Abstract
The vertebrate left–right axis is specified during embryogenesis by a transient organ: the left–right organizer (LRO). Species including fish, amphibians, rodents and humans deploy motile cilia in the LRO to break bilateral symmetry, while reptiles, birds, even-toed mammals and cetaceans are believed to have LROs without motile cilia. We searched for genes whose loss during vertebrate evolution follows this pattern and identified five genes encoding extracellular proteins, including a putative protease with hitherto unknown functions that we named ciliated left–right organizer metallopeptidase (CIROP). Here, we show that CIROP is specifically expressed in ciliated LROs. In zebrafish and Xenopus, CIROP is required solely on the left side, downstream of the leftward flow, but upstream of DAND5, the first asymmetrically expressed gene. We further ascertained 21 human patients with loss-of-function CIROP mutations presenting with recessive situs anomalies. Our findings posit the existence of an ancestral genetic module that has twice disappeared during vertebrate evolution but remains essential for distinguishing left from right in humans.
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Data availability
Data from the phylogenomic analysis are available at https://zenodo.org/record/5188601. Source data are provided with this paper. The remaining data supporting the findings of this study are available from the corresponding author upon reasonable request.
Code availability
Code used for the phylogenomic analysis described in this manuscript is available at https://zenodo.org/record/5188601.
Change history
18 March 2022
A Correction to this paper has been published: https://doi.org/10.1038/s41588-022-01053-8
References
Blum, M. & Ott, T. Animal left–right asymmetry. Curr. Biol. 28, R301–R304 (2018).
Shinohara, K. & Hamada, H. Cilia in left-right symmetry breaking. Cold Spring Harb. Perspect. Biol. 9, a028282 (2017).
Grimes, D. T. Making and breaking symmetry in development, growth and disease. Development 146, dev170985 (2019).
Grimes, D. T. & Burdine, R. D. Left-right patterning: breaking symmetry to asymmetric morphogenesis. Trends Genet. 33, 616–628 (2017).
Sutherland, M. J. & Ware, S. M. Disorders of left–right asymmetry: heterotaxy and situs inversus. Am. J. Med. Genet. C. Semin. Med. Genet. 151C, 307–317 (2009).
Lin, A. E. et al. Laterality defects in the national birth defects prevention study (1998–2007): birth prevalence and descriptive epidemiology. Am. J. Med. Genet. A 164A, 2581–2591 (2014).
Mohapatra, B. et al. Identification and functional characterization of NODAL rare variants in heterotaxy and isolated cardiovascular malformations. Hum. Mol. Genet. 18, 861–871 (2009).
Karkera, J. D. et al. Loss-of-function mutations in growth differentiation factor-1 (GDF1) are associated with congenital heart defects in humans. Am. J. Hum. Genet. 81, 987–994 (2007).
Kosaki, R. et al. Left–right axis malformations associated with mutations in ACVR2B, the gene for human activin receptor type IIB. Am. J. Med. Genet. 82, 70–76 (1999).
Bamford, R. N. et al. Loss-of-function mutations in the EGF-CFC gene CFC1 are associated with human left–right laterality defects. Nat. Genet. 26, 365–369 (2000).
Kosaki, K. et al. Characterization and mutation analysis of human LEFTY A and LEFTY B, homologues of murine genes implicated in left–right axis development. Am. J. Hum. Genet. 64, 712–721 (1999).
Cristo, F. et al. Functional study of DAND5 variant in patients with congenital heart disease and laterality defects. BMC Med. Genet. 18, 77 (2017).
Sempou, E. & Khokha, M. K. Genes and mechanisms of heterotaxy: patients drive the search. Curr. Opin. Genet. Dev. 56, 34–40 (2019).
Perles, Z. et al. A human laterality disorder associated with recessive CCDC11 mutation. J. Med. Genet. 49, 386–390 (2012).
Narasimhan, V. et al. Mutations in CCDC11, which encodes a coiled-coil containing ciliary protein, causes situs inversus due to dysmotility of monocilia in the left–right organizer. Hum. Mutat. 36, 307–318 (2015).
Vetrini, F. et al. Bi-allelic mutations in PKD1L1 are associated with laterality defects in humans. Am. J. Hum. Genet. 99, 886–893 (2016).
Gebbia, M. et al. X-linked situs abnormalities result from mutations in ZIC3. Nat. Genet. 17, 305–308 (1997).
Perles, Z. et al. A human laterality disorder caused by a homozygous deleterious mutation in MMP21. J. Med. Genet. 52, 840–847 (2015).
Guimier, A. et al. MMP21 is mutated in human heterotaxy and is required for normal left–right asymmetry in vertebrates. Nat. Genet. 47, 1260–1263 (2015).
Akawi, N. et al. Discovery of four recessive developmental disorders using probabilistic genotype and phenotype matching among 4,125 families. Nat. Genet. 47, 1363–1369 (2015).
Gros, J., Feistel, K., Viebahn, C., Blum, M. & Tabin, C. J. Cell movements at Hensen’s node establish left/right asymmetric gene expression in the chick. Science 324, 941–944 (2009).
Kajikawa, E. et al. Nodal paralogues underlie distinct mechanisms for visceral left–right asymmetry in reptiles and mammals. Nat. Ecol. Evol. 4, 261–269 (2020).
Aamar, E. & Dawid, I. B. Sox17 and chordin are required for formation of Kupffer’s vesicle and left-right asymmetry determination in zebrafish. Dev. Dyn. 239, 2980–2988 (2010).
Choksi, S. P., Babu, D., Lau, D., Yu, X. & Roy, S. Systematic discovery of novel ciliary genes through functional genomics in the zebrafish. Development 141, 3410–3419 (2014).
Maisonneuve, C. et al. Bicaudal C, a novel regulator of Dvl signaling abutting RNA-processing bodies, controls cilia orientation and leftward flow. Development 136, 3019–3030 (2009).
Beckers, A., Alten, L., Viebahn, C., Andre, P. & Gossler, A. The mouse homeobox gene Noto regulates node morphogenesis, notochordal ciliogenesis, and left right patterning. Proc. Natl Acad. Sci. USA 104, 15765–15770 (2007).
Huang, C.-J., Tu, C.-T., Hsiao, C.-D., Hsieh, F.-J. & Tsai, H.-J. Germ-line transmission of a myocardium-specific GFP transgene reveals critical regulatory elements in the cardiac myosin light chain 2 promoter of zebrafish. Dev. Dyn. 228, 30–40 (2003).
Blum, M. et al. Xenopus, an ideal model system to study vertebrate left-right asymmetry. Dev. Dyn. 238, 1215–1225 (2009).
Szenker-Ravi, E. et al. RSPO2 inhibition of RNF43 and ZNRF3 governs limb development independently of LGR4/5/6. Nature 557, 564–569 (2018).
Hamada, H in Mouse Development (eds Rossant, J & Tam, P. P. L.) Left-right asymmetry, 55–73 (Academic Press, 2002).
Campione, M. et al. The homeobox gene Pitx2: mediator of asymmetric left-right signaling in vertebrate heart and gut looping. Development 126, 1225–1234 (1999).
Logan, M., Pagán-Westphal, S. M., Smith, D. M., Paganessi, L. & Tabin, C. J. The transcription factor Pitx2 mediates situs-specific morphogenesis in response to left-right asymmetric signals. Cell 94, 307–317 (1998).
Ryan, A. K. et al. Pitx2 determines left-right asymmetry of internal organs in vertebrates. Nature 394, 545–551 (1998).
Yoshioka, H. et al. Pitx2, a bicoid-type homeobox gene, is involved in a lefty-signaling pathway in determination of left–right asymmetry. Cell 94, 299–305 (1998).
Nakamura, T. et al. Fluid flow and interlinked feedback loops establish left-right asymmetric decay of Cerl2 mRNA. Nat. Commun. 3, 1322 (2012).
Schweickert, A. et al. The nodal inhibitor Coco is a critical target of leftward flow in Xenopus. Curr. Biol. 20, 738–743 (2010).
Hamada, H., Meno, C., Watanabe, D. & Saijoh, Y. Establishment of vertebrate left–right asymmetry. Nat. Rev. Genet. 3, 103–113 (2002).
Jacob, F. & Monod, J. Genetic regulatory mechanisms in the synthesis of proteins. J. Mol. Biol. 3, 318–356 (1961).
Niehrs, C. & Pollet, N. Synexpression groups in eukaryotes. Nature 402, 483–487 (1999).
Bolkier, Y. et al. Whole-exome sequencing reveals a monogenic cause in 56% of individuals with laterality disorders and associated congenital heart defects. J. Med. Genet. https://doi.org/10.1136/jmedgenet-2021-107775 (2021).
Almagro Armenteros, J. J. et al. SignalP 5.0 improves signal peptide predictions using deep neural networks. Nat. Biotechnol. 37, 420–423 (2019).
Sievers, F. et al. Fast, scalable generation of high-quality protein multiple sequence alignments using Clustal Omega. Mol. Syst. Biol. 7, 539 (2011).
Abdelkhalek, H. B. et al. The mouse homeobox gene Not is required for caudal notochord development and affected by the truncate mutation. Genes Dev. 18, 1725–1736 (2004).
Stauber, M. et al. Identification of FOXJ1 effectors during ciliogenesis in the foetal respiratory epithelium and embryonic left-right organiser of the mouse. Dev. Biol. 423, 170–188 (2017).
Westerfield, M. The Zebrafish Book: A Guide for the Laboratory Use of Zebrafish (Brachydanio Rerio). (Univ. of Oregon Press, 1989).
Yu, X., Ng, C. P., Habacher, H. & Roy, S. Foxj1 transcription factors are master regulators of the motile ciliogenic program. Nat. Genet. 40, 1445–1453 (2008).
Jao, L.-E., Wente, S. R. & Chen, W. Efficient multiplex biallelic zebrafish genome editing using a CRISPR nuclease system. Proc. Natl Acad. Sci. USA 110, 13904–13909 (2013).
Thisse, C. & Thisse, B. High-resolution in situ hybridization to whole-mount zebrafish embryos. Nat. Protoc. 3, 59–69 (2008).
Moreno-Mateos, M. A. et al. CRISPRscan: designing highly efficient sgRNAs for CRISPR-Cas9 targeting in vivo. Nat. Methods 12, 982–988 (2015).
Nakayama, T. et al. Cas9-based genome editing in Xenopus tropicalis. Methods Enzymol. 546, 355–375 (2014).
Hsiau, T. et al. Inference of CRISPR edits from Sanger trace data. BioRxiv https://doi.org/10.1101/251082 (2019).
Belo, J. A. et al. Cerberus-like is a secreted factor with neutralizing activity expressed in the anterior primitive endoderm of the mouse gastrula. Mech. Dev. 68, 45–57 (1997).
Schweickert, A. et al. Cilia-driven leftward flow determines laterality in Xenopus. Curr. Biol. 17, 60–66 (2007).
Rueden, C. T. et al. ImageJ2: ImageJ for the next generation of scientific image data. BMC Bioinf. 18, 529 (2017).
Sbalzarini, I. F. & Koumoutsakos, P. Feature point tracking and trajectory analysis for video imaging in cell biology. J. Struct. Biol. 151, 182–195 (2005).
R Core Team. R: A language and Environment for Statistical Computing. https://www.R-project.org/ (2017).
Bahlo, M. & Bromhead, C. J. Generating linkage mapping files from Affymetrix SNP chip data. Bioinformatics 25, 1961–1962 (2009).
Smith, K. R. et al. Reducing the exome search space for Mendelian diseases using genetic linkage analysis of exome genotypes. Genome Biol. 12, R85 (2011).
Abecasis, G. R., Cherny, S. S., Cookson, W. O. & Cardon, L. R. GRR: graphical representation of relationship errors. Bioinformatics 17, 742–743 (2001).
Abecasis, G. R., Cherny, S. S., Cookson, W. O. & Cardon, L. R. Merlin—rapid analysis of dense genetic maps using sparse gene flow trees. Nat. Genet. 30, 97–101 (2002).
Thiele, H. & Nürnberg, P. HaploPainter: a tool for drawing pedigrees with complex haplotypes. Bioinformatics 21, 1730–1732 (2005).
Lindner, T. H. & Hoffmann, K. easyLINKAGE: a PERL script for easy and automated two-/multi-point linkage analyses. Bioinformatics 21, 405–407 (2005).
Kumar, P., Henikoff, S. & Ng, P. C. Predicting the effects of coding non-synonymous variants on protein function using the SIFT algorithm. Nat. Protoc. 4, 1073–1081 (2009).
Adzhubei, I. A. et al. A method and server for predicting damaging missense mutations. Nat. Methods 7, 248–249 (2010).
Grantham, R. Amino acid difference formula to help explain protein evolution. Science 185, 862–864 (1974).
Pollard, K. S., Hubisz, M. J., Rosenbloom, K. R. & Siepel, A. Detection of nonneutral substitution rates on mammalian phylogenies. Genome Res. 20, 110–121 (2010).
Acknowledgements
We are grateful to all members of the Reversade laboratory for support. We thank Vijay Narasimhan for participating in the initial stages of the zebrafish study. E.S.-R. is supported by a NMRC Open Fund–Young Individual Research Grant (OF-YIRG, #OFYIRG18May-0053). M.B. was supported by DFG grants BL285/xx. T.O. was the recipient of a fellowship of the Landesgraduiertenförderung Baden-Württemberg. B.R. is a fellow of the National Research Foundation (NRF, Singapore) and Branco Weiss Foundation (Switzerland) and an EMBO Young Investigator. This work is supported by an inaugural Use-Inspired Basic Research (UIBR) central fund to B.R. from the Agency for Science & Technology and Research (A*STAR) in Singapore.
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Contributions
E.S.-R., T.O., M.B., P.B. and B.R. designed the study. G.L., V.R., C.T.L., B.E., F.E., B.V., J.A., H.R.C. and C.T.G. performed and supervised evolution genomics analyses. E.S.-R., M.K., W.X.G., Y.L.C., D.K., P.A., S.R. and B.R. performed and supervised the zebrafish functional experiments. T.O. and M.B. performed and supervised the Xenopus experiments. A. Beckers and A.G. performed and supervised the mouse experiments. S.M., P.S., M.F., E.C., A.M., L.D., G.C., S.D.F., C.R., J.-F.D., A. Boland, N.A., R.E., M.R., T.A.-B. and P.B. made clinical diagnoses and collected clinical data and samples. Family 8 from M.R. and T.A.-B. first allowed us to identify CIROP as the causative gene. E.S.-R., A.M.D.B., C.B., S.T., A.Y.-J.N., B.V. and P.B. performed and supervised whole exome sequencing, homozygosity mapping, high throughput cohort resequencing and sequencing analyses. E.S.-R. and B.R. wrote the manuscript with input from S.R., M.B. and P.B.
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Extended data
Extended Data Fig. 1 CIROP, MMP21, C1orf127, DAND5 and PKD1L1 are specifically lost in species with a LRO without motile cilia.
a, Table indicating the presence of the CIROP, MMP21, C1orf127, DAND5 and PKD1L1 genes in metazoan species in relation to the presence or absence of a LRO with motile cilia. It is believed that a LRO with motile cilia evolved in deuterostomia (green), but motile cilia in the LRO disappeared in cetartiodactyla (purple) and sauropsida (red). LRO, left–right organizer; n.d., not determined. b-d, Representation of the CIROP (b), MMP21 (c) and C1orf127 (d) loci in representative jawed-vertebrates whose genomes have been sequenced, showing the loss of the three genes in lizards and turtles (reptilia). Block arrows represent genes with the direction of arrows denoting transcriptional orientation. Orthologous genes are shown with the same color. Black circles represent ends of scaffolds.
Extended Data Fig. 2 Genomic structure of CIROP.
a, Genomic exon–intron structure of four distinct variants of human CIROP (previously known as LMLN2) from Ensembl, and the predicted full-length CIROP as compared to that of other primates and vertebrates as indicated. The alternative exons are color-coded as follows: pink and blue exons are only present in ENST00000637218.2, yellow exon is missing in ENST00000644147.1 and red exon is missing in ENST00000642668.1. Full-length human CIROP corresponds to ENST00000644000.1 with three extra amino acids in the beginning of exon 2 (purple) that are present in ENST00000644147.1 (see b). The UTRs are highlighted in dark gray. b, Genomic region on the UCSC browser containing CIROP with the conservation in vertebrates, showing that the pink and blue exons are not conserved while the yellow and red ones are highly conserved. Zooming in on the intron 1–exon 2 junction region confirms conservation of the three additional amino acids (purple box) present in ENST00000644147.1.
Extended Data Fig. 3 CRISPR–Cas9 knockout of Cirop in zebrafish and Xenopus laevis.
a, Real-time qPCR for cirop relative to actin in zebrafish embryos at indicated stages. cirop expression is only detectable using RNA extracted from tissue at the dorsal posterior side containing the DFCs. Data are mean ± s.e.m. ****P < 0.0001, Two-way ANOVA with Tukey test for multiple comparisons. 50%, 70% and 90% refer to epiboly stages. DFC, dorsal forerunner cells; hpf, hours postfertilization. n = 3 biological triplicates of 30-50 embryos. b, Depiction of genomic and protein structures of zebrafish Cirop. The sequence used is XM_002662823 using a START codon located 168 nucleotides upstream of the original one (light gray). Cirop protein domains are highlighted: the signal peptide (gray), the Zn+ catalytic domain (yellow), the cysteine-rich domain (red), and the transmembrane domain (blue). Yellow line, catalytic domain; green line, met-turn. The site and targeted sequence by the CRISPR gRNA is indicated in purple. Five different alleles were obtained with indicated mutations (green, insertion; red star, deletion), all leading to a frameshift (orange) with an early stop codon (underlined for mutation 1). While all mutations lead to the same LR phenotype in the homozygous state, line 1 was used for further investigation. c, Whole mount in situ hybridization for Cirop in 90% epiboly control and Cirop−/− zebrafish embryos. Scale bar, 0.1 mm. d, Depiction of genomic and protein structures of Xenopus laevis Cirop. A CRISPR gRNA was designed in the beginning of exon 4 (purple), leading to mutations with 92% efficiency. Chr., chromosome; SP, signal peptide; TM, transmembrane domain; wt, wild-type; Xl, Xenopus laevis.
Extended Data Fig. 4 The absence of Cirop does not affect DFCs or midline patterning.
a-c, Analysis of zebrafish embryos injected or not with cirop mRNA by whole mount in situ hybridization (a), real-time qPCR (b) and western blotting (c). Scale bar in a, 0.1 mm. Similar results were obtained in at least 3 independent experiments with 50 embryos each. b, Data are mean ± s.e.m, ****P < 0.0001, two-tailed unpaired t-test. n = 3 biological triplicates of 30-50 embryos. c, Protein extracts from 2 independent experiments are loaded and membrane probed with an anti-cirop antibody. Similar results were obtained with a third independent experiment. d-f, Whole mount in situ hybridization in control and cirop−/− zebrafish embryos for tbxta (d), foxj1a (e), and sox17 (f) at indicated stages. Scale bars, 0.1 mm. Similar results were obtained in at least 3 independent experiments with 50 embryos each.
Extended Data Fig. 5 Mapping analysis of six families with patients presenting with heterotaxy revealed a region of homozygosity on chromosome 14.
a, Individuals of Families 1, 2, 3, 6 and 7 with red symbols in pedigrees were SNP genotyped. Genotypes were used for linkage analysis, which revealed a common region of homozygosity for affected individuals on chromosome 14, including the genomic region of CIROP, with a total Lod_Score of 8.803. Graphical representation obtained using Merlin Autosome. b, Haplotypes obtained with Merlin and presented with Haplopainter revealed a common region of homozygosity on chromosome 14q11.2 delimited by rs17211943 and rs223116 that is 0.72 Mb long. c, Graphical representation of relationships (GRR) showing that parents of Family 3 are actually related. d, Family 8 SNP genotyping of individuals V:2, V:3, V:4 (red in pedigree) revealed a Lod_Score max on chromosome 14q11.2 with SNP rs956163 (pink) for affected individuals. The region of homozygosity shared by V:2 and V:4 is 3.6 Mb long, delimited by rs1114967 and rs724165.
Extended Data Fig. 6 CIROP is conserved among vertebrates.
CIROP proteins alignment highlighting the conservation of the affected amino acids in patients with heterotaxy (purple). CIROP protein domains are highlighted: signal peptide (gray), Zn+ catalytic domain (yellow), cysteine-rich domain (red), and transmembrane domain (blue). Brown C, cysteine switch; dark yellow amino acids, catalytic site (HExxH…H); green M, met-turn; purple, mutated amino acids; Conserv., conservation. Software used, Clustal Omega.
Supplementary information
41588_2021_970_MOESM3_ESM.avi
Supplementary Video 1 Cilia movement in KV of a wt zebrafish embryo. Eight to ten-somite stage zebrafish embryos were dechorionated and mounted in 2% low melting agarose, and KV were imaged with a ×60 water lens from the dorsal posterior end.
41588_2021_970_MOESM4_ESM.avi
Supplementary Video 2 Cilia movement in KV of a cirop−/− zebrafish embryo. Eight to ten somite stage zebrafish embryos were dechorionated and mounted in 2% low melting agarose, and KV were imaged with a ×60 water lens from the dorsal posterior end.
41588_2021_970_MOESM5_ESM.avi
Supplementary Video 3 Movement of endogenous particles in KV of a wt zebrafish embryo. Eight to ten somite stage zebrafish embryos were dechorionated and mounted in 2% low melting agarose, and KV were imaged with a ×60 water lens from the dorsal posterior end.
41588_2021_970_MOESM6_ESM.avi
Supplementary Video 4 Movement of endogenous particles in KV of a cirop−/− zebrafish embryo. Eight to ten somite stage zebrafish embryos were dechorionated and mounted in 2% low melting agarose, and KV were imaged with a ×60 water lens from the dorsal posterior end.
41588_2021_970_MOESM7_ESM.avi
Supplementary Video 5 Analysis of leftward flow in the LRO of Xenopus wild-type and Cirop crispants. Dorsal explants were prepared and fluorescent micro beads added. Time-lapse videos were captured at 2 fps and transformed to gradient-time-trails (GTTs). GTT videos play at about 10× real time and display bead transport from the right to the left side of the LRO. Note that velocity and directionality of bead transport were identical in wt (left) and Cirop crispant (right) specimens.
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Szenker-Ravi, E., Ott, T., Khatoo, M. et al. Discovery of a genetic module essential for assigning left–right asymmetry in humans and ancestral vertebrates. Nat Genet 54, 62–72 (2022). https://doi.org/10.1038/s41588-021-00970-4
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DOI: https://doi.org/10.1038/s41588-021-00970-4
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