The transition zone (TZ) ciliary subcompartment is thought to control cilium composition and signalling by facilitating a protein diffusion barrier at the ciliary base. TZ defects cause ciliopathies such as Meckel–Gruber syndrome (MKS), nephronophthisis (NPHP) and Joubert syndrome1 (JBTS). However, the molecular composition and mechanisms underpinning TZ organization and barrier regulation are poorly understood. To uncover candidate TZ genes, we employed bioinformatics (coexpression and co-evolution) and identified TMEM107 as a TZ protein mutated in oral–facial–digital syndrome and JBTS patients. Mechanistic studies in Caenorhabditis elegans showed that TMEM-107 controls ciliary composition and functions redundantly with NPHP-4 to regulate cilium integrity, TZ docking and assembly of membrane to microtubule Y-link connectors. Furthermore, nematode TMEM-107 occupies an intermediate layer of the TZ-localized MKS module by organizing recruitment of the ciliopathy proteins MKS-1, TMEM-231 (JBTS20) and JBTS-14 (TMEM237). Finally, MKS module membrane proteins are immobile and super-resolution microscopy in worms and mammalian cells reveals periodic localizations within the TZ. This work expands the MKS module of ciliopathy-causing TZ proteins associated with diffusion barrier formation and provides insight into TZ subdomain architecture.
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Reiter, J., Blacque, O. & Leroux, M. The base of the cilium: roles for transition fibres and the transition zone in ciliary formation, maintenance and compartmentalization. EMBO Rep. 13, 608–618 (2012).
Goetz, S. C. & Anderson, K. V. The primary cilium: a signalling centre during vertebrate development. Nat. Rev. Genet. 11, 331–344 (2010).
Blacque, O. E. & Sanders, A. A. Compartments within a compartment: what C. elegans can tell us about ciliary subdomain composition, biogenesis, function, and disease. Organogenesis 10, 126–137 (2014).
Hsiao, Y. C., Tuz, K. & Ferland, R. J. Trafficking in and to the primary cilium. Cilia 1, 4 (2012).
Chih, B. et al. A ciliopathy complex at the transition zone protects the cilia as a privileged membrane domain. Nat. Cell Biol. 14, 61–72 (2011).
Craige, B. et al. CEP290 tethers flagellar transition zone microtubules to the membrane and regulates flagellar protein content. J. Cell Biol. 190, 927–940 (2010).
Garcia-Gonzalo, F. R. et al. A transition zone complex regulates mammalian ciliogenesis and ciliary membrane composition. Nat. Genet. 43, 776–784 (2011).
Hu, Q. et al. A septin diffusion barrier at the base of the primary cilium maintains ciliary membrane protein distribution. Science 329, 436–439 (2010).
Kee, H. L. et al. A size-exclusion permeability barrier and nucleoporins characterize a ciliary pore complex that regulates transport into cilia. Nat. Cell Biol. 14, 431–437 (2012).
Williams, C. L. et al. MKS and NPHP modules cooperate to establish basal body/transition zone membrane associations and ciliary gate function during ciliogenesis. J. Cell Biol. 192, 1023–1041 (2011).
Gilula, N. B. & Satir, P. The ciliary necklace. A ciliary membrane specialization. J. Cell Biol. 53, 494–509 (1972).
Heller, R. F. & Gordon, R. E. Chronic effects of nitrogen dioxide on cilia in hamster bronchioles. Exp. Lung Res. 10, 137–152 (1986).
Roberson, E. C. et al. TMEM231, mutated in orofaciodigital and Meckel syndromes, organizes the ciliary transition zone. J. Cell Biol. 209, 129–142 (2015).
Cevik, S. et al. Active transport and diffusion barriers restrict Joubert syndrome-associated ARL13B/ARL-13 to an Inv-like ciliary membrane subdomain. PLoS Genet. 9, e1003977 (2013).
Huang, L. et al. TMEM237 is mutated in individuals with a Joubert syndrome related disorder and expands the role of the TMEM family at the ciliary transition zone. Am. J. Hum. Genet. 89, 713–730 (2011).
Jauregui, A. R., Nguyen, K. C., Hall, D. H. & Barr, M. M. The Caenorhabditis elegans nephrocystins act as global modifiers of cilium structure. J. Cell Biol. 180, 973–988 (2008).
Williams, C. L., Winkelbauer, M. E., Schafer, J. C., Michaud, E. J. & Yoder, B. K. Functional redundancy of the B9 proteins and nephrocystins in Caenorhabditis elegans ciliogenesis. Mol. Biol. Cell 19, 2154–2168 (2008).
Schouteden, C., Serwas, D., Palfy, M. & Dammermann, A. The ciliary transition zone functions in cell adhesion but is dispensable for axoneme assembly in C. elegans. J. Cell Biol. 210, 35–44 (2015).
Jensen, V. L. et al. Formation of the transition zone by Mks5/Rpgrip1L establishes a ciliary zone of exclusion (CIZE) that compartmentalises ciliary signalling proteins and controls PIP2 ciliary abundance. EMBO J. 34, 2537–2556 (2015).
Baughman, J. M. et al. A computational screen for regulators of oxidative phosphorylation implicates SLIRP in mitochondrial RNA homeostasis. PLoS Genet. 5, e1000590 (2009).
van Dam, T. J., Wheway, G., Slaats, G. G., Huynen, M. A. & Giles, R. H. The SYSCILIA gold standard (SCGSv1) of known ciliary components and its applications within a systems biology consortium. Cilia 2, 7 (2013).
Barker, A. R., Renzaglia, K. S., Fry, K. & Dawe, H. R. Bioinformatic analysis of ciliary transition zone proteins reveals insights into the evolution of ciliopathy networks. BMC Genomics 15, 531 (2014).
Christopher, K. J., Wang, B., Kong, Y. & Weatherbee, S. D. Forward genetics uncovers Transmembrane protein 107 as a novel factor required for ciliogenesis and Sonic hedgehog signaling. Dev. Biol. 368, 382–392 (2012).
Giles, R. H., Ajzenberg, H. & Jackson, P. K. 3D spheroid model of mIMCD3 cells for studying ciliopathies and renal epithelial disorders. Nat. Protoc. 9, 2725–2731 (2014).
Friedland, A. E. et al. Heritable genome editing in C. elegans via a CRISPR-Cas9 system. Nat. Methods 10, 741–743 (2013).
Starich, T. A. et al. Mutations affecting the chemosensory neurons of Caenorhabditis elegans. Genetics 139, 171–188 (1995).
Williams, C. L., Masyukova, S. V. & Yoder, B. K. Normal ciliogenesis requires synergy between the cystic kidney disease genes MKS-3 and NPHP-4. J. Am. Soc. Nephrol. 21, 782–793 (2010).
Valente, E. M. et al. Mutations in TMEM216 perturb ciliogenesis and cause Joubert, Meckel and related syndromes. Nat. Genet. 42, 619–625 (2010).
Kensche, P. R., van Noort, V., Dutilh, B. E. & Huynen, M. A. Practical and theoretical advances in predicting the function of a protein by its phylogenetic distribution. J. R. Soc. 5, 151–170 (2008).
Iglesias, A. et al. The usefulness of whole-exome sequencing in routine clinical practice. Genet. Med. 16, 922–931 (2014).
Shaheen, R. et al. Identification of a novel MKS locus defined by TMEM107 mutation. Hum. Mol. Genet. 24, 5211–5218 (2015).
Nakada, C. et al. Accumulation of anchored proteins forms membrane diffusion barriers during neuronal polarization. Nat. Cell Biol. 5, 626–632 (2003).
Xu, K., Zhong, G. & Zhuang, X. Actin, spectrin, and associated proteins form a periodic cytoskeletal structure in axons. Science 339, 452–456 (2013).
van Dam, T. J. et al. Evolution of modular intraflagellar transport from a coatomer-like progenitor. Proc. Natl Acad. Sci. USA 110, 6943–6948 (2013).
Sanders, A. A., Kennedy, J. & Blacque, O. E. Image analysis of Caenorhabditis elegans ciliary transition zone structure, ultrastructure, molecular composition, and function. Methods Cell Biol. 127, 323–347 (2015).
Hobert, O. PCR fusion-based approach to create reporter gene constructs for expression analysis in transgenic C. elegans. BioTechniques 32, 728–730 (2002).
Arts, H. H. et al. Mutations in the gene encoding the basal body protein RPGRIP1L, a nephrocystin-4 interactor, cause Joubert syndrome. Nat. Genet. 39, 882–888 (2007).
Dawe, H. R. et al. Nesprin-2 interacts with meckelin and mediates ciliogenesis via remodelling of the actin cytoskeleton. J. Cell Sci. 122, 2716–2726 (2009).
York, A. G., Ghitani, A., Vaziri, A., Davidson, M. W. & Shroff, H. Confined activation and subdiffractive localization enables whole-cell PALM with genetically expressed probes. Nat. Methods 8, 327–333 (2011).
This work was financially supported via the European Community’s Seventh Framework Programme FP7/2009 (SYSCILIA grant agreement 241955 to O.E.B., M.A.H., R.H.G. and C.A.J., and Gencodys to M.A.H.), Science Foundation Ireland (11/PI/1037 to O.E.B.), the Dutch Kidney Foundation CP11.18 ‘KOUNCIL’ (to R.H.G.), the GIS-Institut des Maladies Rares (HTS to C.T.-R.), the French Fondation for Rare Disease (to C.T.-R.), the Virgo consortium (FES0908 to M.A.H.), the Netherlands Genomics Initiative (050-060-452, RvdL to M.A.H.), the French Ministry of Health (PHRC national 2010-A01014-35 and 2013 to C.T.-R.), the Fondation pour la Recherche Médicale (DEQ20130326532 to S.S.), the Regional Council of Burgundy (to C.T.-R.), a Sir Jules Thorn Award for Biomedical Research (JTA/09 to C.A.J.), and the UK Medical Research Council (MR/K011154/1 to C.A.J., and MR/K015613/1 to M.P.). We thank the patients and their families for their participation. We also thank the NHLBI GO Exome Sequencing Project and its ongoing studies that produced and provided exome variant calls for comparison: the Lung GO Sequencing Project (HL-102923), the WHI Sequencing Project (HL-102924), the Broad GO Sequencing Project (HL-102925), the Seattle GO Sequencing Project (HL-102926) and the Heart GO Sequencing Project (HL-103010). We thank M. Leroux (Simon Fraser University, Canada), B. Yoder (University of Alabama, USA), the Caenorhabditis elegans Genetics Center (Minnesota, USA), the National Bioresource project (Tokyo, Japan), the International C. elegans gene knockout consortium, and the C. elegans Million Mutation Project for nematode reagents. We are grateful to C. Eggeling and C. Lagerholm (Weatherall Institute of Molecular Medicine and the Wolfson Imaging Center, Oxford, UK) for assistance with STED microscopy, D. Scholz and T. Toivonen (UCD Conway Institute imaging facility, Dublin, IRL) for imaging support, and R. Dijkstra (Scientific Volume Imaging bv, Hilversum, NL) for assistance with STED image deconvolution. We also thank A. Cleasby (Faculty of Biological Sciences, University of Leeds, Leeds, UK) for help with developing the dSTORM technique, B. Chih (Genentech, South San Francisco, CA, USA) for the kind gift of polyclonal anti-TMEM17 and TMEM231 antibodies, and T. McMorrow (University College Dublin, Dublin, Ireland) for the generous gift of the RPTEC/TERT1 cells. We thank D. Rodriguez (Trousseau hospital, Paris) for assistance with analysis of brain MRIs. The dSTORM microscope was generously funded by alumnus M. Beverly, in support of the University of Leeds ‘making a world of difference campaign’.
The authors declare no competing financial interests.
Integrated supplementary information
(a) Frequency histogram of binned mouse gene co-expression scores, derived from weighted analyses of gene expression datasets using a training set of 20 known TZ genes (Supplementary Table 1). This graph is the equivalent of the human gene co-expression dataset presented in Fig. 1a. Frequencies normalised to compare different distributions. Grey hatched; all human genes, yellow; ciliary genes in the SysCilia gold standard21, blue; TZ gene training set. Box-plots display median and quartiles for histogram distributions. (b) Presence and absence of candidate and known TZ genes in 52 eukaryotic species. The presence of orthologues for the 20 TZ training set genes and the five candidate TZ genes were determined by bi-directional best hits using BLAST and PSI-BLAST, as well as custom built hidden Markov models, HHPred, and intermediate sequence searches using PSI-BLAST and TBLASTN. Species are ordered according to their phylogenetic relationship as shown by the phylogenetic tree at the top. The top row indicates which species possess cilia or flagella. Grey columns indicate species lacking a (canonical) TZ. Ciliated species that have lost MKS genes appear to lack well defined Y-shaped linkers22. (c) Model of the four transmembrane helix topology of human TMEM107. Predicted transmembrane regions for TMEM107 and three known TZ proteins (TMEM216, TMEM138, and TMEM17) using TMHMM2.0 (http://www.cbs.dtu.dk/services/TMHMM). Alignment of TMEM107 sequences to the homologous TMEM216, TMEM138, and TMEM17 suggests TMEM107 is homologous to these three TZ proteins (not shown). (d) To model the transmembrane helices we used a standard existing helix obtained from the PDB. We swapped the amino acid side chains one by one using YASARA. The transmembrane topology of TMEM107 was predicted with TMHMM2.0. Helices are ordered anti-clockwise, starting with helix 1 in the right-rear, (bottom to top), helix 2 at the left-rear (top to bottom), helix 3 at the front-left (bottom to top) and helix 4 at the front-right (top to bottom). On the right side the four helices are depicted from a downwards viewpoint. The evolutionary conserved, charged residues (in red, a histidine and an arginine in helix 1, a glutamate in helix 2, a histidine in helix 3 and a glutamate in helix 4) are at the same height in the four helices, suggesting interactions, and therefore a four helix bundle model of the protein’s transmembrane structure. The conserved non-charged residues are in cyan. The mouse Schlei (E125G) mutation (the human equivalent is E131G)23 and the human F106Del and L134Ffs mutations found in this study are indicated by arrows. E45G lies within the extracellular loop between helix 1 and helix 2 and is not depicted here.
Supplementary Figure 2 Expression and localisation analyses of C. elegans and human TMEM107 constructs (wild type and variants).
(a) C. elegans tmem-107 is expressed exclusively in ciliated sensory neurons. Shown are fluorescence images of worms expressing a transcriptional tmem-107p::GFP reporter (P). DiI costain identifies a subset of ciliated neurons, namely 6 amphid cells (ADL, ASH, ASJ, ASK, AWB and ASI (not shown)) and both phasmid cells (PHA/B). Bars; 25 μm (large whole worm panels), 6 μm (small head and tail panels). den; dendrites, cil; cilia. (b) Schematics showing candidate X-box sequences in the promoters of human and nematode TMEM107. (c) DAF-19 RFX transcription factor is required for TMEM-107::GFP expression in C. elegans. Shown are head (left panels) and tail (right panels) regions of N2 (wild-type) and daf-19(m86);daf-12(sa204) double mutant worms expressing a translational tmem-107::gfp transgene (see Fig. 1c). Bars; 6 μm. (d) Analysis of TMEM-107 (wild type and variants) localisation in C. elegans. Shown are fluorescence images of the amphid and phasmid TZ regions (see also bottom schematic) in worms expressing various GFP tagged (C-terminus) TMEM-107 proteins. Top schematic shows the predicted topology of the tetraspan TMEM-107 C. elegans protein and indicates the disrupted domains and sequences. Linker 1 replacement sequence taken from SNG-1, and linker 2 and 3 replacement sequences taken from SPE-38 (see Methods section for further details). The coloured residues denote amino acids mutated in the TMEM107 patients (see Methods section for descriptions). TZ; transition zone. Bars; 1 μm. (e) Analysis of human TMEM107 patient variant protein localisation. Images of hTERT-RPE1 cells expressing GFP-tagged human TMEM107(E45G) or TMEM107(F106del), costained with antibodies for ciliary axonemes (acetylated tubulin; AcTub) and basal bodies (pericentrin). Bars; 10 μm.
Supplementary Figure 3 Sequencing details for the three cases of mutated TMEM107 and clinical details of TMEM107 patient phenotypes.
Integrative genomics viewer data showing: (a) compound heterozygous TMEM107 mutations in case 3 consisting of one frameshift deletion (NM_032354.3: g.8077560delT; p.Leu134Phefs∗8) and one in-frame deletion (NM_032354.3: g.8077890_ 8077893delGAA; p.Phe106del), and (b) homozygous TMEM107 missense variant (NM_183065: g.8079298T > C; p.Glu45Gly) in cases 1 and 2. Clinical details of the three TMEM107-mutated cases are presented in (c), leading to OFDVI and JBTS diagnoses. Cases 1 and 2 had previously been reported40.
(a) Cilium ultrastructure is highly disrupted in tmem-107;nphp-4 double mutants. Low (large panels) and high (small panels) magnification TEM images of cilia from serial cross sections taken from the distal (1), middle (2) and proximal (3) regions of the amphid pore (position of section in pore denoted by numbers in schematic). Wild-type pores consist of 10 ciliary axonemes (only three shown in schematics for simplicity), each consisting of a distal segment (DS; singlet A microtubules), a middle segment (MS; doublet A/B microtubules), a transition zone (TZ; with membrane-microtubule connecting Y-links) and a periciliary membrane compartment (PCMC). In tmem-107(oq100);nphp-4(tm925) double mutants (also harbours the him-5(e1490) mutation linked to nphp-4), multiple axonemes are missing in the middle and distal pore regions, TZ Y-links (Y’s) are reduced or missing, and vesicles frequently accumulate in the TZ and PCMC regions. Also, the majority of double mutant TZs are partially or fully disconnected (undocked) from the ciliary membrane, extending from ectopic anterior positions within the PCMC. In contrast, most or all of the ciliary axonemes are present in tmem-107(oq100) and nphp-4 single mutants. However, nphp-4 worms carrying the tm925 deletion (with or without him-5(e1490)) or the gk529336 nonsense mutation show consistent defects in Y-link integrity and TZs are undocked in two neurons (ADF, ADL). Images are representative of at least 4 analysed amphid pores for all strains except nphp-4(tm925) and nphp-4(gk529336) where 2 pores were analysed. Bars; 200 nm (low magnification images), 100 nm (high magnification images). (b) Compendium of TZ images and associated schematics showing the TZ defects outlined above in (a). Bars; 100 nm. (c) Dye filling assay (DiI) of tmem-107(oq100);nphp-4(tm925) worms transgenically expressing various GFP-tagged TMEM-107 constructs (wild type, E46G, F96del, L120G). Shown are fluorescence images of the head region. Non-transgenic worms are strongly dye-filling defective, whereas dye filling is restored in worms expressing TMEM-107 constructs (wild type or mutant versions). Bars; 10 μm.
(a) FRAP curve and representative time lapse images following quenching of 100% of MKS-2::GFP and TMEM-107::GFP signals at the TZ (boxed region shows the bleached TZ of an amphid channel cilium). Data points represented as mean ± s.d.n = 3 (MKS-2::GFP) or 4 (TMEM-107::GFP) independent experiments. Bar; 500 nm. (b) Raw and deconvolved (decon.) STED and confocal images of C. elegans MKS and NPHP module proteins (GFP-tagged). Bars; 500 nm. (c) Raw and deconvolved (decon.) STED and confocal images of renal RPTEC cells stained for polyglutamylated tubulin (ciliary axonemes; red; confocal only) and either endogenous human RPGRIP1L or TMEM67 (green; confocal and STED). STED imaging reveals that RPGRIP1L and TMEM67 form clusters of discrete signals arranged as a hollow ring at the TZ. Bars; 500 nm. (d) Super-resolution dSTORM microscopy of RPGRIP1L in the ciliary transition zone of human hTERT-RPE1 cells. The loose, tilted ring TZ organisation of RPGRIP1L shown in Fig. 5e (i) and examples from additional cells (ii-iv). dSTORM image reconstruction used 10 nm histogram bins, Gaussian image smoothing in the palm3d reconstruction output and contrast enhancement in FIJI. Dashed circles and ovals circumscribe TZ localisations which form the identified hollow loose ring structure, with discrete clusters of protein denoted by white arrowheads. Localisation density at individual points on the ring varied between samples, and was highest in (i). The distribution of signals in iv deviates significantly from an oval and could represent a partial spiral or helical arrangement. In some images (I, ii), some signal appears to enter the ciliary axoneme (ax) distal to the TZ. Images depth-coded by colour, with the z axis scale bar in nm indicated on the right. Representative bright-field and epifluoresence images from cells stained for RPGRIP1L and TMEM67, with the transition zone acquired and reconstructed in (iv) indicated by the white arrow. Red arrowheads indicate fiducials. Scale bars; 100 nm (dSTORM images; all images identically scaled), 10 μm (bright-field and epifluorescence images).
Red boxes denote the cropped regions shown in Fig. 4e.
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Reconstruction derived from a 200 nm section of a C. elegans amphid channel ciliary TZ. Arrow denotes a Y-link density throughout the tomogram, indicating that the Y-link structures are continuous sheets along the axial plane. Bar; 100 nm. (AVI 5111 kb)
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Lambacher, N., Bruel, AL., van Dam, T. et al. TMEM107 recruits ciliopathy proteins to subdomains of the ciliary transition zone and causes Joubert syndrome. Nat Cell Biol 18, 122–131 (2016). https://doi.org/10.1038/ncb3273
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