Key Points
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A lumen arising de novo creates a space between or within cells where no space existed before.
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De novo lumen formation has been studied in several model systems. Despite apparent differences, there are many cell biological commonalities that now justify a unifying overview.
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A major mechanism of de novo lumen formation is the delivery of membrane material to the apical plasma membrane of the lumen-forming cell, which must be incorporated in a spatially structured manner.
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Lumen formation can be subdivided into three steps: determining the site of lumen initiation, enlarging the apical domain, and the maturation and stabilization of the lumen.
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The major cellular machineries needed for tube construction are cell polarity determinants, vesicle-trafficking systems and the cytoskeleton.
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Although there is currently a good understanding of how material is delivered to the apical membrane during the creation of tubes, an important open question is how the membrane is given its spatial structure.
Abstract
Many organs contain networks of epithelial tubes that transport gases or fluids. A lumen can be generated by tissue that enwraps a pre-existing extracellular space or it can arise de novo either between cells or within a single cell in a position where there was no space previously. Apparently distinct mechanisms of de novo lumen formation observed in vitro — in three-dimensional cultures of endothelial and Madin–Darby canine kidney (MDCK) cells — and in vivo — in zebrafish vasculature, Caenorhabditis elegans excretory cells and the Drosophila melanogaster trachea — in fact share many common features. In all systems, lumen formation involves the structured expansion of the apical plasma membrane through general mechanisms of vesicle transport and of microtubule and actin cytoskeleton regulation.
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References
Jaz´win´ska, A., Ribeiro, C. & Affolter, M. Epithelial tube morphogenesis during Drosophila tracheal development requires Piopio, a luminal ZP protein. Nature Cell Biol. 5, 895–901 (2003).
Keister, M. L. The morphogenesis of the tracheal system of Sciara. J. Morphol. 83, 373–423 (1948).
Shafiq, S. A. Electron microscopy of the development of tracheoles in Drosophila melanogaster. Q. J. Microsc. Sci. 104, 135–140 (1963).
Wolff, J. R. & Bär, T. 'Seamless' endothelia in brain capillaries during development of the rat's cerebral cortex. Brain Res. 41, 17–24 (1972).
Folkman, J. & Haudenschild, C. Angiogenesis in vitro. Nature 288, 551–556 (1980). The first demonstration of angiogenesis in vitro shows that the lumen can develop from isolated capillary endothelial cells.
Bryant, D. M. & Mostov, K. E. From cells to organs: building polarized tissue. Nature Rev. Mol. Cell Biol. 9, 887–901 (2008).
Dong, B. et al. Tube formation by complex cellular processes in Ciona intestinalis notochord. Dev. Biol. 330, 237–249 (2009).
Bayless, K. J., Salazar, R. & Davis, G. RGD-dependent vacuolation and lumen formation observed during endothelial cell morphogenesis in three-dimensional fibrin matrices involves the αvβ3 and α5β1 integrins. Am. J. Pathol. 156, 1673–1683 (2000).
Sacharidou, A., Stratman, A. N. & Davis, G. E. Molecular mechanisms controlling vascular lumen formation in three-dimensional extracellular matrices. Cells Tissues Organs 195, 122–143 (2012).
Connolly, J. O., Simpson, N., Hewlett, L. & Hall, A. Rac regulates endothelial morphogenesis and capillary assembly. Mol. Biol. Cell 13, 2474–2485 (2002).
Tung, J. J., Tattersall, I. W. & Kitajewski, J. Tips, stalks, tubes: notch-mediated cell fate determination and mechanisms of tubulogenesis during angiogenesis. Cold Spring Harb. Perspect. Med. 2, a006601 (2012).
Montesano, R., Matsumoto, K., Nakamura, T. & Orci, L. Identification of a fibroblast-derived epithelial morphogen as hepatocyte growth factor. Cell 67, 901–908 (1991).
Montesano, R., Schaller, G. & Orci, L. Induction of epithelial tubular morphogenesis in vitro by fibroblast-derived soluble factors. Cell 66, 697–711 (1991).
Pollack, A. L., Runyan, R. B. & Mostov, K. E. Morphogenetic mechanisms of epithelial tubulogenesis: MDCK cell polarity is transiently rearranged without loss of cell-cell contact during scatter factor/hepatocyte growth factor-induced tubulogenesis. Dev. Biol. 204, 64–79 (1998).
Blum, Y. et al. Complex cell rearrangements during intersegmental vessel sprouting and vessel fusion in the zebrafish embryo. Dev. Biol. 316, 312–322 (2008).
Lenard, A. et al. In vivo analysis reveals a highly stereotypic morphogenetic pathway of vascular anastomosis. Dev. Cell. 25, 492–506 (2013).
Herwig, L. et al. Distinct cellular mechanisms of blood vessel fusion in the zebrafish embryo. Curr. Biol. 21, 1942–1948 (2011). Demonstrates that lumen formation in the zebrafish vasculature can occur by two distinct mechanisms: membrane invagination and cord hollowing.
Kamei, M. et al. Endothelial tubes assemble from intracellular vacuoles in vivo. Nature 442, 453–456 (2006). Suggests that vascular lumen formation in zebrafish may take place by cell hollowing.
Tanaka-Matakatsu, M., Uemura, T., Oda, H., Takeichi, M. & Hayashi, S. Cadherin-mediated cell adhesion and cell motility in Drosophila trachea regulated by the transcription factor Escargot. Development 122, 3697–3705 (1996).
Uemura, T. et al. Zygotic Drosophila E-cadherin expression is required for processes of dynamic epithelial cell rearrangement in the Drosophila embryo. Genes Dev. 10, 659–671 (1996).
Gervais, L., Lebreton, G. & Casanova, J. The making of a fusion branch in the Drosophila trachea. Dev. Biol. 362, 187–193 (2012).
Kakihara, K., Shinmyozu, K., Kato, K., Wada, H. & Hayashi, S. Conversion of plasma membrane topology during epithelial tube connection requires Arf-like 3 small GTPase in Drosophila. Mech. Dev. 125, 325–336 (2008).
Gervais, L. & Casanova, J. In vivo coupling of cell elongation and lumen formation in a single cell. Curr. Biol. 20, 359–366 (2010). Identifies membrane invagination as the mechanism of subcellular lumen formation in tracheal terminal cells. Shows the role of actin and microtubule cytoskeletons in directing this process.
Lubarsky, B. & Krasnow, M. A. Tube morphogenesis: making and shaping biological tubes. Cell 112, 19–28 (2003).
Buechner, M. Tubes and the single C. elegans excretory cell. Trends Cell Biol. 12, 479–484 (2002).
Khan, L. A. et al. Intracellular lumen extension requires ERM-1-dependent apical membrane expansion and AQP-8-mediated flux. Nature Cell Biol. 15, 143–156 (2013).
Kolotuev, I., Hyenne, V., Schwab, Y., Rodriguez, D. & Labouesse, M. A pathway for unicellular tube extension depending on the lymphatic vessel determinant Prox1 and on osmoregulation. Nature Cell Biol. 15, 157–168 (2013).
Berry, K. L., Bülow, H. E., Hall, D. H. & Hobert, O. A. C. elegans CLIC-like protein required for intracellular tube formation and maintenance. Science 302, 2134–2137 (2003).
JayaNandanan, N., Mathew, R. & Leptin, M. Guidance of subcellular tubulogenesis by actin under the control of a synaptotagmin-like protein and Moesin. Nature Commun. 5, 3036 (2014).
O'Brien, L. E. et al. Rac1 orientates epithelial apical polarity through effects on basolateral laminin assembly. Nature Cell Biol. 3, 831–838 (2001).
Yu, W. et al. Involvement of RhoA, ROCK I and myosin II in inverted orientation of epithelial polarity. EMBO Rep. 9, 923–929 (2008).
Davis, G. & Camarillo, C. W. An α2β1 integrin-dependent pinocytic mechanism involving intracellular vacuole formation and coalescence regulates capillary lumen and tube formation in three-dimensional collagen matrix. Exp. Cell Res. 224, 39–51 (1996).
Davis, G. E. & Bayless, K. J. An integrin and Rho GTPase-dependent pinocytic vacuole mechanism controls capillary lumen formation in collagen and fibrin matrices. Microcirculation 10, 27–44 (2003).
Bayless, K. J. & Davis, G. E. The Cdc42 and Rac1 GTPases are required for capillary lumen formation in three-dimensional extracellular matrices. J. Cell Sci. 115, 1123–1136 (2002). References 32–34 use 3D endothelial cell cultures to show the role of ECM–integrin interactions in lumen formation, identify the roles of RHO GTPases in directing lumen formation and show that coalescence of pinocytic intracellular vacuoles contribute to the lumen.
Koh, W., Mahan, R. D. & Davis, G. E. Cdc42- and Rac1-mediated endothelial lumen formation requires Pak2, Pak4 and Par3, and PKC-dependent signaling. J. Cell Sci. 121, 989–1001 (2008).
Bryant, D. M. et al. A molecular network for de novo generation of the apical surface and lumen. Nature Cell Biol. 12, 1035–1045 (2010). Identifies molecular machineries of vesicular transport required for polarization and delivery of material during lumen formation in MDCK cultures. Together with reference 69, defines the steps of lumen initiation.
Apodaca, G., Gallo, L. I. & Bryant, D. M. Role of membrane traffic in the generation of epithelial cell asymmetry. Nature Cell Biol. 14, 1235–1243 (2012).
Datta, A., Bryant, D. M. & Mostov, K. E. Molecular regulation of lumen morphogenesis. Curr. Biol. 21, R126–R136 (2011).
Martín-Belmonte, F. & Mostov, K. Regulation of cell polarity during epithelial morphogenesis. Curr. Opin. Cell Biol. 20, 227–234 (2008).
Gassama-Diagne, A. et al. Phosphatidylinositol-3,4,5-trisphosphate regulates the formation of the basolateral plasma membrane in epithelial cells. Nature Cell Biol. 8, 963–970 (2006).
Lee, S. & Kolodziej, P. A. The plakin Short Stop and the RhoA GTPase are required for E-cadherin-dependent apical surface remodeling during tracheal tube fusion. Development 129, 1509–1520 (2002).
Lee, M., Lee, S., Zadeh, A. D. & Kolodziej, P. A. Distinct sites in E-cadherin regulate different steps in Drosophila tracheal tube fusion. Development 130, 5989–5999 (2003).
Pollard, T. D., Earnshaw, W. C. & Lippincott-Schwartz, J. Cell Biology (W. B. Saunders Co., 2008).
Zhang, M. & Schekman, R. Unconventional secretion, unconventional solutions. Science 340, 559–561 (2013).
Gálvez-Santisteban, M. et al. Synaptotagmin-like proteins control the formation of a single apical membrane domain in epithelial cells. Nature Cell Biol. 14, 838–849 (2012).
Jung, J.-J. et al. Syntaxin 16 regulates lumen formation during epithelial morphogenesis. PLoS ONE 8, e61857 (2013).
Song, Y., Eng, M. & Ghabrial, A. S. Focal defects in single-celled tubes mutant for Cerebral cavernous malformation 3, GCKIII, or NSF2. Dev. Cell 25, 507–519 (2013).
Wang, Y. et al. Moesin1 and Ve-cadherin are required in endothelial cells during in vivo tubulogenesis. Development 137, 3119–3128 (2010).
Jones, T. A. & Metzstein, M. M. A novel function for the PAR complex in subcellular morphogenesis of tracheal terminal cells in Drosophila melanogaster. Genetics 189, 153–164 (2011).
Schottenfeld-Roames, J. & Ghabrial, A. S. Whacked and Rab35 polarize dynein-motor-complex-dependent seamless tube growth. Nature Cell Biol. 14, 386–393 (2012).
Meder, D., Shevchenko, A., Simons, K. & Füllekrug, J. Gp135/podocalyxin and NHERF-2 participate in the formation of a preapical domain during polarization of MDCK cells. J. Cell Biol. 168, 303–313 (2005).
Martín-Belmonte, F. et al. PTEN-mediated apical segregation of phosphoinositides controls epithelial morphogenesis through Cdc42. Cell 128, 383–397 (2007).
Martín-Belmonte, F. et al. Cell-polarity dynamics controls the mechanism of lumen formation in epithelial morphogenesis. Curr. Biol. 18, 507–513 (2008).
Roland, J. T. et al. Rab GTPase-Myo5B complexes control membrane recycling and epithelial polarization. Proc. Natl Acad. Sci. USA 108, 2789–2794 (2011).
Oshima, K. et al. IKKɛ regulates F actin assembly and interacts with Drosophila IAP1 in cellular morphogenesis. Curr. Biol. 16, 1531–1537 (2006).
Okenve-Ramos, P. & Llimargas, M. Fascin links Btl/FGFR signalling to the actin cytoskeleton during Drosophila tracheal morphogenesis. Development 141, 929–939 (2014).
Boehlke, C. et al. Kif3a guides microtubular dynamics, migration and lumen formation of MDCK cells. PLoS ONE 8, e62165 (2013).
Gloerich, M. et al. Rap2A links intestinal cell polarity to brush border formation. Nature Cell Biol. 14, 793–801 (2012).
Legate, K. R. & Fässler, R. Mechanisms that regulate adaptor binding to β-integrin cytoplasmic tails. J. Cell Sci. 122, 187–198 (2009).
Levi, B. P., Ghabrial, A. S. & Krasnow, M. A. Drosophila talin and integrin genes are required for maintenance of tracheal terminal branches and luminal organization. Development 133, 2383–2393 (2006). Describes the role of ECM–Integrin interaction in stabilization and maturation of the subcellular lumen of tracheal cells.
Buechner, M., Hall, D. H., Bhatt, H. & Hedgecock, E. M. Cystic canal mutants in Caenorhabditis elegans are defective in the apical membrane domain of the renal (excretory) cell. Dev. Biol. 214, 227–241 (1999). Identifies mutations in 12 genes that result in lumen defects in the C. elegans excretory cell. Suggests that the defects result from defective apical membrane morphogenesis.
McKeown, C., Praitis, V. & Austin, J. sma-1 encodes a βH-spectrin homolog required for Caenorhabditis elegans morphogenesis. Development 125, 2087–2098 (1998).
Tong, X. & Buechner, M. CRIP homologues maintain apical cytoskeleton to regulate tubule size in C. elegans. Dev. Biol. 317, 225–233 (2008).
Swanson, L. E. & Beitel, G. J. Tubulogenesis: an inside job. Curr. Biol. 16, R51–R53 (2006).
Wang, S. et al. Septate-junction-dependent luminal deposition of chitin deacetylases restricts tube elongation in the Drosophila trachea. Curr. Biol. 16, 180–185 (2006).
Luschnig, S., Bätz, T., Armbruster, K. & Krasnow, M. A. serpentine and vermiform encode matrix proteins with chitin binding and deacetylation domains that limit tracheal tube length in Drosophila. Curr. Biol. 16, 186–194 (2006).
Bagnat, M. et al. Cse1l is a negative regulator of CFTR-dependent fluid secretion. Curr. Biol. 20, 1840–1845 (2010).
Yang, B., Sonawane, N. D., Zhao, D., Somlo, S. & Verkman, A. S. Small-molecule CFTR inhibitors slow cyst growth in polycystic kidney disease. J. Am. Soc. Nephrol. 19, 1300–1310 (2008).
Ferrari, A., Veligodskiy, A., Berge, U., Lucas, M. S. & Kroschewski, R. ROCK-mediated contractility, tight junctions and channels contribute to the conversion of a preapical patch into apical surface during isochoric lumen initiation. J. Cell Sci. 121, 3649–3663 (2008).
Bagnat, M., Cheung, I. D., Mostov, K. E. & Stainier, D. Y. R. Genetic control of single lumen formation in the zebrafish gut. Nature Cell Biol. 9, 954–960 (2007).
Paul, S. M., Ternet, M., Salvaterra, P. M. & Beitel, G. J. The Na+/K+ ATPase is required for septate junction function and epithelial tube-size control in the Drosophila tracheal system. Development 130, 4963–4974 (2003).
Schottenfeld-Roames, J. & Ghabrial, A. S. Osmotic regulation of seamless tube growth. Nature Cell Biol. 15, 137–139 (2013).
Tsarouhas, V. et al. Sequential pulses of apical epithelial secretion and endocytosis drive airway maturation in Drosophila. Dev. Cell 13, 214–225 (2007).
Jayaram, S. A. et al. COPI vesicle transport is a common requirement for tube expansion in Drosophila. PLoS ONE 3, e1964 (2008).
Förster, D., Armbruster, K. & Luschnig, S. Sec24-dependent secretion drives cell-autonomous expansion of tracheal tubes in Drosophila. Curr. Biol. 20, 62–68 (2010).
Ghabrial, A. S., Levi, B. P. & Krasnow, M. A. A systematic screen for tube morphogenesis and branching genes in the Drosophila tracheal system. PLoS Genet. 7, e1002087 (2011).
Maybeck, V. & Röper, K. A targeted gain-of-function screen identifies genes affecting salivary gland morphogenesis/tubulogenesis in Drosophila. Genetics 181, 543–565 (2009).
Baer, M. M., Bilstein, A. & Leptin, M. A clonal genetic screen for mutants causing defects in larval tracheal morphogenesis in Drosophila. Genetics 176, 2279–2291 (2007).
Baer, M. M., Palm, W., Eaton, S., Leptin, M. & Affolter, M. Microsomal triacylglycerol transfer protein (MTP) is required to expand tracheal lumen in Drosophila in a cell-autonomous manner. J. Cell Sci. 125, 6038–6048 (2012).
Macara, I. G. Parsing the polarity code. Nature Rev. Mol. Cell Biol. 5, 220–231 (2004).
Mellman, I. & Nelson, W. J. Coordinated protein sorting, targeting and distribution in polarized cells. Nature Rev. Mol. Cell Biol. 9, 833–845 (2008).
Tepass, U. The apical polarity protein network in Drosophila epithelial cells: regulation of polarity, junctions, morphogenesis, cell growth, and survival. Annu. Rev. Cell Dev. Biol. 28, 655–685 (2012).
Drubin, D. G. & Nelson, W. J. Origins of cell polarity. Cell 84, 335–344 (1996).
Assémat, E., Bazellières, E., Pallesi-Pocachard, E., Le Bivic, A. & Massey-Harroche, D. Polarity complex proteins. Biochim. Biophys. Acta 1778, 614–630 (2008).
Mellman, I. & Warren, G. The road taken: past and future foundations of membrane traffic. Cell 100, 99–112 (2000).
Hughes, H. & Stephens, D. J. Assembly, organization, and function of the COPII coat. Histochem. Cell Biol. 129, 129–151 (2008).
Rabouille, C. & Klumperman, J. The maturing role of COPI vesicles in intra-Golgi transport. Nature Rev. Mol. Cell Biol. 6, 812–817 (2005).
Cai, H., Reinisch, K. & Ferro-Novick, S. Coats, tethers, Rabs, and SNAREs work together to mediate the intracellular destination of a transport vesicle. Dev. Cell 12, 671–682 (2007).
Hsu, S.-C., TerBush, D., Abraham, M. & Guo, W. The exocyst complex in polarized exocytosis. Int. Rev. Cytol. 233, 243–265 (2004).
Grieve, A. G. & Rabouille, C. Golgi bypass: skirting around the heart of classical secretion. Cold Spring Harb. Perspect. Biol. 3, a005298 (2011).
Andrew, D. J. & Ewald, A. J. Morphogenesis of epithelial tubes: insights into tube formation, elongation, and elaboration. Dev. Biol. 341, 34–55 (2010).
Zheng, Z. et al. LGN regulates mitotic spindle orientation during epithelial morphogenesis. J. Cell Biol. 189 275–288 (2010).
Bañón-Rodríguez, I. et al. EGFR controls IQGAP basolateral membrane localization and mitotic spindle orientation during epithelial morphogenesis. EMBO J. 33, 129–145 (2014).
Davis, G. E., Stratman, A. N., Sacharidou, A. & Koh, W. Molecular basis for endothelial lumen formation and tubulogenesis during vasculogenesis and angiogenic sprouting. Int. Rev. Cell Mol. Biol. 288, 101–165 (2011).
Acknowledgements
The authors thank M. Affolter, S. Caviglia, S. De Renzis, M. Labouesse, S. Luschnig, K. Röper and Y. Schwab for critical comments on the manuscript. Funding was from the DFG (SFB 572 and LE 546/7-1) and EMBO.
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Glossary
- Angiogenesis
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The process of forming new blood vessels by sprouting from already-established vasculature.
- Vasculogenesis
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The formation of new blood vessels from a group of endothelial precursors.
- Pinocytosis
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A mode of endocytosis by which cells non-specifically take up fluid from their surroundings.
- Placodes
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Polarized embryonic epithelial layers from which an organ or a structure later develops, often through the invagination of the placodes.
- Integrin
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A transmembrane protein found as a heterodimer of α- and β-subunits that mediates cell adhesion to the extracellular matrix and to neighbouring cells.
- Apical membrane initiation site
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(AMIS). An early apical domain between cells marking the membrane domain where a lumen will start forming, preceding the formation of a tight-junction-delimited lumen. It is characterized by the accumulation of apical polarity and trafficking proteins.
- Podocalyxin
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A transmembrane glycoprotein that is a major constituent of the glycocalyx, which is found in the kidney.
- Transcytosis
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The transport of vesicles between the apical and basolateral membranes of polarized cells. This requires endocytosis from one membrane domain followed by trafficking of those endosomes to the other domain.
- Pre-apical patch
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(PAP). An apical domain that has developed from the apical membrane initiation site and is found between cells where the lumen has become tight-junction-delimited.
- SNARE
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A protein superfamily that mediates target specificity and membrane fusion in vesicle trafficking. SNAREs can be divided into v-SNAREs (vesicle) and t-SNAREs (target) on the basis of their localization.
- SLP family
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(Synaptotagmin-like protein family). A family of proteins that have a conserved synaptotagmin-like homology domain (SHD) at the amino terminus and that are known to control secretion events.
- Rhabdomere
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A specialized microvillus on the apical surface of a photoreceptor cell that contains the visual pigments.
- Spectrin
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A large heterotetrameric protein of α- and β-subunits that lines the plasma membrane and plays an important part in plasma membrane integrity and cytoskeletal structure.
- Cysteine-rich intestinal protein
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(CRIP). A member of the LIM protein family; it is characterized by its two contiguous zinc-finger domains.
- Cystic fibrosis transmembrane conductance regulator
-
An ion channel that transports chloride across epithelial cell membranes.
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Sigurbjörnsdóttir, S., Mathew, R. & Leptin, M. Molecular mechanisms of de novo lumen formation. Nat Rev Mol Cell Biol 15, 665–676 (2014). https://doi.org/10.1038/nrm3871
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DOI: https://doi.org/10.1038/nrm3871
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