Digging deeper into lymphatic vessel formation in vitro and in vivo
- Benoit Detry†1,
- Françoise Bruyère†1,
- Charlotte Erpicum1,
- Jenny Paupert1,
- Françoise Lamaye3,
- Catherine Maillard1,
- Bénédicte Lenoir1,
- Jean-Michel Foidart1, 2,
- Marc Thiry†3 and
- Agnès Noël†1Email author
© Detry et al; licensee BioMed Central Ltd. 2011
Received: 22 February 2011
Accepted: 24 June 2011
Published: 24 June 2011
Abnormal lymphatic vessel formation (lymphangiogenesis) is associated with different pathologies such as cancer, lymphedema, psoriasis and graft rejection. Lymphatic vasculature displays distinctive features than blood vasculature, and mechanisms underlying the formation of new lymphatic vessels during physiological and pathological processes are still poorly documented. Most studies on lymphatic vessel formation are focused on organism development rather than lymphangiogenic events occurring in adults. We have here studied lymphatic vessel formation in two in vivo models of pathological lymphangiogenesis (corneal assay and lymphangioma). These data have been confronted to those generated in the recently set up in vitro model of lymphatic ring assay. Ultrastructural analyses through Transmission Electron Microscopy (TEM) were performed to investigate tube morphogenesis, an important differentiating process observed during endothelial cell organization into capillary structures.
In both in vivo models (lymphangiogenic corneal assay and lymphangioma), migrating lymphatic endothelial cells extended long processes exploring the neighboring environment and organized into cord-like structures. Signs of intense extracellular matrix remodeling were observed extracellularly and inside cytoplasmic vacuoles. The formation of intercellular spaces between endothelial cells led to tube formation. Proliferating lymphatic endothelial cells were detected both at the tips of sprouting capillaries and inside extending sprouts. The different steps of lymphangiogenesis observed in vivo are fully recapitulated in vitro, in the lymphatic ring assay and include: (1) endothelial cell alignment in cord like structure, (2) intracellular vacuole formation and (3) matrix degradation.
In this study, we are providing evidence for lymphatic vessel formation through tunneling relying on extensive matrix remodeling, migration and alignment of sprouting endothelial cells into tubular structures. In addition, our data emphasize the suitability of the lymphatic ring assay to unravel mechanisms underlying lymphangiogenesis.
The lymphatic vasculature functions as a tissue drainage system and an immunological control system by collecting extravasated fluid, macromolecules and leukocytes from tissues. The lymphatic system is involved in numerous pathologies such as cancer, lymphedema, inflammation and graft rejection [1–5]. It is also implicated in the dissemination of tumor cells to regional lymph nodes which results in poor prognoses of patients with cancers [6, 7]. Reflecting its specialized functions, the lymphatic vasculature displays a distinctive structure. In sharp contrast to blood vessels, the basement membrane of lymphatic vessels is discontinuous or absent. Lymphatic endothelial cells (LEC) display tight junctions and interdigitations, and are connected to the surrounding collagen fibers by anchoring filaments [8–10]. The discovery of specific markers for LECs enabled technical progress in lymphatic vascular biology and greatly promoted lymphatic research [3, 4, 11].
Although mechanisms leading to new blood vessel formation during physiological and pathological processes are well documented, how migrating LEC organized into new lymphatic vessels has long been a mystery. The prevailing view of their origin from the venous system during embryogenesis is supported by studies performed in mouse and zebrafish [12–16]. LEC could also derive from mesenchymal progenitor cells or lymphangioblasts identified in amphibian and birds through a process referred as lymphvasculogenesis [17, 18]. There is an emerging body of work concentrated on attempts to elucidate how to create tubes and generate a complex functional vascular tree [19, 20]. Tube morphogenesis is an important morphogenetic process observed during various developmental and pathological events. Regarding epithelial cells, five putative mechanisms have been proposed for tube formation and include: (1) the wrapping of a cell sheet to form a tube; (2) the budding of cells from a pre-existing tube; (3) the cavitation during which the central cells of a solid spheroidal or cylindral mass of cells are eliminated to create a tube; (4) cord hollowing generating a lumen between aggregated cells or (5) cell hollowing creating intracellular luminal spaces inside a single cell, spanning the length of the cell . Progress in understanding the processes of lumen formation (luminogenesis) has benefited from elegant studies in the zebrafish system [16, 22] and in vitro models of tubulogenesis [23, 24] and of sprouting angiogenesis in 3D extracellular matrix (ECM) environments [25, 26]. For blood vessel formation, it is now widely accepted that blood endothelial cells (BEC) at the tip of the bud (named tip cells) invade the matrix and create a space that can be occupied by a cord of cells without apparent lumen. Behind the tip cell, the so-called "stalk cells" composing the stalk of the sprouting capillary are proliferating and contribute to stalk elongation, as well as to basement membrane deposition . BEC organization along the matrix space generated by migrating cells initiates an extracellular luminal area resulting in the transformation of cord into a tube [19, 20]. Cell hollowing or intracellular vacuolization is an additional mechanism by which individual cells generate vesicles that can interconnect with adjacent cells leading to lumen size increase. In sharp contrast to those major advances made in the field of angiogenesis, little information is available on how LEC migrate and organize into lymphatic vessels during lymphangiogenesis. Although lymphatic vessels are enclosed in a matrix structure mainly composed of collagens, the extent of ECM remodeling in lymphangiogenesis is unclear. A major challenge is the difficulty of establishing appropriate in vivo models and culture systems to enable the dissection of this complex biological process. Recently, several in vivo and in vitro models of lymphangiogenesis have been developed and are useful for exploring the cellular and molecular mechanisms of lymphangiogenesis [13, 28–33].
In the present study, ultrastructural features of neoformed lymphatic vessels have been investigated in two in vivo models and one in vitro 3D culture system: (1) the corneal lymphangiogenic assay induced by thermal cauterization of the mouse cornea ; (2) the lymphangioma model consisting in lymphatic cell hyperplasia induced by intra-peritoneal injection of incomplete Freund's adjuvant [35–37] and (3) the lymphatic ring assay which bridges the gap between in vitro and in vivo systems [38, 39]. We provide innovative morphological data, at the ultrastructural level, demonstrating the pronounced ECM remodeling and intracellular vacuolization during the migration, alignment and organization of channels of sprouting lymphatic cells in vivo. Through Transmission Electron Microscopy (TEM), we show that collagen degradation takes place as an important step for vessel neoformation during lymphangiogenesis.
Induction of lymphangiogenesis in vivo
Ultrastructural features of lymphangiogenesis in vivo
The lymphatic ring assay reproduces in vitro the lymphangiogenic process
Emerging descriptions of cellular and molecular events of tubulogenesis occuring during blood vessel formation have converged on three mechanisms underlying angiogenesis: budding (or sprouting), cord hollowing and cell hollowing [19, 44, 45]. Progress in understanding such angiogenic tube morphogenesis has benefited from 3D culture systems. The present study represents the first ultrastructural description of capillary formation during pathological lymphangiogenesis. In line with the previous descriptions of the angiogenic process, we observed intracellular and extracellular hollowing events. A common feature of the three lymphangiogenic processes studied here is the migration of cells creating spaces that can be occupied by a cord of very thin and elongated cells delimitating a luminal space. In the present study, the involvement of cell proliferation has also been evidenced during cord formation. Cell hollowing or intracellular vacuolization is a mechanism by which individual cells generate vesicles that can enable the cells to interconnect with neighboring cells to form multicellular lumens and tubes [46–48]. Cell vacuolization is a common feature of migrating cells in the three models presented here. Vesicles of various sizes were seen to progressively enlarge and fuse to each other to, in turn, form a large intracellular luminal vesicle. By analogy with the angiogenic process, this space likely fuses with vesicle of adjacent cells to form the lumen of a pre-lymphatic vessel. This concept is supported by the process of cell fusion leading to increased lumen size clearly seen in the lymphangioma both at ultrastructural and histological levels (Figures 1 and 3). The intracellular vacuolization mechanism was initially associated with the morphogenesis of single endothelial cells which had no contact with adjacent cells occurring during the process of vasculogenesis [44, 49, 50]. The intracellular vacuolization has been extensively studied in tubulogenesis assay on 3D matrix leading to the identification of key molecular regulators such as matrix metalloproteinases and small GTPase [46, 47, 51]. In this context, the zebrafish system was suitable to demonstrate the importance of such process in an in vivo context during developmental conditions . The present ultrastructural investigation provides the first evidence of intracellular vacuolization in vivo during lymphangiogenesis. Further investigations are required to give new molecular insights on how this process contributes to lumen formation in lymphatic capillaries. Despite further attempts, we have been unable to set up a real-time visualization of living cells with confocal or two photons microscopes in the lymphatic ring assay.
An exciting advance in the field of angiogenesis came from the finding that several types of specialized endothelial cells (tip cells and stalk cells) are involved in the building of functional blood capillaries. It has been described that lymphatic tip cells expressed more vascular endothelial growth factor receptor-3 (VEGFR3) and neuropilin-2  but the transposition of the new concept of tip/stalk cells from angiogenic sprouts to lymphangiogenic sprouts in terms of cell proliferation is still premature. In order to shed some light on this issue, we have analyzed the proliferation rate of migrating cells in sprouting capillaries, both in vivo in the corneal assay, and in vitro in the lymphatic ring assay. Proliferation assessed by BrdU incorporation was observed both in extending capillaries and at their extremities. In the aortic ring that mimicks the angiogenic process, a quantitative analysis of proliferating cells revealed that none of the tip cells had incorporated BrdU, while 12 ± 5% of the stalk cells were BrdU positive (data not shown). These data suggest that the concept of tip cells defined as non proliferating cells probing the environment can not be extended to the process of lymphangiogenesis and emphasizes differences between the cellular mechanisms underlying lymphangiogenesis and angiogenesis.
Of great interest is our finding that LEC create in vivo, physical spaces within the surrounding collagen rich environment. This is associated with an extensive extracellular matrix remodeling both evidenced extracellular and intracellularly. Long processes extended by LEC were seen to roll up to enclose matrix fragments and create extracellular spaces. The contribution of MMPs in this remodeling process is supported by the inhibition of LEC sprouting achieved by using a synthetic MMP inhibitor. Such observation is in line with our recent identification of the metalloproteinase-2 (MMP2) which displays collagenolytic activity  as a key regulator of lymphangiogenesis . Indeed, the embedding of lymphatic duct fragments issued from MMP2-deficient mice led to impaired LEC sprouting and lymphangiogenic response . The involvement of MMP-driven proteolysis in the lymphangiogenic process is further supported by our previous work using broad spectrum MMP inhibitors in the corneal assay . It is worth noting that intracellular vacuolization and extracellular remodeling are not two exclusive mechanisms (Figure 8). They have been both evidenced in the three distinct in vitro and in vivo models used here and thus likely operate concomitantly during lymphangiogenesis. Altogether, our data emphasize the interest of the lymphatic ring assay to unravel the cellular and molecular mechanisms of lymphangiogenesis. It appropriately recapitulates in vitro the different steps of lymphangiogenesis observed in animal models such as corneal lymphangiogenesis and lymphangioma. The novel emerging panel of in vitro and in vivo models of lymphangiogenesis [13, 30] are suitable to investigate the biology of lymphangiogenesis. This is mandatory for the understanding of several pathological processes such as lymphedema, graft rejection and metastatic dissemination through the lymphatic way.
The present study provides new insights into lymphangiogenic tube formation. It also highlights the suitability of the lymphatic ring assay to investigate lymphangiogenesis associated with different pathological processes.
C57BL/6 mice of either sex, 6 to 8 weeks old, were purchased from Janvier (Saint Berthevin, France). All experimental procedures were performed in accordance to the guidelines of the University of Liège regarding the care and use of laboratory animals.
Corneal lymphangiogenesis was induced by thermal cauterization of the anesthetized central cornea (Unicaïne 0.4%, Thea Pharma, Wetteren, Belgium) by using an ophthalmic cautery (OPTEMP II V, Alcon Surgial, Fort Worth, USA) on mice anesthetised by intraperitoneal injection of ketamine hydrochloride and xylazine (100 mg/kg and 10 mg/kg, respectively) . Seven days later, mice were sacrificed, eyes were removed and corneas were isolated. In some assays, mice were intraperitoneally injected with Bromodeoxyuridine (BrdU, 200 ul) (Merck, Overijse, Belgium), 2 h before sacrifice. Corneas were stained as whole mount after 1 h fixation in ethanol 70%, at room temperature. Whole mounts were blocked in 3% BSA-3% Gloria milk for 1 h and incubated overnight with polyclonal goat anti-mouse LYVE-1 (1/200, R&D System, Abingdon, UK) or mouse anti-BrdU antibody (1/250, Becton Dickinson, Erembodegem, Belgium). After four washes with PBS, corneas were incubated, for 2 h, with Alexa-Fluor 488 coupled rabbit anti-goat antibody (1/200, Molecular Probes, Merelbeke, Belgium) or TRITC coupled rabbit anti-mouse antibody (1/40, Dako, Glostrup, Denmark). Corneas were flatmounted on a microscope slide with Vectashield mounting medium (Vector Laboratories, Burlingame, CA) and visualized by using a fluorescent microscope (AH3-RFCA, Olympus, Hamburg, Germany) or a LeicaTCS SP2 inverted confocal microscope (Leica Microsystems, Wetzl, Germany).
Lymphangioma or lymphatic endothelial hyperplasia was induced by two intraperitoneal injections of incomplete Freund's adjuvant with a 15-day interval, as described [35, 36]. For ethical purposes, buprenorphine injections (0.05 mg/kg) were administered 1 h before and after adjuvant injections, as well as every 12 h during the first 5 days post-injection. After 4 weeks, mice were killed and diaphragms were harvested, fixed in 10% formalin and paraffin embedded. Sections of 4-6 μm thickness were cut and either hematoxylin-eosin stained or immunolabeled by using an anti-LYVE-1 antibody as previously described .
Lymphatic ring assay (LRA)
Thoracic ducts used for lymphatic ring cultures were collected from male and female C57BL/6 mice. Three-dimensional lymphatic ring cultures were carried out as previously described [38, 39]. Ring-shaped explants embedded in rat tail interstitial collagen-I gel cultured in MCDB131 (GIBCO, Merelbeke, Belgium) medium supplemented with either 4% Ultroser (BioSepra, Cergy Saint Cristophe, France) or 10% Fetal Bovine Serum (FBS). Cultures were kept at 37°C in a humidified incubator (HERAcell 150, Heraeus, Hanau, Denmark) under hypoxic conditions (5% O2, 5% CO2 and 90% N2) for 11 days. In some assays, MMP inhibitor (GM6001) was added at indicated doses, in the culture medium at the beginning of the experiment. To assess cell proliferation, rings were incubated with BrdU for 3 hours before fixation and immunostaining. For the immunochemistry of whole mounted rings, cultures were fixed in ethanol 70% for staining with rabbit Lyve-1 antibody (1/600, a kind gift from Kari Alitalo, Finland) or with anti-BrdU antibody (1/250, Becton Dickinson, Erembodegem, Belgium). After washes, lymphatic rings were incubated with FITC coupled swine anti-rabbit antibody (1/40) or FITC conjugated rabbit anti-mouse antibody (1/40, both from Dako, Glostrup, Denmark). For FITC coupled phalloidin labeling (Sigma-Aldrich, Schnelldorf, Germany) rings were fixed in paraformaldehyde (4%) . Nuclei were evidenced by TO-PRO3 and Vectashield Dapi (Molecular Probe, Merelbeke, Belgium). Lymphatic capillaries were visualized under a Leica TCS SP2 confocal microscope (Leica Microsystems, Wetzl, Germany) or a fluorescent microscope (AH3-RFCA, Olympus, Hamburg, Germany). Quantification of LEC sprouting was performed by computerized-assisted method as previously described [38, 39].
Transmission Electron Microscopy (TEM)
Samples (lymphatic ring gels, lymphangioma or cornea) were washed in Sörensen's buffer and fixed for 1 h at 4°C with 2.5% glutaraldehyde in a Sörensen 0.1 M phosphate buffer (pH 7.4) and post-fixed for 30 min with 1% osmium tetroxide. After dehydration in graded ethanol, samples were embedded in Epon. Ultrathin sections obtained with a Reichert Ultracut S ultramicrotome were contrasted with uranyl acetate and lead citrate. Observations were made with a Jeol 100 CX II transmission electron microscope at 60 kV.
The authors acknowledge G. Roland, M. Dehuy, N. Decloux, I. Dasoul, E. Feyereisen, L. Poma and P. Gavitelli for their excellent technical assistance. We are grateful to the help of S. Ormenese (GIGA Imaging plateform). We thank the GIGA plateforms for their contribution. This work was supported by grants from the European Union Framework Program projects (FP7-2007-2011 "MICROENVIMET" No 201279), the Fonds de la Recherche Scientifique Médicale, the Fonds National de la Recherche Scientifique (F.N.R.S., Belgium), the Fondation contre le Cancer, the Fonds spéciaux de la Recherche (University of Liège), the Centre Anticancéreux près l'Université de Liège, the Fonds Léon Fredericq (University of Liège), the D.G.T.R.E. from the « Région Wallonne », the Interuniversity Attraction Poles Programme - Belgian Science Policy (Brussels, Belgium). BD and CE are recipients of a Televie-FNRS grant.
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