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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
Ann Neurol. Author manuscript; available in PMC Apr 1, 2013.
Published in final edited form as:
Published online Mar 23, 2012. doi:  10.1002/ana.23547
PMCID: PMC3334472
NIHMSID: NIHMS353714
Mutations in CIZ1 cause adult-onset primary cervical dystonia
Jianfeng Xiao, MD, PhD,,1 Ryan J. Uitti, MD,2 Yu Zhao, MD, PhD,1 Satya R. Vemula, PhD,1 Joel S. Perlmutter, MD,3 Zbigniew K. Wszolek, MD,2 Demetrius M. Maraganore, MD,4 Georg Auburger, Dr. Med,5 Barbara Leube, Dr. Med,6 Katja Lehnhoff, PhD,5 and Mark S. LeDoux, MD, PhD1*
1Departments of Neurology and Anatomy and Neurobiology, University of Tennessee Health Science Center, Memphis, TN 38163, USA
2Department of Neurology, Mayo Clinic Jacksonville, Jacksonville, FL 32224, USA
3Department of Neurology, Washington University School of Medicine, St. Louis, MO 63110, USA
4Department of Neurology, NorthShore University HealthSystem, Evanston, IL 60201, USA
5Department of Neurology, University Medical School, Frankfurt am Main, Germany
6Institute of Human Genetics, University Hospital of Duesseldorf, Duesseldorf, Germany
*Corresponding author: Mark S. LeDoux, MD, PhD, University of Tennessee Health Science Center, Department of Neurology, 855 Monroe Avenue, Link Building, Suite 415, Memphis, TN 38163, USA Phone: (901) 448-1662 Fax: (901) 448-7440 ; mledoux/at/uthsc.edu
Objective
Primary dystonia is usually of adult onset, can be familial, and frequently involves the cervical musculature. Our goal was to identify the causal mutation in a family with adult-onset, primary cervical dystonia.
Methods
Linkage and haplotype analyses were combined with solution-based whole-exome capture and massively parallel sequencing in a large Caucasian pedigree with adult-onset, primary cervical dystonia to identify a cosegregating mutation. High-throughput screening and Sanger sequencing were completed in 308 Caucasians with familial or sporadic adult-onset cervical dystonia and matching controls for sequence variants in this mutant gene.
Results
Exome sequencing led to the identification of an exonic splicing enhancer mutation in Exon 7 of CIZ1 (c.790A>G, p.S264G) which encodes CIZ1, Cip1-interacting zinc finger protein 1. CIZ1 is a p21Cip1/Waf1-interacting zinc finger protein expressed in brain and involved in DNA synthesis and cell-cycle control. Using a minigene assay, we showed that c.790A>G altered CIZ1 splicing patterns. The p.S264G mutation also altered the nuclear localization of CIZ1. Screening in subjects with adult-onset cervical dystonia identified two additional CIZ1 missense mutations (p.P47S and p.R672M).
Interpretation
Mutations in CIZ1 may cause adult-onset, primary cervical dystonia, possibly by precipitating neurodevelopmental abnormalities that manifest in adults and/or G1/S cell-cycle dysregulation in the mature central nervous system.
Primary generalized dystonias usually begin in childhood, whereas focal dystonias typically present during adulthood. Adult-onset primary dystonias are at least 10-fold more prevalent than early-onset cases.1 Cervical dystonia (CD) or spasmodic torticollis, the most common form of focal dystonia, is characterized by involuntary contractions of the neck muscles, which produce abnormal posturing of the head upon the trunk. Genetic factors are believed to play a major role in the pathogenesis of adult-onset primary dystonia because 10–20% of patients have one or more affected family members.25
Although late-onset primary dystonia has a considerable heritable component, large pedigrees adequately powered for linkage analysis are uncommon and only a few have been described in the literature.69 Moreover, traditional genetic approaches such as linkage analysis have proven to be problematic in focal dystonia given that the characteristic reduced penetrance of mainly early-onset primary dystonias such as DYT1 (TOR1A) and DYT6 (THAP1) tends to be more pronounced in families with adult-onset disease. To date, a single locus (DYT7) for mainly adult-onset CD has been mapped to Chromosome 18p in a German kindred.10
In this study, linkage analysis was combined with whole-exome sequencing to identify the cause of CD in a previously described multigenerational Caucasian pedigree from the United States (Family A, Fig. 1).6,11 In-solution targeted exome capture and massively parallel sequencing was completed for five definitely affected and three unaffected subjects from the kindred. Using an autosomal dominant model, we employed a stepwise filtering process to identify sequence variants (SVs) common to the five definitely affected subjects but absent from at least two of the three unaffected subjects. SVs were filtered under the assumption that the mutation causing this uncommon familial disease is not present in the general population.
FIGURE 1
FIGURE 1
Family A pedigree and linkage analysis. (A) Filled symbols, definitely affected. Half-filled symbols, possibly affected. Symbols with central dots, unaffected carriers. Arrow, proband. CIZ1 genotypes: wild-type (+/+) and heterozygous c.790A>G (more ...)
Cases and samples
Human studies were conducted in accordance with the Declaration of Helsinki, with formal approval from the institutional review boards at each participating study site. All subjects gave written informed consent. Enrollment of patients with primary dystonia and neurologically normal controls was described in previous publications.5,12 All normal controls were examined to exclude dystonia and other neurological disorders. Genomic DNA was extracted from whole blood. All affected subjects had isolated CD with otherwise normal neurological examinations. More specifically, no subject showed clinical evidence of ataxia, spasticity, oculomotor abnormalities, Parkinsonism, or neuropathy.
Linkage analysis
Linkage analysis was performed by genotyping microsatellites at the Genethon facility, Evry, France.13 DNA from members of Family A (6 affected and 10 unaffected) was PCR amplified at short tandem repeat loci (microsatellites) in a multiplexing approach. The amplification products were separated by polyacrylamide gel electrophoresis, transferred to nylon membranes, and visualized by electrochemiluminescence with exposure to radiographic film. Allele sizes were scored visually. Data from 238 microsatellite markers, spaced at approximately equal distances across the 22 autosomes, was analyzed using MERLIN (Multipoint Engine for Rapid Likelihood Inference), version 1.1.2.14 For parametric analysis, heterogeneity logarithm of odds (HLOD) scores were calculated for each marker using an autosomal dominant model with the disease allele frequency set to 0.0001. A nonparametric linkage statistic and associated P values were calculated using the Kong and Cox linear model implemented by MERLIN.
Targeted capture and exome sequencing
DNA samples from five members of Family A with definite CD and three unaffected family members were captured with the SureSelectXT All Exome Kit 50 Mb (Agilent, Santa Clara, CA). In brief, 3 μg of genomic DNA was sheared to yield 100- to 150-bp fragments. Shearing was followed by end repairs and adapter ligation. The DNA-adaptor-ligated fragments were then hybridized to the human exome libraries for 24 h at 65°C. After hybridization, washing, and barcoding, DNA was amplified with emulsion PCR. Enriched DNA fragments were sequenced on a SOLiD 4 system (Applied Biosystems, Carlsbad, CA).
Prior to use of the 50-Mb All Exome Kits, four samples (three definite CD, one unaffected) had been captured with the Agilent SureSelectXT Human All Exon Kit 38 Mb and sequenced independently. Reads from samples that were captured and sequenced twice were combined for downstream analyses. Percentage of exome coverage was based on exons targeted by the 50-Mb All Exome Kit, which incorporates Consensus Coding Sequence (CCDS), NCBI Reference Sequence (RefSeq), and GENCODE annotations.
Read mapping and variant analysis
Color space reads were mapped to the human reference genome (NCBI build 37.1) with NextGENe (SoftGenetics, State College, PA). Using the consolidation and elongation functions of NextGENe, instrument sequencing errors were reduced and sequence reads were lengthened prior to variant analysis. Given that short reads generated by the SOLiD system may not be unique within the genome being analyzed, the condensation tool polished the data for adequate coverage by clustering similar reads with a unique anchor sequence. Using this process, short reads were lengthened and reads with errors were filtered or corrected. The average post-processing read length for all eight samples was 74 nt.
To maximize the probability of detecting the causal SV, all base changes occurring in ≥2 reads in any individual sample were classified as variants for downstream analyses (Supplementary Fig. 1). Variant comparison was done between the five affected and three unaffected subjects to filter the number of SVs to identify the putative causal variant. Variant calls were combined from three independent comparisons of all five definitely affected subjects and two of the three unaffected subjects. The goal of these comparisons was to identify those SVs common to all five affected subjects and absent from at least two of the three unaffected subjects. In this fashion, we reduced the probability of eliminating a true causal variant present in an unaffected carrier. With NextGENe software, all filtering parameters were implemented simultaneously: homozygous SVs, intergenic SVs, deep intronic SVs (≥ 12 nt from splice sites), SVs reported in dbSNP or 1000 Genomes, synonymous SVs, and nonpathogenic nonsynonymous SVs. Additional details of variant analysis are provided in the Supplementary Information.
To validate the results of exome sequencing, primers were designed with Primer3 to amplify regions harboring SVs. PCR products were examined on agarose gels and cleaned with ExoSAP-IT (United States Biochemical, Cleveland, OH) prior to Sanger sequencing in the forward and reverse directions using BigDye Terminator v3.1 chemistry (Life Technologies, Carlsbad, CA) on an Applied Biosystems 3130XL Genetic Analyzer.
CIZ1 and SETX mutation screening
Sanger sequencing and/or high-resolution melting (HRM) were used for mutation screening in 308 subjects with CD and 724 controls. HRM was performed with the LightCycler 480 Real-Time PCR system and High Resolution Master Mix (Roche, Indianapolis, IN) in accordance with our laboratory protocol.5 HRM reactions were performed in either 96- or 384-well plates, using 20 ng of template DNA, 1× HRM Master Mix, 2.5 mM MgCl2, and 200 nM of each primer in a 10-μl reaction volume. Melting curves and difference plots were analyzed by at least two investigators blinded to phenotype. For the 140 samples with shifted melting curves, PCR products were cleaned using ExoSAP-IT and Sanger sequenced in the forward and reverse directions.
In silico analysis of amino acid substitutions and splicing
Missense variants were analyzed with PolyPhen-2,15 SIFT,16 and MutationTaster17 to predict the pathological character of single amino acid mutations. Protein sequence alignment was performed with ClustalW2.18 The possible effects of c.790A>G on CIZ1 splicing were evaluated using a broad array of tools for splice site, exonic enhancer, and silencer site analysis: Relative Enhancer and Silencer Classification by Unanimous Enrichment (RESCUE-ESE19), Putative Exonic Splicing Enhancers/Silencers (PESX20), Human Splicing Finder (HSF21), NetGene2,22 Splice Site Prediction by Neural Network (NNSplice23), MaxEntScan (MES24), ESEfinder,25 and FAS-ESS.26
CIZ1 expression in human brain and lymphoblastoid cell lines
Adult human whole-brain total RNA (FirstChoice Human Brain Reference RNA, 1 mg/ml) was obtained from Ambion (Austin, TX). Fetal human whole brain and adult human cerebral cortex, cerebellum, substantia nigra, and putamen total RNA were purchased from Clontech (Mountain View, CA). Ambion's TRI Reagent was used to isolate RNA from lymphoblastoid cell lines generated from two affected subjects (Family A, III-2 and III-3) and six normal controls, as previously described.12 Reverse transcription was performed with Ambion's RETROscript kit using 500 ng total RNA as template along with both random oligonucleotide and oligo(dT) primers. The reaction mixtures were incubated at 44°C for 1 h and then at 92°C for 10 min. Relative quantitative real-time RT-PCR was performed using the LightCycler 480 with gene-specific primers (Supplementary Table 1) and Universal Taqman probes (Roche) for CIZ1 and the endogenous control (PPID), which encodes 40-kDa peptidyl-prolyl cis-trans isomerase D (cyclophilin D). Quantitative RT-PCR was performed to examine the relative expression of the two major CIZ1 isoforms. cDNA was amplified with primers CIZ1_RE6F and CIZ1_RE9R. The agarose gel was imaged (G:BOX gel imaging system, Syngene, Frederick, MA) and bands were analyzed with ImageJ.
Minigene assay of CIZ1 splicing
To validate the predicted effect of the CIZ1 c.790A>G variant at the donor splice site of Exon 7, we performed a minigene analysis. The wild-type minigene construct was produced by cloning into the exontrap vector pET01 (MoBiTec, Göttingen, Germany). Genomic DNA encompassing Introns 5–9, including Exons 6–9, was amplified with the SequalPrep Long PCR Kit (Invitrogen, Carlsbad, CA). The PCR product and pET01 were digested with XhoI and BamHI at 37°C for 2 h and then 65°C for 20 min. This was followed by ligation of cohesive-ended gel-purified digested PCR products into exontrap vector pET01 with the LigaFast Rapid DNA Ligation System (Promega, Madison, WI). After sequence confirmation, the variant c.790A>G was introduced into the WT-minigene with the QuikChange II XL Site-Directed Mutagenesis Kit (Agilent) to generate a mutant minigene.
Next, HEK 293T (293T) cells were transfected in quadruplicate. Total RNA was extracted with TRI Reagent (Ambion) 72 h after transient transfection. DNA was removed with DNase I (Ambion). Total RNA quality was examined with agarose gel electrophoresis and a NanoDrop ND-1000 spectrophotometer (NanoDrop Technologies, Wilmington, DE). Reverse transcription was performed with Ambion's RETROscript kit using 500 ng total RNA as template. PCR products were examined in a 1% agarose gel, bands were quantified with ImageJ, and the effects of genotype were evaluated with t-tests.
Cellular localization of wild-type and mutant CIZ1
To investigate the effects of p.S264G on the cellular localization of CIZ1, we made constructs that expressed full-length human wild-type and mutant (p.S264G) CIZ1 under CMV promoters. Transient transfections with wild-type and mutant vectors (in quadruplicate, for both) were performed in 293T cells. ImageJ was used to perform particle analysis of GFP-CIZ1 transfected 293T cells. Detailed methods related to the cellular localization of CIZ1 are available in the Supplementary Information.
Linkage analysis and exome sequencing
In the five definitely affected and three unaffected subjects from Family A, sequencing generated 2× or greater coverage for 84.7–92.3% of the exome (Supplementary Table 2). After filtering and elimination of read errors with Sanger sequencing, 23 validated SVs remained (Supplementary Fig. 1). Three of these 23 SVs cosegregated with CD: CIZ1 (c.790A>G and c.1730C>T) and SETX (c.2385_2388delAAAG).
In Family A (Fig. 1), CD showed strongest linkage to microsatellite markers D9S159 and D9S1818 on Chr 9q34 (Fig. 1, Table 1). D9S159 is located 1.34 Mb from CIZ1 on Chr 9q34.11. The marker showing the second highest HLOD score, D9S1818, is located nearby on Chr 9q34.13, 6.11 Mb from CIZ1 and 1.90 Mb from SETX. Haplotype analysis of microsatellite markers on Chr 9 showed that D9S159 cosegregated with mutant CIZ1 (Supplementary Fig. 2).
Table 1
Table 1
Microsatellite Markers Associated with the Highest HLOD Scores
The c.790A>G CIZ1 variant is located within a highly conserved donor splice site in Exon 7 (Fig. 2). In contrast, the c.1730C>T (p.S577F) SV is only partially conserved at the amino acid level and predicted to be damaging by only of one of three prediction programs (Table 2). The SETX variant (c.2385_2388delAAAG, p.I795Ifs*16) is predicted to cause a frameshift and probably induces nonsense-mediated decay.
FIGURE 2
FIGURE 2
Organization of CIZ1 gene, transcripts, and full-length protein. (A) Structure of CIZ1 on the reverse strand of Chr. 9 presented in the 5' to 3' direction showing the location of three identified mutations (c.139C>T, c.790A>G, and c.2015G>T) (more ...)
Table 2
Table 2
In Silico Analysis of Missense Sequence Variants in CIZ1
Mutation screening and in silico analyses
Using 18 pairs of PCR primers placed on flanking intronic and untranslated regions to encompass the coding regions of the 17 CIZ1 exons and at least 50 bp of 5'- and 3'-intronic sequence surrounding each exon (Supplementary Table 1), mutation screening in patients with familial and sporadic CD and neurologically normal controls (Table 3) identified five additional missense SVs (p.P47S, p.P50L, p.Q394E, p.S577F, p.R672M) (Table 2, Supplementary Table 3). In silico analyses with ClustalW2,18 Polyphen-2,15 SIFT,16 and MutationTaster17 indicated that p.P47S (Subject C), p.S264G (Family A), and p.R672M (Subject B) were highly conserved and probably damaging to CIZ1. Moreover, the p.S264G mutation is located within a putative nuclear localization signal (NLStradamus27) and alters a potential phosphorylation site (PredictProtein28 and NetPhos 2.029). Two of the novel variants (p.P50L [Subject D] and p.Q394E [Subjects E and F]) detected during screening are predicated to be benign. Moreover, the p.Q394E variant was also found in 1 of 5378 subjects in the Exome Variant Server database. Similarly, none of the validated nonsynonymous CIZ1 SVs identified in the dbSNP database were highly conserved or predicted to be deleterious by all three prediction programs (Supplementary Table 4).
TABLE 3
TABLE 3
CIZ1 Mutation Screening
Although the SETX variant (c.2385_2388delAAAG) cosegregated with CD in Family A, it was not required for manifestation of CD in Subjects B and C (Supplementary Table 3). With HRM, we did not detect this variant in 724 neurologically normal control subjects.
Effects of c.790A>G on CIZ1 splicing
The Exon 7 mutation in CIZ1 (c.790A>G) may also exert deleterious effects on transcription. RESCUE-ESE,19 PESX,20 and FAS-ESS26 identified an exonic splice enhancer site in wild-type CIZ1 that was disrupted by the c.790A>G mutation. ESEfinder predicted that the mutation abolishes the SF2/ASF and SRp55 binding sites present in the wild-type sequence,30 whereas HSF indicated that the splice and enhancer motif sites were broken in the mutant sequence.21 Similarly, NetGene2 predicted that the c.790A>G variant abolished the Exon 7 donor splice site.22 NNSPLICE's donor site prediction score was reduced from 0.9 in the wild-type sequence to 0.6 in the mutant sequence, whereas MaxEntScan's donor splice site score fell from 8.77 to 4.69 with introduction of the c.790A>G variant.23,24
To examine the predicted effects of the c.790A>G variant at the donor splice site of Exon 7, we performed a minigene assay (Fig. 3). Wild-type and mutant constructs were produced by cloning gDNA into the exontrap vector pET01 (Fig. 3A). Total RNA was extracted 72 h after transient transfection and analyzed with quantitative RT-PCR (Supplementary Fig. 3). Overall, wild-type and c.790A>G constructs produced bands of identical nucleotide sequences but in differing amounts (Fig. 3B). In particular, a splice variant only containing Exons 7 and 8 had significantly more wild-type 293T cells, and c.790A>G cells were barely detectable in an Exon 7-only variant. Alternate splicing of variable Exon 8 was not identified with the pET01 minigene system. While our findings show that c.790A>G exerts notable effects on splicing patterns, the exact in vivo effects of this variant on CIZ1 splicing in humans may show cell type specificity and developmental regulation.
FIGURE 3
FIGURE 3
Minigene assay of CIZ1 splicing. (A) Segment of CIZ1 containing Exons 6, 7, 8, and 9 were cloned into the multiple cloning site (MCS) of the pET01 exontrap vector, which contains a short stretch of a eukaryotic phosphatase gene (E) 3' to the eukaryotic (more ...)
Expression of CIZ1 in brain and lymphoblastoid cell lines
With RT-PCR, we identified two major CIZ1 isoforms in adult and fetal brain (Fig. 2, Supplementary Fig. 4). In adult human brain, both isoforms were present in cerebellum, cerebral cortex, substantia nigra, and putamen. With relative quantitative real-time RT-PCR, CIZ1 expression was greatest in fetal brain and in cerebellum among adult brain regions (Supplementary Table 5). Relative to CIZ1 Isoform 1, Isoform 2 was most strongly expressed in substantia nigra and cerebellum. In lymphoblastoid cell lines, the c.790A>G mutation had no effects on overall expression of CIZ1 and minimal effects on the ratio of Isoform1 to Isoform 2. However, our experiment may have lacked the power to detect an effect of the exonic splicing enhancer mutation on relative expression of Isoforms 1 and 2.
Subnuclear localization of CIZ1
To examine the effects of p.S264G on the subnuclear localization of CIZ1, wild-type and mutant proteins were expressed in 293T cells (Fig. 4). The p.S264G protein did not fail to enter nuclei or form subnuclear foci. However, particle analysis of GFP-CIZ1 transfected cells revealed that p.S264G CIZ1 formed fewer particles per nucleus than the wild-type protein. In comparison to wild-type particles, mutant particles were also larger and encompassed a larger area of the nucleus (Table 4).
FIGURE 4
FIGURE 4
Cellular localization of wild-type and mutant CIZ1 (S264G) in transiently transfected HEK 293T cells. CIZ1 was visualized with a GFP tag or rabbit anti-CIZ1 antibody. In comparison to wild-type CIZ1, average particle size was larger and the number of (more ...)
Table 4
Table 4
Particle Analysis of GFP-CIZ1 Transfected HEK 293T Cells
In a large Caucasian kindred with familial CD, we used linkage analysis and whole-exome sequencing to identify an exonic splicing enhancer mutation in Exon 7 of CIZ1 (c.790A>G, p.S264G). The microsatellite marker D9S159 on Chr 9q34.11 cosegregated with c.790A>G in all affected subjects. CIZ1 encodes Cip1-interacting zinc finger protein 1, a DNA replication factor. The c.790A>G variant was shown to alter splicing of CIZ1 and subnuclear localization of CIZ1. We also showed that both major isoforms of CIZ1 are expressed in motor regions of human brain. Using Sanger sequencing and HRM, 308 subjects with CD and 724 controls were screened for mutations in CIZ1. Two additional pathogenic variants (p.P47S and p.R672M) were identified in subjects with CD, but none were found in controls.
Although supported by several lines of evidence, the association between CIZ1 and CD should be interpreted cautiously. First, we did not identify CIZ1 mutations in a second multiplex kindred with CD. Second, we cannot exclude the possibility that other coding or noncoding variants on Chr 9q34.11 contribute to the pathogenesis of CD. Finally, with the exception of p.S264G, the predicted pathogenicity of all missense variants was limited to in silico analyses. MutationTaster and PolyPhen-2 are associated with accuracy rates of no better than 85.7% and 80.7%, respectively, and the false positive and negative rates for PolyPhen-2 are 21.4% and 16.8%, respectively.17
Although recessive mutations in SETX may be associated with ataxia-ocular apraxia-2 (AOA2; MIM 606002) and occasional subjects with AOA2 may show signs of dystonia, albeit in the lower extremities, all of our manifest CD subjects had otherwise normal neurological examinations without evidence of oculomotor apraxia, ataxia, spasticity, Parkinsonism, neuropathy, or muscle weakness.31 Autosomal dominant missense mutations in SETX have been associated with amyotrophic lateral sclerosis 4 (ALS4; MIM 602433). Because frameshift mutations have not been described in pedigrees with ALS4, it is possible that missense mutations in subjects with ALS4 function via a gain-of-function mechanism. Therefore, the SETX variant identified in Family A is probably benign and not requisite for the expression of CD. However, we cannot exclude the possibility that heterozygous loss-of-function SVs in SETX contribute to the expressivity of CIZ1 mutations.
Primary dystonia may be a neurodevelopmental network disorder of the central nervous system due to dysfunction at one or more nodes of the highly interconnected motor subsystem that includes the cerebellum and basal ganglia.32 The relatively high expression of CIZ1 in fetal brain and cerebellum are compatible with modern theories of dystonia pathobiology. CIZ1 is also expressed in a wide range of non-neural tissues such as spleen, thymus, heart, lung, liver, kidney, stomach, and testis.33CIZ1 harbors at least four alternatively spliced exons.34,35 Exon skipping may be more common in certain cancers, and a CIZ1 isoform due to alternative splicing of Exon 8 may be associated with Alzheimer disease.3436 In the context of CD, splicing defects in CIZ1 may also be critical to the pathogenesis of the c.790A>G variant.
CIZ1 was first identified through its interaction with p21Cip1/Waf1, a cyclin-dependent kinase inhibitor involved in G1/S cell-cycle regulation and cellular differentiation.33 In cell-free systems, CIZ1 is able to promote DNA replication after replication complex formation.37 The C-terminal domain anchors CIZ1 to the nonchromatin nuclear matrix, whereas DNA replication activity resides in the N-terminal half of the protein. Studies of GFP-tagged CIZ1 have shown that formation of subnuclear particles or foci requires both the N- and C-terminal domains.38
We showed that the p.S264G CIZ1 did enter nuclei and form subnuclear foci in cultured cells. However, particle analysis revealed that p.S264G CIZ1 formed larger particles than the wild-type protein. Therefore, p.S264G could exert deleterious effects via a gain-of-function mechanism associated with aberrant DNA synthesis and/or transcription.
Although possibly unique in its association with adult-onset and monosymptomatic focal dystonia, the cellular role and neural localization of CIZ1 are compatible with current themes in dystonia research. Clinically and biologically, CIZ1 shows the greatest similarity to the transcription factors THAP1 and TAF1. THAP1 regulates cell proliferation and G1/S cell-cycle progression39 and THAP1 loss-of-function mutations (DYT6) cause diverse anatomical patterns of childhood and adult-onset dystonia (including CD) with significant intrafamilial phenotypic variability.5,40 While often considered a neurodegenerative disorder, some patients with Lubag (DYT3) manifest isolated focal (including CD) or segmental dystonia for years before the development of Parkinsonism. DYT3 is associated with deficiency of a neuronal isoform of TAF1.41 TAF1 forms part of the TFIID transcriptional complex. Moreover, TAF1 induces G1/S progression by phosphorylating p53 at threonine-55.42 The best characterized primary dystonia, DYT1, is due to mutations in TOR1A and, like DYT6, shows significant intrafamilial variability and is predominantly of childhood onset.43 TorsinA, the protein encoded by TOR1A, is concentrated at the nuclear envelope in its mutant form; it appears to play a role in interactions between the nucleus and cytoskeleton44 and may participate in transcriptional regulation via the TGFβ pathway.45
In addition to significant functional overlap at the cellular level, CIZ1 shows similarities to THAP1 and TOR1A at the systems level. All three genes are robustly expressed in cerebellum, particularly Purkinje cells (Allen Brain Atlas), and recent anatomical studies in genetically engineered DYT1 mouse models suggest that primary dystonia may be a neurodevelopmental abnormality of cerebellar Purkinje cells.46CIZ1 gain-of-function mutations could illicit aberrant transcription during development or activation of the cell cycle in terminally differentiated neurons such as Purkinje cells.47
Supplementary Material
Supp Fig S1-S4 & Table S1-S5
Acknowledgements
At the University of Tennessee Health Science Center, this study was supported by the Neuroscience Institute (to MSL), Dystonia Medical Research Foundation (MSL), NIH grants R01NS048458 (MSL) and R01NS069936 (MSL), and NIH (U54NS065701) Dystonia Coalition Pilot Projects Program. At Washington University School of Medicine, work was supported by the NIH National Institute of Neurological Disease and Stroke grants P30NS05710 (Neuroscience Blueprint Grant) and Clinical Sciences Translation Award RR024992, the American Parkinson's Disease Association (APDA) Advanced Research Center, the Greater St. Louis Chapter of the APDA, the Barnes-Jewish Hospital Foundation (Jack Buck Fund for PD Research and the Elliot H. Stein Family Fund), the Missouri Chapter of the Dystonia Medical Research Foundation and the Murphy Fund. At Mayo Clinic Florida, work was supported by the NIH National Institute of Neurodegenerative Disease and Stroke Morris K. Udall Center of Excellence for Parkinson Disease Research grant (P50NS072187) and NINDS R01 grant (NS057567), the Mayo Clinic Florida Research Committee CR program (MCF 90052030), and the gift from Carl Edward Bolch, Jr., and Susan Bass Bolch (MCF 90052031/PAU 90052) (to RJU and ZKW). ZKW is also partially supported by the NIH/NINDS 1RC2NS070276 and the Dystonia Medical Research Foundation. GA and KL were supported by the BMBF (NGFN-plus). BL was supported by the DFG (German Research Foundation).
The Dystonia Coalition is part of the NIH Rare Diseases Clinical Research Network. Funding and /or programmatic support for this project has been provided by NS067501 from the NIH Office of Rare Diseases Research and the National Institute of Neurological Disorders and Stroke. The views expressed in written materials or publications do not necessarily reflect the official policies of the Department of Health and Human Services; nor does mention by trade names, commercial practices, or organizations imply endorsement by the U.S. Government.
We thank William Taylor for technical assistance with exome sequencing and Jill Searcy, Kate Marshall, and Ling Yan for help collecting clinical information.
Footnotes
Potential Conflicts of Interest
JX, YZ, SV, BL, and KL have none to report. RJU is an Associate Editor for Neurology and receives grant funding from the NIH. JSP receives grant funding from the NIH, Barnes-Jewish Hospital Foundation (Elliot Stein Family Fund and Jack Buck Fund for Parkinson Disease Research), American Parkinson Disease Research Association (APDA), Greater St. Louis Chapter of the APDA, Murphy Fund and Missouri Chapter of the Dystonia Medical Research Foundation; serves on the Scientific Advisory Board for the Dystonia Medical Research Foundation and Editorial Board of Neurology; has received compensation from the East St. Louis Law Firm, and honoraria from the University of Louisville, Toronto Western University, University of Maryland, and American Academy of Neurology. ZKW receives grant funding from the NIH; serves as co-editor-in-chief of Parkinsonism and Related Disorders and regional editor for Neurology, and receives compensation for patents filed by the Mayo Clinic. DMM has received grant funding from the NIH; honoraria from the American Academy of Neurology, Korean Movement Disorders Society, and Journal of Movement Disorders; and patent royalties. GA receives grant support from the DFG, European Union and BMBF. MSL serves on the speakers' bureaus for Lundbeck, Merz, and Teva Neuroscience; serves as an advisor for Merz; serves on the Xenazine Advisory Board for Lundbeck, Inc., and the Botulinum Toxin Type A Advisory Board for Allergan; receives research support from the NIH, Dystonia Medical Research Foundation, and Merz; and receives royalty payments for Animal Models of Movement Disorders (Elsevier).
1. Defazio G, Abbruzzese G, Livrea P, Berardelli A. Epidemiology of primary dystonia. Lancet Neurol. 2004;3:673–678. [PubMed]
2. Duane DD. Familial cervical dystonia, head tremor, and scoliosis: a case report. Adv Neurol. 1998;78:117–120. [PubMed]
3. Grandas F, Elston J, Quinn N, Marsden CD. Blepharospasm: a review of 264 patients. J Neurol Neurosurg Psychiatry. 1988;51:767–772. [PMC free article] [PubMed]
4. Chan J, Brin MF, Fahn S. Idiopathic cervical dystonia: clinical characteristics. Mov Disord. 1991;6:119–126. [PubMed]
5. Xiao J, Zhao Y, Bastian RW, et al. Novel THAP1 sequence variants in primary dystonia. Neurology. 2010;74:229–238. [PMC free article] [PubMed]
6. Uitti RJ, Maraganore DM. Adult onset familial cervical dystonia: report of a family including monozygotic twins. Mov Disord. 1993;8:489–494. [PubMed]
7. Bressman SB, Warner TT, Almasy L, et al. Exclusion of the DYT1 locus in familial torticollis. Ann Neurol. 1996;40:681–684. [PubMed]
8. Munchau A, Valente EM, Davis MB, et al. A Yorkshire family with adult-onset craniocervical primary torsion dystonia. Mov Disord. 2000;15:954–959. [PubMed]
9. Puschmann A, Xiao J, Bastian RW, et al. An African-American family with dystonia. Parkinsonism Rel Disord. 2011;17:547–550. [PMC free article] [PubMed]
10. Leube B, Kessler KR, Goecke T, et al. Frequency of familial inheritance among 488 index patients with idiopathic focal dystonia and clinical variability in a large family. Mov Disord. 1997;12:1000–1006. [PubMed]
11. Jarman PR, del Grosso N, Valente EM, et al. Primary torsion dystonia: the search for genes is not over. J Neurol Neurosurg Psychiatry. 1999;67:395–397. [PMC free article] [PubMed]
12. Xiao J, Zhao Y, Bastian RW, et al. The c.-237_236GA>TT THAP1 sequence variant does not increase risk for primary dystonia. Mov Disord. 2011;26:549–552. [PMC free article] [PubMed]
13. Gyapay G, Morissette J, Vignal A, et al. The 1993-94 Généthon human genetic linkage map. Nat Genet. 1994;7:246–339. [PubMed]
14. Abecasi GR, Cherny SS, Cookson WO, Cardon LR. MERLIN: rapid analysis of dense genetic maps using sparse gene flow trees. Nat Genet. 2002;30:97–101. [PubMed]
15. Adzhubei IA, Schmidt S, Peshkin L, et al. A method and server for predicting damaging missense mutations. Nat Methods. 2010;7:248–249. [PMC free article] [PubMed]
16. Ng PC, Henikoff S. Predicting deleterious amino acid substitutions. Genome Res. 2001;11:863–874. [PubMed]
17. Schwarz JM, Rödelsperger C, Schuelke M, Seelow D. MutationTaster evaluates disease-causing potential of sequence alterations. Nat Methods. 2010;7:575–576. [PubMed]
18. Larkin MA, Blackshields G, Brown NP, et al. Clustal W and Clustal X version 2.0. Bioinformatics. 2007;23:2947–2948. [PubMed]
19. Fairbrother WG, Yeh RF, Sharp PA, Burge CB. Predictive identification of exonic splicing enhancers in human genes. Science. 2002;297:1007–1013. [PubMed]
20. Zhang XH, Chasin LA. Computational definition of sequence motifs governing constitutive exon splicing. Genes Dev. 2004;18:1241–1250. [PubMed]
21. Desmet FO, Hamroun D, Lalande M, et al. Human Splicing Finder: an online bioinformatics tool to predict splicing signals. Nucleic Acids Res. 2009;37:e67. [PMC free article] [PubMed]
22. Brunak S, Engelbrecht J, Knudsen S. Prediction of human mRNA donor and acceptor sites from the DNA sequence. J Mol Biol. 1991;220:49–65. [PubMed]
23. Reese MG, Eeckman FH, Kulp D, Haussler D. Improved splice site detection in Genie. J Comput Biol. 1997;4:311–323. [PubMed]
24. Yeo G, Burge CB. Maximum entropy modeling of short sequence motifs with applications to RNA splicing signals. J Comput Biol. 2004;11:377–394. [PubMed]
25. Cartegni L, Wang J, Zhu Z, et al. ESEfinder: A web resource to identify exonic splicing enhancers. Nucleic Acids Res. 2003;31:3568–3571. [PMC free article] [PubMed]
26. Wang Z, Rolish ME, Yeo G, et al. Systematic identification and analysis of exonic splicing silencers. Cell. 2004;119:831–845. [PubMed]
27. Nguyen Ba AN, Pogoutse A, Provart N, Moses AM. NLStradamus: a simple Hidden Markov Model for nuclear localization signal prediction. BMC Bioinformatics. 2009;10:202. [PMC free article] [PubMed]
28. Rost B, Yachdav G, Liu J. The PredictProtein server. Nucleic Acids Res. 2004;32:W321–326. [PMC free article] [PubMed]
29. Blom N, Gammeltoft S, Brunak S. Sequence and structure-based prediction of eukaryotic protein phosphorylation sites. J Mol Biol. 1999;294:1351–1362. [PubMed]
30. Smith PJ, Zhang C, Wang J, et al. An increased specificity score matrix for the prediction of SF2/ASF-specific exonic splicing enhancers. Hum Mol Genet. 2006;15:2490–2508. [PubMed]
31. Anheim M, Monga B, Fleury M, et al. Ataxia with oculomotor apraxia type 2: clinical, biological and genotype/phenotype correlation study of a cohort of 90 patients. Brain. 2009;132:2688–2698. [PubMed]
32. Neychev VK, Gross RE, Lehericy S, et al. The functional neuroanatomy of dystonia. Neurobiol Dis. 2011;42:185–201. [PMC free article] [PubMed]
33. Mitsui K, Matsumoto A, Ohtsuka S, et al. Cloning and characterization of a novel p21(Cip1/Waf1)-interacting zinc finger protein, ciz1. Biochem Biophys Res Commun. 1999;264:457–464. [PubMed]
34. Rahman FA, Ainscough JF, Copeland N, Coverley D. Cancer-associated missplicing of exon 4 influences the subnuclear distribution of the DNA replication factor CIZ1. Human Mutat. 2007;28:993–1004. [PubMed]
35. Rahman FA, Aziz N, Coverley D. Differential detection of alternatively spliced variants of Ciz1 in normal and cancer cells using a custom exon-junction microarray. BMC Cancer. 2010;10:482. [PMC free article] [PubMed]
36. Dahmcke CM, Buchmann-Moller S, Jensen NA, Mitchelmore C. Altered splicing in exon 8 of the DNA replication factor CIZ1 affects subnuclear distribution and is associated with Alzheimer's disease. Mol Cell Neurosci. 2008;38:589–594. [PubMed]
37. Coverley D, Marr J, Ainscough J. Ciz1 promotes mammalian DNA replication. J Cell Sci. 2005;118:101–112. [PubMed]
38. Ainscough JF, Rahman FA, Sercombe H, et al. C-terminal domains deliver the DNA replication factor Ciz1 to the nuclear matrix. J Cell Sci. 2007;120:115–124. [PubMed]
39. Campagne S, Saurel O, Gervais V, Milon A. Structural determinants of specific DNA-recognition by the THAP zinc finger. Nucleic Acids Res. 2010;38:3466–3476. [PMC free article] [PubMed]
40. Fuchs T, Gavarini S, Saunders-Pullman R, et al. Mutations in the THAP1 gene are responsible for DYT6 primary torsion dystonia. Nat Genet. 2009;41:286–288. [PubMed]
41. Makino S, Kaji R, Ando S, et al. Reduced neuron-specific expression of the TAF1 gene is associated with X-linked dystonia-parkinsonism. Am J Hum Genet. 2007;80:393–406. [PubMed]
42. Li HH, Li AG, Sheppard HM, Liu X. Phosphorylation on Thr-55 by TAF1 mediates degradation of p53: a role for TAF1 in cell G1 progression. Mol Cell. 2004;13:867–878. [PubMed]
43. Ozelius LJ, Hewett JW, Page CE, et al. The early-onset torsion dystonia gene (DYT1) encodes an ATP-binding protein. Nat Genet. 1997;17:40–48. [PubMed]
44. Bragg DC, Armata IA, Nery FC, et al. Molecular pathways in dystonia. Neurobiol Dis. 2011;42:136–147. [PMC free article] [PubMed]
45. Koh YH, Rehfeld K, Ganetzky B. A Drosophila model of early onset torsion dystonia suggests impairment in TGF-beta signaling. Hum Mol Genet. 2004;13:2019–2030. [PubMed]
46. Zhang L, Yokoi F, Jin YH, et al. Altered dendritic morphology of Purkinje cells in Dyt1 DeltaGAG knock-in and Purkinje cell-specific Dyt1 conditional knockout mice. PloS One. 2011;6:e18357. [PMC free article] [PubMed]
47. Wang W, Bu B, Xie M, et al. Neural cell cycle dysregulation and central nervous system diseases. Prog Neurobiol. 2009;89:1–17. [PubMed]