Abstract
Metazoan internal organs are assembled from polarized tubular epithelia that must set aside an apical membrane domain as a lumenal surface. In a global Caenorhabditis elegans tubulogenesis screen, interference with several distinct fatty-acid-biosynthetic enzymes transformed a contiguous central intestinal lumen into multiple ectopic lumens. We show that multiple-lumen formation is caused by apicobasal polarity conversion, and demonstrate that in situ modulation of lipid biosynthesis is sufficient to reversibly switch apical domain identities on growing membranes of single post-mitotic cells, shifting lumen positions. Follow-on targeted lipid-biosynthesis pathway screens and functional genetic assays were designed to identify a putative single causative lipid species. They demonstrate that fatty-acid biosynthesis affects polarity through sphingolipid synthesis, and reveal ceramide glucosyltransferases (CGTs) as end-point biosynthetic enzymes in this pathway. Our findings identify glycosphingolipids, CGT products and obligate membrane lipids, as critical determinants of in vivo polarity and indicate that they sort new components to the expanding apical membrane.
This is a preview of subscription content, access via your institution
Access options
Subscribe to this journal
Receive 12 print issues and online access
$209.00 per year
only $17.42 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
References
Pfeffer, S. R. & Rothman, J. E. Biosynthetic protein transport and sorting by the endoplasmic reticulum and Golgi. Annu. Rev. Biochem. 56, 829–852 (1987).
Mellman, I. & Nelson, W. J. Coordinated protein sorting, targeting and distribution in polarized cells. Nat. Rev. Mol. Cell Biol. 9, 833–845 (2008).
St Johnston, D. & Ahringer, J. Cell polarity in eggs and epithelia: parallels and diversity. Cell 141, 757–774 (2010).
Bryant, D. M. et al. A molecular network for de novo generation of the apical surface and lumen. Nat. Cell Biol. 12, 1035–1045 (2010).
Martin-Belmont, F. & Rodriguez-Foretell, A. E. Acquisition of membrane polarity in epithelial tube formation patterns, signalling pathways, molecular mechanisms, and disease. Int. Rev. Cell Mol. Biol. 274, 129–182 (2009).
Kemphues, K. J., Press, J. R., Morton, D. G. & Chang, N. S. Identification of genes required for cytoplasmic localization in early C. elegans embryos. Cell 52, 311–320 (1988).
Nelson, W. J. Adaptation of core mechanisms to generate cell polarity. Nature 422, 766–774 (2003).
Rodriguez-Boolean, E., Kibitzer, G. & Müsch, A. Organisation of vesicular trafficking in epithelia. Nat. Rev. Mol. Cell Biol. 6, 233–247 (2005).
Di Paolo, G. & De Camilla, P. Phosphoinositides in cell regulation and membrane dynamics. Nature 443, 651–657 (2006).
Lippincott-Schwartz, J. & Phair, R. D. Lipids and cholesterol as regulators of traffic in the endomembrane system. Annu. Rev. Biophys. 39, 559–578 (2010).
Mays, R. W. et al. Hierarchy of mechanisms involved in generating Na/K-ATPase polarity in MDCK epithelial cells. J. Cell Biol. 130, 1105–1115 (1995).
Sprong, H. et al. Glycosphingolipids are required for sorting melanosomal proteins in the Golgi complex. J. Cell Biol. 155, 369–380 (2001).
Hoekstra, D., Maier, O., van der Wouden, J. M., Slimane, T. A. & van Ijzendoorn, S. C. D. Membrane dynamics and cell polarity: the role of sphingolipids. J. Lipid Res. 44, 869–877 (2003).
Degroote, S., Wolthoorn, J. & van Meer, G. The cell biology of glycosphingolipids. Semin. Cell Dev. Biol. 15, 375–387 (2004).
Mayor, S. & Riezman, H. Sorting GPI-anchored proteins. Nat. Rev. Mol. Cell Biol. 5, 110–120 (2004).
Simons, K. & van Meer, G. Lipid sorting in epithelial cells. Biochemistry 27, 6197–6202 (1988).
van Meer, G., Voelker, D. R. & Feigenson, G. W. Membrane lipids: where they are and how they behave. Nat. Rev. Mol. Cell Biol. 9, 112–124 (2008).
Lingwood, D. & Simons, K. Lipid rafts as a membrane-organizing principle. Science 327, 46–50 (2010).
Furukawa, K., Tokuda, N., Okuda, T. & Tajima, O. Glycosphingolipids in engineered mice: insights into function. Semin. Cell Dev. Biol. 15, 389–396 (2004).
Lynch, A. M. & Hardin, J. The assembly and maintenance of epithelial junctions in C. elegans. Front. Biosci. 14, 1414–1432 (2009).
Knight, C. G., Patel, M. N., Azevedo, R. B. & Leroi, A. M. A novel mode of ecdysozoan growth in Caenorhabditis elegans. Evol. Dev. 4, 16–27 (2002).
Baugh, L. R. & Sternberg, P. W. DAF-16/FOXO regulates transcription of cki-1/Cip/Kip and repression of lin-4 during C. elegans L1 arrest. Curr. Biol. 16, 780–785 (2006).
Entchev, E. V. et al. LET-767 is required for the production of branched chain and long chain fatty acids in Caenorhabditis elegans. J. Biol. Chem. 283, 17550–17560 (2008).
Kniazeva, M., Euler, T. & Han, M. A branched-chain fatty acid is involved in post-embryonic growth control in parallel to the insulin receptor pathway and its biosynthesis is feedback-regulated in C. elegans. Genes Dev. 22, 2102–2110 (2008).
Rappleye, C. A., Tagawa, A., Le Bot, N., Ahringer, J. & Aroian, R. V. Involvement of fatty acid pathways and cortical interaction of the pronuclear complex in Caenorhabditis elegans embryonic polarity. BMC Dev. Biol. 3, 8 (2003).
Kuervers, L. M., Jones, C. L., O’Neil, N. J. & Baillie, D. L. The sterol modifying enzyme LET-767 is essential for growth, reproduction and development in Caenorhabditis elegans. Mil. Genet. Genomics 270, 121–131 (2003).
Kniazeva, M., Crawford, Q. T., Seiber, M., Wang, C. Y. & Han, M. Monomethyl branched-chain fatty acids play an essential role in Caenorhabditis elegans development. PLoS Biol. 2, E257 (2004).
Brock, T. J., Browse, J. & Watts, J. L. Fatty acid desaturation and the regulation of adiposity in Caenorhabditis elegans. Genetics 176, 865–875 (2007).
Watts, J. L. & Browse, J. Genetic dissection of polyunsaturated fatty acid synthesis in Caenorhabditis elegans. Proc. Natl Acad. Sci. USA 99, 5854–5859 (2002).
Van Ijzendoorn, S. C. D., Van Der Wouden, J. M., Liebisch, G., Schmitz, G. & Hoekstra, D. Polarized membrane traffic and cell polarity development is dependent on dihydroceramide synthase-regulated sphinganine turnover. Mol. Biol. Cell 15, 4115–4124 (2004).
Chitwood, D. J., Lusby, W. R., Thompson, M. J., Kochansky, J. P. & Howarth, O. W. The glycosylceramides of the nematode Caenorhabditis elegans contain an unusual, branched-chain sphingoid base. Lipids 30, 567–573 (1995).
Ichikawa, S. & Hirabayashi, Y. Glucosylceramide synthase and glycosphingolipid synthesis. Trends Cell Biol. 8, 198–202 (1998).
Leipelt, M. et al. Glucosylceramide synthases, a gene family responsible for the biosynthesis of glucosphingolipids in animals, plants, and fungi. J. Biol. Chem. 276, 33621–33629 (2001).
Marza, E., Simonsen, K. T., Faergeman, N. J. & Lesa, G. M. Expression of ceramide glucosyltransferases, which are essential for glycosphingolipid synthesis, is only required in a small subset of C. elegans cells. J. Cell Sci. 122, 822–833 (2009).
Nomura, K. H. et al. Ceramide glucosyltransferase of the nematode Caenorhabditis elegans is involved in oocyte formation and in early embryonic cell division. Glycobiology 21, 834–848 (2011).
Maier, O., Oberle, V. & Hoekstra, D. Fluorescent lipid probes: some properties and applications (a review). Chem. Phys. Lipids 116, 3–18 (2002).
Hannun, Y. A. & Obeid, L. M. Principles of bioactive lipid signalling: lessons from sphingolipids. Nat. Rev. Mol. Cell Biol. 9, 139–150 (2008).
Varki, A. Essentials of Glycobiology 2nd edn (Cold Spring Harbor Laboratory Press, 2009).
Griffitts, J. S. et al. Glycolipids as receptors for Bacillus thuringiensis crystal toxin. Science 307, 922–925 (2005).
Gerdt, S., Lochnit, G., Dennis, R. D. & Geyer, R. Isolation and structural analysis of three neutral glycosphingolipids from a mixed population of Caenorhabditis elegans (Nematoda:Rhabditida). Glycobiology 7, 265–275 (1997).
Futerman, A. H. & Riezman, H. The ins and outs of sphingolipid synthesis. Trends Cell Biol. 15, 312–318 (2005).
Weisz, O. A. & Rodriguez-Boolean, E. Apical trafficking in epithelial cells: signals, clusters and motors. J. Cell Sci. 122, 4253–4266 (2009).
Chen, C. C. et al. RAB-10 is required for endocytic recycling in the Caenorhabditis elegans intestine. Mol. Biol. Cell 17, 1286–1297 (2006).
Rolls, M. M., Hall, D. H., Victor, M., Stelzer, E. H. & Rapoport, T. A. Targeting of rough endoplasmic reticulum membrane proteins and ribosomes in invertebrate neurons. Mol. Biol. Cell 13, 1778–1791 (2002).
Yamashita, T. et al. A vital role for glycosphingolipid synthesis during development and differentiation. Proc. Natl Acad. Sci. USA 96, 9142–9147 (1999).
Rao, R. P. & Acharya, J. K. Sphingolipids and membrane biology as determined from genetic models. Prostaglandins Other Lipid Mediat. 85, 1–16 (2008).
Schuck, S. & Simons, K. Polarized sorting in epithelial cells: raft clustering and the biogenesis of the apical membrane. J. Cell Sci. 117, 5955–5964 (2004).
Gassama-Diagne, A. et al. Phosphatidylinositol-3,4,5-trisphosphate regulates the formation of the basolateral plasma membrane in epithelial cells. Nat. Cell Biol. 8, 963–970 (2006).
Martin-Belmont, F. et al. PTEN-mediated apical segregation of phosphoinositides controls epithelial morphogenesis through Cdc42. Cell 128, 383–397 (2007).
Haucke, V. & Di Paolo, G. Lipids and lipid modifications in the regulation of membrane traffic. Curr. Opin. Cell Biol. 19, 426–435 (2007).
Simons, K. & Gerl, M. J. Revitalizing membrane rafts: new tools and insights. Nat. Rev. Mol. Cell Biol. 11, 688–699 (2010).
Seamen, E., Blanchette, J. M. & Han, M. P-type ATPase TAT-2 negatively regulates monomethyl branched-chain fatty acid mediated function in post-embryonic growth and development in C. elegans. PLoS Genet. 5, e1000589 (2009).
Wandall, H. H. et al. Egghead and brainiac are essential for glycosphingolipid biosynthesis in vivo. J. Biol. Chem. 280, 4858–4863 (2005).
Mishra, R., Grzybek, M., Niki, T., Hirashima, M. & Simons, K. Galectin-9 trafficking regulates apical-basal polarity in Madin–Darby canine kidney epithelial cells. Proc. Natl Acad. Sci. USA 107, 17633–17638 (2010).
Sampaio, J. L. et al. Membrane lipidome of an epithelial cell line. Proc. Natl Acad. Sci. USA 108, 1903–1907 (2011).
Vance, D. E. & Vance, J. E. Biochemistry of lipids, lipoproteins and membranes 5th edn (Elsevier, 2008).
Gobel, V., Barrett, P. L., Hall, D. H. & Fleming, J. T. Lumen morphogenesis in C. elegans requires the membrane-cytoskeleton linker erm-1. Dev. Cell 6, 865–873 (2004).
Sulston, J. E. & Horvitz, H. R. Post-embryonic cell lineages of the nematode, Caenorhabditis elegans. Dev Biol. 56, 110–156 (1977).
Legouis, R. et al. LET-413 is a basolateral protein required for the assembly of adherens junctions in Caenorhabditis elegans. Nat. Cell Biol. 2, 415–422 (2000).
Kurzchalia, T. V. & Ward, S. Why do worms need cholesterol? Nat. Cell Biol. 5, 684–688 (2003).
Martinez-Alonso, E., Egea, G., Ballesta, J. & Martinez-Menarguez, J. A. Structure and dynamics of the Golgi complex at 15 °C: low temperature induces the formation of Golgi-derived tubules. Traffic 6, 32–44 (2005).
Onelli, E., Prescianotto-Baschong, C., Caccianiga, M. & Moscatelli, A. Clathrin-dependent and independent endocytic pathways in tobacco protoplasts revealed by labelling with charged nanogold. J. Exp. Bot. 59, 3051–3068 (2008).
Brenner, S. The genetics of Caenorhabditis elegans. Genetics 77, 71–94 (1974).
Timmons, L., Court, D. L. & Fire, A. Ingestion of bacterially expressed dsRNAs can produce specific and potent genetic interference in Caenorhabditis elegans. Gene 263, 103–112 (2001).
Curran, S. P. & Ruvkun, G. Lifespan regulation by evolutionarily conserved genes essential for viability. PLoS Genet. 3, e56 (2007).
Hobert, O. PCR fusion-based approach to create reporter gene constructsfor expression analysis in transgenic C. elegans. Biotechniques 32, 728–730 (2002).
Mello, C. C., Kramer, J. M., Stinchcomb, D. & Ambros, V. Efficient gene transfer in C. elegans: extrachromosomal maintenance and integration of transforming sequences. EMBO J. 10, 3959–3970 (1991).
Miller, D. M. & Shakes, D. C. Immunofluorescence microscopy. Methods Cell Biol. 48, 365–394 (1995).
Hall, D. H. Electron microscopy and three-dimensional image reconstruction. Methods Cell Biol. 48, 395–436 (1995).
Sullards, M. C., Wang, E., Peng, Q. & Merrill, A. H. Jr Metabolomic profiling of sphingolipids in human glioma cell lines by liquid chromatography tandem mass spectrometry. Cell Mil. Biol. (Noisy-le-grand) 49, 789–797 (2003).
Acknowledgements
Strains and plasmids were provided by D. Baillie (Fraser University, Burnaby, British Columbia, Canada), A. Croce (IFOM Istituto FIRC di Oncologia Molecolare, Milan, Italy), B. Grant (Rutgers University, Piscataway, New Jersey, USA), K. Kemphues (Cornell University, Ithaca, New York, USA), K. Nehrke (University of Rochester Medical Center, Rochester New York, USA), G. Ruvkun (Massachusetts General Hospital, Harvard Medical School, Boston, Massachusetts, USA), K. Strange (Vanderbilt University Medical Center, Salisbury Cove Maine, USA), J. Simske (Case Western Reserve University School of Medicine, Cleveland, Ohio, USA), S. Mitani (National Bioresource Project Japan) and the Caenorhabditis Genetics Center (NIH Center for Research Resources). We thank G. Ruvkun for the lethal RNAi library, and J. Moore (Avanti Polar Lipids); Mary McKee (MGH Microscopy Core/partially funded by the IBD grant DK43351 and BA DE award DK57521) and K. Nygen; Christopher Crocker; and Edward Membreno for contributions to LC/MS; TEM; illustrations and C. elegans maintenance, respectively. We thank F. Solomon and B. Winckler for critical reading of the manuscript and H. Weinstein and A. Walker for ongoing support. This work was supported by NIH grants HD044589 and GM078653 and a Mattina R. Proctor Award to V.G.
Author information
Authors and Affiliations
Contributions
H.Z. generated and assembled most of the data and contributed to project design, data analysis and writing of the manuscript. N.A. participated in most experiments, carried out the glycosylation screen and contributed to experimental design and data analysis. L.A.K. contributed to the genetic interaction experiments. D.H.H. and J.T.F. contributed to electron microscopy experiments and J.T.F. to writing of the manuscript. V.G. conceived and directed the project, participated in experiments and wrote the manuscript.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing financial interests.
Supplementary information
Supplementary Information
Supplementary Information (PDF 1085 kb)
Supplementary Table 1
Supplementary Information (XLS 113 kb)
Supplementary Table 2
Supplementary Information (XLS 47 kb)
Supplementary Table 3
Supplementary Information (XLS 44 kb)
Supplementary Table 4
Supplementary Information (XLS 39 kb)
Supplementary Table 5
Supplementary Information (XLS 22 kb)
Rights and permissions
About this article
Cite this article
Zhang, H., Abraham, N., Khan, L. et al. Apicobasal domain identities of expanding tubular membranes depend on glycosphingolipid biosynthesis. Nat Cell Biol 13, 1189–1201 (2011). https://doi.org/10.1038/ncb2328
Received:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/ncb2328
This article is cited by
-
Apical–basal polarity and the control of epithelial form and function
Nature Reviews Molecular Cell Biology (2022)
-
Glycolipid-dependent and lectin-driven transcytosis in mouse enterocytes
Communications Biology (2021)
-
Saturated very long chain fatty acid configures glycosphingolipid for lysosome homeostasis in long-lived C. elegans
Nature Communications (2021)
-
Glycosphingolipids and neuroinflammation in Parkinson’s disease
Molecular Neurodegeneration (2020)
-
Wnt-controlled sphingolipids modulate Anthrax Toxin Receptor palmitoylation to regulate oriented mitosis in zebrafish
Nature Communications (2020)