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Lymphangiogenesis Lymphedema and Cancer


Angiogenesis (blood vessel growth), lymphangiogenesis (lymph system growth) are all intrinsically connected with lymphedema and share many of the same genes. We have several pages on both processes.

Pat O'Connor

May 23, 2008


The role of tumor lymphangiogenesis in metastatic spread

(The FASEB Journal. 2002;16:922-934.)
© 2002 FASEB


Ludwig Institute for Cancer Research, Royal Melbourne Hospital, Victoria 3050, Australia

1Correspondence: Ludwig Institute for Cancer Research, Post Office Box 2008, Royal Melbourne Hospital, Victoria 3050, Australia. E-mail:


The high mortality rates associated with cancer can be attributed to the metastatic spread of tumor cells from the site of their origin. Tumor cells invade either the blood or lymphatic vessels to access the general circulation and then establish themselves in other tissues. Clinicopathological data suggest that the lymphatics are an initial route for the spread of solid tumors. Detection of sentinel lymph nodes by biopsy provides significant information for staging and designing therapeutic regimens. The role of angiogenesis in facilitating the growth of solid tumors has been well established, but the presence of lymphatic vessels and the relevance of lymphangiogenesis to tumor spread are less clear. Recently, the molecular pathway that signals for lymphangiogenesis and relatively specific markers for lymphatic endothelium have been described allowing analyses of tumor lymphangiogenesis to be performed in animal models. These studies demonstrate that tumor lymphangiogenesis is a major component of the metastatic process and implicate members of the VEGF family of growth factors as key mediators of lymphangiogenesis in both normal biology and tumors.—Stacker, S. A., Baldwin, M. E., Achen, M. G. The role of tumor lymphangiogenesis in metastatic spread.

Key Words: metastasis • VEGF-C • VEGF-D • VEGF receptors


EACH YEAR IN the United States more than 500,000 people die principally as a result of the metastatic spread of cancer (1 , 2) . Cells from malignant primary tumors spread from their sites of origin to invade local tissue and enter the systemic circulation (3 , 4) . This spread can occur directly into the local tissue or via blood vessels (hematogenous spread) and lymphatics (lymphogenous spread) or by invasion of body cavities such as the pleura or peritoneum. Cells must first invade either blood or lymphatic vessels to enter the circulation. In the case of blood vessels, this requires penetration of the basement membrane and migration through the cellular layers of the vessel (Fig. 1 ). In contrast, it has been proposed that the entry of tumor cells into the lymphatic circulation may be easier due to the nature of lymphatic vessels and that the thin and discontinuous basement membrane of lymphatics (5 6 7) might not provide a significant barrier to entry of tumor cells.

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Figure 1. Lymphatic vessel structure and potential modes of tumor cell dissemination. Lymphatics are thin-walled, low-pressure vessels that collect fluid and cells from the interstitium and return it to the circulation via the thoracic duct. In contrast, blood vessels are subject to high pressure and have a more robust structure with a well-defined basement membrane and supporting cells. Tumor cells leave the primary tumor and spread directly into the surrounding tissue, from which they subsequently invade preexisting blood vessels or lymphatic vessels. Alternatively, the formation of new blood vessels (angiogenesis) or new lymphatic vessels (lymphangiogenesis), induced by tumor-associated angiogenic and/or lymphangiogenic factors, could promote the growth of new vessels into the tumor, providing an escape route for tumor cells (diagram shows single tumor cells leaving the primary mass and entering the lymphatic or vascular system). Cells that invade the lymphatics could find their way into the bloodstream via the thoracic duct (sequential lymphovascular spread) or by the formation of shunts, speculated to exist in tumors, between the vascular system and the lymphatic system. Retrograde spread from blocked lymphatic vessels could be speculated as a means for tumor cells to travel upstream and into the venous system (18) .

Tumor cells can escape from the primary site by entering existing vessels or new vessels actively recruited into the primary tumor (4) . The relative importance of the established vessels vs. the active invasion of a tumor by new blood and lymphatic vessels for the initial metastatic spread of tumor cells is still unclear. Previous studies have established the role of angiogenesis in solid tumor growth (8 9 10 11 12 13 14) , and there is some evidence indicating a direct role for angiogenesis in hematogenous tumor spread (15) . The onset of angiogenesis within small clusters of tumors cells, known as the ‘angiogenic-switch’, is important for blood vessel formation in further development of malignant cells that have already spread from the primary tumor (16 , 17) . However, little is known about the role of tumor lymphangiogenesis (the growth and production of new lymphatic vessels) in the spread of tumors and whether this process is important in the overall context of lymphatic spread (18 , 19) .

Clinical and pathological data point to the spread of solid tumors via the lymphatics as an important early event in metastatic disease (1 , 3) . Detection of tumor cells in lymphatic vessels and regional lymph nodes is a key factor in the staging of human tumors and forms the basis for treatment of regional lymph nodes by surgery and radiation therapy (1 , 3 , 20 , 21) . Although a large body of clinical and pathological evidence points to a major role for the lymphatics in the initial spread of malignant tumors, the exact mechanism whereby tumor cells enter the lymphatic system is uncertain (18 , 22 , 23) .

The role of lymphangiogenesis in promoting the metastatic spread of tumor cells via the lymphatics has been an area that has achieved little publicity in the past decade. This has in part been due to difficulty in studying lymphatic vessels because of their morphology and a lack of lymphatic-specific markers (24) . In reviewing this topic, we will emphasize the progress made in the past 2 years in identifying novel lymphatic markers, as well as lymphangiogenic factors and receptors responsible for the generation and maintenance of the lymphatic system. This will include a summary of the recent clinical and experimental data that support the notion that lymphangiogenic factors influence the spread of tumors and a discussion of the potential therapeutic options for anti-lymphangiogenic treatment of cancer.


The lymphatic system consists of thin-walled, low-pressure vessels, nodes that occur along the course of lymphatic vessels, aggregations of lymphoid tissue, such as the spleen and thymus, and circulating lymphocytes (5 , 6 , 25) . By regulating fluid absorption from the interstitium, harboring macrophages, and providing a passage for lymphocyte trafficking, the lymphatic system maintains plasma volume, prevents increases in tissue pressure, and has an important role in immune system function (26) . Lymphatic vessels are distinct functionally and ultrastructurally from their blood vessel counterparts (27) . Compared to blood vessels, the walls of lymphatic vessels are thinner. This is in part due to the highly attenuated cytoplasm of lymphatic endothelial cells (Fig. 1) (5 , 6 , 27 , 28) . The endothelium of lymphatic vessels contains fewer tight junctions than that of blood vessels and it has been speculated that this may be the cause of the greater permeability of the lymphatic vessels (25) . Lymphatic capillaries have a poorly developed or absent basal lamina and lack associated pericytes (5 , 6) . Their lumens are ~ threefold wider than the lumens of blood capillaries and are more irregularly shaped, appearing collapsed in tissue section (5 , 28 , 29) . Lymphatic vessels are connected to the extracellular matrix by reticular fibers and collagen (5 , 6 , 30) . Upon increases in interstitial fluid and pressure, the connecting tissue fibers become stretched, thereby opening the lymphatic lumen (27 , 29 , 31) . As the lumen widens, the endothelial cells, which overlap under normal conditions, move apart, effectively opening intercellular channels to aid fluid and macromolecular uptake into the lymphatic vessel (5 , 6 , 29 , 30) (Fig. 1) .


The growth of lymphatic vessels, lymphangiogenesis, has received considerable attention in the last 2 years, due to the identification of proteins specifically expressed on lymphatic vessels and the discovery of molecules that can drive lymphatic vessel growth (32 33 34 35 36 37 38) . Vascular remodeling associated with lymphangiogenesis and angiogenesis is likely to involve similar processes, although formal evidence of this assertion has yet to be published. In response to molecular mediators, both lymphatic and vascular endothelial cells proliferate and migrate toward a stimulus as the extracellular matrix is degraded, followed by association of the endothelial cells into tube-like structures (31 , 39) . New production and realignment of the extracellular matrix and controlled apoptosis at appropriate sites are required for blood vascular and lymphatic system formation. Besides using similar processes of remodeling, blood and lymphatic vessels are closely associated in vivo. Blood vascular plexuses often accompany lymphatic vessels (27 , 40) , although the ratio of lymphatic to blood vessels varies depending on tissue type and function (29) . An abundant and neighboring blood supply provides essential nourishment for lymphatic vessels that is needed for adequate function, intrinsic contractility of lymphatic endothelial cells, and an ability to regenerate rapidly when required, processes essential for maintaining fluid balance within an organism (31) . The close association of blood and lymphatic vessels and their coordinated development in vivo suggest that some molecules may control both angiogenesis and lymphangiogenesis (41) .


Progress in understanding lymphangiogenesis has been hampered by the very similar characteristics of blood and lymphatic vessels in tissue section and is confounded by the lack of lymphatic-specific markers (24) . Consequently, visualization of lymphatic vessels in the past was restricted to imaging techniques involving the injection of dyes that are specifically taken up by the lymphatics (reviewed in refs 29 , 31 ). Vital dyes, such as Evans blue, trypan blue, and Patent blue, which are readily taken up by lymphatic but not blood vessels, are less toxic than the materials that were previously used (29) . These dyes and fluorescent conjugates of high molecular weight material such as rhodamine-dextran are now used routinely in animal experiments (37 , 42) . Until recently, immunohistochemical identification of lymphatic vessels was achieved, somewhat unreliably, by comparing staining of pan-endothelial markers with markers of the basal lamina. The pan-endothelial marker PECAM-1/CD31, which is expressed on both blood and lymphatic vessels (43 , 44) , has been used in combination with the basement membrane markers laminin and collagen type IV (45 , 46) . Vessels that reacted with PECAM-1 antibodies but lacked basement membrane staining and red blood cells in their lumens were deemed lymphatic (44 , 47) . Use of the blood vessel-specific marker PAL-E in combination with PECAM-1 has also been useful for identifying lymphatic vessels in human tissue sections (44 , 47) .

More accurate and simplified lymphatic vessel identification has recently been made possible by the discovery of molecules that are specifically expressed by lymphatic endothelium (Table 1) . Vascular endothelial growth factor receptor-3 (VEGFR-3) is predominantly expressed on lymphatic endothelium in normal adult tissues (48 , 49) ; it is also up-regulated on blood vessel endothelium in tumors (44 , 50) and in wound healing (51) . The lymphatic receptor for hyaluronan, LYVE-1, has been reported to be a specific marker of lymphatic vessels (52 , 53) and is thought to function in transporting hyaluronan from the tissue to the lymph (53 , 54) . Antibodies to LYVE-1 have been used to localize the receptor to lymphatic endothelium in normal and tumor tissue (38 , 52) . Although appearing relatively specific for lymphatic endothelium, staining of blood vessels in normal lung tissue has been observed (R. A. Williams, S. A. Stacker, and M. G. Achen, unpublished observations) as has staining of blood vessels in normal hepatic blood sinusoidal endothelial cells (55) . The transcription factor Prox1, although required for lymphatic vessel development and expressed on lymphatic endothelium (56) , is also expressed in other cell types and tissues, including hepatocytes of the liver (57) and lens tissue (58) , and is therefore of limited use immunohistochemically to identify lymphatic vessels. Podoplanin and desmoplakin have been reported to be markers for lymphatic endothelium, but also react with other cell types (59 60 61) . In summary, a more extensive range of markers for lymphatic endothelium is now available that should aid in defining the role of lymphatic vessels in tumor biology.

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Table 1. Markers for discrimination of lymphatics and blood vessels



Recently VEGF-C and VEGF-D, members of the VEGF family of secreted glycoproteins that are ligands for VEGFR-3 (Flt4) (34 , 62 63 64) (Fig. 2 , Table 2 ), have been identified as regulators of lymphangiogenesis in mammals (32 , 42) . Structurally, these protein growth factors differ from the angiogenic growth factor VEGF (65) because of the presence of amino- and carboxyl-terminal propeptides, but retain the central VEGF homology domain (VHD) containing the cystine knot motif that is conserved in all VEGF family members (66 , 67) . Biosynthetic processing of the VEGF-C and VEGF-D polypeptides, by as yet uncharacterized extracellular proteases, results in mature growth factor consisting of dimers of the VHD that bind VEGFR-3 and VEGFR-2 (KDR/Flk-1) with high affinity (68 , 69) . The unprocessed and partially processed forms of VEGF-C and VEGF-D have reduced affinity for both receptors, indicating that processing is important for receptor binding. VEGF-C and VEGF-D are mitogenic for lymphatic and vascular endothelial cells in vitro (34 , 68 , 70) , and VEGF-C (62 , 71) , but not VEGF-D (72) , can induce vascular permeability. Analysis of VEGF-C and VEGF-D function in vivo and in vitro using a range of animal-based assay systems, including the chick chorioallantoic membrane, rabbit cornea assays, and transgenic mouse models, has demonstrated the ability of these factors to drive angiogenesis and lymphangiogenesis (Table 2) (32 33 34 , 37 , 38 , 73 74 75)

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Figure 2. Receptor binding specificities of VEGF family members. VEGF receptors are shown spanning the plasma membrane. VEGFR-1, VEGFR-2, and VEGFR-3 are structurally homologous and consist of seven immunoglobulin homology domains in the extracellular region and a tyrosine kinase domain in the intracellular portion that is interrupted by a tyrosine kinase insert domain. A soluble form of VEGFR-1 exists (125) but is not shown. The extracellular domain of VEGFR-3 is proteolytically cleaved in the fifth immunoglobulin-like domain and the fragments remain associated by disulfide bonds (S-S) (126 , 127) . Neuropilin 1 consists of a short intracellular domain and an extracellular domain containing two complement C1r/s homology domains, two domains with homology to coagulation factors V and VIII, and a single MAM domain (128) . Neuropillin-2, which also binds VEGF, is not shown here. The VEGF family members (represented as dimers) that interact with each receptor are indicated at the top of the figure and are represented in the diagram as dimers bound to the receptors. The biological consequence of signaling through VEGFR-1 is not fully understood whereas activation of VEGFR-2 and VEGFR-3 signals predominately for angiogenesis and lymphangiogenesis, respectively. Nonetheless, it is apparent that VEGFR-2 is also present on lymphatic endothelium and that VEGFR-3 can be expressed on the endothelium of tumor blood vessels. pp = proteolytically processed
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Table 2. Growth factors and receptors involved in lymphangiogenesis

VEGF-C gene expression is induced by a range of growth factors, including platelet-derived growth factor and epidermal growth factor (76) , and by numerous proinflammatory cytokines (77) . Such mediators may be responsible for the induction of VEGF-C gene expression observed in a wide range of human tumors. In contrast to VEGF, expression of VEGF-C is not induced by hypoxia (76) . Expression of the X-linked VEGF-D gene (78) is induced by the transcription factor c-fos (79) and by signaling resulting from cell–cell contact that is dependent on cadherin 11 (80) . As c-fos is induced in a range of human tumors and many tumors are characterized by high cell density, these forms of regulation could induce expression of VEGF-D in tumor cells. VEGF-D has been reported to be regulated by the transcription factor AP-1 in human glioblastoma multiforme (81) .

The receptor for VEGF-C/VEGF-D with specificity for lymphatic endothelium is VEGFR-3 (34 , 62) . This receptor is expressed on venous endothelium at sites of lymphatic vessel sprouting during embryogenesis and, in adults, becomes restricted to lymphatic endothelium (48 , 49) . VEGFR-3 has been shown to play an important role in remodeling and maturation of the primary capillary plexus in the early embryo (82) , and VEGFR-3 mutations have been associated with hereditary lymphedema (83) . Recent studies using a VEGF-C mutant that binds VEGFR-3 but not VEGFR-2 demonstrated that activation of VEGFR-3 is sufficient to induce growth of lymphatics (37) . Studies using a soluble form of the VEGFR-3 extracellular domain expressed as a transgene under the control of the keratin-14 (skin) promotor have demonstrated the requirement for VEGF-C and VEGF-D and, by implication, VEGFR-3 signaling, in lymphangiogenesis (42 , 84) , although the blood vasculature was unaffected (42) . Evidence is now emerging from a range of such studies which suggests that VEGFR-2 is the primary receptor for angiogenesis whereas VEGFR-3 regulates lymphangiogenesis.


The metastatic spread of tumor cells is the underlying cause of most cancer-related deaths (2 , 3) . Clinical and pathological evidence confirms that the metastatic spread of tumors via lymphatic vessels to local/regional lymph nodes is an early event in metastatic disease for many solid human tumors (3 , 20 , 21) . The presence of tumor cells in local lymph nodes is a significant factor in the staging of human tumors and forms the basis for surgical and radiation treatment of regional lymph nodes (3 , 20 , 21 , 84) . More recently, the use of sentinel lymph nodes has developed as a promising method for the diagnosis and staging of such diseases as breast cancer and melanoma (85 86 87) .

Much conjecture exists in the literature regarding the existence of lymphatics within tumors (18 , 22 , 31 , 88) . Until recently, evidence linking the presence of lymphatic vessels in solid tumors with the spread of cancer was not compelling (18) . This was due to the lack of suitable markers to distinguish blood vessels from lymphatic vessels, the difficulty in identifying these vessels by injection techniques, and the poorly defined structure of these vessels. Various reports of the high interstitial pressure in tumors have been used as a theoretical basis for assuming there is a lack of functional lymphatic vessels within the tumor mass (88) . Recent studies using a sarcoma model demonstrated the lack of functional lymphatic vessels in a tumor expressing the lymphangiogenic factor VEGF-C and its receptor, VEGFR-3. The conclusion of that study was that the physical stress exerted by the growing tumor cells caused the collapse of the lymphatic vessels (88) . In contrast, there are many historical reports of lymphatic vessels in solid tumors and anecdotal evidence that many tumors are not edematous in nature, suggesting the existence of functional lymphatic vessels in tumors (31 , 89) . The recent explosion of interest in the development of blood vessels within tumors did for a time overshadow the need for further study of tumor lymphangiogenesis


The discovery of the lymphangiogenic factors VEGF-C and VEGF-D raises the question as to whether these factors are expressed in human cancers and whether this expression is responsible for the ability of tumors to metastasize. It is already clear from a number of studies that members of the VEGF family are expressed in a variety of human tumors (90 91 92 93 94) . VEGF-C and VEGF-D expression has been detected in a range of human tumors including malignant melanoma and lung, breast, colorectal, and gastric carcinomas (see Table 3 ) (91 , 92 , 94) . These studies have used either immunohistochemistry or reverse transcription polymerase chain reaction (RT-PCR) to detect expression of the genes, and these techniques do not take into account the need for proteolytic cleavage to activate the polypeptides. The detection of mRNA or full-length protein may not in all circumstances reflect fully active/mature growth factor. Expression of VEGFR-3 is also an important factor in determining the potential for a lymphangiogenic response. Some studies have shown the coexpression of VEGF-C/D with VEGFR-3 in malignant melanoma (94) and lung cancer (95)

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Table 3. Clinical data showing a relationship between expression of the lymphangiogenic growth factors VEGF-C and VEGF-D and the metastatic spread of tumors

Tumor expression studies (Table 3) have allowed a direct comparison of VEGF-C and VEGF-D expression with clinicopathological factors that relate directly to the ability of primary tumors to spread (e.g., lymph node involvement, lymphatic invasion, secondary metastases, and disease-free survival) (95 96 97 98) . In many instances, these studies demonstrate a statistical correlation between the expression of lymphangiogenic factors and the ability of a primary solid tumor to spread. For example, levels of VEGF-C mRNA in adenocarcinoma of the lung are associated with lymph node metastasis (99) and in breast cancer correlate with lymphatic vessel invasion and shorter disease-free survival (100) . Non-small cell lung cancer showed a significantly increased survival of patients with tumors lacking VEGF-C compared to VEGF-C-positive tumors (101) . Studies by Hashimoto et al. examining samples of cervical cancer demonstrated that the level of VEGF-C mRNA detected by RT-PCR was the sole independent factor influencing pelvic lymph node metastases (102) . The majority of these studies showed significant correlation between VEGF-C levels and the clinical parameters of tumor spread (see Table 3 ), indicating the close association between expression of this lymphangiogenic factor and tumor metastasis. Some studies have suggested that expression of VEGF-D in human tumors is reduced relative to VEGF-C (99 , 103) ; further studies are required to examine this potential difference.

Although studies showing a correlation between expression levels of VEGF-C mRNA and clinical parameters are suggestive of a role for lymphangiogenic factors in promoting tumor spread, no direct demonstration of VEGF-C or VEGF-D involvement had been documented until recently. Studies using expression of VEGF-C and VEGF-D polypeptides in various tumor backgrounds in animal models have provided direct evidence that these factors can indeed promote tumor lymphangiogenesis and spread (Table 4) . Three studies in which full-length VEGF-C was overexpressed in tumor cells showed that the presence of VEGF-C polypeptide induced the formation of lymphatic metastases in regional lymph nodes (see Table 4 ). A study by Skobe et al. showed that expression of VEGF-C in the breast cancer cell line MBA-MD-435 induced increased lymphangiogenesis, but not angiogenesis, in tumors grown in immunocompromised mice (36) . These tumors spread to local lymph nodes and the lung whereas control tumors lacking VEGF-C did not, demonstrating that VEGF-C could drive metastatic spread. Others have used Rip1Tag2 transgenic mice to analyze the activity of overexpressed VEGF-C in a pancreatic ß cell tumor model (35) . In these tumors, VEGF-C promoted the development of peri-tumoral lymphatics and this correlated with an increased rate of metastatic spread to the draining pancreatic lymph nodes. Studies in which the breast tumor cell line MCF-7 expressing recombinant VEGF-C was implanted orthotopically in SCID mice have shown the role played by VEGF-C in promoting tumor growth (104) . Unlike lines expressing VEGF, which showed increased angiogenesis, the VEGF-C expressing lines promoted growth of only tumor-associated lymphatics. Inhibition of this growth by soluble VEGFR-3 protein showed the potential for tumor growth and metastasis to be inhibited with reagents that block VEGFR-3 signaling (104) . Other studies have used the melanoma cell line MeWo to show the effects of VEGF-C overexpression on the metastatic behavior of tumors grown in vivo (105) . These studies showed that MeWo cells expressing VEGF-C induced increased levels of both lymphangiogenesis and angiogenesis, which is in contrast to previous experiments in which VEGF-C was expressed in tumor models. This may reflect the degree of proteolytic processing of the growth factor, and was also seen in an earlier study analyzing the effect of VEGF-D (19 , 38) . This study also reported the recruitment of macrophages into the tumors as a result of overexpression of VEGF-C. This reveals a potential function of VEGF-C and VEGF-D as immune modulators, a role that has already been demonstrated for the related angiogenic factor VEGF (106) .

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Table 4. Experimental data showing a relationship between expression of lymphangiogenic growth factors VEGF-C and VEGF-D and the metastatic spread of tumors

Our study in which cells overexpressing VEGF-D were grown as tumors in SCID/NOD mice showed that VEGF-D was capable of inducing both lymphangiogenesis and the spread of tumor cells to local lymph nodes (38) . This was not seen in tumors expressing the angiogenic factor VEGF, which only induced the formation of additional blood vessels and did not induce spread of the tumor to local lymph nodes. The dependence of these effects on VEGF-D was conclusively demonstrated by the use of a monoclonal antibody (mAb) that blocked the binding of VEGF-D to both VEGFR-2 and VEGFR-3. This antibody inhibited the growth and lymphogenous spread of tumors expressing VEGF-D. In contrast to some of the studies with VEGF-C (36) , overexpression of VEGF-D resulted in an increased angiogenic response in the tumors, presumably via stimulation of VEGFR-2. The difference between the studies could be due to the degree of processing of the full-length forms of the polypeptides in the various models, which may in turn influence the receptor specificity of the growth factors. Identification of the proteases that cleave VEGF-C and VEGF-D will be an important step forward in understanding how these growth factors induce tumor angiogenesis and lymphangiogenesis. It will be important to elucidate the biological function of the propeptides of VEGF-C and VEGF-D and their role in influencing the balance between angiogenesis and lymphangiogenesis. The experimental studies carried out so far do show that the expression of growth factors such as VEGF-C and VEGF-D can directly influence the development of lymphatic vessels within tumors and the rate of metastatic spread. In combination with the expression data derived from various human tumors (Table 3) , these studies allow us to postulate that expression of these growth factors and the proteases that activate them may be critical for determining the metastatic potential of a tumor. However, molecules other than those of the VEGF family are likely to play a role in the development and growth of lymphatic endothelium.


Recent experimental approaches with animal models and analysis of genetic lesions causing hereditary lymphedema in humans have indicated that the VEGF-C/VEGF-D/VEGFR-3 signaling system drives lymphatic hyperplasia and/or lymphangiogenesis during embryonic development (32 , 33 , 37 , 42 , 83 , 107) in adult tissues (108) and in or around tumors (35 , 36 , 38 , 104) . Therefore, manipulation of this pathway offers the opportunity for therapeutic strategies designed to inhibit or stimulate growth of lymphatic vessels in conditions such as lymphedema, cancer and infectious diseases.

Inhibition of lymphangiogenesis
In the context of cancer, it may be beneficial to inhibit lymphangiogenesis so as to reduce the occurrence of lymphogenous metastatic spread. Potential inhibitors of the VEGFR-3 lymphangiogenic signaling pathway include mAbs that block the binding of VEGF-C and VEGF-D to VEGFR-3. A neutralizing VEGF-D mAb that blocks binding to both VEGFR-2 and VEGFR-3 (109) inhibited angiogenesis, lymphangiogenesis, and metastatic spread via the lymphatics in a mouse tumor model that secreted recombinant VEGF-D (38) . Similar studies using neutralizing VEGF-C mAbs have not yet been reported. A mAb against mouse VEGFR-3 was found to block the binding of VEGF-C and presumably VEGF-D. This mAb induced microhemorrhage from tumor blood vessels in a mouse tumor model, although the effects on tumor lymphatics were not analyzed (110) .

An alternative approach to antibodies would be to sequester VEGF-C and VEGF-D with a soluble version of the extracellular domain of VEGFR-3. The potential of this approach is illustrated by a transgenic mouse study in which a soluble form of the ligand binding region of the VEGFR-3 extracellular domain was expressed in the epidermis of the skin (42) . This protein construct consisted of the first three immunoglobulin homology domains of VEGFR-3 fused to the Fc-domain of the human immunoglobulin {gamma} chain. This soluble form of VEGFR-3 inhibited fetal lymphangiogenesis; consequently, these mice developed a lymphedema-like phenotype involving swelling of the feet, edema, and dermal fibrosis (42) . When delivered via an adenovirus, this soluble form of VEGFR-3 blocked the growth of peritumoral lymphatic vessels in a mouse breast cancer model (104) .

An attractive approach for inhibiting the VEGFR-3 signaling pathway would involve identification of orally active small molecules that interfere with the binding of VEGF-C/D to this receptor. Rational design of such compounds will be aided by defining the structure of the complex consisting of the VEGFR-3 ligand binding domain bound to VEGF-C/D. However, peptidomimetic approaches based on the structures of the ligands or receptor, as modeled from the known structure of the VEGF/VEGFR-1 complex (111) , could also be pursued to generate such inhibitors. Small molecule inhibitors of the tyrosine kinase catalytic domain of VEGFR-3 could be useful for blocking this signaling pathway. Tyrosine kinase inhibitors of the closely related VEGFR-2 have shown promise for inhibition of tumor angiogenesis, at least in animal models (112 , 113) . As the catalytic domains of VEGFR-2 and VEGFR-3 are closely related in structure, it is important to test all known VEGFR-2 tyrosine kinase inhibitors for activity against VEGFR-3, although it is already known that one such inhibitor, PTK787/ZK 222584, inhibits both receptors (113) . A series of indolinones has recently been reported that inhibit the kinase activity of VEGFR-3 but not VEGFR-2 (114) .

Inhibitors of the VEGFR-3 signaling pathway may be useful anti-cancer therapeutics via mechanisms other than blocking lymphangiogenesis. For example, Kaposi’s sarcoma (KS) is characterized by the presence of a core of spindle-shaped cells that may be derived from lymphatic endothelium. VEGF-C potently induces proliferation of these cells in vitro (115) and may play a critical role in controlling KS cell growth, migration, or invasion (116) . If this is the case, inhibition of VEGFR-3 signaling may be useful for inhibiting KS progression by acting on tumor cells directly. This may also be true for lymphangiomas and lymphangiosarcomas.

One potential problem that could arise from targeting the VEGFR-3 signaling pathway in cancer is that lymphatic vessel function in normal tissues could be compromised if VEGFR-3 signaling is required for the integrity/function of mature lymphatics. The role of VEGFR-3 in mature lymphatics, which express this marker, is unknown.

Therapeutic lymphangiogenesis
Lymphedema is an impairment of lymph flow from an extremity that may be caused by lymphatic vessel obstruction, ablation, lymphatic insufficiency or dysfunction, or parasitic (filariasis) infection (31 , 117) . Affected areas become swollen and fibrotic as a consequence of accumulation of fluid and insufficient protein and macromolecular uptake by lymph node macrophages (118) . Another common cause of lymphedema is the removal of the entire breast (mastectomy) and axillary lymph nodes (radical mastectomy) for breast cancer treatment (119) . Removal of the axillary lymph nodes impairs fluid clearance from the upper region of the chest and arm (117) . Significant edema of the arm occurs in ~20% of patients who undergo mastectomy and axillary dissection, and is more common in patients who undergo radiotherapy or suffer postoperative infection (117) . Late-onset or secondary edema may occur in radical mastectomy patients as a consequence of infection that affects lymphatic drainage. The use of limbs can be severely affected by lymphedema (117) . It has been proposed that mastectomy patients may benefit from stimulation of lymphangiogenesis in the region of lymph node removal to aid fluid drainage and prevent side effects associated with breast cancer. Therapeutic approaches to achieve this could be based on gene therapy or direct protein application to administer VEGF-C or VEGF-D to affected sites. Alternatively, lymphatic tissue could be transplanted from internal sites to affected skin and VEGF-C/D administered to facilitate lymphangiogenesis from the transplanted tissue. One potential danger associated with such approaches, at least in patients who have had surgery to remove primary tumors and affected lymphatics, is that lymphangiogenesis occurring near the site of tumor removal could facilitate metastatic spread from small islands of remaining tumor. It will be important to test these approaches in appropriate animal models of lymphedema and metastatic spread before proceeding to clinical trials.


The metastatic spread of tumor cells causes the vast majority of cancer deaths. Clinicopathological analyses long ago indicated that lymphatic vessels play a very important role in the metastatic spread of cancer. Therefore, metastatic spread to lymph nodes is considered a prognostic indicator and in part determines the therapeutic approaches used for treatment. Nevertheless, it has not been clear whether tumors induce lymphangiogenesis, which facilitates metastatic spread, or whether such spread occurs via preexisting lymphatics.

Recent studies using mouse tumor models expressing VEGF-C and VEGF-D indicate that these growth factors can induce hyperplasia of peritumoral lymphatics, as well as formation of intratumoral lymphatics (35 , 36 , 38 , 104) , and that these lymphatics facilitate metastatic spread to lymph nodes (35 , 36 , 38) . VEGF-C expression in some human tumors correlates with lymphangiogenesis and dissemination of tumor cells to lymph nodes (120 121 122) . VEGFR-3 can be up-regulated on tumor blood vessels (92) ; a study using a neutralizing VEGFR-3 mAb indicated that signaling via this receptor may be critical for blood vessel integrity in cancer (110) . Therefore, the VEGF-C/VEGF-D/VEGFR-3 signaling system for lymphangiogenesis constitutes a potential new target for development of anti-cancer therapeutics.

The development of approaches to block tumor lymphangiogenesis and treat lymphedema would benefit from the availability of markers specific for lymphatic endothelium. The absence of such markers has been a major problem in the field until recently. However, the advent of lymphatic markers such as VEGFR-3 (44 , 49) , podoplanin (59) , prox-1 (56) , and LYVE-1 (52 , 123) will give researchers a much better opportunity to monitor the effects of potential therapeutics on lymphangiogenesis in tumors and normal tissues. Additional requirements for progress in this field are animal models of lymphangiogenesis, lymphogenous metastatic spread, and lymphedema. Fortunately, progress has been made. A recombinant adenovirus encoding VEGF-C that induces lymphangiogenesis in a rodent model was recently reported (108) and mouse models of lymphedema are now available: the so-called ‘Chy’ mouse, which has an inactivating Vegfr3 mutation in its germline (124) , and another mouse model in which expression of a soluble form of VEGFR-3 in skin blocks fetal lymphangiogenesis (42) . Thus, many of the requisite tools for analysis of therapeutics designed to inhibit or stimulate lymphangiogenesis are now available. Clearly this is a field that will experience rapid progress in the near future.


Note added in proof: Very recent findings have further illustrated the relevance of lymphangiogenesis and VEGF-D to human cancer. Beasley et al. (Cancer Res. 62, 1315–1320, 2002) identified lymphangiogenesis occurring in human head and neck cancer and White et al. (Cancer Res. 62, 1669–1675, 2002) reported that VEGF-D is an independent prognostic indicator for survival in colorectal cancer.

This work was supported by the National Health and Medical Research Council of Australia and the Anti-Cancer Council of Victoria. M.B. is a recipient of the Australian Post-Graduate Research Award and the Anti-Cancer Council of Victoria Post-Doctoral Fellowship. We thank Prof. Tony Burgess for critical analysis of this manuscript and Janna Stickland for assistance with figures



The Formation of Lymphatic Vessels and Its Importance in the Setting of Malignancy


Published 16 September 2002 as 10.1084/jem.20021346

Rockefeller University Press, 0022-1007/2002/9/713/ $5.00
The Journal of Experimental Medicine, Volume 196, Number 6, September 16, 2002 713-718

Michael Detmar and Satoshi Hirakawa

Cutaneous Biology Research Center and Department of Dermatology, Massachusetts General Hospital and Harvard Medical School, Charlestown, MA 02129

Address correspondence to Michael Detmar, CBRC/Department of Dermatology, Massachusetts General Hospital, Building 149, 13th Street, Charlestown, MA 02129. Phone: 617-724-1170; Fax: 617-726-4453; E-mail:

The lymphatic vascular system plays important roles in the maintenance of tissue fluid homeostasis, in the mediation of the afferent immune response, and in the metastatic spread of malignant tumors to regional lymph nodes. It consists of a dense network of blind ending, thin-walled lymphatic capillaries and collecting lymphatics that drain extravasated protein-rich fluid from most organs and transport the lymph via the thoracic duct to the venous circulation (1). Originally discovered as "milky veins" by Gasparo Aselli in the 17th century (2), the mechanisms controlling the normal development of lymphatic vessels and the molecular regulation of their biological function have remained poorly understood in contrast to the rapid progress made in elucidating the formation and molecular control of the blood vascular system (3, 4).

100 yr ago, Florence Sabin proposed that the lymphatic system develops by the sprouting of endothelial cells from embryonic veins, leading to the formation of primitive lymph sacs from which lymphatic endothelial cells then sprout into surrounding organs to form mature lymphatic networks (5, 6). Since these pioneering studies, however, the field of lymphatic research has remained rather neglected, mainly due to the lack of molecular tools to specifically detect and functionally characterize the lymphatic endothelium. The recent identification of several new markers for lymphatic endothelial cells and of lymphatic growth factors and receptors, together with the characterization of genetic mouse models with impaired lymphatic development and/or function, has now led to a "rediscovery" of the lymphatic vascular system and has provided important new insights into the molecular mechanisms that control its development and biological function (7). Importantly, these studies have largely confirmed Sabin's original hypothesis regarding lymphatic development in the mammalian system (Fig. 1).

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Figure 1. Hypothetical model of the distinct steps involved in the embryonic development of the mammalian lymphatic vasculature.


Recently, Wigle and Oliver (8) and Wigle et al. (9) have identified the first gene that is essential for early lymphatic development. Beginning at E9.5 of mouse development, the homeobox gene Prox1 starts to become specifically expressed in a subpopulation of endothelial cells located on one side of the anterior cardinal vein (8). At this stage, the venous endothelium also expresses the hyaluronan receptor LYVE-1, a CD44 homologue (10), and vascular endothelial cell growth factor receptor (VEGFR)-3, a receptor for the lymphangiogenesis factors vascular endothelial growth factor (VEGF)-C and VEGF-D (11). The expression of both of these receptors later becomes restricted to lymphatic endothelium (Fig. 1; reference 12). This is followed by the polarized budding and migration of Prox1+ lymphatic endothelial progenitor cells (8) that progressively down-regulate the expression of blood vascular genes, such as CD34 and laminin (9), and express increasing levels of lymphatic markers such as VEGFR-3 and secondary lymphoid chemokine (CCL21), a ligand for the chemokine receptor CCR7 (13). Importantly, in Prox1 null mice, the budding and sprouting of lymphatic endothelial cells from the veins is arrested around E11.5-E12.0 and they completely lack a lymphatic vascular system (8). With the reported haploinsufficiency effect of Prox1 during the development of the enteric lymphatic system, these findings reveal an essential role of Prox1 during early lymphatic specification and development (8, 9). The exact mechanisms of action of Prox1 during and after the switch from the blood vascular to the lymphatic phenotype remain to be identified. However, recent studies revealed that ectopical expression of Prox1 in primary human blood vascular endothelial cells was sufficient to up-regulate the expression of the lymphatic endothelial cell markers podoplanin and VEGFR-3, and repress the expression of several genes that have been associated with the blood vascular endothelial cell phenotype (14). These results identify Prox1 as a master control gene in the program specifying lymphatic endothelial cell fate (14).

Recent studies in angiopoietin-2–deficient mice suggest an important role of the angiopoietins and their receptor Tie2 for the final developmental steps of lymphatic network patterning (Fig. 1) and lymphatic vessel maturation (14). However, the molecular mechanisms controlling the sprouting of lymphatic endothelial cells from primitive lymph sacs and their migration into adjacent organs and tissues (lymphangiogenesis) have remained unclear. In this issue, Saaristo et al. (15) identify VEGF-C as a potent inducer of lymphatic sprouting and provide experimental evidence that in addition to VEGFR-3, VEGFR-2 may also be required for this process. Previously, the authors had shown that signaling via VEGFR-3 was sufficient to induce hyperplasia of cutaneous lymphatic vessels because transgenic mice with skin-specific overexpression of a mutated VEGF-C (K14-VEGF-C156S) that selectively activates VEGFR-3 developed lymphatic vessel enlargement in the skin (17). In contrast, wild-type VEGF-C activates both VEGFR-3 and, after proteolytic processing, VEGFR-2.

In the study by Saaristo et al., K14-VEGF-C or K14-VEGF-C156S transgenic mice were crossed with VEGFR-3+/LacZ mice in which one allele of VEGFR-3 had been replaced by the LacZ gene, thereby enabling the visualization of lymphatic vessels by X-gal staining. Importantly, whereas VEGF-C156S overexpression mainly caused the enlargement of preexisting lymphatic capillaries, wild-type VEGF-C induced lymphatic vessel sprouting during embryogenesis (16). Similarly, an increased number of cutaneous lymphatic vessels was detected in adult VEGF-C transgenic mice and in adult mice that were intradermally injected with an adenovirus encoding VEGF-C, whereas chronic transgenic delivery of VEGF-C156S or intradermal injection of a VEGF-C156S–encoding adenovirus predominantly induced lymphatic enlargement. Moreover, only VEGF-C but not VEGF-C156S also induced angiogenesis and vascular hyperpermeability in these studies, most likely via interaction with VEGFR-2 on blood vascular endothelium. These results indicate that VEGF-C, through interaction with both VEGFR-3 and VEGFR-2, plays an important role in lymphangiogenesis, i.e., the sprouting of lymphatics from preexisting vessels. This is similar to the effects of VEGF-A in angiogenesis where it induces sprouting of new blood vessels (18, 19). Future studies in mice deficient for VEGF-C or VEGF-D, a related lymphangiogenesis factor with comparable VEGFR binding properties, should reveal whether the activation of VEGFR-3 and VEGFR-2 is only sufficient, as shown here, or also necessary for the induction of lymphangiogenesis during normal embryonic development. Moreover, additional studies are needed to investigate whether or not mesenchymal lymphatic progenitor cells might contribute to embryonic (or postnatal) lymphangiogenesis, as has been recently proposed for the early wing bud development in birds (20).

In addition to providing new insights into the mechanisms directing lymphatic development, this study by Saaristo et al. raises new questions regarding the molecular control of angiogenesis versus lymphangiogenesis. In this study, VEGFR-2 was implicated in the induction of lymphatic sprouting and strong expression of VEGFR-2 was detected on collecting lymphatic vessels. Therefore, one might expect that VEGF, thus far thought to specifically induce blood vascular angiogenesis (21), might also be able to activate lymphatic vessel sprouting via the activation of VEGFR-2. Indeed, VEGFR-2 is expressed by cultured lymphatic endothelial cells (22, 23) and VEGF equally stimulates lymphatic and blood vascular endothelial growth in vitro (23 and unpublished data). Moreover, intradermal injection of a VEGF165-encoding adenovirus into mouse ears resulting in high levels of VEGF expression, potently induced the formation of new lymphatic vessels that persisted for up to 1 yr (Dvorak, H.F., personal communication). In contrast, cutaneous wound healing is associated with up-regulated expression of VEGF (24) and the formation of a richly vascularized granulation tissue that initially contains no or only a few lymphatic vessels (unpublished data). Is the formation of VEGFR-3 and VEGFR-2 heterodimers needed for the efficient formation of lymphatic vessel sprouts, as suggested by Saaristo et al. (16)? Does VEGF need simultaneous activation/binding of VEGFR-1, most likely not expressed by lymphatic endothelium in vivo (unpublished data) but by cultured lymphatic endothelial cells (22), and of VEGFR-2 to exert its angiogenic effects under pathological conditions, as suggested by recent findings in placenta growth factor–deficient mice (21)? Does VEGF, via its vascular permeability–inducing activity, create a tissue environment that is permissive for blood vascular endothelial proliferation and sprouting, but not for lymphangiogenesis despite the activation of VEGFR-2, possibly due to the differential expression of extracellular matrix receptors by lymphatic endothelium? Do the observed effects of adenoviral VEGF expression on lymphangiogenesis represent a physiological response of lymphatic endothelium to increased tissue fluid accumulation, or are they caused by the induction of VEGF-C expression in vascular endothelium as has been reported (25)? Future in vivo and in vitro studies, including gene expression profiling, are needed to address this unresolved discrepancy.

Impaired formation of lymphatic vessels results in insufficient fluid drainage from tissues, leading to chronic lymphedema that is characterized by edematous swelling of the skin, epithelial hyperplasia, dermal fibrosis, delayed tissue repair, and impaired immune response (1). Recently, missense mutations in the VEGFR-3 gene have been detected in some cases of primary congenital lymphedema (Milroy disease), indicating an important role of VEGF-C and/or VEGF-D in the normal development of the human lymphatic system (26). Consequently, a heterozygous inactivating VEGFR-3 mutation was identified in Chy mutant mice that develop cutaneous lymphedema and chylous ascites after birth and may serve as a convenient mouse model for primary lymphedema (27). Importantly, virus-mediated VEGF-C gene therapy stimulated the growth of functional lymphatics in this model (27), indicating the potential applicability of growth factor gene therapy to at least some cases of human lymphedema. However, adenoviral VEGF-C gene therapy also induced blood vascular enlargement and increased vascular permeability via interaction with VEGFR-2, unwanted side effects in the context of clinical antilymphedema therapy (28). Saaristo et al. (16) now provide evidence that these blood vascular side effects were avoided by viral gene transfer of a VEGFR-3–specific mutant form of VEGF-C (VEGF-C156S) to wild-type and Chy lymphedema mice. Remarkably, the authors detected functional cutaneous lymphatic vessels as confirmed by their ability to transport intradermally injected FITC-dextran even 8 mo after the injection of the VEGF-C156S-adeno–associated virus into the ear skin of Chy mutant mice, whereas no changes of blood vascularity were observed.

These findings have potential implications for the development of novel therapies for human lymphedema, and it will be of interest to see whether the intradermal injection of naked VEGF-C156S plasmid cDNA, as previously described for VEGF treatment of peripheral artery disease (29), or of recombinant VEGF-C156S protein will also be able to specifically induce the formation of functional lymphatics, avoiding potential side effects associated with the in vivo application of adenoviral constructs. However, one has to keep in mind that thus far missense mutations of VEGFR-3 have only been detected in a minority of all patients with congenital lymphedema and additional gene mutations are likely responsible for the majority of these cases. The recent identification of inactivating mutations of the FOXC2 gene in the autosomal-dominant disorder lymphedema-distichiasis (30), together with the detection of lymphedema, chylous ascites, or chylothorax in an increasing number of mutant mouse models such as {alpha}9 integrin and angiopoietin-2–deficient mice (15, 31), and the identification of novel lymphatic-specific markers such as Prox1, LYVE-1, and podoplanin (32), suggests the presence of additional disease-specific targets for the future treatment of primary lymphedemas.

Secondary lymphedema is frequently induced by the surgical removal or radiation of lymph nodes in cancer patients, whereas filariasis, a chronic infection with the parasitic worms Brugia malayi or Wuchereria bancrofti, is the leading cause in the developing world. Secondary lymphedema after surgery is associated with the interruption of the normal lymphatic drainage system. Recent studies in an experimental postsurgery lymphedema model, involving the removal of lymphatic vessels from rabbit ears, showed that the injection of VEGF-C protein into the wounded area induced the growth of functional lymphatics along with normalization of the tissue structure (33). Therefore, postsurgical lymphedemas might constitute additional targets for VEGF-C– or VEGF-C165S–based protein or gene therapies. The recent discovery of a direct correlation between experimental tumor-associated lymphangiogenesis and enhanced lymph node metastasis (3437), however, suggests that future studies are warranted to evaluate whether therapeutic regeneration of lymphatic vessels after lymph node removal might increase the risk for enhanced spread of tumor metastases.

Tumor metastasis to regional lymph nodes represents the first step of tumor dissemination in many common human cancers and serves as a major prognostic indicator for the progression of the disease. In contrast to the extensive molecular and functional characterization of tumor angiogenesis (38), i.e., the induction of new blood vessel growth, little is known about the mechanisms through which tumor cells gain entry into the lymphatic system. A widely held view has suggested that lymphatic endothelium only plays a passive role during this process (38) and lymphatic invasion only occurs once stroma-infiltrating tumor cells happen upon preexisting peritumoral lymphatic vessels (Fig. 2 A). However, the recent identification of lymphatic growth factors and receptors, together with the discovery of lymphatic-specific markers and the development of orthotopic cancer metastasis models, have provided important new insights into the formation of tumor-associated lymphatic vessels (7) and their active contribution to lymphatic tumor spread (Fig. 2 B). An increasing number of clinicopathological studies have shown a direct correlation between tumor expression of the lymphangiogenesis factors VEGF-C or VEGF-D and metastatic tumor spread in many human cancers, including cancers of the breast, lung, prostate, cervix, and colon (for review see reference 39), providing circumstantial evidence for the involvement of lymphangiogenesis in tumor progression.

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Figure 2. (A) Traditional model of tumor metastasis via lymphatic and blood vessels. (B) Active lymphangiogenesis model of tumor metastasis.


Several studies in animal tumor models have now provided direct experimental evidence that increased levels of VEGF-C or VEGF-D promote tumor lymphangiogenesis and lymphatic tumor spread to regional lymph nodes and that these effects can be suppressed by blocking VEGFR-3 signaling (3437, 4042). Most of these studies used an antibody against the hyaluronan receptor LYVE-1, a lymphatic-specific CD44 homologue (10, 43), to identify and quantify tumor-associated lymphatic vessels. Although some LYVE-1 expression has been detected in liver sinusoidal endothelial cells that are involved in hyaluronan uptake (44), recent studies applying the combined immunostains of a variety of experimental tumors with antibodies to LYVE-1 and the lymphatic-specific transcription factor Prox1 found that all LYVE-1+ tumor-associated lymphatic vessels also expressed Prox1 (7, 9, 45), confirming the specificity of LYVE-1 expression for lymphatic endothelium.

Despite the accumulated evidence for an active role of VEGF-C– or VEGF-D–induced tumor lymphangiogenesis in cancer metastasis to regional lymph nodes, the existence and biological function of lymphatics within experimental and human tumors has remained controversial. High interstitial pressure within tumors has been proposed to prevent intratumoral lymphatic vessel growth and function as assessed by the lack of lymphatic uptake of tracers that were injected in the vicinity of experimental tumors (46, 47). However, the mechanisms controlling metastatic tumor cell invasion and transport within lymphatic vessels are most likely distinct from those involved in fluid uptake and transport. Indeed, proliferating intratumoral lymphatic vessels have been detected in rapidly progressing tumor xenotransplants and in slowly growing, chemically induced orthotopic squamous cell carcinomas in mice and is associated with lymphatic metastasis (7, 9, 34, 45). Proliferating intratumoral lymphatics have also been found in human head and neck squamous cell carcinomas that were characterized by the correlation of the density of LYVE-1+ tumor-associated lymphatic vessels with the presence of regional lymph node metastasis (48). In contrast, no evidence for tumor lymphangiogenesis was found in invasive breast cancer by the same group of investigators (49). Taken together, these results indicate that active tumor-associated lymphangiogenesis induced by VEGF-C, VEGF-D, or other not yet identified growth factors leads to the proliferation and enlargement of peritumoral and, in some cancers, intratumoral lymphatic vessels, likely enhancing the metastatic spread of many different types of human tumors (Fig. 2 B).

Although the mere increase of lymphatic vessel surface area might simply increase the chance for tumor cell invasion and metastasis, lymphatic endothelial cells probably also play an active role in the chemotactic recruitment and intralymphatic transport of tumor cells. Lymphatic endothelium secretes chemokines such as CCL21 (secondary lymphoid chemokine) that binds to CCR7 (13, 22, 50), leading to chemoattraction and migration of mature dendritic cells from the skin to regional lymph nodes. CCR7 and other chemokine receptors are also expressed by some human cancer cell lines including malignant melanomas and breast cancer cells (51). Importantly, the overexpression of CCR7 in B16 malignant melanoma cells led to a >10-fold increase in the incidence of regional lymph node metastases after injection into the footpad of mice, and treatment with CCL21-blocking antibodies completely prevented metastatic tumor spread to lymph nodes (52). These findings indicate that some tumors might take advantage of preexisting molecular mechanisms designed for the physiological immune response to further their metastatic spread.

After several decades of slow progress, the study of lymphatic vessel formation and its role in malignant disease has now led to the identification of several molecular mechanisms involved in the formation and biological function of lymphatic vessels. Although much has still to be learned about the detailed steps of normal and pathological lymph vessel formation, new targets for innovative therapeutic approaches and new tools for the prognostic evaluation of human cancers are now emerging.

Submitted: August 5, 2002
Accepted: August 13, 2002



Signalling via vascular endothelial growth factor receptor-3 is sufficient for lymphangiogenesis in transgenic mice

Tanja Veikkola, Lotta Jussila, Taija Makinen, Terhi Karpanen, Michael Jeltsch, Tatiana V. Petrova, Hajime Kubo, Gavin Thurston1, Donald M. McDonald1, Marc G. Achen2, Steven A. Stacker2 and Kari Alitalo3

Molecular/Cancer Biology Laboratory and Ludwig Institute for Cancer Research, Haartman Institute, University of Helsinki, PO Box 21 (Haartmaninkatu 3), 00014 Helsinki, Finland, 1Department of Anatomy and Cardiovascular Research Institute, University of California, San Francisco, CA 94143, USA and 2Ludwig Institute for Cancer Research, PO Box 2008, Royal Melbourne Hospital, Parkville, Victoria 3050, Australia 3Corresponding author e-mail: Kari.Alitalo@helsinki.fiT.Veikkola and L.Jussila contributed equally to this work

Received October 13, 2000; revised December 20, 2000; accepted January 29, 2001.

The EMBO Journal, Vol. 20, No. 6 pp. 1223-1231, 2001
© European Molecular Biology Organization


Vascular endothelial growth factor receptor-3 (VEGFR-3) has an essential role in the development of embryonic blood vessels; however, after midgestation its expression becomes restricted mainly to the developing lymphatic vessels. The VEGFR-3 ligand VEGF-C stimulates lymphangiogenesis in transgenic mice and in chick chorioallantoic membrane. As VEGF-C also binds VEGFR-2, which is expressed in lymphatic endothelia, it is not clear which receptors are responsible for the lymphangiogenic effects of VEGF-C. VEGF-D, which binds to the same receptors, has been reported to induce angiogenesis, but its lymphangiogenic potential is not known. In order to define the lymphangiogenic signalling pathway we have created transgenic mice overexpressing a VEGFR-3-specific mutant of VEGF-C (VEGF-C156S) or VEGF-D in epidermal keratinocytes under the keratin 14 promoter. Both transgenes induced the growth of lymphatic vessels in the skin, whereas the blood vessel architecture was not affected. Evidence was also obtained that these growth factors act in a paracrine manner in vivo. These results demonstrate that stimulation of the VEGFR-3 signal transduction pathway is sufficient to induce specifically lymphangiogenesis in vivo.

Keywords: angiogenesis/lymphangiogenesis/vascular endothelial growth factors (VEGFs)/VEGF receptors


Growth of new blood and lymphatic vessels by the processes of angiogenesis and lymphangiogenesis requires the activation of specific signal transduction pathways in endothelial cells. These signals are at least in part mediated by members of the vascular endothelial growth factor (VEGF) family via their receptors (VEGFRs) on the surface of endothelial cells. The members of the mammalian VEGF family known to date are VEGF, placenta growth factor (PlGF), VEGF-B, VEGF-C and VEGF-D. They show significant identity at the level of amino acid sequence, but are strikingly different in terms of their mechanisms of regulation, expression patterns and receptor binding profiles (reviewed by Eriksson and Alitalo, 1999). The prototype VEGF regulates vasculogenesis, haematopoiesis and vascular permeability, and is implicated in many physiological and pathological processes (Ferrara, 1999). Like the VEGFs, the three known VEGFRs are differentially expressed. In adult tissues, VEGFR-1 and VEGFR-2 localize predominantly to blood vascular endothelial cells, whereas VEGFR-3 is expressed mainly in lymphatic endothelia (for references see Veikkola et al., 2000).

VEGF-C and VEGF-D, which are the only known ligands for VEGFR-3, are produced as precursor proteins with N- and C-terminal propeptides flanking the VEGF homology domain (VHD; Joukov et al., 1996; Lee et al., 1996; Orlandini et al., 1996; Yamada et al., 1997; Achen et al., 1998). The secreted factors undergo proteolytic processing, resulting in the cleavage of the propeptides and increased affinity for VEGFR-2 (Joukov et al., 1997; Stacker et al., 1999a). The fully processed or mature forms of VEGF-C and VEGF-D consist of the VHD, which acts as a ligand for both VEGFR-2 and VEGFR-3 (Joukov et al., 1997; Achen et al., 1998). Mature VEGF-C and VEGF-D are mitogenic and chemotactic for endothelial cells in culture and angiogenic in vivo (Lee et al., 1996; Joukov et al., 1997; Achen et al., 1998; Cao et al., 1998; Witzenbichler et al., 1998; Marconcini et al., 1999). Importantly, VEGF-C has been shown to induce lymphangiogenesis in transgenic mouse skin and in mature chick chorioallantoic membrane (Jeltsch et al., 1997; Oh et al., 1997). Replacing the second of the eight conserved, characteristically spaced cysteine residues in the VHD (Cys156) with a serine residue in recombinant VEGF-C resulted in a mutant factor (VEGF-C156S), which is a selective agonist of VEGFR-3 (Joukov et al., 1998). VEGF-C156S induced autophosphorylation of VEGFR-3 but not VEGFR-2 in transfected cells, but its activity in primary endothelial cells was not tested.

Transgenic overexpression of VEGF and VEGF-C has yielded important data on their vascular effects (Jeltsch et al., 1997; Detmar et al., 1998), but these studies have not clarified the roles of specific VEGFRs in angiogenesis versus lymphangiogenesis due to the fact that both VEGF and VEGF-C bind more than one receptor. On the other hand, the in vivo studies of VEGFR function have been hampered by the embryonic death of the knockout mice (Fong et al., 1995; Shalaby et al., 1995; Dumont et al., 1998). We wanted to assess the potential in vivo function of VEGF-D and to determine the role of VEGFR-3-specific signals in lymphangiogenesis. For these studies we generated transgenic mouse strains expressing VEGF-D or VEGF-C156S in the skin under the human keratin 14 (K14) promoter. Here we show that VEGF-D is lymphangiogenic. Importantly, stimulation of only VEGFR-3 by VEGF-C156S was sufficient for generating the hyperplastic lymphatic phenotype, demonstrating that a single receptor tyrosine kinase mediates signals sufficient for lymphatic vascular growth.


Receptor specificity of VEGF-C156S and VEGF-D
We have previously shown that human VEGF-C and VEGF-D bind to soluble human VEGFR-2 and VEGFR-3 (Joukov et al., 1997; Achen et al., 1998) and that VEGF-C156S only binds to human VEGFR-3 (Joukov et al., 1998). Human VEGF-D and VEGF-C156S were assessed for their ability to bind to mouse VEGFRs using soluble fusion proteins where the extracellular domains of mouse VEGFR-2 or VEGFR-3 are fused to the immunoglobulin (Ig) {gamma}-chain Fc domain. VEGF-D was found to bind to both mouse VEGFR-2 and VEGFR-3, whereas VEGF-C156S and mouse VEGF-D bound only to mouse VEGFR-3 (Figure 1A; M.E.Baldwin, B.Catimel, E.Nice, S.Roufail, N.E.Hall, K.L.Stenvers, K.Alitalo, S.A.Stacker and M.G.Achen, in preparation). Therefore, the receptor binding patterns of human VEGF-D and VEGF-C156S are retained in mice. In order to test whether the receptor binding pattern is reflected in VEGFR activation by ligand-induced dimerization in primary dermal microvascular endothelial cells, we stimulated the cells with the purified factors followed by immunoprecipitation of VEGFR-2 and VEGFR-3 and anti-phosphotyrosine immunoblotting analysis. Tyrosyl autophosphorylation of VEGFR-2 was stimulated by VEGF-D but not by VEGF-C156S, whereas VEGFR-3 autophosphorylation was stimulated by both of these ligands (Figure 1B). The differential receptor binding of these factors thus leads to specific receptor activation in primary endothelial cells.

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Fig. 1. Receptor specificity and expression of the transgene-encoded proteins. (A) Receptor binding analysis of metabolically labelled VEGF-C, VEGF-D and VEGF-C156S using soluble human or mouse VEGFR-2 and VEGFR-3. (B) Immunoprecipitation and phosphotyrosine analysis of VEGFR-2 and VEGFR-3 after stimulation of primary dermal microvascular endothelial cells with starvation medium, VEGF-D or VEGF-C156S. The asterisk denotes an intracellular VEGFR-3 precursor, which is not phosphorylated upon stimulation. (C) Transgene mRNA expression in K14-VEGF-D and K14-VEGF-C156S mice, and the levels of VEGFR-2 and VEGFR-3 mRNA were determined by northern blotting and hybridization of total skin RNA with the specific probes. WT, wild type; TG, transgenic mouse.
Generation and analysis of K14-VEGF-D and K14-VEGF-C156S mice
We next wanted to analyse the effects of VEGF-D and VEGF-C156S in vivo in transgenic mice. For this, human VEGF-D and VEGF-C156S cDNAs were cloned into the K14 vector, which directs transgene expression to the basal cells of the epidermis, and the expression constructs were injected into fertilized mouse oocytes. The resulting mouse lines were analysed for the expression of the transgene-encoded growth factors and their receptors, VEGFR-2 and VEGFR-3, by northern blotting of total skin RNA (Figure 1C). While both transgenes were abundantly expressed in the skin, the levels of VEGFR-2 and VEGFR-3 mRNAs were low and seemingly unaffected by transgene expression. K14-VEGF-D and K14-VEGF-C156S mice appeared healthy and their growth and reproductive rates were normal. Histological examination of the skin of both K14-VEGF-D and K14-VEGF-C156S mice revealed large spaces in the upper dermis lacking connective tissue elements, lined by a thin layer of endothelium but devoid of red blood cells (Figure 2). These spaces were reminiscent of the enlarged lymphatic vessels seen in K14-VEGF-C mice (Figure 2C; Jeltsch et al., 1997) and they were often found in the proximity of the hair follicles, where transgene expression was strongest.

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Fig. 2. Lymphatic hyperplasia induced by transgene overexpression. Tissue sections from K14-VEGF-D (A), K14-VEGF-C156S (B), K14-VEGF-C (C) or wild-type (D) skin were immunostained for mouse VEGFR-3. Transgenic skin contains large spaces lined with a VEGFR-3-positive endothelium (A–C), whereas only a few lymphatic capillaries are seen in wild-type skin (D). Scale bar in (D) for (A–D), 50 µm.


Immunohistochemical analysis revealed that the endothelial cells lining the large spaces were positive for the panendothelial marker platelet endothelial cell adhesion molecule-1 (PECAM-1) (not shown). These cells were also positive for VEGFR-3 and for lymphatic vessel endothelial hyaluronan receptor-1 (LYVE-1) (Figure 3), both of which are known to provide relatively specific antigenic markers for the lymphatic endothelium in normal tissues (Jussila et al., 1998; Banerji et al., 1999; Partanen et al., 1999). Interestingly, in K14-VEGF-D mice the endothelium stained weakly positive for VEGFR-2, whereas corresponding structures in K14-VEGF-C156S and wild-type mice were VEGFR-2 negative (Figure 3C, H and M). Very weak staining for von Willebrand factor, and no staining for type IV collagen or smooth muscle actin (data not shown), confirmed the lack of blood endothelial markers, a basal lamina and a pericyte/smooth muscle cell layer, which are all hallmarks of blood vascular structures.

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Fig. 3. Immunohistochemical analysis of transgenic skin. High magnification via Nomarski optics. Note that the VEGFR-3-positive endothelial cells associated with the hair follicles (hf) in both wild-type and transgenic skin are also positive for the lymphatic marker LYVE-1 (A, B, F, G, K and L). Also, the endothelium lining the hyperplastic vessels was positive for VEGFR-2 in the K14-VEGF-D mice (C), whereas the hyperplastic vessels of the K14-VEGF-C156S mice and wild-type lymphatic capillaries were negative (H and M). Hair follicles stained for VEGF-C (D, I and N) and VEGF-D (E, J and O). Scale bar in (O) for (A–O), 15 µm.


To verify transgene expression at the protein level, skin sections were stained using antibodies against VEGF-D and VEGF-C. In the hair follicles of K14-VEGF-D mice the outer root sheath cells stained positive for VEGF-D (Figure 3E, arrow), whereas no staining was observed in the hair follicles of K14-VEGF-C156S or wild-type mice (Figure 3J and O). In K14-VEGF-C156S mice, strong staining for VEGF-C was observed in the same cells (Figure 3I).

Selective hyperplasia of lymphatic, but not blood vessels in transgenic skin
In order to analyse the dermal blood vascular phenotype of K14-VEGF-D and K14-VEGF-C156S mice, whole-mount tissue preparations of transgenic and wild-type ear skin were studied after vascular perfusion with biotin-labelled Lycopersicon esculentum lectin. When injected intravenously, this lectin binds to the surface of the endothelial cells allowing visualization of all blood vascular structures (Thurston et al., 1999). The blood vessels of K14-VEGF-D and K14-VEGF-C156S mice appeared normal in this analysis, as no differences from wild-type vessels could be found (Figure 4A–C). In order to visualize the lymphatic vessels in whole-mount preparations, the transgenic mice were crossed with heterozygous mutant VEGFR-3+/LacZ mice (Dumont et al., 1998). In these mice, one allele coding for VEGFR-3 has been disrupted by insertion of the LacZ coding sequence, allowing the localization of VEGFR-3 expression by staining for ß-galactosidase activity, which gives a blue signal (Dumont et al., 1998). The dermal lymphatic vessels in K14-VEGF-D x VEGFR-3+/LacZ and K14-VEGF- C156S x VEGFR-3+/LacZ compound heterozygous mice were considerably enlarged in comparison with the lymphatic vasculature in VEGFR-3+/LacZ mice (Figure 4D–F). These phenotypes demonstrate that VEGF-D and the VEGFR-3-specific ligand VEGF-C156S induce selective hyperplasia of the lymphatic but not blood vasculature when overexpressed under the K14 promoter.

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Fig. 4. Whole-mount analysis of skin blood and lymphatic vasculature. Blood vessels were visualized by injecting the mice intravenously with biotin-labelled L.esculentum lectin followed by vascular perfusion (AF). Lymphatic vessels were stained blue (arrows) in the skin of K14-VEGF-D and K14-VEGF-C156S mice crossed with heterozygous mutant VEGFR-3+/LacZ mice (D–F). Scale bar in (C) for (A–C), 75 µm; in (F) for (D–F), 65 µm.
Functional analysis of the hyperplastic lymphatic vessels
In order to determine whether the enlarged lymphatic capillaries in K14-VEGF-D and K14-VEGF-C156S skin were functional, we investigated lymphatic fluid uptake and transport in the skin after intradermal injections of TRITC-conjugated dextran, ferritin or Evans Blue dye. When fluorescent dextran was injected into the skin it was rapidly taken up by the superficial lymphatic capillary network in both normal and transgenic mice, as revealed by the fluorescence of these vessels in the tail and ear (Figure 5A–F). In K14-VEGF-D and K14-VEGF-C156S mice the lymphatic capillary network appeared vastly dilated when compared with the wild-type controls. To quantitate the degree of dilation, we measured the average lymphatic capillary diameter as well as the horizontal and vertical mesh sizes of the tail lymphatic capillary network in transgenic and wild-type mice. While the diameter of the lymphatic capillaries in K14-VEGF-D and K14-VEGF-C156S mice was approximately 3-fold larger than in wild-type mice, the horizontal and vertical mesh sizes were not significantly altered (Table I; see also Figure 5A–C).

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Fig. 5. Functional analysis of the lymphatic vessels of normal and transgenic skin. The lymphatic capillaries were visualized by an intradermal injection of TRITC-conjugated dextran into the tail (AC) or ear (DF). Ferritin was used as a marker for lymphatic colloidal transport in histological sections from the ear (arrows in GI). Lymphatic vessels adjacent to the ischiatic vein after injection of Evans Blue into the hind footpads (JL). Scale bar in (C) for (A–C), 170 µm; in (F) for (D–F), 270 µm; in (I) for (G–I), 80 µm; and in (L) for (J–L), 1250 µm. d, diameter; h, horizontal mesh size; v, vertical mesh size.
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Table I. Structural parameters of lymphatic vessel network in the tail
While the lymphatic capillaries of the tail were organized in a regular hexagonal pattern, in the ear they had a more variable architecture. Injections of fluorescent dextran into the ears of wild-type mice showed that normally the lymphatic capillaries are thin and relatively far apart from each other (see Figure 5D). However, in K14-VEGF-D and K14-VEGF-C156S mice the dextran spread more widely and diffusely within the skin. To understand whether the observed pattern reflects lymphatic vessel leakiness and thus spread of the marker thoughout ear interstitium, we injected ferritin intradermally into the ear. Ferritin can be used as a lymph marker and visualized in histological sections by staining with Prussian Blue (Leu et al., 2000). Microscopic observation revealed the blue iron compound to be contained within the lymphatic vessels in both transgenic and wild-type mice, demonstrating that the hyperplastic lymphatic capillaries of K14-VEGF-D and K14-VEGF-C156S mice are not abnormally leaky (Figure 5G–I). Staining of serial sections with endothelial markers revealed the presence of an intact lymphatic endothelium lining the lymphatic spaces in the transgenic mice (data not shown).

The superficial lymphatic capillaries of the skin drain via connecting vessels into the deeper collecting lymphatic channels. The route of lymphatic transport from the skin can be marked by an intradermal injection of Evans Blue dye and by following the appearance of the dye in the deep collecting lymphatic channels. Upon injection of Evans Blue into the hind limb footpads of wild-type, K14-VEGF-D or K14-VEGF-C156S mice the lymphatic vessels on both sides of the ischiatic vein were rapidly stained blue in all of the mice (Figure 5J–L). Therefore, fluid transport from the skin did not appear to be impaired in K14-VEGF-D and K14-VEGF-C156S mice despite the hyperplasia of the superficial lymphatic capillaries.

Paracrine effects of the K14-VEGF-D and K14-VEGF-C156S transgene products
To find out whether transgenic overexpression of VEGF-D and VEGF-C156S had effects beyond the skin, internal organs in the compound heterozygous K14-VEGF-D x VEGFR-3+/LacZ and K14-VEGF-C156S x VEGFR-3+/LacZ mice were stained for ß-galactosidase activity. When the whole-mount lymphatic staining patterns of various internal organs were compared with VEGFR-3+/LacZ mice, no obvious differences were found. As an example, Figure 6A shows the pericardial lymphatic vessels of a K14-VEGF-C156S x VEGFR-3+/LacZ and a VEGFR- 3+/LacZ mouse. Although variable in architecture, there were no consistent differences in these vessels between the normal and transgenic mice.

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Fig. 6. Lack of a visceral phenotype and a short systemic half-life of transgene encoded proteins. (A) Pericardial lymphatic vessels of a K14-VEGF-C156S x VEGFR-3+/LacZ compound heterozygous and a VEGFR-3+/LacZ mouse were analysed by ß-galactoside staining. (B) Relative amounts of VEGF-C, VEGF-D and VEGF in mouse circulation as a function of time after an intravenous injection of recombinant protein. The dashed line denotes 1/2 of the maximum growth factor concentration just after injection.
One explanation for the absence of an internal organ phenotype was to assume that effective concentrations of VEGF-D or VEGF-C156S did not reach these target organs. To determine whether the K14-transgene product can be detected in mouse circulation, we compared serum concentrations of VEGF-D in K14-VEGF-D and wild- type mice of different ages by a specific enzyme-linked immunosorbent assay (ELISA). The antigen could not be detected in any of the samples, although the detection level of this assay as determined by the use of purified recombinant protein was <0.1 pM. In transgenic skin lysates the average VEGF-D concentration was 4.5 pM, whereas no VEGF-D could be detected in the skin of age-matched control mice. As reference tissues, we tested lysates from transgenic and wild-type heart and lung, but no differences between wild-type and transgenic mice were detected.

To measure the systemic half-life of recombinant VEGF-C and VEGF-D proteins, anaesthetized wild-type mice were given an intravenous injection of 1 µg of VEGF-D or VEGF-C, and blood was drawn at different time points after the injection. As determined by ELISA, the half-life of both VEGF-D and VEGF-C in mouse circulation was <15 min (Figure 6B), and by 30 min the factors had been cleared below the assay detection level. For comparison, we also injected recombinant VEGF into the circulation and measured its clearance rate. VEGF disappeared in a similar manner to VEGF-D and VEGF-C with a half-life of ~8 min and complete clearance by 30 min (Figure 6B). Co-injections of VEGF-C and VEGF-D, or VEGF-C together with VEGF, yielded similar clearance rates to those for the single factors (data not shown). As the total blood volume in mice constitutes 5–6% of body weight (1–1.5 ml in mice of 20–25 g), the blood concentrations of the factors after an injection of 1 µg of recombinant protein should range from 16 to 23 nM, i.e. to be well above the reported binding constants of VEGF-D and VEGF-C to VEGFR-2 (560 and 410 pM, respectively) and to VEGFR-3 (200 and 135 pM; Joukov et al., 1997; Stacker et al., 1999a). The injected ligand can thus be considered to saturate the receptor, and the clearance of VEGF-D and VEGF-C from the circulation after factor co-injection was thus probably not due to binding to their specific receptors on blood vessel endothelia. In conclusion, the lymphatic hyperplasia phenotype may be restricted to the skin due to the rapid clearance of the transgene products from the circulation via a receptor-independent mechanism.

Soluble VEGFR-3 blocks the lymphatic hyperplasia in the skin
Recombinant soluble VEGFR-1 or VEGFR-2 can inhibit both physiological and pathological angiogenesis, such as retinal neovascularization, corpus luteum angiogenesis or tumour growth (Aiello et al., 1995; Ferrara et al., 1998; Goldman et al., 1998; Kong et al., 1998; Lin et al., 1998; Takayama et al., 2000). We wanted to test whether the lymphatic hyperplasia in the skin of K14-VEGF-D and K14-VEGF-C156S mice can be neutralized by soluble VEGFR-3. We therefore mated the mice with K14-VEGFR-3-Ig transgenic mice expressing a soluble chimeric protein consisting of the ligand binding portion of the extracellular part of VEGFR-3 joined to the Fc domain of Ig {gamma}-chain (Figure 7A; Makinen et al., 2001). When the skin of the double transgenic mice was examined histologically, lymphatic vessels were no longer seen although transgene expression remained high, as confirmed by northern blotting (Figure 7B and data not shown). Therefore, the soluble VEGFR-3 is capable of inhibiting VEGF-D- and VEGF-C156S-induced lymphatic hyperplasia in vivo.

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Fig. 7. Lymphatic hyperplasia is neutralized in double transgenic mice expressing soluble VEGFR-3-Ig. (A) Schematic presentation of the binding patterns of the transgene-encoded proteins and endogenous VEGF-VEGFR family members in the skin. (B) Immunohistochemical staining of K14-VEGF-D x VEGFR-3-Ig and K14-VEGF-C156S x VEGFR-3-Ig double transgenic skin for VEGFR-3 and LYVE-1 (A–D). Note the lack of lymphatic vessels. Scale bar in (D) for (A–D), 15 µm.


Until now, VEGF-C has been the only growth factor known to target the lymphatic vascular compartment in vivo. VEGF-D is closely related to VEGF-C in structure, and together they are the only known natural ligands for VEGFR-3 (Joukov et al., 1996; Achen et al., 1998). Despite the fact that VEGF-D binds to VEGFR-2 and has been reported to be angiogenic (Marconcini et al., 1999), transgenic overexpression of VEGF-D led to lymphatic hyperplasia but no angiogenesis. In addition, the VEGFR-3-selective mutant factor VEGF-C156S also induced lymphatic hyperplasia, showing that the necessary and sufficient signals for the growth of lymphatic vessels are transduced via VEGFR-3.

It is unlikely that the lymphatic hyperplasia in K14-VEGF-D and K14-VEGF-C156S mice would be a secondary consequence of an increased production of lymphatic fluid in response to increased microvascular permeability, as both VEGF-D and VEGF-C156S have been reported to be inactive in the Miles vascular permeability assay (Joukov et al., 1998; Stacker et al., 1999b). The hyperplasia appeared to result from both increased endothelial cell proliferation in the existing lymphatic vessels and from the formation of additional lymphatic vessels, as we observed greater numbers of enlarged lymphatic capillaries in the ear skin of transgenic mice when compared with wild-type mice. This is consistent with the result from the K14-VEGF-C mice where the lymphatic hyperplasia was associated with an increased endothelial cell proliferation rate (Jeltsch et al., 1997).

The enlarged lymphatic capillaries were located in the upper dermis in association with the hair follicles. This localization correlates well with the published K14 promoter expression in the basal cell layer of the epidermis and in the outer root sheaths of the hair follicles (Byrne et al., 1994). The lymphatic nature of these vessels was confirmed by their staining for both LYVE-1 and VEGFR-3, two antigenic markers for the lymphatic endothelium (Jussila et al., 1998; Banerji et al., 1999). Importantly, blood vessels in K14-VEGF-D and K14-VEGF-C156S mice were VEGFR-3 negative. Our results thus indicate that VEGF-D and VEGF-C156S retain their lymphatic specificity in vivo. Interestingly, the hyperplastic lymphatic capillaries of K14-VEGF-D mice were weakly positive for VEGFR-2, whereas those of K14-VEGF-C156S mice and the lymphatic capillaries of wild-type mice expressed no VEGFR-2. This result may reflect a positive feedback loop where VEGFR-2, which is normally expressed only in trace amounts in the endothelia of the lymphatic capillaries, is upregulated upon ligand binding and receptor activation.

High-resolution, whole-mount imaging techniques were used to study the blood vessel morphology of K14-VEGF-D and K14-VEGF-C156S mice, but no differences between wild-type and transgenic mice were found. Even though VEGFR-3 is expressed in blood vessel endothelia early in embryonic development, by embryonic day 13.5–14.5 when the K14 promoter expression becomes widespread in the skin keratinocytes (Byrne et al., 1994), VEGFR-3 has already been downregulated in blood vessels (Kaipainen et al., 1995). In concert with the VEGFR-3 developmental expression pattern, the binding sites of VEGF and VEGF-C in mouse embryos overlap early in the development, but diverge to target the blood vascular and lymphatic compartments, respectively, as the lymphatic vessels develop (Lymboussaki et al., 1999). VEGFR-3 has been shown to be upregulated on blood vascular endothelium in tumours and in chronic wounds (Partanen et al., 1999; Valtola et al., 1999; Paavonen et al., 2000). In these pathological conditions and in tissue ischaemia, VEGF-C and VEGF-D could thus promote angiogenesis. Tissue proteolytic activity in such processes is also enhanced, possibly contributing to the proteolytic processing of both VEGF-C and VEGF-D and formation of their VEGFR-2 binding forms (Achen et al., 1998; Stacker et al., 1999a). The lack of blood vascular effects of VEGF-D in the present model could thus result from both its incomplete proteolytic processing in the skin and from the low levels of VEGFR-2 present on quiescent blood vascular endothelium (our unpublished data).

The in vivo morphology of the K14-VEGF-D and K14-VEGF-C156S lymphatic capillaries was abnormal. In the tail, the characteristic honeycomb-like pattern was preserved, but the transgenic vessel diameter was approximately three times that of the wild-type mice. In the ears of K14-VEGF-D and K14-VEGF-C156S mice, the superficial lymphatic vascularity was increased in comparison with wild-type mice. The hyperplastic lymphatic capillaries were, however, functional, in that they were capable of liquid uptake and transport to the deep collecting lymphatic vessels. Also, these hyperplastic vessels did not appear to be abnormally leaky, as shown by the ferritin transport experiments.

In contrast to the superficial lymphatic vessels, the collecting vessels and visceral lymphatic vessels of K14-VEGF-D and K14-VEGF-C156S mice were seemingly unaffected by transgene overexpression, and no transgene-encoded growth factors could be detected circulating in the bloodstream. The absence of the growth factors in the circulation possibly results from their distribution to the pericellular matrix of the dermis and from their short systemic half-lives, probably due to clearance by the liver, the spleen and the kidneys, as determined by analysis of iodinated VEGF and VEGF-C injected into rat circulation (K.Paavonen and K.Alitalo, unpublished results). Lack of systemic vascular effects in K14-VEGF-D and K14-VEGF-C156S mice suggests that these molecules could be useful in the development of local gene therapy. Local induction of lymphatic capillary growth is of clinical interest as failure to regenerate lymphatic vessels (e.g. after surgery) results in secondary lymphoedema. Targeted delivery of VEGF-D or VEGF-C156S by gene transfer could perhaps also be used to induce lymphatic regeneration after injury.

Primary lymphoedema, a rare early-onset autosomal dominant disorder of the lymphatic system, was recently linked to mutations in the VEGFR-3 tyrosine kinase domain (Karkkainen et al., 2000). Disruption of VEGFR-3 signalling by receptor inactivating mutations or by soluble VEGFR-3 results in lymphatic hypoplasia, underlining the importance of VEGFR-3-mediated signals for the maintenance of lymphatic function after embryonic development (Karkkainen et al., 2000; Makinen et al., 2001). Other growth factor/receptor families are also likely to participate in the formation and maintenance of the lymphatic vessels. The Tie-1 receptor tyrosine kinase was detected on the hyperplastic lymphatic capillaries of K14-VEGF-C mice (Jeltsch et al., 1997), and targeted disruption of the angiopoietin-2 gene in mice results in a complete absence of the lymphatic vessels (C.Suri, M.Witte and G.Yancopoulos, personal communication). However, our results demonstrate for the first time that VEGFR-3 signalling alone is sufficient for the generation of all the necessary secondary signals for the induction of growth of functional lymphatic vessels in vivo. One important conclusion from these results is thus that specific inhibitors of VEGFR-2 that are used to block angiogenesis (see Veikkola et al., 2000) may not suffice to inhibit lymphangiogenesis and possibly associated metastasis in human tumours (Mandriota et al., 2001; Skobe et al., 2001; Stacker et al., 2001).

Materials and methods

Receptor binding and activation analysis
293T cells were transfected with expression constructs encoding human full-length VEGF-C, VEGF-D or VEGF-C156S using the calcium phosphate precipitation method. Twenty-four hours after transfection the cells were washed twice with phosphate-buffered saline (PBS) and incubated in methionine- and cysteine-deficient MEM for 60 min. The cells were then pulse-labelled for 30 min in medium containing 100 µCi/ml [35S]Met/[35S]Cys (Promix, Amersham), subsequently washed with PBS and chased in Dulbecco’s modified Eagle’s medium supplemented with 0.2% bovine serum albumin (BSA) and 0.2 mM each of non-radioactive cysteine and methionine. After a chasing period of 8 h, the conditioned medium was harvested, supplemented with 1 mM phenylmethylsulfonyl fluoride (PMSF), 4 µg/ml leupeptin and 0.1 U/ml aprotinin, and cleared by centrifugation. Endogenous VEGF was depleted using monoclonal anti-human VEGF antibodies (R&D Systems) and protein G–Sepharose. The binding of VEGF-C, VEGF-D and VEGF-C156S to the human VEGF receptors was assessed by precipitation using soluble recombinant proteins consisting of the first three Ig-like loops of VEGFR-2 or VEGFR-3 fused to the Fc portion of human IgG (Achen et al., 1998). The fusion proteins (200 ng) were incubated with 1 ml of the pulse–chase-labelled conditioned medium at +4°C for 2 h in the binding buffer (0.5% BSA, 0.02% Tween-20 and 1 µg/ml heparin). The complexes were then precipitated with protein A– Sepharose and washed twice with the binding buffer and once with 20 mM Tris pH 7.4. The bound proteins were analysed by SDS–PAGE. The binding of human VEGF-C156S to mouse receptors was tested using 200 ng of recombinant soluble mouse VEGFR-2 (R&D Systems) or 1 ml of conditioned medium from cells transfected with a construct encoding the first three Ig-like loops of mouse VEGFR-3 fused to the Fc portion of IgG.

Receptor stimulation was carried out using passage 4–6 human dermal microvascular endothelial cells (Promocell). Subconfluent cells were starved overnight in microvascular endothelial cell growth medium (Promocell) containing hydrocortisone (1 µg/ml), gentamycin sulfate (50 µg/ml), amphotericin B (50 ng/ml) and 0.2% BSA. The cells were stimulated with 1 µg/ml VEGF-D or VEGF-C156S for 10 min at +37°C, washed twice with ice-cold PBS containing 100 µM sodium orthovanadate and lysed with RIPA buffer (10 mM Tris pH 7.4, 50 mM NaCl, 0.5% sodium deoxycholate, 0.5% NP-40, 0.1% SDS) containing 1 mM sodium orthovanadate, 1 mM PMSF and 0.1 U/ml aprotinin. The lysates were sonicated, clarified by centrifugation at +4°C, and immunoprecipitated with antiserum specific for VEGFR-2 (a kind gift from Lena Claesson-Welsh) or monoclonal antibodies against VEGFR-3 (Jussila et al., 1998). Western blotting analysis was carried out using PY20 phosphotyrosine-specific monoclonal antibodies (Transduction Laboratories) and the ECL method, followed by stripping and analysis using receptor-specific antibodies.

Generation of transgenic mice and northern blot analysis
The cDNAs encoding full-length human VEGF-D and VEGF-C156S were cloned into a human K14 promoter expression cassette (Vassar et al., 1989) and injected into fertilized mouse oocytes of the strain FVB/NIH. Several independent inbred lines of transgenic mice were generated and those lines with high levels of transgene mRNA expression were used for the study. In the transgenic mouse lines used there was only one site of transgene insertion in the genome, and the phenotype had 100% penetrance.

Total RNA from the back skin of wild-type, K14-VEGF-D and K14-VEGF-C156S mice was extracted using the RNeasy kit (Qiagen). RNA (10 µg) was electrophoresed in agarose gels containing formaldehyde, blotted and hybridized with human VEGF-D or VEGF-C156S, mouse VEGFR-2 or mouse VEGFR-3 cDNA probes.

Analysis of blood and lymphatic vessels
For immunohistochemistry, biopsies from the back, ear and internal organs of the transgenic mice were fixed in 4% paraformaldehyde (PFA), dehydrated and embedded in paraffin. After rehydration and microwave treatment for antigen retrieval, 5 µm sections were stained for VEGFR-3 and VEGFR-2 (Kubo et al., 2000), VEGF-C (Joukov et al., 1998), VEGF-D (R&D Systems), LYVE-1 (Banerji et al., 1999), PECAM-1 (Pharmingen), von Willebrand factor (DAKO), {alpha}-smooth muscle actin (Sigma) or collagen IV (Chemicon).

Lectin staining was used to visualize blood vessels in whole-mount tissue preparations (Thurston et al., 1999). One hundred microlitres of 1 mg/ml biotinylated L.esculentum lectin (Sigma) were injected intra venously via the femoral vein into anaesthesized mice and allowed to circulate for 2 min. The mouse was then sacrificed and the tissues were fixed by perfusion with 1% PFA/0.5% glutaraldehyde in PBS. The ears were dissected, washed with PBS, and the cartilage was removed. Bound lectin was visualized by the ABC-DAB peroxidase method, the ears were mounted onto slides and examined by light microscopy. In K14-VEGF-D x VEGFR-3+/LacZ and K14-VEGF-C156S x VEGFR-3+/LacZ compound heterozygous mice blood vessels were visualized as above. Lymphatic vessels were then visualized by incubating the fixed tissues in the ß-galactosidase substrate X-Gal (Sigma) followed by dehydration and mounting.

Studies on lymphatic transport
Microlymphography was performed to visualize the lymphatic network in the ear and tail. TRITC-dextran (mol. wt 2000 kDa; Sigma) was injected into the tip of the tail using a 30 gauge needle, and the progressive staining of the lymphatic network was followed by fluorescence microscopy and photographed.

Ferritin (type I ferritin from horse spleen, mol. wt 480 kDa; Sigma) was injected into the ear. Forty-five minutes after injection the mouse was sacrificed and the ears were prepared for histology. The iron component of ferritin was visualized in histological sections by potassium ferrocyanide/HCl (Prussian Blue) followed by counterstaining.

A bolus of 5 mg/ml Evans Blue (Sigma) in PBS was injected intradermally into the hind footpads of mice. The skin from the lateral surface of the hind limb was removed to expose the ischiatic vein. Transport of the dye through the lymphatic vessels along the ischiatic vein was followed under direct microscopic observation and photographed.

The morphometric parameters of the transgenic and wild-type lymphatic capillaries were determined after visualization by an injection of fluorescent dextran into the tail. Lymphatic capillary diameter, as well as the horizontal and vertical mesh sizes of the lymphatic network, was measured from appropriate photomicrographs, and statistically significant variation from wild type was determined by the Mann–Whitney test.


For determination of VEGF-D from serum samples, MaxiSorp plates (Nunc) were coated overnight at +4°C with 2.5 µg/ml monoclonal anti-human VEGF-D (R&D Systems) in PBS. The wells were blocked with 5% BSA, 0.05% Tween-20 in PBS for 30 min at room temperature (RT). Serum samples were diluted in 5 mg/ml BSA, 0.05% Tween-20 in PBS and incubated in the wells for 1 h at RT, the wells were washed three times with incubation buffer, and 2.5 µg/ml biotinylated rabbit polyclonal anti-VEGF-D antibody (R&D Systems) was added for 1 h at RT. After washes as above, the wells were incubated with ZyMax Streptavidin-Alkaline Phosphatase (Zymed) at 300 ng/ml for 30 min at RT, washed, and the 4-nitrophenyl phosphate (4-NPP) substrate (Roche) at 1 mg/ml in diethanolamine pH 10.3 was added. Optical density was read at 405 nm.

For quantification of VEGF-D in transgenic tissues, age-matched transgenic and wild-type mice were sacrificed and the hair of back skin was removed. Skin, lung and heart were snap-frozen in liquid nitrogen, pulverized using a dismembranator and lysed in 20 mM Tris pH 7.4, 1 mM EDTA, 50 mM NaCl, 50 mM NaF, 1% Triton X-100, 1 mM PMSF, 4 µg/ml leupeptin and 0.1 U/ml aprotinin. Equal amounts of total protein were used for the ELISA, which was performed as above.

The half-lives of the purified factors in blood circulation were estimated by giving anaesthesized mice an intravenous injection of 1 µg of VEGF-D, VEGF-C or VEGF165 in 100 µl of PBS via the femoral vein. Blood was drawn at different time points after the injection and VEGF-D levels were analysed by ELISA as above. For VEGF-C determination, MaxiSorp plates were coated overnight with 15 µg/ml monoclonal anti-human VEGF-D (clone VD4, cross-reacts with VEGF-C; Achen et al., 2000), and 8 µg/ml rabbit anti-human VEGF-C (Joukov et al., 1997) was used for detection. For VEGF determination, Quantikine human VEGF colorimetric sandwich ELISA (R&D Systems) was used.


We thank Eija Koivunen, Pipsa Ylikantola, Riikka Kivirikko, Tapio Tainola, Sanna Karttunen and Sirke Haaka-Lindgren for excellent technical assistance. The K14 expression vector was a kind gift from Dr Elaine Fuchs. Dr David G.Jackson provided the LYVE-1 antibody. This study was supported by grants from the Ida Montini Foundation, the Finnish Cultural Foundation, the Foundation of the Finnish Cancer Institute, the Paolo Fondation, the Finnish Academy of Sciences and the Novo Nordisk Foundation.


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Adenoviral Expression of Vascular Endothelial Growth Factor-C Induces Lymphangiogenesis in the Skin

Molecular Medicine

(Circulation Research. 2001;88:623.)
© 2001 American Heart Association, Inc

Berndt Enholm, Terhi Karpanen, Michael Jeltsch, Hajime Kubo, Frej Stenback, Remko Prevo, David G. Jackson, Seppo Yla-Herttuala, Kari Alitalo

From the Molecular/Cancer Biology Laboratory and Ludvig Institute for Cancer Research (B.E., T.K., M.J., H.K., K.A.), Haartman Institute, University of Helsinki, Finland; Department of Pathology (F.S.), University of Oulu, Oulu, Finland; University of Oxford (R.P., D.G.J.), Molecular Immunology Group, Nuffield Department of Medicine, John Radcliffe Hospital, Headington, Oxford, UK; and A.I. Virtanen Institute and Department of Medicine (S.Y.-H.), University of Kuopio, Kuopio, Finland.

Correspondence to Kari Alitalo, MD, PhD, Molecular/Cancer Biology Laboratory, Biomedicum Helsinki, POB 63 (Haartmaninkatu 8), FIN-00014, Helsinki, Finland. E-mail


Abstract— The growth of blood and lymphatic vasculature is mediated in part by secreted polypeptides of the vascular endothelial growth factor (VEGF) family. The prototype VEGF binds VEGF receptor (VEGFR)-1 and VEGFR-2 and is angiogenic, whereas VEGF-C, which binds to VEGFR-2 and VEGFR-3, is either angiogenic or lymphangiogenic in different assays. We used an adenoviral gene transfer approach to compare the effects of these growth factors in adult mice. Recombinant adenoviruses encoding human VEGF-C or VEGF were injected subcutaneously into C57Bl6 mice or into the ears of nude mice. Immunohistochemical analysis showed that VEGF-C upregulated VEGFR-2 and VEGFR-3 expression and VEGF upregulated VEGFR-2 expression at 4 days after injection. After 2 weeks, histochemical and immunohistochemical analysis, including staining for the lymphatic vessel endothelial hyaluronan receptor-1 (LYVE-1), the vascular endothelial marker platelet–endothelial cell adhesion molecule-1 (PECAM-1), and the proliferating cell nuclear antigen (PCNA) revealed that VEGF-C induced mainly lymphangiogenesis in contrast to VEGF, which induced only angiogenesis. These results have significant implications in the planning of gene therapy using these growth factors.

Key Words: angiogenesis • immunohistochemistry • viruses • vessels • revascularization


Control of the vascular system by modulation of growth factor signaling is essential in attempts to treat diseases such as ischemic cardiovascular disease and cancer.1 2 3 Perhaps the most important family of growth factors involved in the regulation of angiogenesis is the vascular endothelial growth factor (VEGF) family, which includes VEGF, VEGF-B, VEGF-C, VEGF-D, Orf virus–encoded VEGF-E, and the placenta growth factor.4 5 These ligands bind to VEGF receptors (VEGFR)-1, VEGFR-2, and VEGFR-3 with partially overlapping receptor specificities. Both VEGF-C and VEGF-D bind VEGFR-2 and VEGFR-3 but are differentially regulated in cells and in tissues.6 7 8 9 The affinity of VEGF-C and VEGF-D toward their receptors is regulated by proteolytic processing; the affinity of the mature, proteolytically processed forms toward VEGFR-3 is {approx}40 times higher than the affinity toward VEGFR-2.6 10 The importance of VEGFR-3 signals in the vascular system is indicated by targeted mutagenesis of the VEGFR-3 gene, which results in embryonic lethality despite the presence of an intact VEGFR-2.11 The VEGFR-3 gene knockout leads to a disruption of the remodeling of primitive embryonic vasculature into a hierarchy of large and small vessels and results in cardiovascular failure of the embryos. However, in normal adult tissues, VEGFR-3 is largely absent from blood vessel endothelia and remains predominantly expressed in the lymphatic endothelium.9 12 13

Although a variety of angiogenic responses have been shown to be induced by adenoviral expression of VEGF in different mouse tissues,14 the biological functions of VEGF-C in normal adult tissues are thus far less clear. Overexpression of VEGF-C or VEGF in the skin under the keratin 14 promoter induced hyperplasia of lymphatic vessels or angiogenesis, respectively.15 16 17 In addition, recombinant VEGF-C was angiogenic in the early chick chorioallantoic membrane, but it induced exclusively lymphangiogenesis in the differentiated chorioallantoic membrane.18 19 Furthermore, both VEGF and VEGF-C were angiogenic when expressed from a transfected plasmid vector in a rabbit model of hindlimb ischemia.20 VEGF-C expression may thus in general lead to lymphangiogenesis, whereas in early embryonic stages11 or when overexpressed in ischemic tissues, it may stimulate angiogenesis.

The results showing that VEGF-C can induce both angiogenic and lymphangiogenic responses in various settings of gene delivery have raised important questions about the specificity of VEGF-C–induced vascular effects in normal adult tissues. Although in at least some conditions, the goal of proangiogenic gene therapy may be to regenerate all components of the vascular system, in other conditions, such as in secondary lymphedema, only lymphangiogenesis may be desired. In fact, the development of specific lymphangiogenic gene therapy would be an important development for example for the tens of thousands of patients who suffer from lymphedema secondary to axillary evacuation of the lymph nodes or for the millions of patients who develop the disease after filariasis. To resolve questions about the angiogenic versus lymphangiogenic specificity of VEGF-C, we have investigated in the present study the effects of VEGF-C gene transfer on the skin vasculature of adult mice compared with gene transfer of VEGF and ß-galactosidase in the same setting.

Materials and Methods

Generation of Recombinant Adenoviruses Encoding the VEGFs
For the construction of an adenovirus vector encoding VEGF-C, the full-length human VEGF-C cDNA (GenBank accession No. X94216) was cloned under the cytomegalovirus promoter in the pcDNA3 vector (Invitrogen). The SV40-derived polyadenylation signal of the vector was then exchanged for that of the human growth hormone gene, and the transcription unit was inserted into the pAd BglII vector21 as a BamHI fragment. Replication-deficient recombinant E1-E3–deleted adenoviruses were produced in human embryonic kidney 293 cells and concentrated by ultracentrifugation as previously described.22 Recombinant adenoviruses encoding VEGF165 and ß-galactosidase were constructed as previously described.22 23 24 25 26 Adenoviral preparations were confirmed to be free from helper viruses, lipopolysaccharide, and bacteriological contaminants.26

Construction, Expression, and Purification of VEGFR Ig Fusion Proteins
The expression plasmids encoding human VEGFR-1-Ig and VEGFR-3-Ig were constructed by polymerase chain reaction-amplifying the first three Ig homology domains of the extracellular portions of VEGFR-1 and VEGFR-3 with the primer pairs 5'-TCTCGGATCCTCTAGT- TCAGGTTCAAAATT-3' (BamHI site underlined)/5'-GATGAGA-TCTTTATCATATATATGCACTGA-3' (BglII site underlined) and 5'-CCTGGGATCCCTGGTGAGTGGCTACTCCATGAC-3'/5'-GATGAAGAGATCTTCATGCACAATGACCTCGG-3', respectively. The products were cloned into the BglII site of the pMT/BiP · V5 · HisC vector (Invitrogen), and the cDNA coding for the Fc-tail of human IgG1 was cloned in frame with the VEGFR Ig homology domains into the same vector. The expression plasmids were cotransfected with the pCO · Hygro selection plasmid (Invitrogen) into Drosophila S2 cells, and stable cell pools were selected in 150 µg/mL hygromycin B (Calbiochem). The expression of the Ig fusion proteins was induced with 500 µmol/L CuSO4 in serum-free DES medium (Invitrogen) and after 4 days, they were purified from the conditioned medium by protein A affinity chromatography (Amersham Pharmacia). VEGFR-2-Ig was obtained from R&D Systems (catalogue No. 357-KD).

Expression of Recombinant Adenoviral VEGF-C, VEGF, and ß-Galactosidase
Cells (293EBNA) grown in 10% FCS were transfected with pREP7 (Invitrogen) expression vectors encoding VEGF165 or VEGF-C6 , using the calcium phosphate precipitation method or infected by incubation with 2x107 pfu/106 cells (multiplicity of infection=20) of the respective adenoviruses in serum-free medium for 1 hour. The medium was then changed to medium containing 10% FCS, the cells were incubated overnight, and metabolically labeled with 35S-methionine and cysteine (Promix, Amersham) for 6 hours. The media were collected, and labeled VEGF proteins were precipitated using soluble VEGFR-Ig domain fusion proteins. Before VEGF-C precipitation using VEGFR-2-Ig, endogenous VEGF was removed from the supernatants by preadsorption using anti-VEGF monoclonal antibodies (R&D catalogue No. MAB293). The bound proteins were precipitated with protein G Sepharose, washed three times in PBS, dissolved in Laemmli sample buffer, and analyzed by 12.5% or 15% SDS-PAGE. Gels were then dried and analyzed by phosphor-imaging and autoradiography.

Analysis of the Adenovirus-Encoded Transcripts In Vivo
Adenovirus (2x108 pfu) encoding VEGF, VEGF-C, or ß-galactosidase was injected into the tail veins of two C56/Bl6 mice. The mice were sacrificed 4 days later and RNA was extracted from the livers (RNAeasy Kit, Qiagen). Total RNA (15 µg) was subjected to Northern blotting and hybridization with a mixture of 32P-labeled cDNAs specific for VEGF (nucleotides 57 to 639, GenBank accession No. NM003376), VEGF-C (nucleotides 495 to 1661, GenBank accession No. X94216), or LacZ (nucleotides 529 to 977 pBluscript SK+, Stratagene).

All experimental procedures involving laboratory animals were approved by the Helsinki University Ethical Committee and by the Provincial State Office of Southern Finland (permit No. HY 312).

Immunohistochemistry and Morphometry
Recombinant adenovirus or buffer (2x108 pfu) was injected subcutaneously into the backs of C56/Bl6 mice or into the ears of NMRI nude (nu/nu) mice (Harlan). The mice were sacrificed at various time points after injection. Skin from the site of injection was fixed in 4% paraformaldehyde and embedded in paraffin, and 6-µm sections were stained using monoclonal antibodies against VEGFR-2,27 VEGFR-3,28 or polyclonal antibodies against the lymphatic marker LYVE-1, a receptor for hyaluronan and a homologue to the CD44 glycoprotein,29 or mouse platelet–endothelial cell adhesion molecule-1 (PECAM-1) (BD Pharmingen, catalogue No. 01951D), the mouse homologue of the human vascular endothelial antigen CD31. Sections were also stained using polyclonal antibodies against laminin.30 The tyramide signal amplification (TSA) kit (NEN Life Sciences) was used to enhance staining. Negative controls were done by omitting the primary antibodies. Double staining of sections was carried out by first staining sections for proliferating cell nuclear antigen (PCNA) (ZYMED, catalogue No. 93-1143) and subsequently for LYVE-1 and PECAM-1 as detailed above. The results were viewed with an Olympus AX80 microscope and photographed. For quantification, the vessels in the sections were counted using square grids (area=0.16 mm2, x200 magnification), and the mean and probability value were calculated using the Student’s t test. Eight visual fields were quantified in sites of active angiogenesis or lymphangiogenesis in five different ears injected with AdVEGF-C or AdVEGF. For controls, 15 to 20 visual fields in five different ears injected with AdLacZ were quantified. For morphometric quantification of vessel volume, quantitative densitometry of 70 to 80 vessels in 8 to 10 visual fields was performed according to Weibel’s principles using a CAS200 (Becton-Dickinson) automated image analyzer and the proprietary software. Blood vessels were visualized and photographed in situ using a Leica MZ APO microscope.


Expression of VEGF-C and VEGF by Recombinant Adenoviruses In Vitro
To confirm that adenoviral gene transfer of VEGF-C results in secretion of polypeptides that bind to their receptors, 293EBNA cells were infected with the respective adenoviruses. Cells infected with the VEGF-C adenovirus (AdVEGF-C) produced major polypeptides of {approx}29/31 kDa that bound to the VEGFR-3-Ig fusion protein (Figure 1A, top). Under nonreducing conditions, these polypeptides migrated as an {approx}60-kDa band (Figure 1A, lanes NR). In comparison to cell cultures transfected with a VEGF-C plasmid expression vector, very small amounts of the mature, proteolytically processed 21/23 kDa-form of VEGF-C were observed in the culture media of Ad VEGF-C–infected cells, suggesting incomplete proteolytic processing. However, the mature 21/23-kDa species was the predominant VEGF-C form that bound to VEGFR-2-Ig (Figure 1A, bottom), as previously reported.6

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Figure 1. Expression of VEGF-C and VEGF165 by the recombinant adenoviruses in vitro and in vivo. A and B, Receptor binding by adenovirally produced VEGF-C and VEGF in vitro. VEGFR Ig fusion proteins were used to precipitate polypeptides from the media of metabolically labeled 293EBNA cells infected with AdVEGF-C, AdVEGF, or AdLacZ, as indicated. Plasmid controls are indicated as pREP for empty vector backbone, pVEGF-C for the vector encoding VEGF-C, and pVEGF for the vector encoding VEGF. Analysis by SDS-PAGE is shown under reducing and nonreducing (NR) conditions (the exposure in the lower panel in A was about twice as long as the exposure in other panels). Molecular size markers in the figure are indicated in kilodaltons, and arrows indicate bands representing the precipitated VEGF polypeptides. C, Analysis of adenovirus-encoded RNAs in the livers of infected mice. About 15 µg of total RNA extracted from livers of two mice injected with the indicated recombinant adenoviruses was subjected to Northern blotting and hybridization with a mixture of the corresponding cDNA probes. The first lane contains RNA extracted from the liver of an untreated mouse. The blot was stripped and reprobed for ubiquitin mRNA to confirm equal loading. The sizes of the different RNAs are indicated in kilobases on the right.

In an experiment similar to the one outlined above, we also confirmed that adenoviral gene transfer of VEGF results in the expression of VEGF polypeptides that form disulfide-bonded homodimers and bind to VEGFR-1 and VEGFR-2. Figure 1B shows SDS-PAGE analysis of the polypeptides precipitated from the conditioned medium of metabolically labeled AdVEGF-infected cells. A major VEGF polypeptide of {approx}24 kDa and a minor one of {approx}26 kDa are specifically precipitated using VEGFR-1-Ig or VEGFR-2-Ig. The former band comigrated with the major band in similar precipitates from the conditioned media of cells transfected using a plasmid expression vector for hVEGF165. The minor band of 22 kDa in the media of transfected cultures and the 26 kDa-form in cultures infected with the adenovirus probably represent differentially glycosylated polypeptide species. The same bands were also precipitated by monoclonal antibodies against VEGF (data not shown). Under nonreducing conditions, the adenovirally expressed polypeptides migrated in the range of 43 to 45 kDa, indicating a disulfide-stabilized dimeric structure (Figure 1B, lanes NR).

Expression of Adenovirally Encoded VEGF-C and VEGF In Vivo
The expression of VEGF-C and VEGF adenoviruses in vivo was tested by injecting the viruses into the tail veins of C56/Bl6 mice. Because most of the gene expression after intravenous injection of recombinant adenovirus occurs in the liver,31 we extracted RNA from the liver and analyzed it by Northern blotting and hybridization with a combination of probes specific for the adenoviral inserts. As can be seen in Figure 1C, the adenoviruses efficiently express mRNAs of 4.5 and 2.4 kb, encoding VEGF and VEGF-C, respectively, whereas somewhat lower amounts of mRNA of 6.0 kb encoding ß-galactosidase were produced by the control virus. The liver of an uninfected mouse showed no signal.

AdVEGF-C and AdVEGF Stimulate VEGFR Expression
The effects of the adenoviruses in vivo were tested by subcutaneous injection into mouse skin and by analyzing skin sections 4 days later by immunohistochemistry for the VEGF-C receptors VEGFR-2 and VEGFR-3 and for the vascular marker PECAM-1. As can be seen from Figure 2A and from the enclosed insets at higher magnification, adenoviral expression of VEGF-C for 4 days induced the expression of VEGFR-2 and VEGFR-3 in endothelial cells of blood vessels (containing erythrocytes), whereas VEGF gene transfer induced the expression of VEGFR-2 but not VEGFR-3 (Figure 2D). In contrast, the blood vessels in mice injected with AdLacZ (Figure 2B) or PBS (Figure 2C) did not stain for VEGFR-2 or VEGFR-3; only the lymphatic vessels were positive for VEGFR-3 in these mice. Analysis after 2 weeks showed an inflammatory response in all adenovirus-injected samples from the C57/Bl6 mice, confounding immunohistochemical analysis (data not shown). For this reason, we continued our studies in the immunocompromised athymic mice.

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Figure 2. Induction of VEGFR expression in response to adenoviral gene transfer of VEGF-C and VEGF. Sections of skin from mice injected subcutaneously with AdVEGF-C (A), AdVEGF (D), AdLacZ (B), or buffer (C) were stained for VEGFR-3. To evaluate the expression of the different receptors in blood vessels, adjacent sections stained for VEGFR-3, VEGFR-2, and PECAM-1 were observed at x400 magnification (insets). Open arrows indicate vessels staining for VEGFR-3; the upper arrows point to the superficial dermal lymphatic vessels and the lower ones to the blood vessels in the interface of dermis and subcutaneous fat tissue. In panel A, note VEGFR-3 expression in both blood vessels (containing erythrocytes) and lymphatic vessels, whereas in panels B, C, and D, VEGFR-3 expression is restricted to lymphatic vessels.

Lymphangiogenic and Angiogenic Responses to the Adenoviruses
Five ears of three nu/nu mice were injected with each of the adenoviruses. Shown in Figure 3 are AdVEGF-C, AdVEGF, or AdLacZ injection sites of mouse ears photographed in situ 3 days after the injection. As can be seen from this figure, VEGF induced the formation of enlarged, tortuous vessels (Figure 3B, arrows) in contrast to VEGF-C (Figure 3A) or ß-galactosidase (Figure 3C), which did not seem to affect at least the larger blood vessels.

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Figure 3. Comparison of vascular changes in response to VEGF, VEGF-C, and ß-galactosidase in situ. Mouse ears were photographed 3 days after injection with AdVEGF-C (A), AdVEGF (B), AdLacZ (C). Note the prominent enlarged and tortuous blood vessels (arrows) in response to AdVEGF compared with the vasculature in ears injected with AdVEGF-C or AdLacZ.


The adenovirus-injected ears were processed for immunohistochemistry and stained for PECAM-1 and the lymphatic-specific antigen LYVE-1. As can be seen from the LYVE-1 staining shown in Figures 4A and 4B, AdVEGF-C transfer induced the formation of LYVE-1–positive hyperplastic lymphatic vessels (arrows), which did not stain for laminin, a component of the basal laminae of blood vessels (data not shown), whereas AdVEGF (Figure 4D) or AdLacZ (Figure 4C) did not have any effects on the lymphatic vessels. In contrast, AdVEGF induced the formation of blood vessels (Figures 4E and 4F, arrows) whereas the AdLacZ (Figure 4G) did not have any effects on the blood vasculature. The effects of AdVEGF-C on blood vessels were more difficult to evaluate because of the strong lymphangiogenic response. However, there was a small increase of PECAM-1–positive vessels in the AdVEGF-C–injected ears (see Figures 4H and 5B). Some of these may represent newly formed, very weakly PECAM-1–positive lymphatic vessels (Figure 4H, asterisk).

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Figure 4. AdVEGF-C induces lymphangiogenesis and AdVEGF induces angiogenesis in vivo. The ears of nude mice were injected with AdVEGF-C (A and B), AdLacZ (C), or AdVEGF (D) and analyzed immunohistochemically 2 weeks after injection for the lymphatic endothelial-specific antigen LYVE-1. Note the abundant formation of enlarged, hyperplastic lymphatic vessels (arrows) in panel A. In panel B, x400 magnification of the lymphatic vessels shows extensive growth of these vessels around the muscle fibers. Both panels A and B contain blood vessels that are negative for LYVE-1, ie, the two large blood vessels (note erythrocytes) in panel A and the arteriole (asterisk) encroached by a LYVE-1–positive lymphatic vessel in panel B. In contrast, AdVEGF (D) or AdLacZ (C) did not affect the lymphatic vessels (arrows). In AdVEGF-injected mice, PECAM-1 staining revealed angiogenesis as represented by new capillary formation (E; arrows) and at x400 magnification (F). AdLacZ did not have effects on the vasculature (G) whereas AdVEGF-C mainly induced the formation of enlarged, very weakly PECAM-1–positive vessels (marked by asterisks in panel H), which were demonstrated to be lymphatic vessels by LYVE-1 staining (data not shown). Bar=500 µm.


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Figure 5. Quantification of adenovirus-induced vascular responses. The LYVE-1–positive (A) and PECAM-1–positive (B) vessels were counted at sites of adenoviral injection as detailed in Materials and Methods. VEGF-C and VEGF induced distinct lymphangiogenic and angiogenic responses, respectively, whereas a combination of VEGF and VEGF-C did not markedly potentiate either lymphangiogenic or angiogenic responses. C and D, Quantification of the volume of vessels staining for LYVE-1 and of strongly PECAM-1–staining vessels 2 weeks after injection. Error bars are ±2 SD.

Quantitatative Analysis of the Adenovirus-Induced Lymphatic and Blood Vessels
As can be seen from the results of counting the LYVE-1–positive and strongly PECAM-1–staining vessels with lumens in Figure 5A, AdVEGF-C induced an {approx}4-fold increase (P<0.01) of lymphatic vessel density (Figure 5A) whereas VEGF induced a 2-fold increase (P<0.01) of blood vessel density (Figure 5B). The combination of AdVEGF and AdVEGF-C did not significantly (P>0.5) potentiate either of these responses. VEGF-C increased the total volume of the LYVE-1–positive vessels by 7.5-fold (P<0.01) (Figure 5C), whereas VEGF increased the volume of the blood vessels by 5.7-fold (P<0.01) (Figure 5D).

Endothelial Cell Proliferation in Lymphangiogenesis Induced by VEGF-C
As can be seen in Figure 6A and at higher magnification in Figures 6B and 6C, sequential staining for both LYVE-1 and PCNA revealed that the lymphatic vessels in AdVEGF-C–injected ears contained proliferating lymphatic endothelial cells. For example, the lymphatic endothelial cells surrounding a small arteriole in Figure 6C stain for PCNA (closed arrowhead), whereas the blood vascular endothelial cells do not (open arrowhead). Figure 6D shows PCNA-positive nuclei in the wall of a blood vessel in an ear injected with AdVEGF. In contrast, the lymphatic vessels in the ears injected with AdVEGF or AdLacZ did not stain for PCNA. Approximately 30% (n=50) of the nuclei in the lymphatic vessels formed in response to VEGF-C stained positive for LYVE-1, whereas the proportion of PCNA-positive nuclei in blood vessels in ears injected with AdVEGF was only 6% (n=50). This low figure may reflect the fact that the peak in endothelial cell proliferation in the blood vessels occurs earlier during angiogenesis induced by VEGF.14

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Figure 6. VEGF-C induces endothelial cell proliferation in the lymphatic vessels. Shown are sections sequentially stained for both PCNA and LYVE-1. Arrowheads indicate PCNA-positive and open arrowheads PCNA-negative nuclei. In panel A and at higher magnification in panels B and C, the lymphatic vessels formed in response to VEGF-C contain several PCNA-positive nuclei. Note the PCNA-negative nuclei in the arteriole surrounded by two lymphatic vessels containing PCNA-positive nuclei in panel C. Gene transfer of VEGF had no effect on the lymphatic vessels as can be seen in panel D. However, blood vessels in these sections contained PCNA-positive nuclei, in contrast to the ß-galactosidase control shown in panel E, in which the blood and lymphatic vessels were negative. Panels A and E also contain numerous PCNA-positive nuclei in hair follicles as indicated by an asterisk in panel A. Bar=500 µm in panel A and 100 µm in panels B through E.


The present study shows that VEGF-C expressed subcutaneously by adenoviral gene transfer induces proliferation and enlargement of lymphatic vessels in a process that we refer to herein as lymphangiogenesis. VEGF-C also strongly upregulates VEGFR-2 and VEGFR-3 expression in blood vessels. In contrast, adenoviral gene transfer of VEGF induced VEGFR-2 upregulation in the endothelial cells of blood vessels and angiogenesis, as described earlier.14 24

The vessel density in foci of lymphatic vessel formation in the ears infected with AdVEGF-C increased {approx}4-fold in comparison to ears injected with AdVEGF, AdLacZ, or buffer control as measured by quantification of LYVE-1–positive vessels. The lack of smooth muscle cells around the vessels and erythrocytes within the vessels generated in 2 weeks was in accordance with the lymphatic vessel morphology. Furthermore, these vessels did not stain for laminin, a component of the basal laminae (data not shown). The density of strongly PECAM-1–positive vessels in the ears infected with AdVEGF increased {approx}2-fold compared with ears infected with AdVEGF-C, AdLacZ, or buffer. It may also be noted that LYVE-1 expression was not upregulated in blood vessels in AdVEGF-induced angiogenesis (eg, see Figure 3A). Thus, the response to AdVEGF-C was primarily lymphangiogenic, whereas very little angiogenesis was seen, unlike in the experiments in which plasmid expression vectors were used in ischemic rabbit muscle.20

In cell culture, the majority of the adenovirally produced VEGF-C consisted of the partially processed 29/31-kDa form, which binds VEGFR-3 but only very weakly to VEGFR-2.6 In our in vivo assay in normal dermis, this could be the predominant form, whereas in ischemic tissue the 21-kDa form of VEGF-C, which has a higher binding affinity toward VEGFR-2, may predominate because of increased expression of VEGF-C processing enzymes in the latter. A major difference between our assay conditions and those used in experiments using ischemic hindlimb as a target for plasmid delivery is the presence of abundant amounts of endogenous VEGF induced by hypoxia in the latter. However, at least in our initial experiments, simulation of such conditions by coinjection of AdVEGF and AdVEGF-C did not result in a substantial potentiation of the angiogenic response.

The mechanisms of lymphangiogenesis in adult tissues have not been elucidated. The generation of lymphatic vessels could in principle require endothelial cell sprouting from or splitting of preexisting lymphatic vessels or blood vessels, in situ differentiation of endothelial cells, or recruitment and lymphatic differentiation of endothelial precursor cells, as has been described in other models.32 33 34 In embryos, lymphatic vessels are mainly formed by the process of sprouting from certain venous structures, although in the avian species, mesenchymal precursor cells called lymphangioblasts also exist.35 36 We do not yet know the mechanisms of lymphangiogenesis in the adult, but the present results are compatible with the process of sprouting lymphatic vessels from preexisting ones and perhaps splitting of such enlarged lymphatic vessels that we observed in the AdVEGF-C–treated ears. The upregulation of VEGFR-2 and VEGFR-3 in blood vessels in response to VEGF-C raises the interesting possibility that endothelial cells in blood vessels could also participate in lymphangiogenesis by the process of migration and transdifferentiation. Such upregulation of both VEGF-C receptors in the blood vascular endothelium should also be considered when using gene therapy in the setting of tissue ischemia.

It has been shown that the angiogenic response induced by AdVEGF is a highly dynamic process involving the initial formation of mother vessels and endothelial glomeruloid bodies.14 Thus, our analysis at the 2-week time point does not reveal the kinetics of possible transient blood vessel responses. The responses to VEGF-C in blood vessel endothelia, which upregulate both receptors for VEGF-C, remain to be characterized. Therapeutic angiogenesis ultimately requires the induction of entire vascular structures consisting of arteries, veins, and lymphatics. Thus, proangiogenic therapy could consist of different growth factors that cover the entire genetic program for the induction of new vessels.37 Our studies in transgenic mouse embryos and newborn mice have revealed that the developing lymphatic vasculature is dependent on VEGF-C for survival signals and when the embryonic tissues are deprived of such signals by blocking both VEGF-C and VEGF-D, the forming lymphatic vessels regress by specific lymphatic endothelial apoptosis (T. Makinen et al, unpublished observations, 2000). Therefore, further studies are needed to determine the long-term effects of the transient viral expression of VEGF-C, whether this results in permanent and functional lymphatic vasculature and whether stable changes of the blood vasculature can also be observed.


This study was supported by grants from the Sigrid Juselius Foundation, the Finska Lakaresallskapet, the Finnish Academy, the University of Helsinki Hospital (TYH 8105), the State Technology Development Center, and EU Biomed Program BMH-98-3380. Dr Jackson is supported by the Medical Research Council and by a grant (00-311) from the Association for International Cancer Research


Original received October 23, 2000; revision received February 2, 2001; accepted February 2, 2001.



Role of Lymphangiogenesis in Cancer

  1. Address reprint requests to Sudha S. Sundar, MPhil, MRCOG, Cheltenham General Hospital, 9 Northcroft, The Park, Cheltenham, Gloucestershire GL50 2NL, United Kingdom; e-mail:


Regional lymph node metastasis is a common event in solid tumors and is considered a marker for dissemination, increased stage, and worse prognosis. Despite rapid advances in tumor biology, the molecular processes that underpin lymphatic invasion and lymph node metastasis remain poorly understood. However, exciting discoveries have been made in the field of lymphangiogenesis in recent years. The identification of vascular endothelial growth factor ligands and cognate receptors involved in lymphangiogenesis, an understanding of the embryology of the mammalian lymphatic system, the recent isolation of pure populations of lymphatic endothelial cells, the investigation of lymphatic metastases in animal models, and the identification of markers that discriminate lymphatics from blood vessels at immunohistochemistry are current advances in the field of lymphangiogenesis, and as such are the main focus of this article. This review also evaluates evidence for lymphangiogenesis (ie, new lymphatic vessel formation in cancer) and critically reviews current data on the prognostic significance of lymphatic vascular density in tumors. A targeted approach to block pathways of lymphangiogenesis seems to be an attractive anticancer treatment strategy. Conversely, promotion of lymphangiogenesis may be a promising approach to the management of treatment-induced lymphedema in cancer survivors. Finally, the implications of these developments in cancer therapeutics and directions for future research are discussed.


Lymphangiogenesis: A Potential New Therapy for Lymphedema

Jan 2012

Cooke JP.


Stanford Cardiovascular Institute, Stanford, CA.


At the level of the capillaries, the systemic circulation loses about 2-4 liters of fluid and about 100g of protein into the interstitium daily. This ultrafiltrate of the systemic capillaries is returned to the circulatory system by the lymphatics. The lymphatic vasculature is highly specialized to perform this service, beginning with the blind-ended lymphatic capillaries. These vessels are highly permeable to protein, fluid and even cells, due to fenestrations in their basement membrane, and discontinuous button-like junctions rather than tight intercellular junctions as observed in the systemic capillaries(1). The lymphatic capillaries merge into collectors and larger lymphatic conduits that are invested with vascular smooth muscle (capable of contracting and propelling lymph forward) and valves for unidirectional flow. These conduits merge at lymph nodes, delivering antigens to the immune cells and serving as an early warning system of pathogen invasion. The lymph nodes drain into conduits that ultimately merge into the thoracic duct which empties into the left subclavian vein.


9-Cis Retinoic Acid Promotes Lymphangiogenesis and Enhances Lymphatic Vessel Regeneration: Therapeutic Implications of 9-Cis Retinoic Acid for Secondary Lymphedema

Jan 2012

Choi I, Lee S, Chung HK, Lee YS, Kim KE, Choi D, Park EK, Yang D, Ecoiffier T, Monahan J, Chen W, Aguilar B, Lee HN, Yoo J, Koh CJ, Chen L, Wong AK, Hong YK.


1 University of Southern California Keck School of Medicine, Los Angeles, CA;



The lymphatic system plays a key role in tissue fluid homeostasis and lymphatic dysfunction due to genetic defects or lymphatic vessel obstruction can cause lymphedema, disfiguring tissue swellings often associated with fibrosis and recurrent infections without available cures to date. In this study, retinoic acids (RAs) were determined to be a potenttherapeutic agent that is immediately applicable to reduce secondary lymphedema.


We report that RAs promote proliferation, migration and tube formation ofcultured lymphatic endothelial cells (LECs) by activating FGF-receptor signaling. Moreover, RAs control the expression of cell-cycle checkpoint regulators such as p27(Kip1), p57(Kip2) and the aurora kinases through both an Akt-mediated non-genomic action and a transcription-dependent genomic action that is mediated by Prox1, a master regulator of lymphatic development. Moreover, 9-cisRAwas found to activate in vivo lymphangiogenesis in animals based on mouse trachea, matrigel plug and cornea pocket assays. Finally, we demonstrate that 9-cisRA can provide a strong therapeutic efficacy in ameliorating the experimental mouse tail lymphedema by enhancing lymphatic vessel regeneration.


These in vitro and animal studies demonstrate that 9-cisRA potently activates lymphangiogenesis and promotes lymphatic regeneration in an experimental lymphedema model, presenting it as a promising novel therapeutic agent to treat human lymphedema patients.


Lymphedema People Angiogenesis Related Pages:


Angiogenesis and Cancer

Angiogenesis and Cancer Control

Angiogenesis Inhibitors and Cancer


Lymphedema People Lymphangiogenesis Related Pages:

The Formation of Lymphatic Vessels and Its Importance in the Setting of Malignancy

Lymphangiogenesis Lymphedema and Cancer

Lymphangiogenesis and Gastric Cancer

Lymphangiogenesis in Head and Neck Cancer

Lymphangiogenesis and Kaposi's Sarcoma VEGF-C

Lymphangiogenesis in Wound Healing

A model for gene therapy of human hereditary lymphedema

VEGFR-3 Ligands and Lymphangiogenesis (1)

VEGFR-3 Ligands and Lymphangiogenesis (2)

VEGFR-3 Ligands and Lymphangiogenesis (3)

Vascular Endothelial Growth Factor; VEGF


VEGF-D is the strongest angiogenic and lymphangiogenic effector

Inhibition of Lymphatic Regeneration by VEGFR3

VEGFR3 and Metastasis in Prostate Cancer


Lymphedema People Genetics, Research, Lymphangiogenesis, Angiogenesis Forum


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Lymphatic Disorders Support Group @ Yahoo Groups

While we have a number of support groups for lymphedema... there is nothing out there for other lymphatic disorders. Because we have one of the most comprehensive information sites on all lymphatic disorders, I thought perhaps, it is time that one be offered.


Information and support for rare and unusual disorders affecting the lymph system. Includes lymphangiomas, lymphatic malformations, telangiectasia, hennekam's syndrome, distichiasis, Figueroa
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Page Updated: Dec. 24, 2011