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Mutations in DZIP1L, which encodes a ciliary-transition-zone protein, cause autosomal recessive polycystic kidney disease

Nature Genetics volume 49, pages 1025–1034 (2017)Cite this article

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Abstract

Autosomal recessive polycystic kidney disease (ARPKD), usually considered to be a genetically homogeneous disease caused by mutations in PKHD1, has been associated with ciliary dysfunction. Here, we describe mutations in DZIP1L, which encodes DAZ interacting protein 1-like, in patients with ARPKD. We further validated these findings through loss-of-function studies in mice and zebrafish. DZIP1L localizes to centrioles and to the distal ends of basal bodies, and interacts with septin2, a protein implicated in maintenance of the periciliary diffusion barrier at the ciliary transition zone. In agreement with a defect in the diffusion barrier, we found that the ciliary-membrane translocation of the PKD proteins polycystin-1 and polycystin-2 is compromised in DZIP1L-mutant cells. Together, these data provide what is, to our knowledge, the first conclusive evidence that ARPKD is not a homogeneous disorder and further establish DZIP1L as a second gene involved in ARPKD pathogenesis.

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Figure 1: Mapping of a new ARPKD locus on 3q22.1–q23 and identification of DZIP1L mutations.
Figure 2: Dzip1lwpy/wpy-mutant embryos show multiple dysmorphic features.
Figure 3: Dzip1lwpy/wpy mice display progressive cystic kidney disease and hepatic defects after at least four crosses to a CD1 background.
Figure 4: DZIP1L localization overlaps with basal-body, centrosome and transition-zone markers.
Figure 5: Dzip1l is required for ciliary differentiation in the zebrafish kidney tubule.
Figure 6: DZIP1L associates with the ciliary-transition-zone protein SEPT2.
Figure 7: Loss of DZIP1L affects the localization of PC1 and PC2 to the ciliary membrane.

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References

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Acknowledgements

The authors thank the patients and their families for their cooperation and interest in the study; M.T. Toh and Y.T. Koh for technical assistance; W.I. Goh of the Institute of Medical Biology's Microscopy Unit for assistance with super-resolution microscopy; J. Lefevre and N. Hamilton for advice on statistical analyses; and C. Cortés, M. Pitt, F. Olsson, L. Zhao, L. Wilkinson and P. Karaith Oliva for assistance, insightful discussion and advice. The authors also thank C. Has (University Medical Center Freiburg) for kindly providing control fibroblasts; S. Somlo, M. Ma and K. Dong (Yale University) for Pkd1- and Pkd2-mutant cells; L. Lei (Nanyang Technological University) for the RPE-1 cell line stably expressing Arl13b–GFP; L. Pelletier (Lunenfeld-Tanenbaum Research Institute) for basal-body and transition-zone-protein cDNA clones; and R. Witzgall (Institute for Molecular and Cellular Anatomy, University of Regensburg), G. Wu (Center of Translational Cancer Research and Therapy, Beijing) and R. Rohatgi (Stanford University) for antibodies. We also thank the staff of the University of Queensland (UQ) QBP animal house for assistance with mouse husbandry and the Australian Phenomics Facility for maintaining mice throughout the screen. Confocal microscopy at UQ was carried out at the Institute for Molecular Bioscience Dynamic Imaging Facility for Cancer Biology, which is funded through the generous support of the Australian Cancer Research Foundation. We also acknowledge the Australian Microscopy & Microanalysis Facility (AMMRF) at the Centre for Microscopy and Microanalysis at UQ. M.H., S.N., V.F. and C.B. are supported as employees of Bioscientia/Sonic Healthcare; C.W. was supported as a recipient of a University of Queensland Vice-Chancellor's Senior Research Fellowship; and S.R. is supported as a Senior Principal Investigator at the Institute of Molecular and Cell Biology, Singapore. M.H.L. is supported as a Senior Principal Research Fellow of the Australian National Health and Medical Research Council (NHMRC). F.H. is supported as the Warren E. Grupe Professor. This work was supported by grants from the German Research Fund (DFG) to K.Z. and C.B., the DFG Collaborative Research Centre (SFB) KIDGEM 1140 and the Federal Ministry of Education and Research (BMBF, 01GM1515C) to C.B., the Australian NHMRC (APP1045464) to C.W., the National Institutes of Health NIH (DK068306) to F.H. and the Agency for Science, Technology and Research (A*STAR) of Singapore to W.H. and S.R. This paper is dedicated to the memory of Markus Nauck, who recently passed away.

Author information

Author notes

  1. Vicki Metzis

    Present address: Francis Crick Institute, London, UK

  2. Shubha Vij

    Present address: Temasek Life Sciences Laboratory Limited, National University of Singapore, Singapore

  3. Melissa H Little

    Present address: Murdoch Children's Research Institute and Department of Pediatrics, University of Melbourne, Melbourne, Victoria, Australia

  4. Peter Papathanasiou

    Present address: Department of Materials, Imperial College London, London, UK

  5. Hao Lu, Maria C Rondón Galeano and Elisabeth Ott: These authors contributed equally to this work.

  6. Carol Wicking and Carsten Bergmann: These authors jointly directed this work.

Authors and Affiliations

  1. Institute of Molecular and Cell Biology, Singapore

    Hao Lu, P Jaya Kausalya, Shang Yew Tay, Shubha Vij, Walter Hunziker & Sudipto Roy

  2. Institute for Molecular Bioscience, The University of Queensland, Brisbane, Queensland, Australia

    Maria C Rondón Galeano, Geraldine Kaeslin, Vicki Metzis, Andrew D Courtney, Melissa H Little, Andrew C Perkins & Carol Wicking

  3. Department of Medicine IV, Medical Center–University of Freiburg, Faculty of Medicine, University of Freiburg, Freiburg, Germany

    Elisabeth Ott, Carina Kramer, Daniel Epting & Carsten Bergmann

  4. Institute of Human Genetics, RWTH Aachen University, Aachen, Germany

    Nadina Ortiz-Brüchle, Nadescha Hilger, Klaus Zerres & Carsten Bergmann

  5. Center for Human Genetics, Bioscientia, Ingelheim, Germany

    Milan Hiersche, Steffen Neuber, Valeska Frank & Carsten Bergmann

  6. John Curtin School of Medical Research, Australian National University, Acton, Australian Capital Territory, Australia

    Robert Tunningley, Belinda Whittle & Peter Papathanasiou

  7. Division of Pediatric Nephrology, University Children's Hospital Center for Child and Adolescent Medicine, Heidelberg University Hospital, Heidelberg, Germany

    Elke Wühl

  8. Department of Pediatric Nephrology, University Children's Hospital Essen, Essen, Germany

    Udo Vester

  9. Institute of Pathology, MHH University Medical School Hannover, Hannover, Germany

    Björn Hartleben

  10. Mater Research Institute, Faculty of Medicine and Biomedical Sciences, The University of Queensland, Woolloongabba, Queensland, Australia

    Andrew C Perkins

  11. Institute of Medical Biology, A*STAR, Singapore

    Graham D Wright

  12. Department of Physiology, Yong Loo Lin School of Medicine, National University of Singapore, Singapore

    Walter Hunziker

  13. Singapore Eye Research Institute, Singapore

    Walter Hunziker

  14. Department of Medicine, Boston Children's Hospital, Harvard Medical School, Boston, Massachusetts, USA

    Heon Yung Gee & Friedhelm Hildebrandt

  15. Department of Pharmacology, Brain Korea 21 PLUS Project for Medical Sciences, Yonsei University College of Medicine, Seoul, Republic of Korea

    Heon Yung Gee

  16. Department of Pediatrics and Communicable Diseases, University of Michigan, Ann Arbor, Michigan, USA

    Edgar A Otto

  17. Department of Pediatrics, Yong Loo Lin School of Medicine, National University of Singapore, Singapore

    Sudipto Roy

  18. Department of Biological Sciences, National University of Singapore, Singapore

    Sudipto Roy

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  1. Hao Lu
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  2. Maria C Rondón Galeano
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Contributions

H.L., P.J.K., S.Y.T., S.V. and S.R. performed the zebrafish mutant and morphant analyses, protein interaction studies, localization experiments with human fibroblasts and validation of antibodies to PCs. W.H. supervised the protein interaction studies. E.O., C.K. and D.E. performed the zebrafish morpholino analyses. M.C.R.G., G.K., V.M. and A.D.C. performed the mouse analyses, and M.C.R.G. and G.K. performed the localization experiments with mouse cells. R.T., P.P. and A.C.P. performed the screen that identified the mouse mutant, and B.W. was involved in mapping and identifying the mouse mutation. G.D.W. performed the super-resolution microscopy experiments. M.H.L. assisted with design and analysis of mouse kidney experiments. N.O.-B., N.H., V.F. and S.N. performed the human mutation analysis. S.N., V.F., M.H., H.Y.G., E.A.O., F.H. and C.B. carried out the WES data processing and analyses. S.N., V.F., E.W., U.V., H.Y.G., K.Z., F.H. and C.B. recruited and clinically characterized the study subjects and collected samples. B.H. performed histologic evaluation of the data. S.R., C.W. and C.B. conceived the project, designed and supervised the experiments, analyzed and interpreted the data and wrote the manuscript. All authors reviewed the final manuscript.

Corresponding authors

Correspondence to Sudipto Roy, Carol Wicking or Carsten Bergmann.

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Competing interests

The authors declare no competing financial interests.

Integrated supplementary information

Supplementary Figure 1 Pedigree and DZIP1L mutation data.

Top: Pedigrees and sequence electropherograms of the patients described in this study depicting the respective homozygous DZIP1L mutations. Bottom: Renal ultrasound data of patients with DZIP1L mutations demonstrating bilaterally enlarged, hyperechogenic kidneys with poor cortico-medullary differentiation and multiple tiny cysts characteristic of ARPKD. (a-e) Longitudinal sections of right (a, patient B16-3; d, patient A3533-1; e, patient B8031-1) and left (b,c, patient B16-2) kidney with a large number of echogenic spots and tiny, predominantly medullary cysts throughout the kidney (“pepper-salt pattern”). (f,g) Transverse (f) and longitudinal (g) section of the left kidney of patient B8031-1 with multiple small cysts and echogenic spots. Color doppler with multiple twinkling phenomena identifying these spots as small calcifications (g).

Supplementary Figure 2 Sequence alignment of human, mouse and zebrafish DZIP1L proteins.

Multiple sequence alignment of human (Accession number: NM_173543.2), mouse (Accession number: NM_028258.4) and zebrafish DZIP1L proteins (Accession number: NM_001037227.1) using Clustal W method. Amino acid identity is shaded (black). The zinc finger (red) and coiled-coil domains (blue) are highlighted.

Supplementary Figure 3 Characterization of Dzip1lwpy/wpy mice.

(a) Sanger sequencing scans of the wild-type (top) and homozygous Dzip1lwpy/wpy mutation (bottom), and the location of the stop codon in the predicted protein (on the right). Asterisks mark the mutated base, arrow marks location of the stop codon. (b) Dzip1lwpy/wpy mutant embryos show evidence of altered hedgehog signalling in the limb. Indicated markers analysed by wholemount in situ hybridisation in 11.5 dpc limbs show expanded ectopic expression of Hoxd13, Grem1 and Gli1 into the anterior of the Dzip1lwpy/wpy mutant forelimb bud (arrows in b). Anterior is to the top in all limb images. Similar data were obtained in hindlimbs (not shown). n = limbs from 3 independent embryos. (c) IF analysis with a battery of markers to hedgehog-dependent neural populations reveals no consistent difference between wild-type control and Dzip1lwpy/wpy mutant sections through the neural tube of 10.5 dpc embryos at the level of the forelimb. Similar data were obtained with sections at hindlimb level (not shown). n = multiple sections from 3 independent embryos for all markers except Shh (n=2). Scale bar in b = 200μm; c = 100μm. (d) qRT-PCR analysis of Gli1 transcript levels shows that, compared to Dzip1l+/+ MEFs, Dzip1lwpy/wpy MEFs fail to significantly upregulate HH signalling in response to SAG treatment (n=4 cell lines derived from independent embryos, cells treated in three separate experiments; each data point is the average of 3 separate culture wells). Statistical analysis based on separate unpaired two-tailed Student’s t-tests for Dzip1l+/+ t(6)=14.91, ****p<0.0001, and Dzip1lwpy/wpy t(6)=0.6943, p=0.5135, ns, not significant, +/-SAG treatment. Error bars, s.e.m. All data presented are from embryos on a C57BL/6:C3H mixed background.

Supplementary Figure 4 Examples of histological analyses of liver sections.

(a-f) Elastica van Gieson (EvG) and (g-l) Silver staining of liver sections from Dzip1l+/+ (a-c; g-j) and Dzip1lwpy/wpy (d-f; j-l) mice at P21, following at least 4 crosses to a CD1 background. Arrows mark bile ducts, showing an excess of bile ducts surrounding blood vessels in Dzip1lwpy/wpy mutant mice. Scale bar = 50μm.

Supplementary Figure 5 Maternal expression of dzip1l, efficacy of the dzip1l-splice morpholinos and generation of the dzip1l-deletion allele.

(a) The maternal expression of dzip1l mRNA at the 64-cell stage of zebrafish embryonic development. Zygotic expression is shown for the 10 somite stage. acta1b expression was used as loading control. (b) Splice morpholino (e5i5MO) efficiently interferes with dzip1l pre-mRNA splicing. RT-PCR with primers binding to exon2 and exon10, expected band of about 650 bp is visible from cDNA of wild-type embryos (arrow). A smaller band of 350 bp (arrowhead) is visible from cDNA of the morphants. Sequencing of this band showed deletion of exon 5-7. The lower band (asterisk) is a non-specific band. acta1b expression was used as loading control. (c), Splice morpholino (i1e2MO) interference with dzip1l pre-mRNA splicing. RT-PCR with primers binding to exon1 and exon3, expected band of about 680 bp is visible from cDNA of wild-type embryos (arrow) and significantly reduced in cDNA of morphant embryos. Wild-type control band was confirmed by sequencing, lower bands are unspecific. (d) Splice morpholino (i4e5MO) interferes with dzip1l pre-mRNA splicing. RT-PCR with primers binding to exon2 and exon10, expected 870 bp band is visible in cDNA of wild-type embryos and missing in morphant embryos (arrow). eef1a1l1 expression was used as loading control. (e) Schematic diagram of CRISPR/Cas9 mediated genomic modification of the dzip1l locus. Two guide RNAs, targeting exon2 and exon13, were designed to excise the intervening genomic DNA, to create a deletion allele. (f) PCR screening of CRISPR/Cas9 mediated dzip1l deletion among F0 founder fish with primers binding to exon2 and exon14. Fish 1, 2 and 3 did not carry the dzip1l deletion allele. Fish 4 and 5 were identified as dzip1l deletion allele carriers with the expected PCR product of 350 bp (arrow). Sequencing of this band showed the expected deletion. (g) Three otoliths in the otic vesicle of a 24 hpf maternal-zygotic dzip1l mutant embryo (n = 10 embryos).

Supplementary Figure 6 dzip1l zebrafish morphants display ciliopathy-associated phenotypes.

(a) Wholemount of a 20 hpf zebrafish embryo showing dzip1l expression in the nasal placode (short arrow) and pronephric duct (long arrow). (b) dzip1l expression in MCC precursors in the pronephric duct at 24 hpf (arrows). Inset shows expression in individual MCC precursors. (c) dzip1l expression in the nasal placode at 24 hpf (arrows). (d) Wild-type control embryo at 3 dpf. (e) Two otoliths in the otic vesicle of a wild-type embryo at 3 dpf. (f) dzip1l translation blocking morpholino (dzip1l MO) injected embryo at 3 dpf with shortened, slightly curved body, hydrocephalus (arrow) and reduced retinal size (arrowhead) (g) Three otoliths in the otic vesicle of a dzip1l morpholino (i4e5) injected embryo at 3 dpf. (h) Wild-type glomeruli (upper left panel) and cystic dilated glomeruli in dzip1l MO injected embryos (lower left panel, white asterisk) at 48 hpf. (i) Incidence of glomerular cysts and otolith deposition defects in control (co MO), translation blocking (TB MO) and splice site (i1e2 MO and i4e5 MO) morpholino injected Tg(wt1b::GFP) embryos at 48 hpf (for glomerular analysis n = 425 embryos for translation blocking morpholino, n = 349 embryos (i1e2) and n = 384 embryos (i4e5) for splice morpholinos; t(13)=5.68, ***p=0.0008 (i1e2 MO), t(17)=8.36 and t(14)=8.27, ****p <0.0001 (TB and i4e5 MO); for otolith analysis n = 243 embryos for translation blocking morpholino, n = 236 embryos (i1e2) and n = 326 embryos (i4e5) for splice morpholinos; t(7)=5.42, **p=0.0011 (TB MO), t(6)=14.93, ****p<0.0001(i1e2 MO), t(7)=4.34, **p=0.003 (i4e5MO). Statistical analysis based on an unpaired two-tailed Student`s t-test. Error bars, s.e.m. (j) Co-injection of zebrafish dzip1l mRNA (ddzip1l) can rescue dzip1l morpholino caused phenotypes; n = 463 embryos for morpholino and n = 313 embryos for mRNA co-injected embryos; t(9)=9.06, ****p<0.0001. Scale bars in a-g = 100μm; h = 50μm; each data point represents a single experiment.

Supplementary Figure 7 Analysis of protein and transcript levels in DZIP1L-mutant cells.

a,b) Staining with a C-terminal DZIP1L antibody (Sigma; green) reveals no signal in Dzip1lwpy/wpy MEFs (arrows in b), consistent with the truncated nature of the protein; (c,d,) Staining with an antibody raised to the entire DZIP1L protein (Abnova; green) reveals a residual level of truncated protein in Dzip1lwpy/wpy MEFs (arrows in d). The cilium is marked with acetylated-α-tubulin and γ-tubulin in (a,b; magenta) and IFT88 in (c,d; magenta). (e) Based on an unpaired two-tailed Student’s t-test, qRT-PCR revealed significantly reduced but not absent Dzip1l transcript levels in Dzip1lwpy/wpy MEFs relative to Dzip1l+/+ wild-type MEFs, t(6)=5.905, **p=0.001 (n=4, consisting of 3 cell lines derived from independent embryos, analysed across two experiments (separate RNA preparations from one cell line analysed in both experiments); each data point is the average of 3 separate culture wells). Dzip1 levels were not altered, t(6)=0.385, p=0.7135, ns, not significant, to compensate for reduced Dzip1l. Error bars, s.e.m. (f) DZIP1L (magenta) localizes at the transition zone in control human dermal fibroblasts, but is not detectable in DZIP1L mutant dermal fibroblasts from individual B155 (p.Gln155*) with the C-terminal Sigma antibody (arrows in g). Ciliary axonemes and basal bodies were labelled with anti-acetylated tubulin and γ-tubulin antibodies, respectively (green). (h) Immunoblotting with anti-DZIP1L antibody (Abnova) revealed a band corresponding to full length DZIP1L in control cell lysate (arrow) but not in lysate from DZIP1L mutant human fibroblasts. Scale bars in a-d = 2.5μm; g,h = 1μm.

Supplementary Figure 8 Dzip1lwpy/wpy-mutant embryos show defects in ciliogenesis ex vivo but not in vivo.

(a) Staining for cilia in serum-starved MEFs revealed a defect in the ability of isolated Dzip1lwpy/wpy mutant cells to ciliate relative to wild-type cells. Image and quantification based on ARL13B marking the cilium (red, arrows in a), but a significant reduction was also seen with acetylated-α-tubulin staining (n = 6 cell lines, each from an independent embryo; between 100-540 cells counted per cell line (1434 Dzip1l+/+ and 1837 Dzip1lwpy/wpy cells in total), across two separate experiments). (b) Imaging and quantification of cilia in 11.5dpc forelimb buds following staining with acetylated-α-tubulin revealed no significant difference in cilia frequency (n = forelimb from 3 independent embryos, with 9 sections analysed per limb, >1900 cells counted per limb). Note the final cilia/nuclei ratio in limb sections will not necessarily represent the actual percentage of ciliated cells in the tissue, but is a relative value across the samples (see online methods for details). Statistical analysis based on an unpaired, two-tailed Student’s t-test. (a) t(10)=10.87, ****p<0.0001; (b) t(4)=0.63, p=0.5624, ns, not significant. Error bars, s.e.m. Scale bar in a,b = 10μm. All data presented are from embryos on a C57BL/6:C3H mixed background.

Supplementary Figure 9 DZIP1L specifically associates with the ciliary-transition-zone protein SEPT2.

(a) Co-IP experiments from HEK293T cells transfected with expression constructs for epitope-tagged SEPT2 (C-terminal HA) and DZIP1L, CEP89, B9D1, MKS1 and NPHP1 (N-terminal Myc), reveal a specific interaction between SEPT2 and DZIP1L but none of the other proteins analysed. (b) Co-IP with DZIP1L (C-terminal HA) and SEPT2, CEP89, B9D1, MKS1 and NPHP1 (N-terminal Myc) confirmed the DZIP1L-SEPT2 interaction, and showed that none of the other proteins analysed interact with DZIP1L. Co-IP experiments were repeated three times.

Supplementary Figure 10 Validation of the 7e12 antibody to PC1.

(a,b) PC1 antibody (mAb 7e12 raised to the N-terminal leucine rich repeat region of PC1 (24-180aa), aliquots obtained from both Abcam and Santa Cruz Biotechnology – see online methods for details) detected overexpressed PC1 protein (green) in IMCD3 cells. (c-j) PC1 staining (green) on the ciliary membrane in Pkd1 or Pkd2 heterozygote control cells, and lack of staining in null mutant cells, with the PC1 7e12 antibody preparation indicated. Acetylated-α-tubulin (magenta) marks the ciliary axoneme. (k,l) Quantification of the percentage of ciliated cells with PC1 along the axoneme in Pkd1 or Pkd2 heterozygous control cells. No homozygous mutant cells analysed displayed detectable PC1 staining along the axoneme. Note that PC2 is required for ciliary trafficking of PC1, so axonemal PC1 staining would not be expected in Pkd2 null cells. Cilia were scored from two independent experiments. Approximately 100 ciliated cells were analysed in each experiment. The values are not compared statistically. Error bars, s.e.m. Scale bars = 1μm. See online methods for details of the Pkd1 and Pkd2 cell lines used for these validation experiments.

Supplementary Figure 11 Validation of antibody to PC2 and localization of SMO in Dzip1lwpy/wpy MEFs.

(a, b) PC2 staining (green) on the ciliary membrane in Pkd2 heterozygote control but not null mutant cells with the PC2 antibody (MV12). Acetylated-α-tubulin (magenta) marks the ciliary axoneme. (c) Quantification of the percentage of ciliated cells with PC2 along the axoneme in Pkd2 heterozygote and homozygote mutant cells. Statistical analysis based on an unpaired two-tailed Student’s t-test, t(5)=2.57,**p=0.0033. Cilia were scored in three experiments, with cells from two independent coverslips analysed for each experiment (approximately 100 ciliated cells scored on each coverslip). (d,e) In response to treatment with the HH signalling agonist SAG, SMO (green) localizes along the axoneme (acetylated-α-tubulin, magenta) in both Dzip1l+/+ and Dzip1lwpy/wpy MEFs. (f) quantification shows no difference in SMO localization between the two genotypes (n=4 MEF cell lines derived from individual embryos, analysed across two experiments; 28-91 cilia scored per cell line, total 299 Dzip1l+/+ cilia and 137 Dzip1lwpy/wpy cilia scored). Statistical analysis based on an unpaired two-tailed Student’s t-test, t(6)=0.0292, p=0.9776, ns, not significant. Error bars, s.e.m. Scale bars in a,b = 1μm; d,e = 2μm.

Supplementary Figure 12 Comparison of REVEL and CADD pathogenicity-prediction scores between DZIP1L and PKHD1.

A higher score indicates a more likely pathogenic effect of nucleotide exchanges at the corresponding position. For DZIP1L, red bars denote SNVs reported in this study, and green bars denote missense or loss-of-function variants found in ExAC. Height of green bars corresponds to the ExAC allele frequency in percent. (a) REVEL score. The x-axis refers to the coding DNA position of the longest transcript of the respective gene (DZIP1L = ENST00000327532; PKHD1 = ENST00000371117). Exon numbers and bounds are drawn above the axis. The y-axis displays scores for all possible amino acid changes at the corresponding position, averaged with a scope of 4 bases to the left and right by loess smoothing (b) CADD score. The x-axis refers to the coding DNA position of the longest transcript of the respective gene (DZIP1L = ENST00000327532; PKHD1 = ENST00000371117). Exon numbers and bounds are drawn above the axis. The y-axis displays scores for all possible amino acid changes at the corresponding position, averaged with a scope of 3 bases to the left and right by application of a Daniell kernel function.

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Supplementary Figures 1–12 and Supplementary Tables 1 and 3. (PDF 2700 kb)

Supplementary Table 2

DZIP1L Y2H screen. (XLS 26 kb)

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Lu, H., Galeano, M., Ott, E. et al. Mutations in DZIP1L, which encodes a ciliary-transition-zone protein, cause autosomal recessive polycystic kidney disease. Nat Genet 49, 1025–1034 (2017). https://doi.org/10.1038/ng.3871

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