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<record><TEI><teiHeader><fileDesc><titleStmt><title xml:lang="en">Smooth muscle–endothelial cell communication activates Reelin signaling and regulates lymphatic vessel formation</title><author><name sortKey="Lutter, Sophie" sort="Lutter, Sophie" uniqKey="Lutter S" first="Sophie" last="Lutter">Sophie Lutter</name></author><author><name sortKey="Xie, Sherry" sort="Xie, Sherry" uniqKey="Xie S" first="Sherry" last="Xie">Sherry Xie</name></author><author><name sortKey="Tatin, Florence" sort="Tatin, Florence" uniqKey="Tatin F" first="Florence" last="Tatin">Florence Tatin</name></author><author><name sortKey="Makinen, Taija" sort="Makinen, Taija" uniqKey="Makinen T" first="Taija" last="Makinen">Taija Makinen</name></author></titleStmt><publicationStmt><idno type="wicri:source">PMC</idno><idno type="pmid">22665518</idno><idno type="pmc">3373399</idno><idno type="url">http://www.ncbi.nlm.nih.gov/pmc/articles/PMC3373399</idno><idno type="RBID">PMC:3373399</idno><idno type="doi">10.1083/jcb.201110132</idno><date when="2012">2012</date><idno type="wicri:Area/Pmc/Corpus">004042</idno><idno type="wicri:explorRef" wicri:stream="Pmc" wicri:step="Corpus" wicri:corpus="PMC">004042</idno></publicationStmt><sourceDesc><biblStruct><analytic><title xml:lang="en" level="a" type="main">Smooth muscle–endothelial cell communication activates Reelin signaling and regulates lymphatic vessel formation</title><author><name sortKey="Lutter, Sophie" sort="Lutter, Sophie" uniqKey="Lutter S" first="Sophie" last="Lutter">Sophie Lutter</name></author><author><name sortKey="Xie, Sherry" sort="Xie, Sherry" uniqKey="Xie S" first="Sherry" last="Xie">Sherry Xie</name></author><author><name sortKey="Tatin, Florence" sort="Tatin, Florence" uniqKey="Tatin F" first="Florence" last="Tatin">Florence Tatin</name></author><author><name sortKey="Makinen, Taija" sort="Makinen, Taija" uniqKey="Makinen T" first="Taija" last="Makinen">Taija Makinen</name></author></analytic><series><title level="j">The Journal of Cell Biology</title><idno type="ISSN">0021-9525</idno><idno type="eISSN">1540-8140</idno><imprint><date when="2012">2012</date></imprint></series></biblStruct></sourceDesc></fileDesc><profileDesc><textClass></textClass></profileDesc></teiHeader><front><div type="abstract" xml:lang="en"><p>Reelin signaling is activated by communication between the two cell types of the collecting lymphatic vessels and promotes smooth muscle cell recruitment, which is necessary for lymphatic vessel morphogenesis and function.</p></div></front><back><div1 type="bibliography"><listBibl><biblStruct><analytic><author><name sortKey="Aplin, A C" uniqKey="Aplin A">A.C. 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<front><journal-meta><journal-id journal-id-type="nlm-ta">J Cell Biol</journal-id><journal-id journal-id-type="iso-abbrev">J. Cell Biol</journal-id><journal-id journal-id-type="hwp">jcb</journal-id><journal-title-group><journal-title>The Journal of Cell Biology</journal-title></journal-title-group><issn pub-type="ppub">0021-9525</issn><issn pub-type="epub">1540-8140</issn><publisher><publisher-name>The Rockefeller University Press</publisher-name></publisher></journal-meta><article-meta><article-id pub-id-type="pmid">22665518</article-id><article-id pub-id-type="pmc">3373399</article-id><article-id pub-id-type="publisher-id">201110132</article-id><article-id pub-id-type="doi">10.1083/jcb.201110132</article-id><article-categories><subj-group subj-group-type="heading"><subject>Research Articles</subject><subj-group><subject>Article</subject></subj-group></subj-group></article-categories><title-group><article-title>Smooth muscle–endothelial cell communication activates Reelin signaling and regulates lymphatic vessel formation</article-title><alt-title alt-title-type="short">Reelin in lymphatic development</alt-title></title-group><contrib-group><contrib contrib-type="author"><name><surname>Lutter</surname><given-names>Sophie</given-names></name></contrib><contrib contrib-type="author"><name><surname>Xie</surname><given-names>Sherry</given-names></name></contrib><contrib contrib-type="author"><name><surname>Tatin</surname><given-names>Florence</given-names></name></contrib><contrib contrib-type="author" corresp="yes"><name><surname>Makinen</surname><given-names>Taija</given-names></name></contrib></contrib-group><aff><institution>Lymphatic Development Laboratory, Cancer Research UK London Research Institute, London WC2A 3LY, England, UK</institution></aff><author-notes><corresp>Correspondence to Taija Makinen: <email>taija.makinen@cancer.org.uk</email></corresp></author-notes><pub-date pub-type="ppub"><day>11</day><month>6</month><year>2012</year></pub-date><volume>197</volume><issue>6</issue><fpage>837</fpage><lpage>849</lpage><history><date date-type="received"><day>31</day><month>10</month><year>2011</year></date><date date-type="accepted"><day>2</day><month>5</month><year>2012</year></date></history><permissions><copyright-statement>© 2012 Lutter et al.</copyright-statement><copyright-year>2012</copyright-year><license license-type="openaccess"><license-p>This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see <ext-link ext-link-type="uri" xlink:href="http://www.rupress.org/terms">http://www.rupress.org/terms</ext-link>). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at <ext-link ext-link-type="uri" xlink:href="http://creativecommons.org/licenses/by-nc-sa/3.0/">http://creativecommons.org/licenses/by-nc-sa/3.0/</ext-link>).</license-p></license></permissions><self-uri xlink:role="icon" xlink:type="simple" xlink:href="JCB_201110132_thumb.gif"></self-uri><abstract abstract-type="precis"><p>Reelin signaling is activated by communication between the two cell types of the collecting lymphatic vessels and promotes smooth muscle cell recruitment, which is necessary for lymphatic vessel morphogenesis and function.</p></abstract><abstract><p>Active lymph transport relies on smooth muscle cell (SMC) contractions around collecting lymphatic vessels, yet regulation of lymphatic vessel wall assembly and lymphatic pumping are poorly understood. Here, we identify Reelin, an extracellular matrix glycoprotein previously implicated in central nervous system development, as an important regulator of lymphatic vascular development. Reelin-deficient mice showed abnormal collecting lymphatic vessels, characterized by a reduced number of SMCs, abnormal expression of lymphatic capillary marker lymphatic vessel endothelial hyaluronan receptor 1 (LYVE-1), and impaired function. Furthermore, we show that SMC recruitment to lymphatic vessels stimulated release and proteolytic processing of endothelium-derived Reelin. Lymphatic endothelial cells in turn responded to Reelin by up-regulating monocyte chemotactic protein 1 (MCP1) expression, which suggests an autocrine mechanism for Reelin-mediated control of endothelial factor expression upstream of SMC recruitment. These results uncover a mechanism by which Reelin signaling is activated by communication between the two cell types of the collecting lymphatic vessels—smooth muscle and endothelial cells—and highlight a hitherto unrecognized and important function for SMCs in lymphatic vessel morphogenesis and function.</p></abstract></article-meta></front><body><sec><title>Introduction</title><p>The lymphatic vasculature initially develops as a primitive plexus of identical vessels, all expressing the hyaluronan receptor lymphatic vessel endothelial hyaluronan receptor 1 (LYVE-1). This plexus is extensively remodeled to give rise to the functional lymphatic vasculature, which consists of two distinct vessel types: lymphatic capillaries (also called initial lymphatics) and collecting lymphatic vessels (<xref ref-type="bibr" rid="bib17">Jurisic and Detmar, 2009</xref>; <xref ref-type="bibr" rid="bib34">Tammela and Alitalo, 2010</xref>). Only blind-ended lymphatic capillaries retain expression of LYVE-1, and their distinctive morphology is ideally suited to their function; loose, overlapping cell–cell junctions, discontinuous basement membrane (BM), and wide lumens allow efficient absorption of protein-rich fluid from the interstitium. The fluid is further drained into the collecting lymphatic vessels, which develop luminal valves and a BM, and recruit smooth muscle cells (SMC), to assist in the unidirectional propulsion of lymph (<xref ref-type="bibr" rid="bib17">Jurisic and Detmar, 2009</xref>; <xref ref-type="bibr" rid="bib34">Tammela and Alitalo, 2010</xref>).</p><p>In the blood vessels, the vascular BM is an important structural component, which is formed predominantly of Collagen IV, laminin, nidogen/entactin, and heparan sulfate proteoglycans (<xref ref-type="bibr" rid="bib31">Sasaki et al., 2004</xref>). It helps to keep the vessel wall structure in shape and withstand mechanical forces generated by pulsatile blood flow and vasoconstriction, which regulates blood flow and pressure. In addition to its structural importance, more recent studies have revealed a complex functionality of the vascular BM, including involvement in endothelial cell (EC) migration, proliferation, survival, signal transduction, and vessel morphogenesis (<xref ref-type="bibr" rid="bib2">Aszódi et al., 2006</xref>). In addition, because the vascular BM is produced by and shared between ECs and SMCs, it directly interacts with both cell types to modulate their behavior and communication. For example, endothelial-derived ECM can stimulate proliferation of SMC in vitro (<xref ref-type="bibr" rid="bib6">Figueroa et al., 2004</xref>). Similarly, Heparan-sulfate (HS), deposited by ECs, can support mural cell attachment on the vessels in vivo; however, the mural cells require cell-autonomous HS to efficiently polarize and migrate to the nascent blood vessels (<xref ref-type="bibr" rid="bib33">Stenzel et al., 2009</xref>). ECM molecules can also participate in storing, masking, presenting, or sequestering growth factors, thereby modulating their bioavailability and activity (<xref ref-type="bibr" rid="bib14">Hynes, 2009</xref>). Furthermore, during development, the vascular BMs undergo compositional and structural alterations, including conformational changes and proteolytic processing of specific matrix components, which can provide important angiogenic or anti-angiogenic cues (for review see <xref ref-type="bibr" rid="bib18">Kalluri, 2003</xref>).</p><p>In contrast to the vascular BM, little is known of the composition and importance of the ECM of lymphatic vessels. Here, we identify Reelin as a lymphatic-specific matrix molecule and demonstrate its important function in the formation of functional collecting lymphatic vessels. In addition, we uncover a unique mechanism by which Reelin signaling is activated via communication between the two cell types that form the collecting vessels: endothelial and SMCs. The specific defects displayed by the <italic>Reln</italic>-deficient mice further highlight a hitherto unrecognized important function of SMCs in lymphatic vessel morphogenesis and function.</p></sec><sec sec-type="results"><title>Results</title><sec><title>Characterization of collecting vessel formation and lymphatic vascular wall assembly</title><p>To establish a timeline and a defined set of markers of lymphatic collecting vessel differentiation, we investigated SMC recruitment to dermal lymphatic vessels using antibodies against α-smooth muscle actin (α-SMA), desmin, and NG2, the markers of SMCs and pericytes; and against LYVE-1, a marker of immature lymphatic vessels and lymphatic capillaries. The developing vasculature was analyzed in mouse ear skin, which allows whole-mount visualization of vessels that grow postnatally into the ear to form a primary vascular plexus, which is subsequently remodeled into a functional vessel network consisting of lymphatic capillaries and collecting vessels (<xref ref-type="bibr" rid="bib25">Mäkinen et al., 2005</xref>). During the first postnatal week, the expanding dermal lymphatic vasculature of the ear was devoid of SMCs and the endothelia was uniformly positive for LYVE-1 (unpublished data). At postnatal day 12 (P12), LYVE-1 was down-regulated in specific areas (<xref ref-type="fig" rid="fig1">Fig. 1 A</xref>). Although the first α-SMA–positive SMCs could be found loosely attached to the lymphatic vessels at this stage, LYVE-1 down-regulation did not correlate with their presence (<xref ref-type="fig" rid="fig1">Fig. 1 A</xref> and not depicted). At P14, developing collecting vessels acquired SMCs, which were found predominantly in areas of low LYVE-1 expression (<xref ref-type="fig" rid="fig1">Fig. 1 B</xref>). By P16, lymphatic collecting vessels were identified by their prominent SMC coverage and absent LYVE-1 expression (<xref ref-type="fig" rid="fig1">Fig. 1 C</xref>). NG2, a marker of vascular pericytes, was not expressed in lymphatic SMCs (<xref ref-type="fig" rid="fig1">Fig. 1 D</xref>), whereas desmin-positive SMCs were found only in lymphatic vessels that had acquired features of mature collecting vessels, such as prominent SMC coverage, luminal valves, and complete down-regulation of LYVE-1 expression (<xref ref-type="fig" rid="fig1">Fig. 1, E–G</xref>). Desmin and α-SMA were coexpressed in lymphatic SMCs (<xref ref-type="fig" rid="fig1">Fig. 1 H</xref>).</p><fig id="fig1" position="float"><label>Figure 1.</label><caption><p><bold>SMC recruitment and ECM deposition during collecting vessel formation.</bold> (A–H) Immunofluorescence staining of wild-type ear skin at the indicated stages (P12, P14, P16, or P21) with antibodies against LYVE-1 (green, A–C and E–G), α-SMA (red, A–D and H), desmin (red, E–G; green, H), PECAM-1 (blue, A–C and E–G), and Podoplanin (blue, D). Arrows in A indicate areas of LYVE-1 down-regulation before SMC recruitment. The arrow in D indicates an NG2-positive blood vessel, and the arrowhead in D indicates an NG2-negative collecting vessel. P, postnatal day. (I–L) Immunofluorescence staining of wild-type ear skin at P14 (I, I′, K, and K′) and P16 (J, J′, L, and L′) with antibodies against Collagen IV (green I–J′), Laminin-α5 (green K–L′), α-SMA (red), and LYVE-1 (blue). Arrows show Laminin-α5 expression in luminal valves and arrowheads indicate expression of Laminin-α5 by SMC. The broken line in K outlines the LYVE-1<sup>−</sup> collecting vessel. The asterisks in K and L show LYVE-1<sup>+</sup> lymphatic capillaries. Bars, 50 µm. (M) Schematic timeline of dermal collecting lymphatic vessel differentiation.</p></caption><graphic xlink:href="JCB_201110132_Fig1"></graphic></fig><p>Another characteristic feature of collecting lymphatic vessels is the presence of vascular BM. To investigate if matrix deposition can serve as a marker of collecting vessel differentiation, we examined the expression of Collagen IV and Laminin-α5 in the developing vessels. At P14, Collagen IV was evident in LYVE-1–positive vessels that had no SMCs (<xref ref-type="fig" rid="fig1">Fig. 1, I and I′</xref>), which suggests its deposition by ECs. At later stages, Collagen IV was found predominantly around collecting vessels (<xref ref-type="fig" rid="fig1">Fig. 1, J and J′</xref>), with lower levels in capillaries (not depicted; <xref ref-type="bibr" rid="bib27">Pflicke and Sixt, 2009</xref>). Conversely, Laminin-α5 expression was not detectable in prospective collecting lymphatic vessels before SMC recruitment, except in luminal valves (<xref ref-type="fig" rid="fig1">Fig. 1, K and K′</xref>; <xref ref-type="bibr" rid="bib3">Bazigou et al., 2009</xref>). In more matured collecting vessels, Laminin-α5 staining highlighted the SMCs around collecting vessels (<xref ref-type="fig" rid="fig1">Fig. 1, L and L′</xref>), which suggests that these cells are its main source. However, in the blood vasculature, Laminin-α5 is produced by ECs (<xref ref-type="bibr" rid="bib42">Wu et al., 2009</xref>), and additional expression by lymphatic endothelium that may be induced upon SMC contact cannot be excluded. In summary, these results suggest that both endothelial and SMCs contribute to the production of matrix components of the lymphatic BM. In addition, deposition of ECM may provide the earliest sign of collecting vessel specification (<xref ref-type="fig" rid="fig1">Fig. 1 M</xref>).</p></sec><sec><title>Identification of Reelin as a lymphatic endothelial-specific matrix molecule</title><p>To further characterize lymphatic vessel wall assembly, we attempted to identify new specific components of the lymphatic BM. We dissected mesenteric collecting lymphatic vessels, arteries, and veins of P7 mice and identified differentially expressed genes using Affymetrix expression arrays. The reliability of the dataset was demonstrated by the identification of several known lymphatic, arterial, and venous markers that were up- or down-regulated in the appropriate datasets (<ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp1">Table S1</ext-link>). 14 genes involved in ECM formation, composition, or degradation were up-regulated more than twofold in the lymphatic vessels in comparison to the blood vessels (<ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp2">Tables S2</ext-link> and <ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp3">S3</ext-link>). Preferential expression of Efemp1, Spock3, and Nid2 in lymphatic in comparison to blood vessels was further confirmed by immunofluorescence of mouse skin or Western blot analysis of primary ECs (Fig. S1, A–D). Among the identified genes, we focused on Reelin that was found as being expressed almost fivefold higher in the lymphatic vessels compared with the veins (Table S2). Previously, specific Reelin immunoreactivity was reported in the developing lymphatic vessels in rat and human (<xref ref-type="bibr" rid="bib30">Samama and Boehm, 2005</xref>). We further confirmed lymphatic specific expression of Reelin in vivo in murine embryonic (<xref ref-type="fig" rid="fig2">Fig. 2, A and A′</xref>) and postnatal (<xref ref-type="fig" rid="fig2">Fig. 2, B–E′</xref>; and Fig. S1, E–F′) lymphatic vasculature of the skin and mesentery. The apparent expression in arteries that was detected in microarray profiling (Table S2) was likely caused by contaminating nerve tissue expressing Reelin (Fig. S1, E–E‴).</p><fig id="fig2" position="float"><label>Figure 2.</label><caption><p><bold>Reelin is a lymphatic endothelial-specific matrix molecule that shows differential distribution in lymphatic capillaries and collecting vessels.</bold> (A and A′) Immunofluorescence of E16 dorsal skin for Reelin (green), LYVE-1 (red), and PECAM-1 (blue). Arrowheads and arrows indicates Reelin-positive and -negative lymphatic and blood vessels, respectively. (B–G′) Immunofluorescence staining of P6 mesentery (B–C′) and P21 ear-skin (D–G′), with antibodies against Reelin (green) and α-SMA (red). Blue staining (in B) is for PECAM-1 and (in D–E′) for LYVE-1. Boxed areas in D and E are magnified in D′ and E′, showing Reelin expression only. Vessel borders are highlighted by broken lines in B′, C′, D′, E′, and F′). Tissue was unpermeabilized in F–G′ to detect extracellular proteins, and the fluorescent signal was amplified using TSA technology. The arrowhead in F indicates weak Reelin immunoreactivity outside capillary LECs. Bars: (A, D, E, F and G) 50 µm; (B and C) 500 µm; (D’ and E’) 20 µm.</p></caption><graphic xlink:href="JCB_201110132_Fig2"></graphic></fig></sec><sec><title>Differential localization of Reelin in mature lymphatic capillaries and collecting vessels</title><p>Immunofluorescence for Reelin in the postnatal mesenteric and dermal lymphatic vasculature revealed intense perinuclear and punctuated staining in the blind-ended lymphatic capillaries (<xref ref-type="fig" rid="fig2">Fig. 2, B, B′, D, and D′</xref>). In contrast, the collecting vessels showed a weaker, diffuse Reelin immunoreactivity (<xref ref-type="fig" rid="fig2">Fig. 2, C, C′, E, and E′</xref>). To test if the differential protein distribution was related to different subcellular localization of Reelin, we used nonpermeabilized tissue, together with tyramide signal amplification (TSA) procedure, to allow specific staining of extracellular proteins. Under these conditions, low Reelin immunoreactivity was detected in the lymphatic capillaries (<xref ref-type="fig" rid="fig2">Fig. 2, F and F′</xref>), whereas intense, patchy labeling of collecting lymphatic vessels was observed (<xref ref-type="fig" rid="fig2">Fig. 2, G and G′</xref>). Particularly strong immunoreactivity for extracellular Reelin (<xref ref-type="fig" rid="fig2">Fig. 2, G and G′</xref>), but not for total Reelin protein (Fig. S1, E and E′), was observed around lymphatic valves, which suggests that Reelin is preferentially deposited in the valve regions, possibly by binding to specific matrix components that are enriched in these areas. Co-staining with cell compartment–specific markers suggested that intracellular Reelin in lymphatic capillaries resided predominantly in the endoplasmic reticulum (Fig. S1, G and H), similar to what has been reported in neurons (<xref ref-type="bibr" rid="bib35">Tinnes et al., 2011</xref>). Together, these data suggest that Reelin is secreted by the lymphatic ECs (LECs) into the ECM of the collecting vessels, but remains intracellular in the endothelia of lymphatic capillaries.</p></sec><sec><title>SMC contact activates Reelin signaling by stimulating its secretion from lymphatic endothelia and proteolytic processing</title><p>We conjectured that the secretion of Reelin into the ECM could be related to SMC recruitment in the collecting vessels. To test this, we analyzed Reelin expression and secretion by the LECs that were either cultured alone or together with SMCs. Lymphatic endothelial-specific expression of Reelin was maintained in vitro, as assessed by quantitative PCR (qPCR) analyses of RNA extracted from primary human LECs, blood vascular ECs (BECs), and SMCs (<xref ref-type="fig" rid="fig3">Fig. 3 A</xref>). In addition, immunoprecipitation and Western blot analysis using an antibody that recognizes an epitope at the N terminus of Reelin (<xref ref-type="fig" rid="fig3">Fig. 3 B</xref>) confirmed that only LECs produced Reelin protein (<xref ref-type="fig" rid="fig3">Fig. 3 C</xref>). Analysis of the medium from LECs revealed secretion of full-length 450-kD Reelin polypeptide that was partially processed by cleavage within the C terminus (<xref ref-type="fig" rid="fig3">Fig. 3 B</xref>; <xref ref-type="bibr" rid="bib16">Jossin et al., 2007</xref>) to give rise to a smaller fragment of 370 kD (<xref ref-type="fig" rid="fig3">Fig. 3 D</xref>). When LECs were co-cultured with SMCs, secretion of full-length Reelin into the medium was increased, with a corresponding increase in the production of the 370-kD fragment, whereas the production of Reelin was not affected (<xref ref-type="fig" rid="fig3">Fig. 3, D and E</xref>). In addition, a second processing event in the N terminus resulted in the formation of a 180-kD fragment (<xref ref-type="fig" rid="fig3">Fig. 3, B and 3D</xref>) and a central Reelin fragment, which is capable of binding and activation of the canonical Reelin receptors ApoER2 and VLDLR (<xref ref-type="bibr" rid="bib16">Jossin et al., 2007</xref>). Incubation of supernatant from LECs with SMCs confirmed that SMCs can mediate the second N-terminal processing event (<xref ref-type="fig" rid="fig3">Fig. 3 F</xref>). No increase in Reelin secretion or processing was observed when LECs were cultured in the conditioned medium collected from the SMCs, or on extracellular matrices produced by fibroblasts (unpublished data), which suggests that the effect was not mediated by an SMC-derived secreted protein. Together, these results demonstrate that SMC contact stimulates the secretion of Reelin from the LECs and its proteolytic processing, which suggests that SMC recruitment to the collecting lymphatic vessels activates Reelin signaling.</p><fig id="fig3" position="float"><label>Figure 3.</label><caption><p><bold>Reelin secretion and processing is stimulated by SMC contact.</bold> (A) qPCR analysis showing relative <italic>RELN</italic> expression in three independent samples of LEC, BEC, and HUVSMC and two samples of HAoSMC. Mean ± SEM is plotted (error bars). (B) Schematic of Reelin structure, showing antibody binding sites (asterisks), cleavage sites, and products. The arrows and arrowheads indicate polypeptides that are visualized in Western blotting: arrow, full-length protein; arrowhead, C-terminally cleaved polypeptide; open arrowhead, N- and C-terminally cleaved polypeptide. Adapted from <xref ref-type="bibr" rid="bib16">Jossin et al. (2007)</xref>. (C and D) Analysis of whole-cell lysate (C) and medium (D) of LEC and SMC cultured either alone or together by immunoprecipitation and Western blotting with antibodies to Reelin. The arrows indicates full-length Reelin, arrowheads indicate cleavage products. Analysis of whole cell lysate in C by Western blotting with antibodies to CD31/PECAM-1 (for LEC) and α-SMA (for SMC) show that equivalent numbers of cells were used. (E) Quantification of Reelin protein levels, normalized to CD31/PECAM-1, in whole cell lysate (black bars) and culture medium (gray bars). Data represent mean (<italic>n</italic> = 3) ± SEM (error bars). (F) Analysis of LEC-conditioned medium alone or after incubation with SMC. Note the cleavage product of 180 kD only in the presence of SMC (open arrowhead).</p></caption><graphic xlink:href="JCB_201110132_Fig3"></graphic></fig></sec><sec><title>Defective collecting lymphatic vessel formation in Reelin-deficient mice</title><p>To investigate the physiological function of Reelin in lymphatic vasculature, we analyzed the dermal lymphatic vessels in Reelin (encoded by <italic>Reln</italic>)-deficient <italic>Reeler</italic> mice (<xref ref-type="bibr" rid="bib5">D’Arcangelo et al., 1995</xref>). The mature dermal lymphatic vasculature is composed of three types of vessels: collecting vessels, precollectors, and capillaries. Precollectors share features of both capillaries and collecting vessels; in most cases they are LYVE-1–positive vessels that contain luminal valves but no SMCs. They often abruptly branch off the main LYVE-1–negative collecting vessel (<xref ref-type="fig" rid="fig4">Fig. 4 A</xref>, see also <xref ref-type="fig" rid="fig4">Fig. 4 F</xref>). However, in some cases, transition from one type of vessel to another is more gradual; individual α-SMA–positive cells can be found attached on vessels that contain some LYVE-1<sup>+</sup> ECs (unpublished data). Notably, however, SMCs are rarely found directly attached to LYVE-1<sup>+</sup> LECs, or on uniformly LYVE-1<sup>+</sup> vessels. Because the definition of precollecting vessel is ambiguous, in this study we considered only two vessel categories that we defined as collecting vessels based on the presence of α-SMA<sup>+</sup> SMCs and as capillaries based on positive LYVE-1 immunoreactivity.</p><fig id="fig4" position="float"><label>Figure 4.</label><caption><p><bold>Reduced SMC coverage and defective collecting lymphatic vessel differentiation in <italic>Reln</italic>-deficient mice.</bold> (A, B, and D–I) Immunofluorescence of ear skin from wild-type (A, D, F, and H) and <italic>Reln<sup>−/−</sup></italic> (B, E, G, and I) mice for the indicated proteins. For A and B, individual images (projections of confocal z stacks) were aligned to trace the entire collecting vessel from the proximal (Q1) to distal (Q4) end, and the overlapping blood vasculature was blacked out for clarity (original images are shown in <ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp4">Fig. S2</ext-link>). Broken lines outline the collecting vessels. (C) LYVE-1 immunoreactivity in wild-type and <italic>Reln<sup>−/−</sup></italic> vessels (mean ± SEM [error bars]; <italic>n</italic> = 6 vessels). (J) SMC coverage in vessel quarters (mean ± SEM; <italic>n</italic> ≥ 12 vessels). (K) Quantification of widest and narrowest vessel points and the difference between the two (<italic>n</italic> ≥ 12 vessels). The boxes represent interquartile range, lines indicate mean, and whiskers indicate minimum and maximum values. *, P < 0.05; **, P < 0.01; ***, P < 0.005. Bars: (A and B) 200 µm; (D–I) 50 µm.</p></caption><graphic xlink:href="JCB_201110132_Fig4"></graphic></fig><p>We observed an apparently normal LYVE-1–positive dermal lymphatic capillary network in the ears of the <italic>Reln</italic>-deficient mice (<ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp5">Fig. S2, A and B</ext-link>). To allow unambiguous distinction of collecting vessels, we traced the entire vessel from the proximal to the distal end of the ear. In comparison to LYVE-1–negative collecting vessels in wild-type mice, <italic>Reln</italic> mutant vessels maintained an abnormally high level of LYVE-1 (<xref ref-type="fig" rid="fig4">Fig. 4, A–C</xref>; and Fig. S2, E and F). In addition, <italic>Reln</italic> mutants showed reduced SMC coverage when compared with wild-type vessels (<xref ref-type="fig" rid="fig4">Fig. 4, D–G and J</xref>). Although a similar proximal-to-distal decrease was observed, in each vessel quarter the SMC coverage was significantly lower in <italic>Reln</italic> mutant compared with the wild type (<xref ref-type="fig" rid="fig4">Fig. 4 J</xref>). The early differentiation of dermal collecting vessels appeared normal in <italic>Reln</italic> mutants, as indicated by deposition of Collagen IV and formation of luminal valves (Fig. S2, G–H′; and <xref ref-type="fig" rid="fig4">Fig. 4, H and I</xref>); however, deposition of SMC-derived Laminin-α5 was strongly decreased (Fig. S2, I–J′). In addition, <italic>Reln</italic> mutants displayed irregular lymphatic vessel diameter, with abnormal constrictions and swellings (<xref ref-type="fig" rid="fig4">Fig. 4, B and K</xref>). These defects were not secondary to nervous system defects because normal patterning of the nerves was found in <italic>Reln</italic> mutant ears (Fig. S2, C and D). In contrast to the dermal collecting lymphatic vessels, the large collecting vessels of the mesentery appeared normal in <italic>Reln</italic>-deficient mice, as indicated by normal morphology and SMC coverage (<ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp6">Fig. S3</ext-link>). In summary, these data show that <italic>Reln</italic> deficiency leads to specific defects in dermal collecting lymphatic vessel formation.</p></sec><sec><title>Impaired lymphatic vessel function in Reelin mutant mice</title><p>To assess if the observed morphological defects affect lymphatic function in <italic>Reln</italic> mutants, we investigated the uptake and transport of large-molecular-weight fluorescent dextran–injected intradermally in the ear. In wild-type mice, the dye was quickly taken up into the vessels surrounding the injection site (<xref ref-type="fig" rid="fig5">Fig. 5 A</xref>). However, except for in the immediate vicinity of the injection site, dye was rarely seen in LYVE-1–positive capillaries, whereas the collecting vessels were strongly positive (<xref ref-type="fig" rid="fig5">Fig. 5 B</xref>). In contrast, in <italic>Reln<sup>−/−</sup></italic> mice we frequently observed no dye in vessels near the injection site (<xref ref-type="fig" rid="fig5">Fig. 5 C</xref>), which suggests impaired lymphatic function. When dye was taken up, it was retained in LYVE-1–positive vessels (<xref ref-type="fig" rid="fig5">Fig. 5 D</xref>). We also observed dye at the vessel edges in the mutant ear (<xref ref-type="fig" rid="fig5">Fig. 5 D</xref>), which may be indicative of dye leakage from the vessel and retention within the BM, further suggesting functional deficiency in the collecting vessels of these mice. It is possible that discontinuous coverage of SMC around collecting lymphatic vessels may aid solute exchange and therefore increase collecting vessel “leakiness” (<xref ref-type="bibr" rid="bib32">Scallan and Huxley, 2010</xref>). To further assess the potential impairment of lymphatic function, we used the Evans blue method to allow quantitative measurement of dye content of the inguinal lymph nodes after hind limb footpad injection. Control mice showed significantly more dye in the lymph nodes when compared with the <italic>Reln<sup>−/−</sup></italic> mice (<xref ref-type="fig" rid="fig5">Fig. 5 E</xref>). In addition, we performed real-time imaging of the uptake of indocyanine green (ICG) dye by lymphatic vessels by measuring the increase in the fluorescent signal in the vessel over time after injection of the dye into the hind limb footpad. Abnormal dye uptake was observed in the <italic>Reln</italic> mutant mice compared with their control littermates (<xref ref-type="fig" rid="fig5">Fig. 5 F</xref>). All except one <italic>Reln</italic>-deficient mouse displayed reduction in the rate of lymphatic flow, whereas one showed an abnormally fast dye uptake. In control littermates, the dye reached the inguinal lymph nodes within seconds (<xref ref-type="fig" rid="fig5">Fig. 5 G</xref> and <ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp7">Fig. S4 A</ext-link>). In contrast, in most <italic>Reln</italic> mutants, no ICG dye was detectable in the lymph nodes at the end of 5 min of imaging (<xref ref-type="fig" rid="fig5">Fig. 5 G</xref> and Fig. S4 B). In spite of impaired lymphatic function, no apparent edema or ascites formation could be observed in the mutant mice (unpublished data).</p><fig id="fig5" position="float"><label>Figure 5.</label><caption><p><bold>Functional impairment in <italic>Reln<sup>−/−</sup></italic> collecting lymphatic vessels.</bold> (A–D) P21 wild-type (A and B) and <italic>Reln<sup>−/−</sup></italic> (C and D) ear skin, injected with FITC-dextran (green) and costained with antibodies against α-SMA (red; A and C) and LYVE-1 (blue, A and C; and red, B and D). The broken line in A and C indicates the injection site, and arrows point to adjacent lymphatic vessels. Arrows in B and D point to LYVE-1–positive capillaries, and the arrowhead in B indicates an LYVE-1–negative collecting vessel filled with FITC-dextran. The broken line in B outlines the collecting vessel and its connection to LYVE-1<sup>+</sup> capillaries (not outlined). (E) Quantification of Evans blue dye content in inguinal lymph nodes of control and <italic>Reln<sup>−/−</sup></italic> mice after 5 min of injection into the hind limb footpad. Values from individual lymph nodes and mean values are plotted (error bars; <italic>n</italic> = 11 [<italic>Ctrl</italic>] and 6 [<italic>Reln<sup>−/−</sup></italic>] mice). (F) Efficiency of fluorescent signal, representing efficiency of lymphatic flow, in a defined region of a vessel after ICG injection into the hind limb footpad over time. (G) Representative images of control and <italic>Reln<sup>−/−</sup></italic> hind limbs after ICG dye injections at three time points (excluding a 6–8-s delay before recording was started). In control but not in <italic>Reln<sup>−/−</sup></italic> mice, the dye reached the inguinal lymph node (asterisks). Bars: (A–D) 100 µm; (G) 0.5 cm.</p></caption><graphic xlink:href="JCB_201110132_Fig5"></graphic></fig></sec><sec><title>Reelin induces production of SMC recruitment factor MCP-1 in LECs via a noncanonical signaling pathway</title><p>In the nervous system, Reelin mediates its functions via two members of the low-density lipoprotein receptor family, VLDLR and ApoER2 (<xref ref-type="bibr" rid="bib37">Trommsdorff et al., 1999</xref>). Both receptors were expressed in cultured primary SMCs and LECs (<ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp8">Fig. S5 A</ext-link>). In addition, VLDLR was expressed in lymphatic endothelia in vivo (Fig. S5, B–C′). This suggests that either cell type—LEC or SMC—could therefore respond to Reelin to regulate SMC recruitment during collecting vessel formation. However, although LEC-conditioned medium promoted SMC migration in a transwell assay, this was not affected by a function-blocking antibody to Reelin (<xref ref-type="fig" rid="fig5">Fig. 5 E</xref> and S5 D). Furthermore, Reelin did not stimulate migration of SMCs (Fig. S5 D), which suggests that Reelin is not the LEC-derived factor responsible for SMC recruitment. We therefore tested a possibility that Reelin acts in an autocrine fashion in LECs to induce production of SMC recruitment factors. Indeed, we found a transient increase in the expression of <italic>MCP1/CCR2</italic>, a known regulator of SMC recruitment and migration (Fig. S5 D; <xref ref-type="bibr" rid="bib28">Religa et al., 2005</xref>; <xref ref-type="bibr" rid="bib23">Ma et al., 2008</xref>; <xref ref-type="bibr" rid="bib1">Aplin et al., 2010</xref>), in LECs upon Reelin stimulation (<xref ref-type="fig" rid="fig6">Fig. 6, A and B</xref>), whereas the expression of four other genes implicated in SMC recruitment was not affected (<xref ref-type="fig" rid="fig6">Fig. 6, A and B</xref>). Increased Angiopoietin-2 expression that was induced similarly by control and full-length Reelin containing 293 supernatant was likely caused by a high level of VEGF produced by these cells (<xref ref-type="bibr" rid="bib19">Kärpänen et al., 2006</xref>), which has previously been shown to stimulate Ang-2 expression in LECs (<xref ref-type="bibr" rid="bib39">Veikkola et al., 2003</xref>). Notably, <italic>MCP1</italic> expression showed only a modest increase upon stimulation with the central Reelin fragment (<xref ref-type="fig" rid="fig6">Fig. 6 A</xref>), which suggests that the canonical Reelin signaling pathway was not responsible for this effect. In agreement, analysis of <italic>Vldlr</italic> and <italic>Apoer2</italic> double mutant mice revealed normal lymphatic vasculature (<xref ref-type="fig" rid="fig5">Figs. 5 G</xref>; <xref ref-type="fig" rid="fig6">Fig. 6, C and D</xref>; and Fig. S5 F). In addition, mouse mutants lacking Dab1, an intracellular adaptor molecule activated in response to Reelin signaling, showed normal lymphatic vessels (<xref ref-type="fig" rid="fig6">Fig. 6 E</xref> and Fig. S5 H). All parameters measured were normal (<xref ref-type="fig" rid="fig6">Fig. 6, F–H</xref>), with the exception that the <italic>Vldlr<sup>−/−</sup></italic>;<italic>Apoer2<sup>−/−</sup></italic> vessels showed a modest decrease in SMC coverage in the distal-most quarter (<xref ref-type="fig" rid="fig6">Fig. 6 F</xref>). In addition, lymphatic function was not impaired in these mice (Fig. S5 I). In summary, these results suggest that Reelin acts in an autocrine fashion in the LECs to regulate SMC recruitment and collecting vessel differentiation (<xref ref-type="fig" rid="fig7">Fig. 7</xref>). Interestingly, unlike in the nervous system, the canonical signaling pathway is not required for Reelin signaling in lymphatic vessels, which suggests the involvement of alternative receptors.</p><fig id="fig6" position="float"><label>Figure 6.</label><caption><p><bold>LECs respond to Reelin via noncanonical signaling pathway.</bold> (A and B) Fold change in expression of SMC recruitment genes after Reelin stimulation. Note the strong increase in <italic>MCP1</italic> expression at 4 and 8 h in cells stimulated with full-length Reelin containing supernatant from HEK 293 cells only. The mean ± SEM of three experiments is plotted (error bars). *, P < 0.05; **, P < 0.001. (C–E) Immunofluorescence staining of P21 ear skin from wild-type (C), <italic>Apoer2<sup>−/−</sup>;Vldlr<sup>−/−</sup></italic> (D), and <italic>Dab1<sup>Scm</sup></italic> (E) mice using LYVE-1 (green), α-SMA (red), and Podoplanin (blue) antibodies. Individual images were aligned to trace the vessel from the proximal to the distal end, and overlapping blood vasculature was blacked out for clarity. A segment from the middle section of each vessel is displayed (original images are shown in <ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1" id="supp9">Fig. S5</ext-link>). (F) SMC coverage in vessel quarters (mean ± SEM [error bars]; <italic>n</italic> ≥ 6 vessels per genotype). *, P = 0.04. (G) Quantification of the widest and narrowest vessel points and the difference between the two. Boxes represents interquartile range, lines indicates mean, and whiskers indicate minimum and maximum values. (H) LYVE-1 immunoreactivity in collecting vessels (mean ± SEM [error bars]; <italic>n</italic> ≥ 6 vessels). All differences in F–H were nonsignificant, except where indicated. Bars, 200 µm.</p></caption><graphic xlink:href="JCB_201110132_Fig6"></graphic></fig><fig id="fig7" position="float"><label>Figure 7.</label><caption><p><bold>Model of Reelin signaling during collecting lymphatic vessel formation.</bold> Reelin (red stars) is produced by LEC (gray cells). Upon SMC (blue cell) contact, Reelin is secreted from LEC (red cells) and cleaved to release diffusible fragments (red gradient). Reelin can stimulate expression of <italic>MCP1</italic> in LECs to enhance further SMC recruitment.</p></caption><graphic xlink:href="JCB_201110132_Fig7"></graphic></fig></sec></sec><sec sec-type="discussion"><title>Discussion</title><p>Although proteins of the vascular BM are known to play diverse and essential roles in the maintenance of both the structural and biological function of the vessel, the composition and function of the lymphatic BM remains poorly characterized. Furthermore, differences in the BM surrounding different types of lymphatic vessels have been largely ignored, yet they are likely important because of the distinct functions of lymphatic capillaries and collecting vessels in mediating fluid absorbance and transport, respectively (<xref ref-type="bibr" rid="bib17">Jurisic and Detmar, 2009</xref>; <xref ref-type="bibr" rid="bib34">Tammela and Alitalo, 2010</xref>). In this study, we identify Reelin as the first lymphatic specific ECM protein and demonstrate its important function in the formation of functional collecting vessels.</p><p>Reelin is a large glycoprotein previously implicated in the development of the nervous system. Our analyses revealed that Reelin is additionally widely expressed in the lymphatic vasculature, both during development and in adult tissues. Although Reelin was produced by LEC in both the capillaries and collecting vessels, only the latter showed efficient secretion. Consistent with this, <italic>Reln</italic>-deficient mice had normal lymphatic capillaries but exhibited specific dermal collecting vessel defects, characterized by reduced SMC coverage and high expression of lymphatic capillary marker LYVE-1. Interestingly, SMCs were frequently found in direct contact with LYVE-1<sup>+</sup> LECs in the mutant but not in wild-type vessels. Notably, however, the largest collecting vessels, such as mesenteric lymphatic vessels that acquire more elaborate and continuous smooth muscle coverage, were apparently normal in <italic>Reln</italic> mutants, which suggests that there are different requirements for the formation of vessels of different calibers. We also noted a difference in the initiation of collecting vessel maturation in these two tissues. Although the developing dermal collecting vessels underwent a gradual increase in SMC coverage and concomitant decrease in LYVE-1 expression in a proximal-to-distal direction over a period of several days, the mesenteric lymphatic vessels rapidly acquired SMCs along their entire length, which was only later followed by down-regulation of LYVE-1 expression. These observations further suggest that the mechanisms regulating collecting vessel formation may differ depending on the caliber of the vessel and tissue environment.</p><p>Previous data on various mouse models lacking key components required for mural cell (MC) recruitment attest to their importance in the formation and maintenance of functional blood vessels (<xref ref-type="bibr" rid="bib12">Hellström et al., 2001</xref>). However, because of embryonic lethality, none of the MC-deficient mice described so far have allowed the investigation of the importance of lymphatic SMC in vivo. Our study describes for the first time a mouse model that shows a lymphatic-specific deficiency in SMCs, which occur in the absence of blood vascular or other major lymphatic defects. Concurrent with reduced SMC coverage, the <italic>Reln</italic>-deficient collecting vessels were enlarged and displayed irregular vessel diameters. Such defects are consistent with phenotypes described for pericyte-deficient blood vessels (<xref ref-type="bibr" rid="bib12">Hellström et al., 2001</xref>), which suggests that lymphatic SMC may have similar roles in the maintenance of vessel stability to those of the blood vascular system. However, important functional differences between lymphatic and blood vascular SMC exist. Although blood flow is generated by the beating heart and the vascular SMCs are involved in regulating vascular tone, the lymphatic vessels themselves must generate the force required for lymph propulsion (for review see <xref ref-type="bibr" rid="bib40">von der Weid and Zawieja, 2004</xref>). This action is dependent on lymphatic SMCs, which, unlike the vascular SMCs, are able to undergo both tonic and phasic contractions (<xref ref-type="bibr" rid="bib41">Wang et al., 2009</xref>). The impaired lymphatic function may imply that, in addition to the observed morphological abnormalities, <italic>Reln</italic> deficiency can directly affect SMC contractility and lymph pumping. These findings may have clinical significance, as congenital lymphedema has been observed in some human patients carrying mutations in the <italic>RELN</italic> gene (<xref ref-type="bibr" rid="bib13">Hong et al., 2000</xref>).</p><p>In spite of intense studies during the past decade, the exact cellular function of Reelin has remained unclear. In early corticogenesis, Reelin stimulates neuronal migration by attracting leading processes and inducing their branching and attachment to the ECM (<xref ref-type="bibr" rid="bib26">Ogawa et al., 1995</xref>; <xref ref-type="bibr" rid="bib36">Tissir and Goffinet, 2003</xref>; <xref ref-type="bibr" rid="bib7">Franco et al., 2011</xref>). In contrast, at later stages, when the migrating neurons reach their destination at the marginal zone of the cortex, Reelin acts as a stop signal by inducing neuronal detachment from radial glial scaffold and stabilization of the cytoskeleton (<xref ref-type="bibr" rid="bib11">Hartfuss et al., 2003</xref>; <xref ref-type="bibr" rid="bib4">Chai et al., 2009</xref>; <xref ref-type="bibr" rid="bib43">Zhao and Frotscher, 2010</xref>). These opposite functions of Reelin may be explained, at least in part, by binding and activation of different receptors: VLDLR and ApoER2 (<xref ref-type="bibr" rid="bib10">Hack et al., 2007</xref>). Proteolytic processing by matrix metalloproteinases may also play an important role in regulating diffusion and receptor-binding properties of Reelin (<xref ref-type="bibr" rid="bib22">Lambert de Rouvroit et al., 1999</xref>; <xref ref-type="bibr" rid="bib16">Jossin et al., 2007</xref>; <xref ref-type="bibr" rid="bib35">Tinnes et al., 2011</xref>). Although the central part of Reelin binds VLDLR and ApoER2, the N-terminal region mediates protein aggregation and has been suggested to regulate signaling by anchoring the protein to the ECM (<xref ref-type="bibr" rid="bib38">Utsunomiya-Tate et al., 2000</xref>; <xref ref-type="bibr" rid="bib21">Kubo et al., 2002</xref>; <xref ref-type="bibr" rid="bib43">Zhao and Frotscher, 2010</xref>). Because the cleavage products generated in LEC/SMC cultures were identical to those documented in the nervous system (<xref ref-type="bibr" rid="bib16">Jossin et al., 2007</xref>), the same mechanism of processing is likely. However, although functionally important (<xref ref-type="bibr" rid="bib16">Jossin et al., 2007</xref>), it is unclear how the processing is regulated in vivo and whether the individual cleavage products have specific, or even opposite functions. Interestingly, mechanisms where proteolytic processing produces protein fragments that exhibit opposing effects on angiogenesis to those of the parental molecule have been described for a subclass of vascular ECM proteins (for review see <xref ref-type="bibr" rid="bib18">Kalluri, 2003</xref>).</p><p>In the developing nervous system, Reelin is produced mainly by Cajal-Retzius neurons in the cortex, where it instructs newly born neurons to assume their position and orientation (<xref ref-type="bibr" rid="bib5">D’Arcangelo et al., 1995</xref>; <xref ref-type="bibr" rid="bib36">Tissir and Goffinet, 2003</xref>). During this process, the neurons are intimately apposed to radial glial cells that form a scaffold along which the neuroblasts migrate, and both cell types rely on Reelin signaling to direct correct patterning of the cortex (<xref ref-type="bibr" rid="bib11">Hartfuss et al., 2003</xref>; <xref ref-type="bibr" rid="bib36">Tissir and Goffinet, 2003</xref>; <xref ref-type="bibr" rid="bib20">Keilani and Sugaya, 2008</xref>; <xref ref-type="bibr" rid="bib43">Zhao and Frotscher, 2010</xref>). In the lymphatic system, we found that Reelin was produced exclusively by ECs, while its release and proteolytic processing was stimulated by the adjacent SMCs. This suggests a mechanism by which Reelin signaling is activated via communication of the two cell types that form the collecting vessels. We further found that LECs, in turn, respond to Reelin by up-regulating the expression of monocyte chemotactic protein 1 (MCP1), a factor known to promote SMC migration and proliferation (<xref ref-type="bibr" rid="bib28">Religa et al., 2005</xref>; <xref ref-type="bibr" rid="bib23">Ma et al., 2008</xref>; <xref ref-type="bibr" rid="bib1">Aplin et al., 2010</xref>). This suggests an autocrine function of Reelin in the LECs in controlling the expression of factors involved in SMC recruitment. Interestingly, although the canonical Reelin receptors VLDLR and ApoER2 were expressed in the lymphatic vasculature, <italic>Apoer2;Vldlr</italic> double mutant mice did not recapitulate the phenotype observed in <italic>Reln</italic>-deficient lymphatic vessels. Normal lymphatic development in <italic>Apoer2;Vldlr</italic> double mutant mice, which display an identical neurological phenotype compared with <italic>Reln</italic> mutants (<xref ref-type="bibr" rid="bib37">Trommsdorff et al., 1999</xref>), further indicates that the collecting vessel defects in <italic>Reln</italic>-deficient mice are not secondary to nervous system defects or the failure of these mice to gain weight, but instead suggests involvement of alternative Reelin receptors.</p><p>In summary, we have demonstrated an unexpected function of Reelin in regulating the formation of functional larger-caliber lymphatic vessels. Our results have uncovered a mechanism by which a specific signaling pathway is activated by communication between the two cell types of lymphatic vessels—EC and SMC—and demonstrate for the first time that SMCs play an important role in regulating lymphatic morphogenesis and function.</p></sec><sec sec-type="materials|methods"><title>Materials and methods</title><sec><title>Antibodies</title><p>The antibodies used for immunofluorescence were rat antibodies to mouse platelet EC adhesion molecule 1 (PECAM-1; MEC3.1, BD); rabbit antibodies to mouse Laminin-α5 (provided by L. Sorokin, University of Münster, Münster, Germany; <xref ref-type="bibr" rid="bib29">Ringelmann et al., 1999</xref>), Desmin (Abcam), Collagen IV (AbD Serotec), GFP (Invitrogen), LYVE-1 (ReliaTech), Erp57 (Abcam), and Golph4 (Abcam); rabbit antibody to human Efemp1 (Abcam); hamster antibodies to mouse PECAM-1 (Millipore) and Podoplanin (Developmental Studies Hybridoma Bank); goat antibodies to mouse Integrin-α9, Reelin, and VLDLR (all from R&D Systems); mouse antibody to mouse acetylated α-tubulin (Abcam; used in conjunction with mouse-on-mouse blocking reagents [Vector Laboratories]); and Cy3-conjugated mouse antibody against α-SMA (Sigma-Aldrich). Secondary antibodies conjugated to Cy2, Cy3, or Cy5 were obtained from Jackson ImmunoResearch Laboratories, Inc. Antibodies used for Western blot analysis were rabbit antibodies to human Efemp1 (Abcam), Nidogen2 (Immundiagnostik), and Spock3 (Abcam), and mouse antibody to human α-tubulin (Sigma-Aldrich).</p></sec><sec><title>Mice</title><p><italic>Reeler</italic> (<italic>Reln<sup>rl</sup></italic>) and <italic>Dab1<sup>scm</sup></italic> (<italic>A/A-Dab1<sup>scm</sup>/J</italic>) mice were obtained from The Jackson Laboratory. <italic>Apoer2<sup>−/−</sup>;Vldlr<sup>−/−</sup></italic> mice were obtained by crossbreeding of mice lacking <italic>Apoer2</italic> or <italic>Vldlr</italic>, which carry an insertion of a Neomycin cassette to replace exons 17 and 18, or into exon 5, leading to deletion of part of the coding sequence and disruption of the reading frame, respectively (<xref ref-type="bibr" rid="bib8">Frykman et al., 1995</xref>; <xref ref-type="bibr" rid="bib37">Trommsdorff et al., 1999</xref>). Both lines were provided by J. Herz (University of Texas Southwestern Medical Center, Austin, TX). <italic>Vegfr3<sup>lz</sup></italic> mice in which the first coding exon of <italic>Vegfr3</italic> gene was replaced by a lacZ cassette were provided by K. Alitalo (University of Helsinki, Helsinki, Finland). The mice were maintained on C57BL/6J background except for <italic>Dab1<sup>scm</sup></italic>, which were on C3HeB/FeJ* DC/Le background. All three mutant strains lacking components of the Reelin signaling pathway exhibited an ataxic gait and tremors starting around two weeks of age. In addition, homozygous <italic>Reeler</italic> and <italic>Apoer2<sup>−/−</sup>;Vldlr<sup>−/−</sup></italic> mutant mice, but not <italic>Dab1<sup>scm</sup></italic> mutants, were smaller in size, weighing ∼50–60% of the weight of the littermate controls at three weeks of age when most analyses were done. All mouse experiments were approved by the UK Home Office.</p></sec><sec><title>Microarray analysis</title><p>To identify lymphatic-specific ECM molecules, mesenteric arteries, veins, and lymphatic vessels were dissected from P7 wild-type (C57BL/6J) pups. Samples from nine animals (in two litters) were pooled. Total RNA was extracted and RNase-treated using an RNeasy Micro kit (QIAGEN) as per the manufacturer’s instructions. 100 ng of RNA from two independent samples of each vessel was amplified using WT-Ovation Pico RNA Amplification System (NuGEN) and used for expression profiling using the Mouse exon 1.0 ST Array (Affymetrix), according to the manufacturer’s instructions. The array data were processed using robust multiarray analysis (<xref ref-type="bibr" rid="bib15">Irizarry et al., 2003</xref>) to generate gene-level signal estimates for core level probe sets. The arrays were normalized using quantile normalization and antigenomic probes used to model the background. Quantification was run using apt-1.10.0 from Affymetrix. Before statistical analysis, all Affymetrix control probe sets were removed. Gene changes were selected using a linear model where the standard errors were corrected using empirical Bayes shrinkage (fdr < 0.05). To identify lymphatic-specific transcriptional changes, we applied a nestedF procedure across the three contrasts, selecting genes significant in only the lymphatic groups (fdr < 0.05). Analysis was performed in Bioconductor (<xref ref-type="bibr" rid="bib9">Gentleman et al., 2004</xref>) using Limma. Genes that were expressed at least twofold higher in different vessel types were searched in Uniprot database to identify proteins involved in ECM function or structure.</p></sec><sec><title>Immunostaining and X-Gal staining</title><p>For whole-mount immunostaining, tissue was fixed in 4% paraformaldehyde, permeabilized in 0.3% Triton X-100 in PBS, and blocked in PBST plus 3% milk. Primary antibodies were incubated at 4°C overnight in blocking buffer. After washing in PBSTx, the samples were incubated with fluorescence-conjugated secondary antibodies in blocking PBSTx plus 1% milk before further washing and mounting in Mowiol. Reelin staining in Figs. S1 (E–F′) and <xref ref-type="fig" rid="fig2">Fig. 2 (F–G′)</xref>, and VLDLR staining in Fig. S5 (B–C′) were amplified using the TSA Fluorescein Tyramide Reagent Pack (PerkinElmer). For VLDLR staining (Fig. S5, B–C′), goat anti–mouse VLDLR antibody (R&D Systems) was pre-absorbed with a <italic>Vldlr<sup>−/−</sup></italic> ear in TNB buffer (0.1 M Tris-HCl, pH 7.5, 0.15 M NaCl, and 0.5% blocking buffer [supplied in kit]) overnight before resuming normal amplification and staining protocol with a wild-type or mutant ear, as detailed above. Immunostaining for <xref ref-type="fig" rid="fig2">Fig. 2 (F–G′)</xref> was performed in the absence of Triton X-100. Staining for β-galactosidase activity in the lymphatic vessels in <italic>Vegfr3<sup>lz</sup></italic> mice was done using the substrate X-Gal (5-bromo-4-chloro-3-indolyl-β-<sc>d</sc>-galactopyranoside), and the images were acquired with a stereomicroscope (MZ16F) equipped with camera (DFC420C) and Microsystems software (all from Leica).</p></sec><sec><title>Image acquisition and quantitative analyses</title><p>Confocal images for <xref ref-type="fig" rid="fig2">Figs. 2 (B and C)</xref>, <xref ref-type="fig" rid="fig3">3 (G and H)</xref>, <xref ref-type="fig" rid="fig6">6 (C–E)</xref>, S3 (A–D), and S5 (F–H), were acquired at room temperature using a confocal microscope (LSM 710; Zeiss Imager.Z2 and Zen 2009 software; Carl Zeiss) with a Plan-Apochromat 20×/0.8 NA air objective lens (<xref ref-type="fig" rid="fig6">Fig. 6, C–E</xref>; and Fig. S5, F–H) or DIC Plan-Apochromat 40×/1.3 NA water immersion objective lens (<xref ref-type="fig" rid="fig2">Fig. 2, B and C</xref>; <xref ref-type="fig" rid="fig3">Fig. 3, G and H</xref>; and Fig. S3, A–D). All other confocal images were acquired using an LSM 510 microscope (Axioplan 2 and LSM 510 version 3.2 software; all from Carl Zeiss), with Plan-Apochromat 10×/0.45 NA (Fig. S2, A–D) or 20×/0.75 NA (<xref ref-type="fig" rid="fig4">Fig. 4, A and B</xref>; and Fig. S2, E and F) air objective lenses, or Korr C-Apochromat 63×/1.2 NA (Fig. S1, G and H) or 40×/1.2 NA (all other images) water immersion objective lenses. All confocal images, except Fig. S1 (G and H), represent maximum projection images of confocal z stacks. For visualization of collecting vessels from the proximal to distal end of the ear, images were captured along the length of three randomly chosen collecting lymphatic vessels from <italic>Reln<sup>−/−</sup></italic> (<italic>n</italic> = 5) and wild-type (<italic>n</italic> = 4) mice. The same analysis was performed for <italic>Apoer2<sup>−/−</sup>;Vldlr<sup>−/−</sup></italic>, <italic>Dab1<sup>Scm</sup></italic>, and their respective littermate controls (<italic>n</italic> = 3 per each genotype, two vessels per mouse). The images were aligned and processed using Photoshop CS4 or CS5 (Adobe). Thresholding and quantification of the whole-vessel images were performed using MetaMorph Imaging software (Molecular Devices). An inclusive threshold was applied to the α-SMA channel so that the background was captured by, while all SMC were excluded from, the threshold. The length of the vessel was measured and divided into quarters. A region was drawn around each vessel segment, excluding overlapping blood vessels. The SMC coverage was calculated and subtracted from 100%. A similar thresholding technique was used to quantify the percentage of LVYE-1 immunofluorescence. The line tool in MetaMorph was used to measure the width of the widest and narrowest vessel points.</p></sec><sec><title>Analysis of lymphatic vessel function</title><p>For qualitative analysis of lymphatic vessel function, FITC-dextran (8 mg/ml in PBS; Sigma-Aldrich) was injected intradermally into the ears of anaesthetized mice and analyzed by confocal microscopy. For quantitative analysis, 2 µl of ICG dye (1% in dH<sub>2</sub>O; Sigma-Aldrich) was injected into the footpads of anaesthetized mice inside the IVIS Spectrum Whole Body Imaging System (Caliper Life Sciences). Image acquisition and analysis was performed with Living Image 3 software (Caliper Life Sciences) using the following parameters: field of view 6.6, binning 8, and F-stop 2. 70 images were taken with a 2-s exposure time and no delay between acquisitions, which resulted in an image every 3–4 s. The efficiency of the fluorescent signal was measured in a region of interest (ROI) created around a small area of the lymphatic vessel above the knee. The mean efficiency against time was plotted using Prism 5 software, using 4 s as an interval. For the quantitative Evans blue assay, 2 µl Evan’s blue dye (2%; Sigma-Aldrich) was injected into the hind limb footpad. After 5 min, the mice were sacrificed and inguinal lymph nodes were dissected. Both inguinal lymph nodes were weighed together and a mean value was used to calculate dye content in each of them separately. Each lymph node was placed in 5 µl formamide at 60°C for 24 h, followed by an absorbance reading at 620 nm. Evans blue concentration was calculated using a standard curve and presented as dye content per gram of tissue.</p></sec><sec><title>Cell culture</title><p>Human LECs, isolated from primary dermal microvascular EC cultures (PromoCell; <xref ref-type="bibr" rid="bib24">Mäkinen et al., 2001</xref>), human umbilical venous SMCs (HUVSMCs; TCS Cellworks), and aortic SMCs (PromoCell) were cultured in endothelial or SMC growth medium (Promocell). For co-culture experiments, LEC were plated on fibronectin-coated plates at 50% confluency in IgG-stripped EC growth medium 4 h before an equal number of HUVSMC were added to the culture. Medium was collected and the cells were lysed 2 d after plating. Supernatant from HEK 293 cells transfected with pCrL (full-length mouse Reelin cDNA; a gift from Tom Curran, Joseph Stokes Jr. Research Institute, The Children’s Hospital of Philadelphia, Philadelphia, PA) and untransfected control cells was concentrated 25× using an Amicon ultra-15 centrifugal filter unit (Sigma-Aldrich) and used for stimulation of LEC or SMC at dilution 1:20.</p></sec><sec><title>Reelin stimulation</title><p>LEC were starved overnight in serum-free EC growth medium, then supplemented with 0.2% BSA, hEGF-5, and HC-500, followed by stimulation with full-length Reelin supernatant from HEK 293 cells (diluted 1:20), control supernatant (diluted 1:20), or recombinant central Reelin fragment (R&D Systems) for 4, 8, or 24 h. RNA was extracted using RNeasy Mini kit (QIAGEN) according to the manufacturer’s instructions and used for qPCR analysis using the following Taqman probes (Applied Biosystems): <italic>PDGFB</italic>, Hs00966522_m1; <italic>ANGPT2</italic>, Hs01048042_m1; <italic>TGFβ1</italic>, Hs00998133_m1; <italic>HBEGF</italic>, Hs00181813_m1; and <italic>CCL2/MCP-1</italic>, Hs00234140_m1 and <italic>GAPDH</italic>; Hs99999905_m1.</p></sec><sec><title>Western blot analysis</title><p>Cell growth medium was collected and clarified by centrifugation, and the cells were lysed in Triton X-100 lysis buffer (50 mM Tris, pH 7.5, 120 mM NaCl, 10% glycerol, 1% Triton X-100) supplemented with Complete Protease inhibitors (Roche). Cell lysate and growth medium were subjected to immunoprecipitation and/or Western blot analysis using mouse antibodies against human Reelin (EMD) and PECAM-1 (Dako), or Cy3-conjugated mouse anti–α-SMA (Sigma-Aldrich). Quantification was performed using ImageJ, and the ratio of Reelin in the lysate and medium, normalized to CD31/PECAM-1, was calculated for LEC/SMC co-culture relative to LEC cultured alone.</p></sec><sec><title>qPCR</title><p>RNA from LEC, BECs, human aortic SMCs (HAoSMCs), and HUVSMC was reverse transcribed using Superscript II Reverse transcription (Invitrogen) and random hexamers (Promega). cDNA was amplified by real-time qPCR (ABI 7900HT) using Platinum SYBR Green qPCR super mix-UDG with ROX reagent (QIAGEN). Each primer was used at a concentration of 0.4 µM. Cycling conditions were as follows: step 1, 2 min at 50°C; step 2, 10 min at 95°C; step 3, 15 s at 95°C; and step 4, 1 min at 60°C. Steps 3 and 4 were repeated 40 times. Data from the reaction was collected and analyzed using the complementary computer software. Relative quantifications of gene expression were normalized to expression of <italic>GAPDH</italic>. The primers were: <italic>RELN</italic>, 5′-GTTTTGCAAATTGGGAGCAT-3′ and 5′-CCGTTGTTGACGCTGTATTG-3′; <italic>GAPDH</italic>, 5′-GAAGGTGAAGGTCGGAGTC-3′ and 5′-GACAAGCTTCCCGTTCTCAG-3′; <italic>PECAM1</italic>, 5′-GAGAGGACATTGTGCTGCAA-3′ and 5′-GGGTTTGCCCTCTTTTTCTC-3′; and <italic>LYVE1</italic>, 5′-GCTTTCCATCCAGGTGTCAT-3′ and 5′-AGCCTACAGGCCTCCTTAGC-3′.</p></sec><sec><title>Transwell migration assay</title><p>Migration assay was performed using 6.5-mm Transwell with 8 µm polycarbonate membrane inserts (Corning) in a 24-well plate. The lower face of the membrane was coated with 1 µg/ml fibronectin (Sigma-Aldrich) before use. Confluent HUVSMC were serum-deprived in SMC growth medium containing 0.2% BSA for 22 h. 1 × 10<sup>5</sup> cells were seeded in the upper chamber and allowed to migrate toward EC growth medium (control) or conditioned medium collected from LECs over a 48-h period, full-length Reelin supernatant from HEK 293 cells (diluted 1:20), control supernatant (diluted 1:20), 10 ng/ml MCP-1 (R&D Systems), or 3 ng/ml PDGF-BB (R&D Systems) over a 23-h period. Reelin function was inhibited using blocking antibody against Reelin (CR-50, 50 µg/ml; MBL). Cells on the upper face of the membrane were removed and migrated cells on the lower face were fixed and stained with 0.5% crystal violet solution in 25% methanol. Images of the filters were acquired with a stereomicroscope (MZ16 F) equipped with a camera (DFC420C) and Microsystems software (all from Leica). After photographing, the incorporated dye was solubilized in 0.2 ml sodium citrate solution, followed by absorbance reading at 550 nm.</p></sec><sec><title>Statistical analysis</title><p>P-values were calculated using unpaired two-tailed Student’s <italic>t</italic> tests.</p></sec><sec><title>Online supplemental material</title><p>Fig. S1 shows the identification of extracellular matrix–associated proteins that are preferentially expressed in lymphatic vessels. Fig. S2 shows the characterization of <italic>Reeler</italic> phenotype in the skin. Fig. S3 shows the characterization of <italic>Reeler</italic> phenotype in the mesentery. Fig. S4 shows real-time imaging of lymphatic function in 3-wk-old control and <italic>Reln</italic>-deficient mice. Fig. S5 shows the involvement of Reelin signaling pathway components in lymphatic development and the effect of Reelin on SMC migration. Table S1 presents the validation of microarray data for known markers of lymphatic vessels, arteries, and veins. Table S2 shows genes associated with extracellular matrix that were differentially expressed in lymphatic and blood vessels. Table S3 shows the full dataset that was used for the validation of microarray hits. Online supplemental material is available at <ext-link ext-link-type="uri" xlink:href="http://www.jcb.org/cgi/content/full/jcb.201110132/DC1">http://www.jcb.org/cgi/content/full/jcb.201110132/DC1</ext-link>.</p></sec></sec><sec sec-type="supplementary-material"><title>Supplementary Material</title><supplementary-material id="PMC_1" content-type="local-data"><caption><title>Supplemental Material</title></caption><media mimetype="text" mime-subtype="html" xlink:href="supp_197_6_837__index.html"></media><media xlink:role="associated-file" mimetype="application" mime-subtype="pdf" xlink:href="supp_jcb.201110132_JCB_201110132_sm.pdf"></media><media xlink:role="associated-file" mimetype="application" mime-subtype="vnd.ms-excel" xlink:href="supp_jcb.201110132_JCB_201110132_TableS3.xls"></media></supplementary-material></sec></body><back><ack><p>We thank Joachim Herz for <italic>Apoer2</italic> and <italic>Vldlr</italic> mice, Michael Frotscher and Hans Bock for sending these mice to us, Kari Alitalo for <italic>Vegfr3<sup>lz</sup></italic> mice, Lydia Sorokin for Laminin-α5 antibodies, and Tom Curran for the pCrl plasmid. We thank the CRUK GeneChip Service Facility at Cancer Research UK Paterson Institute for performing the microarray experiments, Phil East for analysis of microarray data, Francois Lassailly for help with whole-body imaging, and the animal unit staff at the London Research Institute for animal husbandry.</p><p>This work was supported by Cancer Research UK (S. Lutter, S. Xie, F. Tatin, and T. Makinen) and EMBO Young Investigator Program (T. Makinen).</p></ack><fn-group><fn><p><def-list><title>Abbreviations used in this paper:</title><def-item><term>α-SMA</term><def><p>α-smooth muscle actin</p></def></def-item><def-item><term>BEC</term><def><p>blood vascular endothelial cell</p></def></def-item><def-item><term>BM</term><def><p>basement membrane</p></def></def-item><def-item><term>EC</term><def><p>endothelial cell</p></def></def-item><def-item><term>HAoSMC</term><def><p>human aortic smooth muscle cell</p></def></def-item><def-item><term>HUVSMC</term><def><p>human umbilical venous smooth muscle cell</p></def></def-item><def-item><term>ICG</term><def><p>indocyanine green</p></def></def-item><def-item><term>LEC</term><def><p>lymphatic EC</p></def></def-item><def-item><term>LYVE-1</term><def><p>lymphatic vessel endothelial hyaluronan receptor 1</p></def></def-item><def-item><term>MCP1</term><def><p>monocyte chemotactic protein 1</p></def></def-item><def-item><term>PECAM-1</term><def><p>platelet EC adhesion molecule 1</p></def></def-item><def-item><term>qPCR</term><def><p>quantitative PCR</p></def></def-item><def-item><term>SMC</term><def><p>smooth muscle cell</p></def></def-item><def-item><term>TSA</term><def><p>tyramide signal amplification</p></def></def-item></def-list></p></fn></fn-group><ref-list><ref id="bib1"><mixed-citation publication-type="journal"><person-group person-group-type="author"><name><surname>Aplin</surname><given-names>A.C.</given-names></name><name><surname>Fogel</surname><given-names>E.</given-names></name><name><surname>Nicosia</surname><given-names>R.F.</given-names></name></person-group><year>2010</year><article-title>MCP-1 promotes mural cell recruitment during angiogenesis in the aortic ring model</article-title>. <source>Angiogenesis.</source><volume>13</volume>:<fpage>219</fpage>–<lpage>226</lpage><pub-id pub-id-type="doi">10.1007/s10456-010-9179-8</pub-id><pub-id pub-id-type="pmid">20571857</pub-id></mixed-citation></ref><ref id="bib2"><mixed-citation publication-type="journal"><person-group person-group-type="author"><name><surname>Aszódi</surname><given-names>A.</given-names></name><name><surname>Legate</surname><given-names>K.R.</given-names></name><name><surname>Nakchbandi</surname><given-names>I.</given-names></name><name><surname>Fässler</surname><given-names>R.</given-names></name></person-group><year>2006</year><article-title>What mouse mutants teach us about extracellular matrix function</article-title>. <source>Annu. 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