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Molecular and Cellular Biology, June 2008, p. 4026-4039, Vol. 28, No. 12
0270-7306/08/$08.00+0 doi:10.1128/MCB.02062-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Division of Experimental Oncology 2, Department of Molecular Oncology and Translational Research, CRO-IRCCS, Aviano, Pordenone, Italy,1 IGM-CNR, Unit of Bologna c/o IOR, Bologna, Italy,2 Mouse Genetics Laboratory, Department of Histology Microbiology and Medical Biotechnologies, University of Padua, Padua, Italy,3 Department of Biomedical Sciences and Technologies, University of Udine, Udine, Italy,4 MATI Center of Excellence, University of Udine, Udine, Italy5
Received 16 November 2007/ Returned for modification 25 December 2007/ Accepted 27 March 2008
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Lymphatic and blood endothelial cells express different lineage-specific molecules involved in the regulation of their biological functions that are frequently used as distinguishing markers, such as vascular endothelial growth factor receptor 3 (VEGFR-3) (22), podoplanin (3), Prox-1 (44), LYVE-1 (1, 34), neuropilin 2 (46), CCL21 (24), and desmoplakin (12). Recently, a comparative microarray analysis of gene expression profiles of lymphatic and blood endothelial cells identified previously unknown lymphatic lineage genes, including macrophage mannose receptor 1, plakoglobin, the chemokine CCL20, the integrin
9β1 (19, 32), and EMILIN1 (33).
EMILIN1 is an ECM glycoprotein associated with elastic fibers (4, 6) and composed of an N-terminal cysteine-rich domain and the EMI domain (11), followed by a coiled-coil structure, a short collagenous stalk, and a C-terminal gC1q domain (7). EMILIN1 is particularly abundant in the walls of large blood vessels, such as the aorta (10), and has been implicated in multiple functions. EMILIN1 is involved in elastogenesis and in the maintenance of blood vascular cell morphology (48). It interacts with the
4β1 integrin through the gC1q1 domain (36) and has strong adhesive and migratory properties for different cell types (10, 36, 37). EMILIN1, via the EMI domain, regulates pro-transforming growth factor beta (TGF-β) maturation and is involved in blood pressure homeostasis (47).
To directly investigate the physiological function of EMILIN1 in lymphatic vessels, we studied the effects of its absence in mice that had targeted deletions in the Emilin1 gene (48). Here, we report that EMILIN1 is highly expressed by LECs in vitro and that it colocalizes with lymphatic vessels in several mouse tissues. Importantly, Emilin1 deficiency results in hyperplasia and enlargement of lymphatic vessels and in a significant reduction of anchoring filaments compared to those of wild-type (WT) mice. The lymphatic vessels of Emilin1–/– mice are functionally altered. We found that lack of EMILIN1 leads to a mild lymphedema associated with inefficient lymph drainage and increased leakage. In addition, Emilin1–/– mice develop larger lymphangiomas than their WT littermates. Altogether, these findings demonstrate an important role of EMILIN1 in the structure-function relationship of lymphatic vessels and identify EMILIN1 as a lymphangiogenesis modulator.
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Mouse procedures and cell culture. Procedures involving animals and their care were conducted according to institutional guidelines in compliance with national laws (D.Lgs. no. 116/92). Emilin1–/– mice (CD1 and C57BL/6 strains) were generated as previously described by Zanetti et al. (48). For lymphangioma induction, the protocol published by Mancardi et al. (27) was followed using BALB/c (Harlan Italy S.r.L., Udine, Italy) and CD1 mice. Briefly, the mice were intraperitoneally injected twice, with a 15-day interval, with 200 µl of emulsified (1:1 with PBS) incomplete Freund's adjuvant (Sigma, St. Louis, MO). Hyperplastic vessels were isolated from the liver and diaphragm after day 30 and treated with 0.5 mg/ml collagenase A (Roche Diagnostics, Monza, Italy), and the resulting single-cell suspension was cultured as previously described (27). After 7 to 10 days of culture, subconfluent cells (lymphangioma-derived endothelial cells [LAECs]) were recovered with trypsin/EDTA and characterized by immunocytochemistry.
Human microvascular endothelial cells dermal lymphatic-neonatal (HMVEC-dLyNeo), HMVEC lung lymphatic (HMVEC-LLy), and the media optimized for their growth (EGM-2 MV) were purchased from Cambrex Bio Science (Verviers, Belgium); bEnd3, a mouse endothelioma cell line derived from brain capillaries, was from ATCC (Manassas, VA). Human umbilical vein endothelial cells (HUVEC) were isolated from three to five normal umbilical cord veins by collagenase digestion following the standard procedure previously described (25).
Immunofluorescence and whole-mount staining. Mouse tissues were excised, embedded in OCT (Kaltek, Padova, Italy), snap-frozen, and stored at –80°C. Cryostat sections (7 µm) were air dried at room temperature and kept at –80°C wrapped in aluminum foil. Before being used, the sections were equilibrated at room temperature, hydrated with phosphate-buffered saline (PBS) for 5 min, and fixed with PBS-4% paraformaldehyde (PFA) for 15 min. Then, the sections were permeabilized (with PBS, 1% bovine serum albumin, 0.1% Triton X-100, 2% fetal calf serum) for 5 min and saturated with the blocking buffer (PBS, 1% bovine serum albumin, and 2% serum) for 30 min. The primary antibodies were then incubated at room temperature for 1 h, followed by three 5-min washes in PBS and secondary-antibody incubation for 1 h. Multiple staining was performed using a combination of differently conjugated secondary antibodies: Alexa Fluor 488, Alexa Fluor 568, and Alexa Fluor 633 (Molecular Probes, Eugene, OR). Nuclei were visualized with propidium iodide or ToPro, both from Molecular Probes.
For whole-mount staining, mice were anesthetized with 0.4 g Avertin (Sigma)/kg of body weight, perfused with PBS PFA 1%, and sacrificed. Tissue specimens of interest were finely minced and fixed in 4% PFA for 18 h at 4°C. Then, the blocking solution (PBS, 0.3% Triton X-100, and 5% serum) was added for 8 h at 4°C, followed by primary-antibody incubation for 18 h at 4°C. After five washes with PBS 0.3% Triton X-100, the secondary antibody was added, and the specimens were incubated for 18 h at 4°C. Images were acquired with a Leica TCS SP2 confocal system (Leica Microsystems Heidelberg, Mannheim, Germany), using Leica confocal software.
Electron microscopy. Skin fragments were dissected from 12-week-old mice, fixed with 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) overnight, washed in 0.1 M sodium cacodylate buffer overnight, and treated with 2% tannic acid in 0.1 M sodium cacodylate buffer as previously described (48). All samples were dehydrated with ethanol and embedded in Epon E812. Ultrathin sections were obtained from several blocks, stained with lead citrate and uranyl acetate, and observed in a Philips EM 400 transmission electron microscope operated at 100 kV.
In vitro tube formation assay. To assess the ability of mouse LAECs to form vessel-like structures in vitro, Matrigel (8.8 mg/ml; BD Biosciences, Erembodegem, Belgium) was added to the wells of a 96-well plate in a volume of 50 µl and allowed to solidify for 30 min at 37°C. Once it was solid, 2 x 105 LAECs were seeded in each well in 200 µl of EGM-2 MV medium and incubated at 37°C for 6 h. Tube formation was visualized with a camera-equipped inverted Nikon ECLIPSE TS100 microscope.
Computer-assisted morphometric analyses. To quantitatively evaluate vessel density and diameter, LYVE-1-, podoplanin-, and MMRN2-stained cryostat sections were analyzed using a Leica TCS SP2 confocal system. On the acquired images, computer-assisted morphometric analyses were performed using ImageJ software (http://rsb.info.nih.gov).
In vivo lymphatic-vessel function analysis. (i) Lymph drainage analysis. To evaluate the lymph flow of WT and Emilin1–/– mice from CD1 and C57BL/6 strains, a modified version of the Miles assay, published by Sugaya et al. (39), was applied. Briefly, a 3% solution of Evans blue dye (Sigma) in PBS was injected (1 µl/g) into the footpads of anesthetized mouse hind limbs. To quantify the lymph flow, draining local and distal lymph nodes were harvested 30 min after injection. The accumulated dye was extracted after the lymph nodes were incubated in formamide (Sigma) overnight at 55°C, and it was quantified spectrophotometrically at 620 nm (GENios Plus; TECAN Italia S.r.L.).
(ii) Lymphatic-vessel leakage assay. To evaluate lymphatic-vessel leakage of WT and Emilin1–/– mice, another version of the Miles assay (38), modified by us, was performed. Evans blue dye (3%) was injected (1 µl/g) into the footpads of hind limbs, and after 5 min, an inflammatory agent, mustard oil (Sigma), diluted to 5% in mineral oil (Sigma), was intradermally injected into the ventral skin. As a control, mineral oil only was injected. After 30 min, the animals were sacrificed and the leakage was detected as a blue spot on the underside of the skin. Evans blue was quantified as described above by excising the portion of skin affected by the extravasation and the control skin.
(iii) Intravital lymphangiography. For intravital lymphangiography, 1 µl of a 1% solution of Evans blue dye in PBS was injected intradermally at the inner surface of the rim of the ear. Mouse ear lymphatic vessels were photographed 1, 3, and 5 min after the dye injection.
RNA extraction and RT-PCR. Total cellular RNA was isolated from lymphatic and blood endothelial cells using Trizol (Invitrogen, Milan, Italy) according to the manufacturer's protocol. Reverse transcription (RT) reactions were performed with 1 µg of total RNA using AMV Reverse Transcriptase (Promega Italia, Milan, Italy). RNA was reverse transcribed into first-strand cDNA using random hexamer primers. The primers for the PCR amplification of mouse EMILIN1 were 5'-TGTGCCTAGGGTAGCATTTTC-3' and 5'-GAGGCTGAAGACGCCCAGAG-3'. The size of the amplification product was 320 bp. Taq DNA polymerase was obtained from Roche (Monza, Italy). RT-PCR amplification of a 700-bp fragment of β-actin cDNA served as a positive internal control. Amplification products were resolved on 2% agarose gels stained with ethidium bromide.
Quantitative real-time PCR. Real-time PCR was carried out on an ABI PRISM 7900 HT sequence detection system (Applied Biosystems, Warrington, United Kingdom) using the Power Sybr Green PCR Master Mix kit (Applied Biosystems). The calculated amount of EMILIN1 mRNA was normalized to the endogenous reference control mRNA β-actin. All primers were designed with Primer3 software (Universal Probe Library Assay Design Center, Applied Science, Roche, Monza, Italy) and were as follows: for human EMILIN1, 5' CAGTGTCCCCAAAGCATCAT 3' and 5'CACTCCATGTCGGTCACTG T 3'; for human β-actin, 5' CCAACCGCGAGAAGATGA 3' and 5' CCAGAGGCGTACAGGGATAG 3'. The results were analyzed with SDS 2.1 software (Applied Biosystems).
Statistical analysis. The statistical significance of the results was determined by using the unpaired Student's t test. A P value of <0.05 was considered significant.
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FIG. 1. Human and mouse LECs express high levels of EMILIN1 in vitro. (A) Characterization of human lung and dermal neonatal LECs (HMVEC-LLy and HMVEC-dLyNeo). The positive staining for the lymphatic specific markers, LYVE-1, Prox-1, VEGFR-3, podoplanin, and CD31, is green. Nuclei were stained by ToPro (blue). Scale bar, 45 µm. (B) Quantitative RT-PCR was performed on mRNA extracted from LECs (HMVEC-LLy and HMVEC-dLyNeo, at passage 5) and blood endothelial cells (HUVEC, at passage 4). The relative EMILIN1 (E1) expression versus β-actin was quantified using the SDS 2.1 program. The analysis confirmed abundant EMILIN1 expression by LECs, revealing a threefold increase in HMVEC-LLy (**, P < 6 x 10–6) and a twofold increase in HMVEC-dLyNeo (*, P < 0.006) compared with HUVEC. HMVEC-LLy produced higher levels of EMILIN1 then did HMVEC-dLyNeo ( , P < 0.006). (C) Comparative RT-PCR analysis of EMILIN1 mRNA levels in mouse LAECs and in bEnd3 (a mouse endothelioma cell line). (D) Immunofluorescence staining of EMILIN1 (green) shows its abundant production and extracellular deposition by LAECs (left) compared to bEnd3 cells (right). Nuclei were stained red by propidium iodide. Scale bar, 10 µm.
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FIG. 2. EMILIN1 is expressed in association with lymphatic vessels. (A to D) Cryostat sections of normal mouse tissues doubly stained with anti-EMILIN1 (green) and anti-LYVE-1 (red) antibodies. In all mouse tissues and organs examined, EMILIN1 was uniformly distributed in the stroma. (A) In the skin, EMILIN1 staining colocalizes with LYVE-1-positive lymphatic vessels surrounding hair follicles (arrowheads; scale bar, 75 µm). (B) In the small intestine, EMILIN1 colocalizes with LYVE-1-positive lacteals and submucosal lymphatic vessels (arrowheads; scale bar, 75 µm). (C and D) At higher magnification, in the lung and lymph nodes, it is more evident that EMILIN1 is distributed at the abluminal surfaces of LECs (arrows; scale bars, 45 µm). In the lymph node, EMILIN1-positive fibers connecting LECs to the surrounding ECM are evident (asterisks).
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FIG. 3. Hyperplasia and enlarged lymphatic vessels in Emilin1-deficient mice. (A and B) Immunofluorescence staining of mouse skin (A) and small intestine (B) for podoplanin (red), LYVE-1 (blue), and MMRN2 (green) revealed a higher number of enlarged lymphatic vessels in the skin (A, bottom) and in the intestines (B, bottom) of Emilin1–/– mice (n = 10) than in their WT littermates (n = 10) (A and B, top). (C and D) Vessel counts per field. In a double-blind study, transversally oriented cryostat sections were observed at x40 magnification, and the vessel density was evaluated in random fields. The number (mean ± standard error [SE]) of LYVE-1-positive vessels was significantly increased in the skin (C) (**, P < 1.5 x 10–7) and in the intestine (D) (*, P < 0.0002) of Emilin1–/– mice. The numbers (mean ± SE) of MMRN2-positive blood vessels were not significantly different in the two mouse genotypes. (E) Immunofluorescence analysis of mouse skin for the proliferation marker Ki67 (green) and for podoplanin (red) showed a higher number of proliferating LECs in Emilin1–/– (n = 5) than in WT (n = 5) mice. Representative images are shown above the graphs; the arrows indicate Ki67-positive LECs. The count of Ki67-positive cells per lymphatic vessel in mouse skin cryostat sections revealed a threefold-higher percentage of Ki67-positive lymphatic vessels (bottom left) and a significant increase in Ki67-positive cells per lymphatic vessel (bottom right) (mean ± SE; *, P = 0.015) in Emilin1–/– mice compared to their WT littermates. Ki67 quantification analysis was performed, examining 49 WT and 55 Emilin1–/– mouse lymphatic vessels. (F and G) A computer-assisted image analysis (ImageJ software) confirmed that the diameters (mean ± SE) of lymphatic vessels were significantly increased in the skin (F) (**, P < 6 x 10–6) and in the intestine (G) (*, P = 0.03) of Emilin1–/– mice. E1, EMILIN1. Scale bars, 75 µm.
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FIG. 4. Hyperplasia and enlarged lymphatic vessels in lymph nodes of Emilin1-deficient mice. (A) Representative immunofluorescence images of mouse inguinal lymph node cryostat sections stained for LYVE-1 (red) and MMRN2 (green). Scale bars, 300 µm. (B) ImageJ software analysis revealed that the relative area occupied by lymphatic vessels in lymph nodes of Emilin1–/– (E1–/–; n = 9) mice was significantly higher (mean ± standard error [SE]; *, P < 2 x 10–6) than in those of their WT littermates (n = 9). (C) Higher-magnification images show dilated LYVE-1-positive vessels (right). Scale bars, 75 µm. (D) The mean value ± SE of the diameters (ImageJ software analysis) of lymphatic vessels is reported. *, P < 2 x 10–7.
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FIG. 5. Abnormal lymphatic-vessel morphology in Emilin1-deficient mice. (A) Whole-mount immunofluorescence staining with LYVE-1 shows an irregular morphology of ear lymphatic vessels. The white asterisks indicate buds on lymphatic vessels. Scale bars, 300 µm. (B to D) LYVE-1 diaminobenzidine-peroxidase whole-mount staining of submucosal (B) (scale bars, 500 µm) and subserosal (C and D) (scale bars, 500 µm) lymphatic vessels. The black asterisks indicate dysmorphic structures.
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FIG. 6. Lymphangioma induction in Emilin1-deficient mice. (A) Representative images show that lymphangioma plaques were more numerous and larger in Emilin1–/– mice (the arrowheads indicate the white plaques that developed on the liver and diaphragm surfaces). (B) Lymphangioma development. Class I corresponds to few and small lymphangioma plaques, class III to numerous and large plaques, and class II to an intermediate situation. (C) Lymphatic-vessel density. Lymphangioma cryostat sections were stained with LYVE-1, and in a double-blind study, lymphatic-vessel densities were evaluated as low, medium, and high. (D) Quantitative ImageJ analysis. The average fluorescence intensity confirmed that lymphatic-vessel density was significantly increased in Emilin1–/– (E1–/–) and Emilin1+/– (E1+/–) mouse lymphangiomas compared with those of their WT littermates (*, P < 0,04; **, P < 6 x 10–5). No significant differences were observed for MMRN2-positive blood vessels. The error bars indicate standard deviations. (E) Immunofluorescence analysis of cryostat sections of mouse lymphangiomas. LYVE-1-positive lymphatic vessels (red; note that liver sinusoids are also positive) and MMRN2-positive blood vessels (green). L, liver; T, tumor. Scale bars, 75 µm. (F) Whole mounts of diaphragm plaques of WT and Emilin1–/– mice stained with LYVE-1. Scale bars, 300 µm. (G) Proliferation rate. WT and Emilin1–/– LECs isolated from mouse lymphangiomas (LAECs) were seeded onto glass coverslips and grown in EBM plus 2% fetal bovine serum. After 72 h, Ki67-positive nuclei per field were counted at x60 magnification. *, P < 0.006. (H) Tube formation assay. WT and Emilin1–/– LAECs were seeded onto Matrigel and allowed to form tube-like structures for 6 h. The images were acquired with a camera-equipped inverted microscope (x10 original magnification).
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Emilin1 deficiency affects LEC anchorage. To investigate the lymphatic defects caused by Emilin1 deficiency at the ultrastructural level, transmission electron microscopy analysis was performed on WT and Emilin1–/– mouse skin specimens. WT lymphatics of skin displayed an irregular lumen with frequent abluminal endothelial cell projections into the ECM. The abluminal plasma membranes of the endothelial cells showed a discontinuous basal lamina and bundles of microfibrils, i.e., the anchoring filaments. These radiated from the membranes of LECs and reached small elastin fibrils (with diameters ranging from 150 to 500 nm) (Fig. 7A); in the connection region, the LEC membrane appeared to be modified and the subcortical associated cytoskeleton was characterized by the presence of thin filaments (Fig. 7A, inset). Emilin1–/– lymphatic vessels showed differences concerning the abluminal surfaces of the endothelial cells: rare endothelial cell projections into the ECM were detected, giving a smooth appearance to the abluminal surface, and the number of bundles of anchoring filaments was significantly reduced with respect to the WT (Fig. 7B and C) (P < 0.005). Another frequent finding was the presence of abnormal intercellular junctions. WT LECs showed characteristic overlapping junctions (Fig. 7D), whereas Emilin1–/– LECs frequently presented multiple overlapping contacts. These junctions did not originate from an interdigitation but were the result of an extended overlap among several LECs that appeared thin and tightly packed and developed adherens junctions (Fig. 7E). These aspects were never detected in WT animals.
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FIG. 7. EMILIN1 deficiency affects LEC anchorage. Transmission electron microscopy analysis of WT (A and D) and Emilin1–/– (B and E) skin lymphatic vessels. (A) WT LECs display characteristic bundles of anchoring filaments (arrows), which extend from the abluminal side of the plasma membrane into the adjoining connective tissue (inset, arrowheads). (B) Emilin1–/– LECs show reduced numbers of tufts of anchoring filaments (arrow). (C) Quantitative analysis of the bundles of anchoring filaments in WT and Emilin1–/– skin lymphatics (n = 10). The values express means ± standard deviations per 100 µm and show a significant reduction in the number of Emilin1–/– lymphatic vessels (P < 0.005). (D) Normal lymphatic endothelium showing typical overlapping junctions (arrows) between adjacent LECs. (E) Abnormal overlapping junctions in an Emilin1–/– lymphatic vessel involving several LECs (arrows); the LECs in these areas appeared thin and packed and developed adherens junctions. Scale bar, 200 nm; inset scale bar, 60 nm.
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FIG. 8. Impaired lymphatic function in Emilin1-deficient mice. (A) Evans blue dye accumulation in inguinal lymph nodes 30 min after dermal injection into the footpad; the dye is barely detectable in Emilin1–/– mice (right), whereas the inguinal lymph node is easily visualized in WT mice (left). The images were acquired with a camera-equipped dissection microscope. (B) Evans blue dye contents (mean ± standard deviation [SD]) in inguinal and axillary lymph nodes expressed as percentages of the amount (ng/mg) of dye in WT lymph nodes after dermal injection into the footpad (*, P < 1.5 x 10–6; **, P < 8 x 10–7). E1–/–, Emilin1–/–. (C) Lymph leakage. Spots of Evans blue extravasation in mustard oil-treated mouse skin. (D) Quantification (mean ± SD) of Evans blue dye content in treated (*, P < 0.0085) and untreated (**, P < 6 x 10–5) skin. (E) Intravital lymphangiography. To visualize lymphatic vessels, 2 µl of 1% Evans blue dye was intradermally injected into the rims of the ears of WT (n = 5) and Emilin1–/– (n = 5) mice. At 1, 3, and 5 min, the ears were photographed with a camera-equipped dissection microscope. One representative experiment is shown. The arrows indicate the major leakage areas in Emilin1–/– mouse ears; the arrowheads indicate the extension of the leakage (bottom). (Right) The irregular morphology of Emilin1–/– mouse lymphatic vessels is evidenced by processed images. (F) Peripheral edema. Hind limbs of WT and Emilin1–/– mice of equal weight were compared.
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Next, intravital lymphangiography 1 min after Evans blue dye injection demonstrated that markedly leaky lymphatic vessels were already visualized in Emilin1–/– (n = 5) mouse ears (Fig. 8E, bottom), especially near the site of injection. After 3 and 5 min, the extent of leakage progressively increased in Emilin1–/– mouse skin, and also, the more distal lymphatic vessels presented signs of dye extravasation (Fig. 8E, bottom). On the other hand, leakage was never observed in WT (n = 5) mice (Fig. 8E, top). Of note, the irregular and tortuous pattern of Emilin1–/– mouse lymphatic vessels compared with those of their WT littermates (Fig. 8E, processed images [right]). In accord with lymphatic-vessel defects, Emilin1–/– mice showed swelling of the paws, indicating mild lymphedema (Fig. 8F), whereas the formation of ascites was never observed in Emilin1-deficient mice.
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The present study extended, by quantitative RT-PCR, previous microarray data showing that LECs express abundant EMILIN1 mRNA (33). We detected a twofold and a threefold increase of EMILIN1 mRNA relative levels, compared to HUVEC, in human dermal neonatal and lung microvascular LECs, respectively. This difference is in agreement with the phenotypic heterogeneity of LECs isolated from different organs and different segments of the lymphatic vasculature, as previously reported (17). Moreover, RT-PCR and immunofluorescence analyses showed much higher EMILIN1 production and deposition in mouse lymphatic cells than in blood endothelial cells of mouse origin. Consistent with the absence of a basement membrane in vivo, LECs secrete very little ECM in comparison to blood endothelial cells (15); thus, the abundant EMILIN1 production in vitro suggested a potential involvement of this protein in the structure and function of lymphatic vessels.
EMILIN1 was expressed in all mouse tissues in association with lymphatic vessels, and it frequently colocalized with the abluminal surfaces of LECs. Moreover, EMILIN1-positive fibers were observed radiating from LECs to the surrounding ECM. Considering the association of EMILIN1 with elastic fibers, its expression in the lymphatic perivascular area indicates that EMILIN1 represents a component of the lymphatic fibrillar elastic apparatus previously described (18). This apparatus is formed by three hierarchical components disposed concentrically around lymphatic capillaries: the oxytalan fibers, identified as fibrillin microfibrils (35), connected directly to LECs and representing the real anchoring filaments; then elaunin fibers; and, more distant from the vessel, elastic fibers (18). It has been postulated that this apparatus is highly sensitive to interstitial stresses. An increase in the interstitial fluid volume exerts tension on LECs, widening the capillary lumen and opening the overlapping cell junctions, thus facilitating lymph formation (40). EMILIN1 involvement in this delicate physiologic mechanism was demonstrated by a comparative study between WT and Emilin1–/– mice that highlighted the fact that Emilin1 deficiency induces defects in lymphatic vascular structure and function. Emilin1–/– mice display hyperplasia and enlargement of dermal, as well as visceral, lymphatic vessels, and frequently these vessels present a tortuous and irregular pattern. Moreover, LECs of Emilin1–/– mice showed in an ultrastructural examination a significant reduction in the number of anchoring filaments and abnormal multiple overlapping intercellular junctions. Lymphatic-vessel defects were associated in Emilin1–/– mice with impaired lymph drainage, enhanced lymph leakage, and mild lymphedema. These findings indicate that the absence of EMILIN1 causes defective ECM anchorage of LECs and, consequently, dilation of the lymphatic-vessel lumen and reduced responsiveness to interstitial pressure variations. The resulting abnormal lymphatic function is more likely the consequence of enhanced lymph leakage than of defective lymph transport by collecting lymphatic vessels. In this view, the main phenotype characteristic of Emilin1 null mice is in lymph formation. Notably, the phenotype displayed by Emilin1–/– mice is the first abnormal lymphatic phenotype associated with deficiency of an ECM protein, and thus, it represents a useful tool to demonstrate the supposed fundamental role of anchoring filaments and the lymphatic perivascular elastic apparatus, as well as of the surrounding ECM, in lymphatic-vessel function.
In contrast to other recently described lymphatic-lineage gene-targeting mouse models that in most cases are embryonic or perinatal lethal (reviewed in reference 41), homozygous disruption of the Emilin1 gene induces a mild phenotype, indicating that the protein does not play a key role in the developmental processes of the lymphatic vasculature. On the contrary, it seems to be involved in regulation of the growth of lymphatic vessels and in maintenance of their integrity, a fundamental requirement for an efficient lymphatic-system function. Malfunctions of the lymphatic system rarely result in human life-threatening diseases. The most common disorder is lymphedema, which derives from the failure of lymph transport (23). Lymphedema may be an inherited disease caused by mutations identified in genes encoding VEGFR-3 (45), FOXC2 (14, 16), and Sox18 (20) or it may be acquired and occur after obstruction or damage of lymphatic vessels (23). Considering the lymphatic phenotype displayed by Emilin1–/– mice, we suppose that Emilin1 deficiency may resemble early stages of acquired lymphedema. Tissue inflammation following injury, exposure to radiation, or infection may induce the release of proteolytic enzymes that degrade EMILIN1 and render lymphatic vessels nonresponsive to the changes in the interstitium and therefore may cause an acute lymphatic insufficiency. This hypothesis is supported by preliminary unpublished data about EMILIN1-specific degradation by several enzymes abundantly present in the inflammation microenvironment. Moreover, Negrini and colleagues have recently demonstrated that fragmentation and disorganization of ECM components in the lung lead to interstitial and eventually severe edema (29).
Finally, the observations that Emilin1–/– mice display lymphatic hyperplasia and develop larger lymphangiomas associated with an increased lymphatic vessel density than their WT littermates suggest a complementary role of EMILIN1 in this context as a lymphangiogenesis modulator. This hypothesis is supported by the finding that threefold more Ki67-positive nuclei colocalizing with podoplanin-positive LECs were found in samples obtained from Emilin1–/– mice than in those from WT mice. Thus, EMILIN1 may play for LECs a regulator function similar to that of thrombospondins (TSPs) for blood endothelial cells: both TSP-1 and TSP-2 were reported to inhibit angiogenesis in vivo, contributing to the normal quiescence of blood vasculature (2, 21, 42, 43). Alternatively, the increased lymphatic-vessel density of Emilin1–/– mice may be a consequence of the higher levels of the active form of TGF-β1, since pro-TGF-β1 maturation is not adequately regulated by the absence of EMILIN1 in these mice (47). This growth factor may promote lymphangiogenesis directly, since LECs express high levels of the TGF-β1 coreceptor endoglin (31), or indirectly as a VEGF-C inducer (13). Further studies will be necessary to unveil the precise underlying molecular mechanisms.
In conclusion, the present study identifies EMILIN1 as an important component of the lymphatic perivascular elastic apparatus and demonstrates its involvement in the structure-function relationship of lymphatic vessels, as well as in lymphangiogenesis. Notably, this is the first abnormal lymphatic phenotype associated with the deficiency of an ECM protein, and we suggest that it represents a novel model of lymphatic dysfunction.
We thank Maria Teresa Mucignat for technical assistance.
Published ahead of print on 14 April 2008. ![]()
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