by the Ethics Committee of Xiangyang No.1. People’s Hospital, Hubei (2019GCP065). Informed consent was obtained from all participants. We obtained 5 mL of peripheral venous blood from each participant, which was stored in ethylenediaminetetraacetic acid anticoagulant tubes and frozen at −20°C until subsequent use.

The proband was a 29-year-old man engaged in medical work. His parents were not consanguineous. The patient was hospitalized because of symptom progression, including difficulty climbing stairs, shaking while walking, sudden falls, vague speech, and a choking cough. The patient had an abnormal walking posture 1 year prior, which manifested as unsteady gait and dysarthria. No mental retardation and urinary incontinence were observed. Neurological examination found that the patient had visual impairment, speech scanning, and intentional tremor symptoms (online suppl. File 1: a video of the proband’s phenotype; for all online suppl. material, see https://doi.org/10.1159/000534692). His intelligence level was normal. Moreover, he showed abnormal manifestations in the upper limbs and a positive finger-to-nose test. Plain brain magnetic resonance imaging (MRI) revealed cerebellar atrophy without obvious pontine atrophy. Specifically, the MRI scan revealed a decrease in the volume of the bilateral cerebellar hemispheres, expansion of the fourth ventricle, and widening of the cerebellar cistern groove. Further, MRI perfusion showed a bilateral decrease in cerebellar hemisphere perfusion (Fig. 1c). The local perfusion of the right cerebellar hemisphere was greater than that of the left; contrastingly, the bilateral cerebral hemispheres showed symmetrical perfusion. Neuroelectrophysiological examination revealed significantly abnormal brainstem auditory evoked potentials and somatosensory evoked potentials. The bilateral lower limbs showed significantly abnormal somatosensory evoked potentials, as well as normal motor and sensory nerves, F-waves, and H-reflexes. The rehabilitation evaluation results were as follows: modified Barthel Index score, 40 (0–40, heavily dependent); Berg Balance Scale score, 31 (range: 0–56); and Tinetti Balance and Gait Scale score, 6 (range: 0–28). His father, aunt, grandmother, and granduncle had similar symptoms; however, their ages at onset were different (Table 1). There was an obvious family history in the pedigree. Based on the MRI findings and clinical phenotypes, the patient was diagnosed with SCA, balance and coordination dysfunction, dysphagia, and dysarthria.

Genomic DNA was extracted from the whole blood of each affected member (Fig. 1). DNA from the proband and his parents (IV-2, III-5, III-6) were sequenced by whole-exome sequencing (WES). The Roche KAPA HyperExome chip and MGISEQ-2000 sequencing platform (BGI) were used for in-solution enrichment of coding exons and flanking intronic sequences following the manufacturer’s instructions. We used cutadapt (v1.15) to trim adaptor sequences at the tail of the sequencing reads; subsequently, the sequencing reads were aligned to the human reference genome (UCSC hg19) using BWA (v0.7.15). Duplicate reads were marked using Picard (v2.4.1). Qualimap [10] (v2.2.1) was used to calculate the base quality metrics, genome mapping rate, and coverage of the targeted regions. We performed base quality score recalibration, indel realignment, and variant (SNV & InDel) calling according to the best practice protocol of the Genome Analysis Toolkit (v3.8). Variant filtering was performed using a fine-tuned in-house script. Pass filter variants were annotated using the Pubvar variant annotation engine (www.pubvar.com) and Variant Effect Predictor [11]. Variant models for dominant and recessive inheritance were separately identified in genetic analysis. The exclusion criteria for the variants were as follows: maximum population frequency >0.01, low genotype confidence, or prediction as benign by the following five algorithms: SIFT [12], PolyPhen 2 [13], MetaSVM [14], MCAP [15], and MutationTaster [16]. The pathogenicity of the candidate causative mutations was graded using InterVar (1.0.8) [17] according to the American College of Medical Genetics and Genomics (ACMG) guidelines [18]. Next, we performed polymerase chain reaction (PCR) and bidirectional Sanger sequencing of DNA obtained from the family members to verify the gene mutations identified through WES. PrimerBLAST (https://www.ncbi.nlm.nih.gov/tools/primer-blast/) was used to design primers containing the mutation site. The primers used were as follows: CACNA1A_F, TCA​CAG​TCC​AGG​GTC​ACT​CAG​CA; CACNA1A_R, CAG​CCA​AGA​TGG​GAA​ACA​GCA; CCDC88C_F, TGGCGAGGGCGTTTGTC; CCDC88C_R, GGA​GGG​ACT​ATG​GAG​ACA​GAA​CG; ZNF142_F, TTC​TGT​TTC​TTC​TGG​CTC​CCT​G; and ZNF142_R, CCT​CCA​CAA​ATC​CTC​CAT​CCT​T. The Online Analysis Tools, South China Rare Diseases Data Center (RDDC, https://rddc.tsinghua-gd.org/), and SWISS-MODEL (https://swissmodel.expasy.org/) were used to predict the effect of this mutation.

The minigene splicing assay was performed for the splicing mutation. The wild-type (WT) and mutant-type (MT) forms of the minigene constructs, encompassing CCDC88C exons 20–22 and partial introns 20–21 (the middle part was deleted because introns 20–21 were too large), were amplified from genomic DNA of the proband Ⅳ-2. The primers were designed through seamless cloning to amplify the genomic DNA of heterozygous CCDC88C (NM_001080414:c.3636-4 A>G); additionally, the three gene fragments for insertion were obtained. The amplification primers were as follows: AF, 5′-AAG​CTT​GGT​ACC​GAG​CTC​GGA​TCC​GTG​GAG​AAC​TCC​ACG​CTG​AGT​TCC​CAG​A-3′; AR, 5′-TAT​AAC​TCT​TAG​CCT​CTC​CTG​AGA​CCC​TCT​CCT​GAG​AC-3′; BF, 5′-CAG​GAG​AGG​CTA​AGA​GTT​ATA​AAG​TTT​CTG​CCC​TAC​AG-3′; BR, 5′-GAT​AAA​GTG​TGC​CCT​CAG​CTG​GGA​AGG​TGC​GCA​TCC​AG-3′; CF, 5′-AGC​TGA​GGG​CAC​ACT​TTA​TCT​CAC​GAG​CTG​AAA​TTA​GA-3′; and CR, 5′-TTA​AAC​GGG​CCC​TCT​AGA​CTC​GAG​CTC​ACA​GTG​GTT​GTC​CAG​CTT​GGT​CAG​C-3′ (AF/AR, BF/BR, and CF/CR, a set of primers). AF and CR contained the BamHI and XhoI restriction sites, respectively. The amplified products were cloned into the pMini-CopGFP vector (Beijing Hitrobio Biotechnology Co., Ltd.) using a Seamless Cloning and Assembly Kit (Vazyme, C112-02, Nanjing, China) and transformed into Escherichia coli. Single clone was selected for plasmid extraction and the WT and MT forms of minigene plasmids were verified by Sanger sequencing. We used the following identification primers: globin intron-F (5′-GAT​ATA​CAC​TGT​TTG​AGA​TGA​GGA-3′) and CCDC88C-CX-1R (5′-CAC​TGG​GAG​CCA​TGT​CAA​GG-3′). Human embryonic kidney 293T cells were incubated in Dulbecco’s modified Eagle’s medium supplement with 10% fetal bovine serum (HyClone) and incubated at 37°C and 5% CO₂. The recombinant plasmids were transiently transfected into 293T cells with Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA). After cells cultured for 48 h, total RNA was isolated using Trizol Reagent (Thermo Fisher, USA) and phenol-chloroform extraction. RT-PCR was performed using the HiScript II First-Strand cDNA Synthesis Kit (Vazyme, Nanjing, China). We used the following RT-PCR primers: MiniRT-F (5′-GGC​TAA​CTA​GAG​AAC​CCA​CTG​CTT​A-3′) and CCDC88C-RT-R (5′-CTC​ACA​GTG​GTT​GTC​CAG​CTT​G-3′). The RT-PCR products were analyzed through agarose gel electrophoresis and Sanger sequenced to identify the effect of the mutation on splicing. Detailed CCDC88C minigene splicing assay report is included in the supplementary materials (online suppl. file 2: verification of in vitro splicing of CCDC88C gene mRNA).

We identified three gene heterozygous mutations, including CACNA1A (NM_001127221:c.3467 A>T, p.Asn1156Ile), CCDC88C (NM_001080414:c.3636-4 A>G), and ZNF142 (NM_001105537:c.2584 C>T, p.Arg862Cys), in the proband and his father. To verify the three mutations, bidirectional Sanger sequencing was performed on ten individuals (III-3, Ⅲ-4, Ⅲ-5, III-6, III-8, III-9, IV-1, IV-2, IV-3, Ⅳ-4, Fig. 1a) on the pedigree. Sanger sequencing revealed that only the CCDC88C gene mutation completely co-segregated with the SCA phenotype (Fig. 1; online suppl. Fig. 1–3; File 3). This CCDC88C heterozygous mutation (NM_001080414:c.3636-4 A>G) is novel. It has not been reported or listed in the Human Gene Mutation Database, Exome Variant Server, 1,000 Genomes database, ClinVar, and dbSNP. Its frequency is less than 0.001 in gnomAD and ExAC. SCA40 (OMIM:616053) is associated with this gene. Using SpliceAI [19] and NetGene2 (v2.42, http://www.cbs.dtu.dk/services/NetGene2) to predict CCDC88C (NM_001080414:c.3636-4 A>G) site, it was found that this mutation can affect a splice site; therefore, it can be considered as a harmful mutation. Additionally, this variation was classified as likely pathogenic based on the ACMG guidelines (PS2+PP1_Strong +PP3+PP4).

According to the in silico analysis of RDDC, there are two splice variants in CCDC88C (NM_001080414:c.3636-4 A>G) gene (online suppl. Fig. 4; File 3). After SWISS-MODEL prediction, no “tropomyosin alpha-1 chain (structure of human cardiac thin filament in the calcium-free state)” and “flagellar-associated protein 59 (radial spoke 2 stalk, IDAc, and N-DRC attached with doublet microtubule)” were found in the mutation type.

1. We performed a minigene assay to further characterize the splicing effects of this mutation. In the agarose gel electrophoresis of the RT-PCR products, both WT and mutant strains produced visible bands (Fig. 2c). Sanger sequencing revealed that the normal splice isoform originated from the RNA extracted from human embryonic kidney 293T cells transfected with WT plasmid (Fig. 2a). Notably, the mutation led to insertion of CAG bases (c.3636-1_3636-3 insCAG); however, Sanger sequencing revealed double peaks after insertion (Fig. 2b). To verify the existence of the bimodal pattern, we performed cloning and Sanger sequencing of special fragments. Adopting the principle of seamless cloning, the RT-PCR product of CCDC88C-MT recovered from agarose gel was amplified using primer CCDC88C-AF and CCDC88C-CR to obtain the target insertion gene fragment. The target gene fragments were cloned into the pMini CopGFP vector. Then the plasmid constructs were transformed into E. coli and monoclonal RNA extraction, reverse transcription, and Sanger sequencing have been carried out (online suppl. file 2). The transformed E. coli contained the normal (WT) and abnormal splicing isoforms (Fig. 2d). Minigene analysis suggested that the c.3636-4 A>G substitution led to an additional splice receptor (AG) site in intron 20, which led to the emergence of two isoforms (Fig. 2e; online suppl. file 2). According to minigene assay, the mutation resulted in the insertion of a serine (Ser) between amino acids 1,211 and 1,212 of the WT protein. The mutation site just formed a new AG structure. During the intron splicing process, the three bases behind it were retained. Interestingly, this alternative splicing is only partially functional as the WT splice variant is also detectable in HEK293 cells expressing the plasmid carrying the CCDC88C(A>G) sequence.

Our proband had an obvious family genetic history and presented with lower limb ataxia during college. This SCA phenotype was found to be caused by a novel splice mutation, CCDC88C (NM_001080414:c.3636-4 A>G), which results in the insertion of a serine residue between 1,211 and 1,212 amino acid loci of the original WT protein. It was detected in 4 affected individuals in the family and was absent in 6 unaffected individuals. The proband was diagnosed with SCA40 (OMIM:616053) based on the clinical manifestations and genetic testing results. Given the unavailability of specific tissues and cells, we used an in vitro minigene assay to evaluate the splicing effect of this mutation. Our results showed that HEK293 cells transfected with the mutant plasmid contained both the normal (WT) and abnormal splicing isoforms. These results suggest that the insertion mutations in CCDC88C crucially involved in the pathological process of SCA40 may be through a dominant-negative effect on the WT CCDC88C protein; however, further studies are warranted to elucidate the underlying molecular mechanism.

Our patient was diagnosed with SCA40, which is a rare subtype of SCA, and presented with progressive ataxia, peripheral nerve damage, and pyramidal tract signs. A Chinese family member carried a missense mutation c.G1391A (p.R464H), presented with occult onset and slow progression after the age of 40 years. The first symptom was cerebellar ataxia, followed by a vertical gaze disorder and mild spastic paraplegia. MRI examination revealed pontocerebellar atrophy. The patient required crutches and a wheelchair for approximately 10 and 20 years, respectively, after disease onset [9]. The age at onset of the proband and his sister in the Polish family was 49 and 33 years, respectively, which is consistent with our findings of differences in the age of onset among individuals in the same pedigree carrying the same CCDC88C mutation (c.127 G > A). The initial symptoms were tremor, mild ataxia, Parkinsonism, active tendon reflexes, and cognitive impairment; however, activity was adversely limited at >20 years following onset. Imaging did not reveal pontocerebellar atrophy [20]. Another heterozygous mutation (c.1886G>A) in CCDC88C was identified in a Shandong Chinese pedigree with SCA40. The age at onset of the proband was 61 years; however, the other family members who were carriers of the heterozygous mutation did not present with the clinical phenotype [21]. Yahia et al. [22] reported a case on a Sudanese patient with childhood-onset spastic paraparesis without cerebellar signs caused by a missense mutation (c.1993G > A) in the CCDC88C gene. Boros, F. et al. [23] reported a female patient carried a missense mutation (p.R203W) in CCDC88C who developed late-onset ataxia, dysmetria, and intention tremor. Taken together, these findings suggest that the clinical manifestations of SCA40 may be related to race, gene mutation sites, and other factors [24]. The aforementioned gene mutation is involved in the disease through its regulation of c-Jun N-terminal kinase (JNK) and caspase-3 in the apoptotic signaling pathway. Specifically, this gene mutation causes JNK hyperphosphorylation, upregulates caspase-3 activity, and induces apoptosis [9, 22]. However, studies have also shown that A mutations do not affect c-Jun N-terminal kinase 1 (JNK1) phosphorylation, nor do they trigger apoptosis signals, only a small-scale activation of the JNK pathway [23].

In conclusion, we identified a novel heterozygous mutation, CCDC88C (NM_001080414:c.3636-4 A>G), in the splice region of eight affected family members of a four-generation Chinese kindred. This mutation led to the insertion of CAG bases (c.3636-1_3636-3 insCAG). Further studies are warranted to elucidate the molecular mechanisms of CCDC88C and thus identify potential therapeutic targets for limiting the deleterious effects of mutations. Nevertheless, our findings expand the mutational spectrum and clinical phenotype of the CCDC88C gene and further confirm its pathogenic role in SCA40.

The authors are grateful to the family members for their participation in the studies.

All procedures in studies involving human participants were performed in accordance with the ethical standards of the institutional and/or national research committee and with the 1964 Helsinki Declaration and its later amendments or comparable ethical standards. The study received ethical approval by the Ethics Committee of Xiangyang No.1. People’s Hospital, Hubei, China (NO. 2019GCP065). All authors and patients consented to participate in the study. Written informed consent was obtained for participation in this study.

The authors have no relevant financial or nonfinancial interests to disclose.

This investigation was supported by the Experimental Animal Resources Development and Utilization Project of Hubei Province of China (Grant No. 2020DFE025), the Open Project of Hubei Key Laboratory of Wudang Local Chinese Medicine Research (Grant numbers WDCM2021), and the Scientific and Technological Project of Xiangyang City of Hubei Province (Grant No. 2020YL28).

All authors contributed to the study conception and design. M.S. and S.C. led the clinical management of the patient and parents. M.S., D.L., and S.C. performed the literature survey, wrote the manuscript, and edited the manuscript. S.C. and Y.L. performed the sample recruitment, whole-exome and Sanger sequencing, and data analysis. All authors read and approved the final manuscript.

All relevant data are within the paper, including raw data from WES. Further information is available from the corresponding authors on request. All raw data related to our study have been submitted to the NCBI Sequence Read Archive (SRA) (SRA RunSelector: https://www.ncbi.nlm.nih.gov/bioproject/PRJNA817397). The accession number is PRJNA817397. Reference datasets, such as the human reference genome (UCSC hg19) and transcription number (NM_001080414.3), are provided in the “Whole-Exome Sequencing and Sanger Sequencing Validation” section. Data are not publicly available due to ethical reasons. Further inquiries can be directed to the corresponding author.