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Generation of specialized blood vessels via lymphatic transdifferentiation


The lineage and developmental trajectory of a cell are key determinants of cellular identity. In the vascular system, endothelial cells (ECs) of blood and lymphatic vessels differentiate and specialize to cater to the unique physiological demands of each organ1,2. Although lymphatic vessels were shown to derive from multiple cellular origins, lymphatic ECs (LECs) are not known to generate other cell types3,4. Here we use recurrent imaging and lineage-tracing of ECs in zebrafish anal fins, from early development to adulthood, to uncover a mechanism of specialized blood vessel formation through the transdifferentiation of LECs. Moreover, we demonstrate that deriving anal-fin vessels from lymphatic versus blood ECs results in functional differences in the adult organism, uncovering a link between cell ontogeny and functionality. We further use single-cell RNA-sequencing analysis to characterize the different cellular populations and transition states involved in the transdifferentiation process. Finally, we show that, similar to normal development, the vasculature is rederived from lymphatics during anal-fin regeneration, demonstrating that LECs in adult fish retain both potency and plasticity for generating blood ECs. Overall, our research highlights an innate mechanism of blood vessel formation through LEC transdifferentiation, and provides in vivo evidence for a link between cell ontogeny and functionality in ECs.

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Fig. 1: Stepwise formation of the AF vasculature.
Fig. 2: Lymphatic vessels give rise to the adult AF vasculature through transdifferentiation.
Fig. 3: The functional specialization of AF vessels is linked to their cellular origins.
Fig. 4: Vascularization of regenerating AFs recapitulates developmental programs.

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Data availability

Sequencing data generated for this study are available at the Gene Expression Omnibus under accession number GSE197161. Reagents and source data are available from the corresponding authors on request.


  1. Augustin, H. G. & Koh, G. Y. Organotypic vasculature: from descriptive heterogeneity to functional pathophysiology. Science 357, eaal2379 (2017).

    PubMed  Google Scholar 

  2. Petrova, T. V. & Koh, G. Y. Organ-specific lymphatic vasculature: from development to pathophysiology. J. Exp. Med. 215, 35–49 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  3. Semo, J., Nicenboim, J. & Yaniv, K. Development of the lymphatic system: new questions and paradigms. Development 143, 924–935 (2016).

    Article  CAS  PubMed  Google Scholar 

  4. Gutierrez-Miranda, L. & Yaniv, K. Cellular origins of the lymphatic endothelium: implications for cancer lymphangiogenesis. Front. Physiol. 11, 577584 (2020).

    Article  PubMed  PubMed Central  Google Scholar 

  5. Parichy, D. M., Elizondo, M. R., Mills, M. G., Gordon, T. N. & Engeszer, R. E. Normal table of postembryonic zebrafish development: staging by externally visible anatomy of the living fish. Dev. Dyn. 238, 2975–3015 (2009).

    Article  PubMed  PubMed Central  Google Scholar 

  6. Marí-Beffa, M. & Murciano, C. Dermoskeleton morphogenesis in zebrafish fins. Dev. Dyn. 239, 2779–2794 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  7. Vogel, W. O. P. & Claviez, M. Vascular specialization in fish, but no evidence for lymphatics. Z. Naturforsch. 36, 490–492 (1981).

    Article  Google Scholar 

  8. Steffensen, J. F., Lomholt, J. P. & Vogel, W. O. P. In vivo observations on a specialized microvasculature, the primary and secondary vessels in fishes. Acta Zool. 67, 193–200 (1986).

    Article  Google Scholar 

  9. Olson, K. R. Secondary circulation in fish: anatomical organization and physiological significance. J. Exp. Zool. 275, 172–185 (1996).

    Article  Google Scholar 

  10. Jensen, L. D. E. et al. Nitric oxide permits hypoxia-induced lymphatic perfusion by controlling arterial-lymphatic conduits in zebrafish and glass catfish. Proc. Natl Acad. Sci. USA 106, 18408–18413 (2009).

    Article  CAS  ADS  Google Scholar 

  11. Rummer, J. L., Wang, S., Steffensen, J. F. & Randall, D. J. Function and control of the fish secondary vascular system, a contrast to mammalian lymphatic systems. J. Exp. Biol. 217, 751–757 (2014).

    CAS  PubMed  Google Scholar 

  12. Karpanen, T. & Schulte-Merker, S. in Methods in Cell Biology (eds. Detrich, H. W. et al.) Vol. 105, 223–238 (Academic, 2011).

  13. Jung, H. M. et al. Development of the larval lymphatic system in zebrafish. Development 144, 2070–2081 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  14. Gancz, D., Perlmoter, G. & Yaniv, K. Formation and growth of cardiac lymphatics during embryonic development, heart regeneration, and disease. Cold Spring Harb. Perspect. Biol. 12, a037176 (2019).

    Article  CAS  Google Scholar 

  15. Potente, M. & Makinen, T. Vascular heterogeneity and specialization in development and disease. Nat. Rev. Mol. Cell Biol. 18, 477–494 (2017).

    Article  CAS  PubMed  Google Scholar 

  16. Yaniv, K. et al. Live imaging of lymphatic development in the zebrafish. Nat. Med. 12, 711–716 (2006).

    Article  CAS  PubMed  Google Scholar 

  17. Dunwoodie, S. L. The role of hypoxia in development of the mammalian embryo. Dev. Cell 17, 755–773 (2009).

    Article  CAS  PubMed  Google Scholar 

  18. Akiva, A. et al. On the pathway of mineral deposition in larval zebrafish caudal fin bone. Bone 75, 192–200 (2015).

    Article  CAS  PubMed  Google Scholar 

  19. Bennet, M. et al. Simultaneous Raman microspectroscopy and fluorescence imaging of bone mineralization in living zebrafish larvae. Biophys. J. 106, L17–L19 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Zhao, L. et al. Notch signaling regulates cardiomyocyte proliferation during zebrafish heart regeneration. Proc. Natl Acad. Sci. USA 111, 1403–1408 (2014).

    Article  CAS  ADS  PubMed  PubMed Central  Google Scholar 

  21. Kobayashi, I. et al. Jam1a–Jam2a interactions regulate haematopoietic stem cell fate through Notch signalling. Nature 512, 319–323 (2014).

    Article  CAS  ADS  PubMed  PubMed Central  Google Scholar 

  22. Azimi, M. S. et al. Lymphatic-to-blood vessel transition in adult microvascular networks: a discovery made possible by a top-down approach to biomimetic model development. Microcirculation 27, e12595 (2020).

    PubMed  Google Scholar 

  23. Johnson, N. C. et al. Lymphatic endothelial cell identity is reversible and its maintenance requires Prox1 activity. Genes Dev. 22, 3282–3291 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Ma, W. & Oliver, G. Lymphatic endothelial cell plasticity in development and disease. Physiology 32, 444–452 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Chen, C.-Y. et al. Blood flow reprograms lymphatic vessels to blood vessels. J. Clin. Invest. 122, 2006–2017 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  26. Chen, J. et al. Acute brain vascular regeneration occurs via lymphatic transdifferentiation. Dev. Cell 56, 3115–3127 (2021).

    Article  CAS  PubMed  Google Scholar 

  27. Kim, J. et al. Impaired angiopoietin/Tie2 signaling compromises Schlemm’s canal integrity and induces glaucoma. J. Clin. Invest. 127, 3877–3896.

  28. Corada, M. et al. Sox17 is indispensable for acquisition and maintenance of arterial identity. Nat. Commun. 4, 2609 (2013).

    Article  ADS  CAS  PubMed  Google Scholar 

  29. Gancz, D. et al. Distinct origins and molecular mechanisms contribute to lymphatic formation during cardiac growth and regeneration. eLife 8, e44153 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Nicenboim, J. et al. Lymphatic vessels arise from specialized angioblasts within a venous niche. Nature 522, 56–61 (2015).

    Article  CAS  ADS  PubMed  Google Scholar 

  31. Hen, G. et al. Venous-derived angioblasts generate organ-specific vessels during zebrafish embryonic development. Dev. Camb. Engl. 142, 4266–4278 (2015).

    CAS  Google Scholar 

  32. Keren-Shaul, H. et al. MARS-seq2.0: an experimental and analytical pipeline for indexed sorting combined with single-cell RNA sequencing. Nat. Protoc. 14, 1841–1862 (2019).

    Article  CAS  PubMed  Google Scholar 

  33. Moon, K. R. et al. Visualizing structure and transitions in high-dimensional biological data. Nat. Biotechnol. 37, 1482–1492 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  34. Street, K. et al. Slingshot: cell lineage and pseudotime inference for single-cell transcriptomics. BMC Genom. 19, 477 (2018).

    Article  CAS  Google Scholar 

  35. Wolf, F. A. et al. PAGA: graph abstraction reconciles clustering with trajectory inference through a topology preserving map of single cells. Genome Biol. 20, 59 (2019).

    Article  PubMed  PubMed Central  Google Scholar 

  36. Harikumar, A. et al. Embryonic stem cell differentiation is regulated by SET through interactions with p53 and β-catenin. Stem Cell Rep. 15, 1260–1274 (2020).

    Article  CAS  Google Scholar 

  37. Zhou, X. et al. HMGB2 regulates satellite-cell-mediated skeletal muscle regeneration through IGF2BP2. J. Cell Sci. 129, 4305–4316 (2016).

    Article  CAS  PubMed  Google Scholar 

  38. Garza-Manero, S. et al. Maintenance of active chromatin states by HMGN2 is required for stem cell identity in a pluripotent stem cell model. Epigenetics Chromatin 12, 73 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Bao, X. et al. CSNK1a1 regulates PRMT1 to maintain the progenitor state in self-renewing somatic tissue. Dev. Cell 43, 227–239 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. Jerafi-Vider, A. et al. VEGFC/FLT4-induced cell-cycle arrest mediates sprouting and differentiation of venous and lymphatic endothelial cells. Cell Rep. 35, 109255 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  41. Shin, M. et al. Vegfc acts through ERK to induce sprouting and differentiation of trunk lymphatic progenitors. Development 143, 3785–3795 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. Singh, S. P., Holdway, J. E. & Poss, K. D. Regeneration of amputated zebrafish fin rays from de novo osteoblasts. Dev. Cell 22, 879–886 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Silvent, J. et al. Zebrafish skeleton development: high resolution micro-CT and FIB-SEM block surface serial imaging for phenotype identification. PLoS ONE 12, e0177731 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  44. Das, R. N. & Yaniv, K. Discovering new progenitor cell populations through lineage tracing and in vivo imaging. Cold Spring Harb. Perspect. Biol. 12, a035618 (2020).

    Article  CAS  PubMed  Google Scholar 

  45. Vogel, W. O. P. Zebrafish and lymphangiogenesis: a reply. Anat. Sci. Int. 85, 118–119 (2010).

    Article  PubMed  Google Scholar 

  46. Gur-Cohen, S. et al. Stem cell-driven lymphatic remodeling coordinates tissue regeneration. Science 366, 1218–1225 (2019).

    Article  CAS  ADS  PubMed  PubMed Central  Google Scholar 

  47. Louveau, A. et al. CNS lymphatic drainage and neuroinflammation are regulated by meningeal lymphatic vasculature. Nat. Neurosci. 21, 1380–1391 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  48. Da Mesquita, S. et al. Functional aspects of meningeal lymphatics in ageing and Alzheimer’s disease. Nature 560, 185–191 (2018).

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  49. Pavlov, V. et al. Hydraulic control of tuna fins: a role for the lymphatic system in vertebrate locomotion. Science 357, 310–314 (2017).

    Article  CAS  ADS  PubMed  PubMed Central  Google Scholar 

  50. Oliver, G., Kipnis, J., Randolph, G. J. & Harvey, N. L. The lymphatic vasculature in the 21st century: novel functional roles in homeostasis and disease. Cell 182, 270–296 (2020).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  51. Pawlak, J. B. et al. Lymphatic mimicry in maternal endothelial cells promotes placental spiral artery remodeling. J. Clin. Invest. 129, 4912–4921.

  52. Song, E. et al. VEGF-C-driven lymphatic drainage enables immunosurveillance of brain tumours. Nature 577, 629–630 (2020).

    Article  CAS  Google Scholar 

  53. Jin, S.-W., Beis, D., Mitchell, T., Chen, J.-N. & Stainier, D. Y. R. Cellular and molecular analyses of vascular tube and lumen formation in zebrafish. Development 132, 5199–5209 (2005).

    Article  CAS  PubMed  Google Scholar 

  54. Matsuoka, R. L. et al. Radial glia regulate vascular patterning around the developing spinal cord. eLife 5, e20253 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  55. Spoorendonk, K. M. et al. Retinoic acid and Cyp26b1 are critical regulators of osteogenesis in the axial skeleton. Development 135, 3765–3774 (2008).

    Article  CAS  PubMed  Google Scholar 

  56. Shin, J., Poling, J., Park, H.-C. & Appel, B. Notch signaling regulates neural precursor allocation and binary neuronal fate decisions in zebrafish. Development 134, 1911–1920 (2007).

    Article  CAS  PubMed  Google Scholar 

  57. Davison, J. M. et al. Transactivation from Gal4-VP16 transgenic insertions for tissue-specific cell labeling and ablation in zebrafish. Dev. Biol. 304, 811–824 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  58. Avraham-Davidi, I. et al. ApoB-containing lipoproteins regulate angiogenesis by modulating expression of VEGF receptor 1. Nat. Med. 18, 967–973 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  59. White, R. M. et al. Transparent adult zebrafish as a tool for in vivo transplantation analysis. Cell Stem Cell 2, 183–189 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  60. Villefranc, J. A., Amigo, J. & Lawson, N. D. Gateway compatible vectors for analysis of gene function in the zebrafish. Dev. Dyn. 236, 3077–3087 (2007).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  61. Hesselson, D., Anderson, R. M., Beinat, M. & Stainier, D. Y. Distinct populations of quiescent and proliferative pancreatic beta-cells identified by HOTcre mediated labeling. Proc. Natl Acad. Sci. USA 106, 14896–14901 (2009).

    Article  CAS  ADS  PubMed  PubMed Central  Google Scholar 

  62. Suster, M. L., Abe, G., Schouw, A. & Kawakami, K. Transposon-mediated BAC transgenesis in zebrafish. Nat. Protoc. 6, 1998–2021 (2011).

    Article  CAS  PubMed  Google Scholar 

  63. Dahlem, T. J. et al. Simple methods for generating and detecting locus-specific mutations induced with TALENs in the zebrafish genome. PLoS Genet. 8, e1002861 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  64. Han, Y. et al. Vitamin D stimulates cardiomyocyte proliferation and controls organ size and regeneration in zebrafish. Dev. Cell 48, 853–863 (2019).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  65. Oehlers, S. H. et al. Interception of host angiogenic signalling limits mycobacterial growth. Nature 517, 612–615 (2015).

    Article  CAS  ADS  PubMed  Google Scholar 

  66. Lyubimova, A. et al. Single-molecule mRNA detection and counting in mammalian tissue. Nat. Protoc. 8, 1743–1758 (2013).

    Article  CAS  PubMed  Google Scholar 

  67. Schindelin, J. et al. Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682 (2012).

    CAS  PubMed  Google Scholar 

  68. Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 550 (2014).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  69. Zhu, A., Ibrahim, J. G. & Love, M. I. Heavy-tailed prior distributions for sequence count data: removing the noise and preserving large differences. Bioinformatics 35, 2084–2092 (2019).

    Article  CAS  PubMed  Google Scholar 

  70. Hoffman, D. et al. A non-classical monocyte-derived macrophage subset provides a splenic replication niche for intracellular Salmonella. Immunity 54, 2712–2723 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  71. Manco, R. et al. Clump sequencing exposes the spatial expression programs of intestinal secretory cells. Nat. Commun. 12, 3074 (2021).

    Article  CAS  ADS  PubMed  PubMed Central  Google Scholar 

  72. Kult, S. et al. Bi-fated tendon-to-bone attachment cells are regulated by shared enhancers and KLF transcription factors. eLife 10, e55361 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  73. McFarland, A. P. et al. Multi-tissue single-cell analysis deconstructs the complex programs of mouse natural killer and type 1 innate lymphoid cells in tissues and circulation. Immunity 54, 1320–1337 (2021).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  74. Satija, R., Farrell, J. A., Gennert, D., Schier, A. F. & Regev, A. Spatial reconstruction of single-cell gene expression data. Nat. Biotechnol. 33, 495–502 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  75. van Dijk, D. et al. Recovering gene interactions from single-cell data using data diffusion. Cell 174, 716–729 (2018).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  76. Wolf, F. A., Angerer, P. & Theis, F. J. SCANPY: large-scale single-cell gene expression data analysis. Genome Biol. 19, 15 (2018).

    Article  PubMed  PubMed Central  Google Scholar 

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We thank H. Raviv, L. Shen, D. Robinson and A. Solomon for technical assistance; K. Kumari for assistance with image analysis and illustrations; G. Almog, R. Hofi, A. Glozman and A. Tatarin for animal care; M. Biton for insights into scRNA-seq analyses; S. Kapishnikov for assistance with µCT experiments; and all of the members of the Yaniv laboratory for discussion, technical assistance and continuous support. This work was supported in part by the European Research Council (818858) to K.Y., the Minerva Foundation (712610) to K.Y. and the H&M Kimmel Institute for Stem Cell Research (Weizmann Institute). Research in the Yaniv laboratory is generously supported by a research grant from the Estate of Mady Dukler, a research grant from Madame Olga Klein – Astrachan and the Estate of Emile Mimran (SABRA program). K.Y. is the incumbent of the Enid Barden and Aaron J. Jade Professorial Chair in Memory of Canter John Y. Jade and is the Director of the Aharon Katzir-Katchalsky Center. R.N.D. was supported by an EMBO long-term fellowship (ALTF 1532-2015), an Edith and Edward F. Anixter Postdoctoral Fellowship and a senior postdoctoral fellowship by the Weizmann Institute of Science. W.H. was supported by the Deutsche Forschungsgemeinschaft (HE4585/4-1) and by the North Rhine-Westphalia ‘return fellowship’. R.A. is supported by the European Research Council (756653) and the Israel Science Foundation (1890/17). K.D.P. acknowledges support from the National Institutes of Health (R35 HL150713, R01 HD105033). All illustrations shown in the figures (except for Fig. 1i) and Extended Data figures were obtained from BioRender.

Author information

Authors and Affiliations



R.N.D. designed and conducted experiments, analysed data and co-wrote the manuscript. I.B., N.M., G.L. and Y.T. conducted experiments and data analyses. S.S. and Y.T. performed bioinformatics analyses. M.B. and J.N. generated transgenic lines. Y.H. and K.D.P. generated knockin zebrafish and analysed data. D.H. and R.E.-A. assisted with smFISH experiments. W.H. provided transgenic lines. R.A. supervised scRNA-seq data analyses. K.Y. directed the study, designed experiments, analysed data and co-wrote the paper with input from all of the authors.

Corresponding authors

Correspondence to Rudra N. Das or Karina Yaniv.

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The authors declare no competing interests.

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Nature thanks Dominic Gruen, Oliver Stone and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

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Extended data figures and tables

Extended Data Fig. 1 Morphological and molecular characterization of anal and dorsal fins formation.

a-d′, mrc1a:EGFP and prox1a:RFP colocalization specifically highlights lymphatic vessels in zebrafish larvae. a, Bright-light image of 14 dpf zebrafish trunk, with boxes demarking dorsal (green), medial (red) and ventral (blue) areas depicted in b-d. b,c,d, show prox1a (red) and mrc1a (green) transgene expression, b′,c′,d′ depict colocalization channel (yellow). e-e′, intravascularly injected Qdot705 (cyan) are detected within blood vessels but not in mrc1a/prox1a co-labelled lymphatics (yellow). f-f″, Trunk blood and lymphatics vessels are differentially visualized in kdrl:BFP;mrc1a:EGFP;prox1a:RFP juvenile fish. mrc1a/prox1a colocalization (f′) labels lymphatics, while kdrl highlights blood vessels (f″, blue). g-j, Bright-light images of AF development with yellow arrowheads indicating important stage-specific features, such as mesenchyme condensation (g, arrowheads), appearance of rays (h, arrowheads), ray growth (i, arrowheads) and formation of joints (j, arrowheads). AF is first detected when the larva reaches ~5.2 mm in length (~15-17 dpf). k-m, Triple Tg(prox1a:RFP;mrc1a:EGFP;kdrl:BFP) juvenile zebrafish, whose prox1a:RFP;mrc1a:EGFP colocalization channel is shown in Fig. 1c–g. n-n″, LVC mediated vascularization of DF. n, Schematic showing the position of the DF. n′,n″, Confocal images of Tg(prox1a:RFP;mrc1a:EGFP;kdrl:BFP) animals show contribution of LVC and BVC during DF formation. n′ shows all channels, n″ shows prox1a:RFP;mrc1a:EGFP colocalization channel along with kdrl:BFP. Scale bars, 20 µm (e′), 30 µm (b′,c′,d′,f′,g,k,k′), 50 µm (h,i,l,m,n′), 100 µm (j), 150 µm (a). DL, Dorsal lymphatic, SL, Spinal lymphatic, DIL, Deep intersegmental lymphatic, TD, Thoracic duct, CCL, Collateral cardinal lymphatic, LL, Lateral lymphatic, SIL, Superficial intersegmental lymphatic. Panel n was created using BioRender.

Extended Data Fig. 2 Molecular characterization of the AF vascular component.

a-a″, Confocal image of Tg(mrc1a:GFP;prox1a:RFP) larva showing connection (a″, arrowhead) of AF lymphatic sprout to the TD and CCL (a″, arrows). White dashed lines demarcate the developing AF; a″ depicts colocalization channel. b-d, Expression of lyve1b:dsRed through stages I-III of AF development shown along with kdrl:GFP+ BVC. Arrows in b point to the LVC sprouts at stage I. e-j, smFISH analysis of blood and lymphatic marker expression at stages I (e,f,h) and II (g) of AF development. e-h, prox1a mRNA puncta (magenta) are detected only in mrc1a:GFP+ LVC, while fli1a puncta (yellow) marks both the kdrl:GFP+ BVC and mrc1a:GFP+ LVC (e,e′,h). Blue dashed lines enclose the LVC and white box demarcates the insets shown in e′. Illustrations indicate the imaged ROI (black dashed box). I,j, Abundance of prox1a mRNA (i) and fli1a mRNA (j) (nAFs-LVC = 10; nAFs-BVC = 12) in LVC vs BVC. Boxes show 25th to 75th percentiles, line depicts median and whiskers show the full range of the data points. k-k″, Distribution of calcein in early stage III AF (k), following intramuscular injection in the trunk. k′,k″, High-magnification of yellow box in k, showing calcein (green) within the lumen of a prox1a+ LVC. Arrows point to the bony rays labelled by calcein. l-l′, Hypoxyprobe staining in stage II AF (l) and its vehicle (PBS) injected control (l′). Yellow dashed lines indicate the area of the main lymphatic vessel in the AF. m,n, Stage IV AF in prox1a (m) and mrc1a (n) transgenic fish, whose colocalization channel is depicted in Fig. 1k. o, Intact fli1a:dsRed+ vasculature is present in stage IV AFs. p,p′, Intravascularly injected Qdot705 are detected in all vessels (fli1a:dsRed+) of the adult AF. Scale bars, 20 μm (k,k′,l,l′), 30 μm (a,b,c), 50 μm (d), 100 μm (p,p′), 200 μm (m), 500 μm (o). TD, Thoracic duct, CCL, Collateral cardinal lymphatic, PCV, Posterior cardinal vein.

Extended Data Fig. 3 A platform for long-term EC lineage tracing.

a, Schematic representation of the flibow construct used to drive expression of brainbow in ECs. Cre recombinase allows expression of different fluorescent proteins (mCerulean or EYFP) in non-recombinant (expressing tdTomato) animals. b, Schematic representation of the hsp70l:creERT2;flibow lineage tracing protocol used in this study. c-g, Representative examples of induced recombination. Combined heat- and 4-OHT treatment allowed clear identification of 27 larvae (out of 54 screened) displaying differentially labelled ECs. c′, Normalized intensity plotted across a range of emitted wavelength (x-axis) from 5 PCV-ECs (c, numbered arrowheads), depicting how ECs display a unique spectral signature, based on the levels of expression of the 3 fluorophores. 4-OHT (d, 61/65) or heat treatment (e, 49/50) alone did not induce recombinant outcomes. d′ and e′ show ‘spectral signatures’ from 9 ECs from each group, demonstrating non-recombinant outcome depicted by the presence of tdTomato emission signal only. f, No recombination is detected in adult AFs treated with 4-OHT (22/22) or heat (g, 56/57) alone, and raised through adulthood. h-h″, Confocal images of flibow larvae showing distinct labelling of lymphatic components in the trunk starting from 24 h after Cre induction. h, ECs in the early TD of 3.5 dpf embryo labelled in green (arrowheads), are detected 24 h after heat and 4-OHT treatments. h′-h″, Reiterative imaging of the same embryo at 4.5 dpf (h′) and 5.5 dpf (h″) shows the growing TD composed of green ECs (arrowheads). i-j′, Assessment of the stability of the fluorophore ratios after Cre induction. i,i′, Average normalized intensity from a ‘switched’ clonal population (n = 4) measured at 24, 48 and 72 h post treatment (hpt), showing slight deviation with time (i′). Intensities corresponding to each fluorophore are depicted separately for the 3 time-points and show a decay in tdTomato signal, that contributes to the deviation in the spectral signature shown in i. j,j′, Similar measurements carried out on ‘switched’ clonal ECs from another sample at 16, 18, 22, 29 days post treatment (dpt), shows a negligible deviation between the channels (j), along with stable intensity proportions between the fluorophores (j′). Scale bars, 15 μm (c), 30 μm (d,e,h,h′,h″), 300 μm (f,g). PCV, Posterior Cardinal vein; DA, Dorsal Aorta; EC, endothelial cell; 4-OHT, 4-hydroxytamoxifen. Panels a, b were created using BioRender.

Extended Data Fig. 4 Lineage tracing of BVC and LVC in growing AF.

a, Summary of hsp70l:creERT2;flibow experiments. b-c′, Confocal images of flibow fish AF (Fish #26) showing a green labelled-blood vessel sprout generating the BVC in stage I AF (b,b′, arrowhead, green). Clonally related ECs are detected in the dorso-anterior AF of the same animal at adult stages (c, arrowheads). Initial lymphatic sprout (b,b″, arrows, blue) in contrast, expands throughout the entire fin (c′). AF is demarcated by yellow dashed lines, asterisks in c indicate signal from unrelated non-EC autofluorescence. d-e, Flibow lineage tracing of BVC. Confocal images of the kdrl:creERT2;flibow stage III (d) and adult AF (e) showing distinct labelling of BVC component near the trunk-fin junction (arrows). LVC-derived ray vessels display non-recombinant tdTomato signal. White dashed box indicates region shown in the inset. The lack of fluorescent signal in the middle of AF is due to obstruction by the pigmented stripes in non-casper fish. f-f′, Confocal images of the kdrl:creERT2;β-actin2:loxP-BFP-loxP-DsRed adult AF showing contribution of BVC (arrows) to AF vasculature. When kdrl:creERT2 mediated recombination was induced between 3.5-5.5 dpf (nswitch line = 40, nflibow = 66), 100% of the labelled clones found in the AF (nswitch line = 10, nflibow = 18), were restricted to the BVC. Yellow dashed lines indicate the boundaries of the AF. Scale bars, 10 μm (b), 30 μm (d), 200 μm (c′), 300 μm (e,f).

Extended Data Fig. 5 Loss of lymphatic fate is associated with elevated expression of blood EC markers.

a-b, Scatterplot showing rlog expression of genes (a), in PROX1 suppressed HDLECs, that are upregulated (dark green), downregulated (light green) or did not change significantly (grey). Genes of interest are annotated in the plot. Plots (b) showing mean normalized expression of selected genes upregulated following PROX1 suppression in HDLECs (siPROX1). The box in the plots show 25th to 75th percentiles, the line shows the median and the whiskers show the full range of the data points. (****, p-adj value < 0.0001; ***, p-adj value < 0.001; *, p-value < 0.05, Wald test). c-g, Confocal images of Tg(sox17GFP) showing expression of sox17 across stages I-III AFs. c-c′, Initially, no sox17GFP expression is detected in lyve1b+ sprouts. d-g, Single GFP channel images of Fig 2h–k showing gradual enrichment of sox17 expression. h-h′, flt1_9a:EGFP does not label prox1a:RFP+ differentiated lymphatic vessels (TD) in 13 dpf larval trunk. i-j′, Gradual increase of flt1_9a:EGFP expression at stage III shown along with lyve1b:dsRed+ (i) and prox1a:RFP+ (j). Arrowheads in single GFP channel point to flt1_9a+ newly expressing ECs within the LVC plexus. k, Confocal image of Tg(flt1:YFP) showing specific expression in the BVC in early stage III AF (arrowheads). l-m″, Gradual expression of kdrl in sox17GFP + vessels of AF. l-l′, LVC (lyve1b:dsRed) and BVC (kdrl:BFP) shown together along with transdifferentiated LVC-ECs (sox17GFP) in stage II AF. The transdifferentiating LVC-ECs (l′, arrows) do not show expression of kdrl, which at this stage is restricted to BVC (l′, arrowheads; l″). m-m′, At the onset of stage IV (m), the growing sox17GFP + vessels, start expressing kdrl (m′, arrows). BVC-ECs are sox17GFP - but kdrl+ (m′, arrowheads; m″). Scale bars, 30 µm (h,c,l), 50 µm (i,j), 100 µm (k). TD, Thoracic duct.

Extended Data Fig. 6 Expression of lymphatic and blood vessel-specific genes across the different scRNA-seq cell clusters.

a-c, UMAP (shown in Fig. 2m) plots with embedded imputed expression of selected marker genes. a, Blood vessel markers specifically enriched in the BVC. b, Lymphatic markers specific for LEC cluster. c, Blood vessel markers enriched in LVC1 cluster (transdifferentiated LECs). d, Partition graph abstraction (PAGA) analysis of LVC clusters. e, UMAP showing the trajectory across the clusters (black line) following Slingshot analysis. f-g, PHATE (shown in Fig. 2p) plots with embedded imputed expression of selected genes.

Extended Data Fig. 7 Sox17 misexpression results in suppression of lymphatic fate.

a-b′, sox17 misexpression results in loss of lymphatic marker expression in the AF. Stage IV AF (demarcated by yellow dashed lines) showing expression of lyve1b:dsRed, at the dorsal trunk-fin junction of the AF (a), which is lost following mosaic expression of UAS sox17 in hsp70l:Gal4 animals (b, asterisks) (N = 2, n = 65). a′,b′ show the corresponding bright-light images. c,d, Confocal images of a 4.5 dpf Tg(fli1a:Gal4; lyve1b:dsRed) embryo showing the TD (c, arrowheads) that is lost (d, asterisks) following mosaic overexpression of UAS sox17. e-g, Confocal images of a 7 dpf larva with mosaic overexpression of UAS:sox17 in Tg(fli1a:Gal4) embryos showing complete (f, asterisks) or partial (g, arrowheads point to TD, asterisks denote absent TD segments) disruption in TD formation (N = 2, ninjected embryos=234, nununinjected embryos = 87). Scale bars, 20 µm (c,d), 50 µm (e,f,g), 100 µm (a,b). TD, Thoracic duct; DA, Dorsal Aorta; PCV, Posterior cardinal vein.

Extended Data Fig. 8 Different blood flow pattern and connectivity with the systemic circulation in WT and flt4−/− animals.

a-b′, Different tracers are readily detectable in the LVC of Stage III AFs. Distribution of Qdot705 (a,a′, cyan) and FITC Dextran (MW = 500 kDa) (b-b′, yellow) in stage III AF. LVC is labelled by mrc1a/prox1a colocalization (a,a′, yellow) or prox1a (b, red); BVC is labelled by kdrl (blue) in b,b′. c-c′, Appearance of flt1_9a:GFP in the LVC (arrowheads) is detected in animals with intravascularly injected Qdot705 (cyan) before the tracer has entered the LVC compartment. Arrows point to BVC expression of flt1_9a:GFP. d-e′. Early stages of AF development, before the connection between AF and DA is established, shown through fli1a:dsRed and sox17GFP expression analysis. d, sox17+ ECs are not detected in stage I AF but are present in the trunk, near the DA. e-e′, Stage II AF depicting sox17+ sprouts (arrowheads) extending dorsally from the AF. DA-derived sox17+ sprouts extend ventrally but remain within the trunk (arrow). f-h, Connection of LVC-derived plexus with the trunk vasculature is established through LVC sprouts that extend dorsally to reach the trunk. f-f′, Stage II AF (f) show transdifferentiating LECs (lyve1b+;sox17+, arrowheads) sprouting dorsally towards the trunk. Arrow points to a trunk sox17+ vessel that has not entered the AF. f′ shows high magnification of single channel from f, to clearly depict the extending sprouts (arrowheads). g-g″, Microangiography with Qdot705 depicts patent connections established between the DA and stage III AF (not in the image) through a LVC-derived vessel (g′,g″, lyve1b+, white arrowheads). h., Lineage tracing through Tg(hsp70l:creERT2;flibow) animal depicting distinct labelling of BVC (crimson/violet mosaic, arrows) and LVC (green, arrowheads) and all their subsequent progenies. The LVC vessels sprout dorsally to connect with the trunk circulation. Scale bars, 20 µm (c′), 30 µm (d,e,f,g), 50 µm (a,b).

Extended Data Fig. 9 In absence of lymphatics the AF vasculature derives from trunk BECs.

a-a′, Trunk lymphatics (mrc1a+, a) are absent in flt4−/−mutants (a′). The only mrc1a+ structure visible in the trunk of flt4−/− is the PCV. b-e, The DF in flt4−/−mutants is vascularized by BECs. b-c, Early sprouting event in developing DFs in stage matched WT and flt4−/− animals, showing lyve1b+ LVC sprouts in WT (b) that are missing in flt4−/− (c). d-e, Juvenile DF is fully vascularized by LVC in WT and BECs in flt4−/−. f-h, NTR–MTZ ablation of LVC in Tg(prox1a:Gal4;UAS NTR-mCherry;kdrl:GFP) animals, results in kdrl+ BVC-derived vascularization of the AF (f), while DMSO (vehicle) treated animals develop a normal LVC-derived AF plexus (g,h). f shows the status of kdrl+ vessels in the AF, 3 days post first MTZ treatment. The vehicle treated AFs are shown at 5 days (g) and 15 days (h) after the first DMSO treatment. i-j, Lineage analysis of AF ECs in flt4−/−. i, Stage I AF of flt4−/−;hsp70l:creERT2;flibow fish (Fish #A) showing distinctly labelled clones (green, arrowheads) in BVC (connected to PCV and cloacal vessels). j, Adult AF vasculature of the same fish shown in m, bears clones of matching identity (arrowheads). Scale bars, 30 µm (c,f), 40 µm (g), 50 µm (a,a′,b,d,e,h,i), 300 µm (j).

Extended Data Fig. 10 Functional characterization of AF vessels in WT and flt4−/−.

a,b, Differential erythrocyte flow in WT vs. flt4−/− animals. Bright-light intensity profile from the lumen of fin ray vessels in WT (a) or flt4−/− (b) fish (nWT=3, nflt4−/−=3) showing different modes of erythrocyte flow in WT (no flow, intermittent flow, continuous flow) (a), vs. continuous erythrocyte flow in flt4−/− mutants (b). c, Quantification of erythrocyte-mediated intensity change (spikes) events (nwt=10; nflt4−/− =9). d-e, Differences in erythrocyte flow in WT vs. flt4−/− juvenile fish (stage IV). Bright-light intensity profile (d) from the lumen of fin ray vessels in WT vs flt4−/− are shown, along with the quantification of erythrocyte-mediated intensity change (spikes) events (e) (nwt=6; nflt4−/− =4). The box in the plots (c,e) show 25th to 75th percentiles, the line in the middle shows the median and the whiskers show the full range of the data points. f-h″, Confocal imaging of erythrocytes in the AF is shown with gata1a:dsRed, in the background of pan-endothelial fli1a:GFP. k-l, Confocal images of entire AFs, showing gata1a+ cells majorly accumulated in the anterior BVC in WT animals (f, arrow), as opposed to widespread distribution of gata1a+ cells in all AF vessels (g) in flt4−/−. h-h″, High magnification images of single rays from f and g depicting the presence or absence of gata1a+ cells inside the lumen of WT LVC derived AF ray vessels (h), WT BVC vessels (h′) and flt4−/− AF ray vessels (h″). i, Schematic illustration of the different functional properties of AF vessels originating via LEC transdifferentiation vs. BEC angiogenesis. j, 3D rendered image of a µCT scan of adult AF, showing the external rays (white) and the internal radials (green). k, Cross-section of the radials depicting the size and morphological defects (arrowhead) detected in flt4−/− as compared to WT siblings. Yellow dashed lines enclose one set of radials. l, Quantification of the MR length is plotted (nwt = 15; nflt4−/− = 15). Scale bars, 300 µm (f,g). ***p < 0.0005. Panel i was created using BioRender.

Extended Data Fig. 11 Lineage tracing of regenerating AF vasculature.

a-b, mrc1a:EGFP labelled LVC growth along osx:mCherry labelled bones, as detected in 5 dpa (a) and 7 dpa (a′,a″) AFs. Inset (a″) shows mrc1a channel only to visualize the LECs (arrowheads) growing along a single regenerating ray. b, Status and extent of BVC (kdrl:EGFP+, arrowheads) coverage in a 5dpa regenerating AF shown in contrast to LVC (prox1a:RFP+). c, flt1_9a:GFP expression is detected in prox1a:RFP+ LECs of the regenerating AF plexus at 4 dpa (arrowheads). d-e, Clonal identity of AF vessels before and after regeneration traced in the same animal. d, Dissected AF from an adult hsp70l:creERT2;flibow fish, showing distinctly labelled clones (green) throughout the fin except for the anterior-most rays. e, Regenerated AF in same fish shown in d, at 20 dpa. ECs of same clonal identity (green), populate the regenerating vasculature recapitulating the distribution seen in d. f, Proportion of new clones (blue) in AF regenerated vascular plexus under different experimental setups. No YFP/mCerulean expression was detected in the uninduced controls during the regeneration process (nheat-shock only = 29; n4-OHT only = 32). g, Regenerated AF in adult kdrl:creERT2;flibow fish at 20dpa, shows distinctly labelled BVC (arrowheads) at the anterior trunk-fin junction of the AF. Scale bars, 20 µm (c), 100 µm (a,a″), 300 µm (g), 500 µm (d,e). dpa, days post amputation.

Supplementary information

Supplementary Information

Supplementary Methods and Supplementary References.

Reporting Summary

Supplementary Table 1

Description of the parameters used for staging the different phases of AF.

Supplementary Table 2

List of hsp70l:creERT2;flibow animals traced for lineage analyses. Information of the stages that were imaged and EC subsets that combinations of YFP/mCerulean are provided.

Supplementary Table 3

Full list of DEGs for each of the clusters. Average log2-transformed fold change (avg_log2FC) and adjusted P value (p_val_adj) for each DEGs is listed. Five separate worksheet tabs are provided, each presenting the DEGs for a particular cluster.

Supplementary Table 4

Sequence of the smFISH probe library used for detection of prox1a and fli1a RNA.

Supplementary Video 1

Thoracic-duct-derived lymphatic sprouts colonize the developing AF. 3D volume rendered image depicting the mrc1a:EGFP+ and prox1a:RFP- PCV (green) and the mrc1a:EGFP,prox1a:RFP double-positive lymphatic vessels (yellow) that penetrate the stage I AF. Scale bar, 50 μm.

Supplementary Video 2

Erythrocyte content in BVC and LVC. Time-lapse video showing erythrocytes (gata1a:dsRed) and mrc1a:EGFP+ vessels at stage I (a) and stage II (b). The yellow arrowheads indicate the LVC sprouts/plexus and the white arrows indicate the erythrocyte flow occurring within the BVC (not labelled). Asterisks indicate gata1a+ stationary cells that are not erythrocytes. The video is being played at 10 fps. cl, cloaca; PCV, posterior cardinal vein.

Supplementary Video 3

Erythrocyte flow in stage IV AFs of WT and flt4−/− animals. Time-lapse bright-light video depicting erythrocyte flow along a fin ray in a WT (left) and flt4−/− (right) stage IV AF. The yellow arrows indicate the vessels. Images are shown for a 5 min recording, with a frame interval of 0.152 s and the video is played at 50 fps.

Supplementary Video 4

Erythrocyte flow in AFs of WT and flt4−/− adult animals Time-lapse bright-light video depicting erythrocyte flow along a fin ray in a WT (left) and flt4−/− (right) adult fish AF. The yellow arrows indicate the vessels. Images are shown for a 5 min recording, with a frame interval of 0.152 s and the video is played at 50 fps.

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Das, R.N., Tevet, Y., Safriel, S. et al. Generation of specialized blood vessels via lymphatic transdifferentiation. Nature 606, 570–575 (2022).

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