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Cancer Research 67, 5211, June 1, 2007. doi: 10.1158/0008-5472.CAN-06-4676
© 2007 American Association for Cancer Research
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Cell, Tumor, and Stem Cell Biology

Distinctive Features of Angiogenesis and Lymphangiogenesis Determine Their Functionality during De novo Tumor Development

Alexandra Eichten1, William C. Hyun3 and Lisa M. Coussens1,2,3

1 Department of Pathology, 2 Cancer Research Institute, and 3 Comprehensive Cancer Center, University of California at San Francisco, San Francisco, California

Requests for reprints: Lisa M. Coussens, Department of Pathology and Comprehensive Cancer Center, University of California at San Francisco, 2340 Sutter Street, N-221, Box 0875, San Francisco, CA 94115. Phone: 415-502-6378; Fax: 415-514-0878; E-mail: coussens{at}cc.ucsf.edu.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Blood and lymphatic vasculature are essential components of all organs, responsible for maintaining organ fluid dynamics and tissue homeostasis. Although both vessel systems are composed of similar lineages of endothelial cells whose crude functions include fluid and cell transport, each system also possesses distinctive physiologic properties, enabling their distinctive functions in tissues. The role of hematogenous vasculature and development of angiogenic blood vessels during cancer development is well established; however, the role of lymphangiogenesis and structural/functional alterations occurring within lymphatic vessels during cancer development are incompletely understood. To assess premalignant versus malignant alterations in blood and lymphatic vasculature associated with squamous epithelial skin carcinogenesis, we assessed architectural and functional features of both vascular systems using a mouse model of de novo carcinoma development. We report that, as vasculature acquires angiogenic and/or lymphangiogenic properties, angiogenic blood vessels become leaky in premalignant tissue and at peripheries of carcinomas, where enlarged lymphatic capillaries efficiently drain increased tissue fluid, thereby maintaining tissue hemodynamics. In contrast, central regions of carcinomas exhibit elevated tissue fluid levels, compressed lymphatic lumina, and decreased vascular leakage, thus indicating impaired hemodynamics within solid tumors. Together, these data support the notion that therapeutic delivery of anticancer agents is best realized in premalignant tissues and/or at the peripheries of solid tumors where hemodynamic forces support drug delivery. Strategies to normalize intratumoral hemodynamics would therefore enhance therapeutic delivery to otherwise poorly accessible central regions of solid tumors. [Cancer Res 2007;67(11):5211–20]


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Blood and lymphatic vessels comprise two interdependent vascular networks in all tissues. Whereas blood vessels deliver blood cells, plasma proteins, and oxygen to tissues, lymphatics, composed of lymph-forming capillaries and collecting vessels, continually regulate interstitial fluid pressure by draining interstitial fluid and debris to maintain tissue homeostasis. When tissues are acutely damaged, activation of both vascular systems occurs as part of innate repair programs; once complete, both systems return to their homeostatic states. In contrast, sustained activation of one or both vascular networks is associated with some chronic disorders, such as rheumatoid arthritis (1) and psoriasis (2, 3), and contributes to disease pathogenesis. Cancer development is similarly associated with chronic activation of blood vasculature (i.e., angiogenesis) in premalignant and malignant tissues (4). Activation of angiogenic vasculature in premalignant tissue is characterized by increased proliferation of vascular endothelial cells (VEC) and sprouting of new immature leaky blood vessels from preexisting vascular beds (5). Increased leakage of plasma proteins from immature angiogenic vessels leads to increased interstitial fluid content, lymph formation, and drainage via lymphatic vessels back into the blood circulation (6). High interstitial fluid pressure (IFP), which forms a barrier to transcapillary transport, results in inefficient delivery of therapeutic drugs from vasculature into tumor stroma (7). It has been postulated that high IFP, common to many solid tumors, is in part a result of inefficient clearance of tissue fluid by lymphatic vessels (7). This postulate is supported by histochemical studies evaluating lymphatic architecture and diminished lumen diameters in archival human carcinomas and murine xenograft tumors (8–12). Studies evaluating functional changes in the status of lymphatic endothelial cells (LEC) and/or lymphatic vessels compared with VECs and blood vessels in premalignant tissues have not been well described.

To critically examine distinctive versus common physiologic and functional properties of blood versus lymphatic vasculature that accompany and/or contribute to cancer development, we used a transgenic mouse model of de novo epithelial squamous cell carcinogenesis (e.g., K14-HPV16 mice; ref. 13). HPV16 mice progress through well-defined premalignant stages before de novo carcinoma development, mirroring histopathologic stages observed during human cervical carcinogenesis (14). HPV16 mice develop hyperplastic skin lesions with 100% penetrance by 1 month of age that focally progress to dysplasia by 3 to 6 months (13). Precursor dysplasias undergo malignant conversion into varying grades of squamous cell carcinoma (SCC) in skin in 50% of mice that metastasize to regional lymph nodes with a 30% frequency (13, 15). Angiogenic vasculature is first evident in premalignant hyperplasias, development of which is linked to infiltration of innate immune cells (e.g., mast cells, granulocytes, and macrophages; refs. 15–18). Using this model, we assessed molecular, histopathologic, and functional variables of blood and lymphatic vasculature to reveal their distinctive physiologic properties at each stage of neoplastic development.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Animal Husbandry, Genotype, and Histopathologic Analyses
All animals were maintained within the University of California at San Francisco (UCSF) Laboratory for Animal Care barrier facility according to Institutional Animal Care and Use Committee procedures. HPV16 transgenic mice (19), preparation of tissue sections (ear skin) for histologic examination, and characterization of neoplastic stages based on H&E histopathology and cytokeratin intermediate filament expression have been described previously (13, 15, 20). Paraffin-embedded tissue sections were fixed by immersion in 10% neutral-buffered formalin, dehydrated through graded ethanol and xylenes, embedded in paraffin, cut with a Leica 2135 microtome into 5-µm-thick sections, and histopathologically examined following H&E staining and immunoreactivity of keratin intermediate filaments. Hyperplastic lesions were identified by a 2-fold increase in epidermal thickness and an intact granular cell layer with keratohyalin granules. Dysplastic lesions were characterized based on basal and spinous cell layers with hyperchromatic nuclei representing greater than half of the total epidermal thickness and incomplete terminal differentiation of keratinocytes. SCC was identified by abundance of abnormal mitotic figures and an invasive loss of integrity in epithelial basement membrane with clear development of malignant cell clusters proliferating in the dermis. SCCs were graded as has been described previously (17).

Flow Cytometry
Single cell suspensions were prepared from ear (n = 4) and tumor (n = 7) tissue as described previously (21). Cells were incubated for 10 min at 4°C with rat anti-mouse CD16/CD32 monoclonal antibody (mAb; BD Biosciences) at a 1:200 dilution in PBS/bovine serum albumin (BSA) to prevent nonspecific antibody binding. Subsequently, cells were washed and incubated for 20 min with phycoerythrin-conjugated anti-mouse CD31 (1:200; eBioscience) and anti-mouse podoplanin hybridoma supernatant (mAb clone 8.1.1, 1:1000; Developmental Studies Hybridoma Bank, University of Iowa). Cells were washed twice with PBS/BSA and incubated with Alexa Fluor 647–conjugated anti-hamster antibody (1:1,000; Molecular Probes) for 20 min. After two washes with PBS/BSA, 7-aminoactinomycin D (1:10; BD Biosciences) was added to discriminate between viable and dead cells. Data acquisition was done on a FACSCalibur using CellQuestPro software (BD Biosciences), and data analysis was done using FlowJo software (Tree Star, Inc.). Data shown represent the mean ± SE. P values of <0.05 were considered to be statistically significant.

Immunohistochemistry
Immunohistochemistry on paraffin-embedded tissue sections. Tissue sections were deparaffinized, briefly washed in PBS, and blocked for 15 min in blocking buffer (5.0% normal goat serum/2.5% BSA/PBS). Primary antibodies were diluted in 0.5x blocking buffer: rat anti-mouse CD31 (1:50; PharMingen), Syrian hamster anti-mouse podoplanin (1:200, clone 8.1.1; Developmental Studies Hybridoma Bank, University of Iowa), guinea pig anti-mouse cytokeratin pan (1:100; Progen Biotechnik GmbH), and rabbit anti-mouse lymphatic vessel endothelial receptor-1 (LYVE-1; 1:200; Upstate). Sections were incubated with primary antibody overnight at 4°C in a humidified chamber followed by three brief washes in PBS and subsequently incubated with secondary antibodies Alexa Fluor 488–conjugated anti-rat antibody (1:500; Molecular Probes), Alexa Fluor 633–conjugated anti-guinea pig antibody (1:500; Molecular Probes), Alexa Fluor 594–conjugated anti-hamster antibody (1:500; Molecular Probes), or Alexa Fluor 594/488–conjugated anti-rabbit antibody (1:500; Molecular Probes) for 2.0 h at room temperature in a humidified chamber. After five 3-min washes in PBS, the fluorescently stained sections were mounted in ProLong Gold mounting medium (Molecular Probes). Fluorescence was visualized using a laser scanning confocal microscope LSM510 META (Carl Zeiss MicroImaging, Inc.) and analyzed with Zeiss LSM Image Examiner. Ear tissue (n = 4/time point) and tumor tissue (n = 5) were assessed for lymphatic luminal status and blood and lymphatic vessel density by counting the number of vessels in five random high-power fields of vision per time point per tissue, tumor center, and periphery.

Immunohistochemical detection of proliferating endothelial cells. Fresh bromodeoxyuridine (BrdUrd; Sigma) suspended in water (1.0 mg/mL) was given to mice in drinking water ad libitum and changed every 2nd day for a total of 8 days. For fluorescent angiography, animals were tail vein injected with FITC-conjugated Lycopersicon esculentum (tomato) lectin (2.0 mg/mL; Vector Laboratories) 3.0 min before cardiac perfusion with the following reagents in chronological order: 15 mL of 1% paraformaldehyde/0.5% glutaraldehyde, 7.0 mL PBS, 15 mL PBS containing 1.0% BSA, and 15 mL of 4.0% paraformaldehyde at a constant flow rate of 7.0 mL/min. Tissue was harvested and fixed in 10% zinc-buffered formalin for 8 to 16 h at 4°C before further processing followed by dehydration through graded alcohols and xylene and embedding in paraffin. Paraffin sections (5 µm thick) were cut using a Leica 2135 microtome. Blood vasculature was detected by fluorescent angiography (22), and immunohistochemical analysis was done detecting lymphatic vessels (LYVE-1 reactivity), keratin-expressing epithelial cells (pan-keratin reactivity), proliferating nuclei (BrdUrd incorporation), and all nuclei as follows: Sections were deparaffinized, incubated in 2.0 N HCl for 1.0 h at room temperature, rinsed in PBS, and blocked for 30 min in blocking buffer (5.0% normal goat serum/2.5% BSA/PBS). Primary antibodies were diluted in 0.5x blocking buffer: guinea pig anti-mouse cytokeratin pan (1:100), biotinylated anti-BrdUrd (1:50; clone Br-3; Caltag), and rabbit anti-mouse LYVE-1 (1:200). Sections were incubated with primary antibody overnight at 4°C followed by three brief washes in PBS and subsequently incubated with secondary antibodies (all 1:500; Molecular Probes) Alexa Fluor 633–conjugated anti-guinea pig antibody, Alexa Fluor 546-streptavidin–conjugated antibody, and Alexa Fluor 594–conjugated anti-rabbit antibody for 2.0 h at room temperature in a humidified chamber. After five 3-min washes in PBS, the fluorescently stained sections were subjected to a nuclear staining with SYTO62 (0.7 µmol/L in H2O; Molecular Probes) for 10 min and then mounted in ProLong Gold mounting medium. Fluorescence was visualized using a laser scanning confocal microscope LSM510 META and analyzed with a Zeiss LSM Image Examiner. A minimum of 100 endothelial cell nuclei in control (n = 4), premalignant ear (n = 4), and tumor tissue (n = 7) was identified in tissue sections based on their morphologic protrusion into the vessel lumen, and the percentage of proliferating nuclei was determined.

Fluorescent Angiography and Immunohistochemistry of Whole-Mount Ear Tissue and Thick Frozen Tumor Sections
Whole-mounted ear tissue (n = 3 per time point) was prepared as described previously (23). Briefly, mice were anesthetized with a 2% isofluorane/98% oxygen mixture and 100 µL of FITC-conjugated L. esculentum (tomato) lectin (2.0 mg/mL) were injected into the tail vein and allowed to circulate for 3.0 min followed by cardiac perfusion with 35 mL of PBS-buffered 4% paraformaldehyde (pH 7.4; ref. 22). Ears were harvested, ventral and dorsal aspects were separated, and cartilage was removed from ventral aspects, which were then immersion fixed in PBS-buffered 4% paraformaldehyde (pH 7.4) overnight at 4°C and subjected to the following staining procedure under exclusion of light: Ventral aspects of ears were rinsed briefly in PBS containing 0.3% Triton X-100 (PBS-T) and blocked in PBS-T containing 3% goat serum overnight at 4°C. Tissue was subsequently incubated with primary rabbit anti-mouse LYVE-1 antibody (1:2,000; Upstate) diluted in PBS-T over two nights at 4°C. After five washes in PBS-T (5–10 min each) and a 2- to 3-h blocking incubation with PBS-T containing 3% goat serum at 4°C, tissue was incubated overnight at 4°C with a Alexa Fluor 594–conjugated anti-rabbit antibody (1:500; Molecular Probes). Tissue was then washed 10 times for 5 min at 4°C and mounted in ProLong Gold mounting medium.

Tumors (n = 5) were dissected, flash frozen in OCT compound, and cut into 200 µm sections and subsequently rinsed briefly in PBS-T and blocked in PBS-T containing 3% goat serum for 2.0 h at room temperature. Tissue was subsequently incubated with primary rabbit anti-mouse LYVE-1 antibody (1:2,000) diluted in PBS-T overnight at 4°C. After three 3-min washes in PBS-T, tissue was incubated for 3 h at room temperature with Alexa Fluor 594–conjugated anti-rabbit antibody (1:500). After three 3-min washes in PBS-T, tissue was mounted in ProLong Gold mounting medium. Fluorescence was visualized using a laser scanning confocal microscope LSM510 META and analyzed with a Zeiss LSM Image Examiner (Carl Zeiss MicroImaging).

Intravital Perfusion of Lymphatics
Intravital perfusion of lymphatics was modified from what has been described by Nagy et al. (24). Mice were anesthetized with a 2% isofluorane/98% oxygen mixture and cradled in a transparent acrylic resin mold (Syndicate Sales, Inc.). Ears (n = 6 per time point) were mounted flat on a resin support, held in place by silicone vacuum grease, and viewed in a dissecting microscope (Olympus). Colloidal carbon (Higgins non-waterproof black drawing ink; Sanford) was diluted 1:1 in PBS, filtered through a 0.22-µm filter, and injected i.d. through a prepulled borosilicate glass micropipette with an inner diameter of 1.0 µm (World Precision Instruments) attached to a 100 µL Hamilton syringe (World Precision Instruments). The micropipette was repeatedly injected into the dermis of the peripheral ear until a lymphatic vessel was entered. Colloidal carbon was then slowly injected. For lymphatic vessel diameter analysis, ear tissue (n = 3 per time point) was subsequently harvested and fixed overnight in acetone at 4°C. After fixation, the tissue was cleared in toluene 48 h at room temperature and mounted in Permount (Fisher Scientific). For photography, a digital camera (Nikon Coolpix 950) was used and the images were analyzed using Openlab software (Improvision). For in vivo analysis of colloidal carbon clearance, images were captured using a LUMAR microscope (Carl Zeiss MicroImaging) within 1.0 min of injection and again after 20 min. Both ears of negative littermate (–LM) and 4-month HPV16 mice (n = 3 each) were analyzed.

Miles Assay
Mice were anesthetized with a 2% isofluorane/98% oxygen mixture, and Evans blue dye (30 mg/kg in 100 µL PBS; Sigma-Aldrich) was injected into the tail vein. In some experiments, after 1 min, 30 µL of 5% mustard oil (phenyl isothiocyanate, 98%; Sigma-Aldrich) diluted in mineral oil (Sigma-Aldrich) or mineral oil as control were applied to the dorsal and ventral surfaces of the ear; the application process was repeated 15 min later. Evans blue dye was allowed to circulate for 30 min. Anesthetized mice were subsequently cardiac perfused with 1% paraformaldehyde in 0.05 mol/L citrate buffer (pH 3.5). Ears (n = 6 per time point; n = 3 per mineral oil/mustard oil treatment) were removed, blotted dry, and weighed. Tumors (n = 4) were dissected and cut into seven pieces representing six peripheral sections and center. Premalignant nontumor tissue was derived from neck, chest, and abdomen of non–tumor-bearing HPV16 animals. Evans blue dye was extracted from ears, nontumor tissue, and tumor pieces in 1.0 mL formamide for 48 h at 60°C and measured spectrophotometrically at 610 nm in a SpectraMax 340 (Molecular Devices). Data are expressed as mean ± SE. P values of <0.05 were considered to be statistically significant.

Tissue Fluid Determination
Mice were euthanized, and ears (n = 6 per time point) and/or tumors (n = 4) were removed and weighed (wet weight). Tumors were dissected into six peripheral sections and tumor centers. Premalignant nontumor tissue was derived from neck, chest, and abdomen of non–tumor-bearing HPV16 animals. Tissue was snap frozen, lyophilized, and reweighed (dry weight). The difference between wet and dry weight reflects fluid tissue component, although dry weight reflects solid tissue component. Data are expressed as mean ± SE. P values of <0.05 were considered to be statistically significant.

In vivo High Molecular Dextran Injection
Functionality of initial lymphatics was measured as uptake and drainage of interstitial fluid containing high molecular weight (2,000,000 Da) dextran conjugated to the fluorophore tetramethylrhodamine isothiocyanate (TRITC; 10 mg/mL in PBS; Sigma) following i.d. injection. Drainage of TRITC-dextran was monitored in –LM (n = 6) and 4-month HPV16 (n = 6) ears over 20 min using a LUMAR microscope.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Endothelial composition of blood and lymphatic vasculature during neoplastic progression. To evaluate the endothelial cell composition of premalignant versus malignant tissues (skin) during squamous carcinogenesis, we determined the relative percentage of endothelial cells at each stage of neoplastic progression in HPV16 mice, compared with wild-type negative littermate (–LM) controls, by assessing expression of the pan-endothelial cell adhesion molecule CD31 (25), present on both VECs and LECs, albeit at varying levels of expression (26), using flow cytometry of single cell suspensions (Fig. 1A ). We found a progressive increase in the relative percentage of CD31+ endothelial cells at each distinct premalignant stage (Fig. 1A). Carcinomas on the other hand contained a lower percentage of endothelial cells by comparison with premalignant tissues (Fig. 1A), although tumor vasculature possessed angiogenic characteristics (16).


Figure 1
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  		Figure 1. Increased endothelial cell content in neoplastic tissue. A, flow cytometric analysis of normal, premalignant, and carcinoma tissue detecting the pan-endothelial cell (EC) marker CD31. Percentage CD31+ endothelial cells out of the total number of cells analyzed. B, distinction of VECs from LECs by flow cytometry based on CD31 and podoplanin expression. Inset, example plot from dysplastic (4 mo old) HPV16 ear tissue biopsy. C, immunohistochemical detection of CD31 and lymphatic-specific markers podoplanin and LYVE-1 distinguishes between CD31Hi/podoplanin–/LYVE-1– blood vessels (open arrow) and CD31Lo/podoplanin+/LYVE-1+ lymphatic vessels (solid arrow). e, epidermis; d, dermis; c, cartilage. Dashed line, basement membrane. Bar, 50 µm. *, P ≤ 0.05, two-tailed unpaired nonparametric Mann-Whitney U test.

 
To determine the hematogenous versus lymphatic composition of these endothelial cell populations, we further used flow cytometry of single cell suspensions and quantitatively examined endothelial cells expressing CD31 and the mucin-type transmembrane glycoprotein podoplanin (Fig. 1B). Histochemically, podoplanin-positive endothelial cells are also LYVE-1 positive (Fig. 1C) and thus allow distinction between CD31Hi/podoplanin– VECs from CD31Lo/podoplanin+ LECs. The average difference in CD31 signal intensity between CD31Hi VECs and CD31Lo LECs was 28%. A progressive increase in both VEC and LEC populations was found during premalignant progression; however, at the tumor stage, VECs represented a larger proportion of the total CD31+ cells (Fig. 1B).

To determine the degree to which VECs and LECs exhibited proliferative behavior at each neoplastic stage, we evaluated endothelial cell proliferation in blood and lymphatic vessels as a function of BrdUrd incorporation. BrdUrd analysis of endothelial cells was limited to vessels with open lumina so as to distinguish BrdUrd-positive endothelial cells [whose nuclei protrude into vessel lumina (Fig. 2A )] from BrdUrd-positive mural cells (whose nuclei protrude toward interstitia). BrdUrd-positive CD31Hi/LYVE-1– VECs and CD31Lo/LYVE-1+ LECs were not observed in skin sections of –LM mice, reflecting the quiescent nature of endothelial cells in homeostatic tissue (Fig. 2B). In contrast, BrdUrd-positive VECs and LECs were readily identified throughout premalignant progression albeit with distinct profiles (Fig. 2A and B). Whereas BrdUrd-positive VEC percentages steadily increased at each distinct premalignant stage (Fig. 2B), percentages of premalignant BrdUrd-positive LECs remained constant, albeit higher than in –LM skin (Fig. 2B). In carcinomas, open lumen blood and lymphatic vessels possessed distinct BrdUrd-positive endothelial cell distributions dependent on (a) SCC grade (lower-grade, well-differentiated SCC-I versus higher-grade, moderate-poorly differentiated SCC-II) and (b) SCC location (tumor center versus tumor periphery). Open lumen blood and lymphatic vessels were identified at peripheries of both SCC-I and SCC-II carcinomas and in centers of SCC-I (Fig. 2A and B). In the center of SCC-IIs, open lumens were only found in blood vessels and not in lymphatic vessels (Fig. 2B). Analysis of proliferating endothelial cells in vessels with open lumens in carcinomas revealed an increase in both BrdUrd-positive VECs and LECs (Fig. 2B) compared with precursor premalignant lesions. VECs and LECs both evidenced increased presence of BrdUrd-positive endothelial cells in tumor centers and peripheries of SCC-I (Fig. 2A and B); however, in SCC-II, proliferating VECs were found in tumor centers and in the periphery, whereas proliferating LECs were limited to tumor periphery (Fig. 2A and B).


Figure 2
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  		Figure 2. VEC and LEC proliferation in premalignant and carcinoma tissue. A, quiescent and proliferating (red staining) endothelial cell nuclei (solid arrows) in blood (green staining) and lymphatic (yellow staining) vessels in premalignant and malignant (white staining for cytokeratin-positive keratinocytes; open arrow) tissue. Bar, 50 µm. e, epidermis; d, dermis. B, quantitative analysis of proliferating VECs and LECs in –LM, premalignant, and carcinoma tissue. Proliferating LECs were identified in the periphery and center of well-differentiated grade 1 SCCs (SCC-I) but limited to periphery of less-differentiated grade 2 SCCs. Absence of open lumen lymphatic vessels SCC-II centers precluded analysis of LECs in that locale. *, P ≤ 0.05, two-tailed unpaired nonparametric Mann-Whitney U test. Dashed line, basement membrane. Blue staining, SYTO62-nuclear counterstain.

 
Blood and lymphatic vessel density and diameter during neoplastic progression. To determine whether increased proliferation of endothelial cells in blood or lymphatic vessels yielded increased vessel densities and sprouting of new vessels or alternatively only resulted in increased girth (diameter) of preexisting vessels, we assessed blood and lymphatic vessel diameters and density (number of vessels per random high-power field of vision) at each neoplastic stage (Fig. 3 ). Using fluorescent angiography to assess blood vessel diameter in –LM and histopathologically staged neoplastic lesions, we found that blood vessel diameters significantly increased at each premalignant stage (Fig. 3A). In SCC-I and SCC-IIs, blood vessel diameter was similar in both tumor centers and periphery (Fig. 3A). Assessment of LYVE-1+ lymphatic capillary diameter similarly revealed a progressive increase during premalignant progression, albeit less dramatic compared with blood vessels (Fig. 3A). Because collecting lymphatic vessels express LYVE-1 at low levels, we used intralymphatic perfusion of colloidal carbon to assess their diameter in –LM and premalignant tissue and observed a progressive increase in diameter in premalignant tissue (Fig. 3B). Lymphatic vessel diameters in carcinomas at SCC-I peripheries were similar to late dysplastic tissues, whereas lymphatic vessels at SCC-II peripheries were further dilated (Fig. 3A). LYVE-1+ lymphatic vessel diameters in SCC-I centers were decreased by comparison with peripheries but were also significantly greater than those found in the center of SCC-IIs (Fig. 3A), thus indicating that proliferation of endothelial cells results in vessels with enhanced diameters in both vessel systems in premalignant and SCC tissues.


Figure 3
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  		Figure 3. Vessel architecture and diameter during neoplastic progression. A, quantitative assessment of blood vessel diameter following fluorescent angiography (green staining) and lymphatic capillary diameter based on immunodetection of LYVE-1 (red staining) in whole-mount tissue sections of biopsies from –LM ear, premalignant ear, and carcinomas. IHC, immunohistochemistry. Bar, 50 µm. B, assessment of collecting lymphatic vessels following intralymphatic perfusion with colloidal carbon (black staining). Red-colored vessels, reveal visibility of blood vessels. Bar, 1.0 mm. C, quantitative analysis of vessel density of CD31Hi/LYVE-1– blood (open arrows) and CD31Lo/LYVE-1+ lymphatic (solid arrows) vessel structures per random microscopic field of vision (FOV) in –LM, premalignant, and tumor tissue. Bar, 50 µm. *, P ≤ 0.05, two-tailed unpaired nonparametric Mann-Whitney U test.

 
To evaluate the sprouting activity of new vessels as a result of increased VEC and/or LEC proliferation, we assessed vessel densities by quantitatively examining CD31Hi/LYVE-1– (blood) and CD31Lo/LYVE-1+ (lymphatic) vessels in tissue sections (Fig. 3C). We found significant increases in the number of CD31Hi/LYVE-1– blood vessels per field of vision at each premalignant stage (Fig. 3C). In carcinomas, the number of CD31Hi/LYVE-1– blood vessels remained comparable with late dysplastic tissue independent of localization or SCC grade (Fig. 3C). In contrast, the progressive increase in lymphatic vessel diameter was not accompanied by increased vessel density as shown by constant numbers of CD31Lo/LYVE-1+ lymphatic vessels in premalignant tissues and in periphery and centers of SCC-I and periphery of SCC-IIs (Fig. 3C). CD31Lo/LYVE-1+ lymphatic vessels were decreased in SCC-II centers (Fig. 3C). Together, these data indicate that proliferation of endothelial cells in the two vessel systems results in distinct phenotypic responses. Whereas blood vessels increase in diameter and density, lymphatic vessels increase only in diameter without an increase in density, thus reflecting increased sprouting of new blood vessels (i.e., classic angiogenesis) as opposed to increased girth without sprouting of lymphatic vessels.

Altered tissue fluid homeostasis and lymphatic functionality in neoplastic tissue. Altered proliferation and/or diameter of either vessel system could variably alter their physiologic functions in tissue and thereby alter tissue hemodynamics and/or fluid content. To assess functionality of blood vessels, we evaluated their perfusability on fluorescein-conjugated L. esculentum injection (Fig. 3A) and their leakage characteristics using a modified Miles assay (Fig. 4A ; ref. 27). These studies revealed increased extravasation of Evans blue tracer in hyperplastic and dysplastic HPV16 tissue compared with –LM skin (Fig. 4A). In spite of the increased density and diameter of perfused, thus functional blood vessels in tumor tissue (Fig. 3A and C), we found decreased vascular leakage in carcinomas where distinct extravasation characteristics were found in tumor centers versus periphery (Fig. 4A).


Figure 4
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  		Figure 4. Functional interdependence of blood and lymphatic vessels during neoplastic progression. A, plasma protein extravasation and blood vessel leakiness in –LM, premalignant, and tumor tissue as assessed by Miles assay. B, quantification of tissue fluid in –LM, premalignant, and tumor tissue. C, induction of acute vascular leakage following mustard oil (MO) application in ear tissue of –LM and dysplastic HPV16 mice results in significantly increased vascular leakage, 16.5-fold (P ≤ 0.0001, two-tailed unpaired nonparametric Mann-Whitney U test) and 3.5-fold (P = 0.0043, two-tailed unpaired nonparametric Mann-Whitney U test), respectively. MinO, mineral oil. D, mustard oil–induced leakage results in significantly increased tissue fluid in –LM controls (8.0%) and premalignant tissue (8.0%). *, P ≤ 0.05, two-tailed unpaired nonparametric Mann-Whitney U test.

 
To examine whether increased extravasation of plasma proteins resulted in increased tissue fluid content, we evaluated the relationship between blood vessel leakage (Fig. 4A) and lymphatic drainage by determining tissue fluid amounts at each neoplastic stage (Fig. 4B) and found that tissue fluid in premalignant HPV16 ear tissue was significantly higher than –LM tissue and further increased in tumor peripheries, with the central regions of SCCs having the highest fluid content (Fig. 4B). Because a major function of lymphatics is to drain excess tissue fluid, one interpretation of these data would be that the drainage capacity of premalignant lymphatics had been exceeded due to lymphatic dysfunction perhaps reflected by tissue fluid accumulation. To access this hypothesis, we further induced vascular leakage acutely by topical application of mustard oil (compared with mineral oil) and assessed Evans blue dye leakage into dysplastic HPV16 ear skin versus –LM. As expected, mustard oil application further increased leakage of plasma proteins out of blood vessels in both –LM and HPV16 skin, 16.5- and 3.5-fold, respectively (Fig. 4C); however, these increases resulted in similar accumulations of tissue fluid content in –LM (8%) and dysplastic HPV16 (8%) tissue (Fig. 4D), indicating that clearance of tissue fluid by premalignant lymphatics is not at maximum capacity. Thus, whereas lymphatics in acutely activated homeostatic tissue are more efficient at clearing excess tissue fluid, functional lymphatics in premalignant tissue maintain tissue fluid dynamics to a similar degree.

To directly assess lymphatic functionality in premalignant tissue, we i.d. injected high molecular weight, fluorescently labeled dextran (TRITC-dextran) tracer and monitored uptake and drainage kinetics by lymphatic capillaries over time in –LM and dysplastic ear skin (Fig. 5A ). Lymphatic capillaries were rapidly labeled by tracer in both control and HPV16 ears, indicating rapid uptake. Twenty minutes following injection, tracer-containing capillaries were detectable, indicating continuous lymph formation, thus showing their functional status.


Figure 5
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  		Figure 5. Functionality of lymphatic vasculature during premalignant progression. A, functionality of initial lymphatics (arrows) was assessed following i.d. injection of high molecular weight (2 MDa) TRITC-conjugated (red staining) dextran in –LM and dysplastic HPV16 ears and clearance monitoring over a 20-min time period. B, draining capability of collecting lymphatic vessels in –LM and dysplastic HPV16 ear tissue was assessed by intralymphatic injection of colloidal carbon (black staining) and clearance monitoring over 20 min. Colloidal carbon in the lymphatic vasculature (arrow) was completely drained after 20 min.

 
Functionality of collecting lymphatic vessels in premalignant tissue was also evaluated following intralymphatic injection of colloidal carbon (Fig. 5B). Colloidal carbon is excluded by lymphatic capillaries during lymph formation due to its large molecular weight.4 Direct injection of colloidal carbon into dermal lymphatics of –LM and dysplastic ear skin revealed efficient drainage of tracer from collecting lymphatic vessels in both tissues within 20 min (Fig. 5B), thus showing their similar functionality. Together, these analyses reveal functionality of lymphatic capillaries and collecting vessels in premalignant tissues based on their ability to efficiently clear tracer and fluid (Fig. 5A and B) through their open lumina (Fig. 3A and B). Direct analysis of lymphatic functionality in tumor tissue was hampered by technical difficulties doing direct intralymphatic colloidal carbon injection as well as analysis of interstitial drainage using smaller molecular weight dyes (colorimetric or fluorescent) readily taken up by functional lymphatics. Despite these failed attempts to analyze lymphatic functionality in tumor tissue, the implication that open luminal status is associated with vessel functionality in premalignant tissue, in combination with our assessment of tissue fluid content in tumor centers versus tumor periphery (Fig. 4B), leads us to conclude that lymphatics with open lumina in premalignant tissue, at peripheries of SCCs and centers of grade 1 SCCs (Fig. 3C), retain functionality where they efficiently clear interstitial fluid, as opposed to tumor centers of less-differentiated SCCs where lymphatic lumens are predominantly compressed and exhibit diminished fluid drainage capacity (Fig. 3C, bottom right), a characteristic feature of poorly differentiated carcinomas (28–30).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Data presented herein indicate that blood and lymphatic vasculature undergo distinct physiologic and architectural alterations during de novo neoplastic progression accompanying carcinoma development. Whereas endothelial cells of each network acquire proliferative capabilities as neoplasia ensues, resultant phenotypes of blood versus lymphatic vessels are distinct. Whereas blood vessels undergo classic angiogenic changes (e.g., increased endothelial cell proliferation, vessel diameter, density, and leakage), lymphatic vessels evidence no alterations to support the notion that sprouting of new vessels occurs. Instead, the increased level of proliferating LECs present in lymphatic vessels enhances diameter of vessels that efficiently clear interstitial fluid in premalignant tissues and peripheries of carcinomas, but not in their centers, areas where interstitial fluid content remains high. Together, these studies show that neoplasia-associated angiogenesis, as opposed to lymphangiogenesis, is a distinct vascular process even when initiated by similar physiologic stimuli.

Endothelial cell proliferation. Endothelial cells in quiescent tissues divide approximately once every 2 to 3 years if unstimulated (31). During cancer development, however, it is well established that local tissue levels of the proangiogenic factor vascular endothelial growth factor (VEGF)-A increase and, as a result, enhance proliferation of VECs and sprouting of new blood vessels from preexisting vascular beds (5). In contrast, lymphangiogenesis (e.g., induction of LEC proliferation) in human cancers (32, 33) correlates instead with increased tissue levels of VEGF-C and VEGF-D (34–36). Accordingly, exogenous expression of VEGF-C or VEGF-D in experimental murine tumor xenograft models induces LEC proliferation and subsequent lymphangiogenesis (37, 38), whereas exogenous VEGF-A expression induces not only an angiogenic response but also lymphangiogenesis in nontumor (24) and tumor tissue (39). It is not clear, however, if LEC responses to increased VEGF-A levels represent a direct intrinsic response of LECs or, instead, an indirect physiologic response resulting from increased tissue fluid due to increased leakage of plasma proteins from newly formed immature angiogenic blood vessels.

In this study, we found evidence of angiogenesis as well as lymphangiogenesis in premalignant tissues of HPV16 mice and in emergent malignant carcinomas. Patterns of LEC proliferation varied from that observed with VECs, thus indicating distinct responses between the two endothelial cell subpopulations to proangiogenic and/or prolymphangiogenic growth factors present in their microenvironment. Whereas VEGF-A mRNA levels have been reported to remain constant during premalignant angiogenesis in a UV-induced murine skin carcinoma model (40), enhanced VEC proliferation in HPV16 mice is paralleled by progressive increases in VEGF-A mRNA expression (41) and protein levels (42). About LEC proliferation, our analyses indicate that, although LECs in premalignant tissue evidence enhanced proliferative status compared with homeostatic tissue, their proliferation does not significantly increase until malignant conversion occurs. Because we have observed increased mRNA levels of VEGF-C in fully malignant carcinomas (data not shown), our interpretation of these data is that LECs respond modestly to increased VEGF-A levels in premalignant tissue but more are responding to increased microenvironmental stress and need to clear fluid from premalignant tissue resultant from leaky immature angiogenic blood vessels.

Vascular sprouting. VEGF-A is a major proangiogenic factor well documented to induce VEC proliferation and vascular sprouting (43). Sprouting of the lymphatic vasculature, however, depends on VEGF-C in experimental chick chorioallantoic membrane assays, wound healing, and embryonic development (44–46). Recent studies in murine tumor models (transgenic and xenograft models) provide evidence that increased levels of VEGF-C and/or VEGF-D promote tumor lymphangiogenesis, including lymphatic vessel sprouting (47–50). In contrast to these experimental tumor models, no increase in lymphatic vessel density was observed in HPV16 tissues, indicating that LEC proliferation in a de novo tumor model lacking exogenous expression of VEGF-C and/or VEGF-D does not result in lymphatic vessel sprouting but instead contributed to increased diameter (girth) of existing lymphatics to enable rapid clearance of tissue fluid resulting from increased leakage of plasma proteins from angiogenic blood vessels.

Tissue fluid dynamics in premalignant and SCC tissue. Blood and lymphatic vessels are functionally interdependent vascular systems (6) that together regulate tissue fluid dynamics and influence tissue homeostasis. In a UV-induced skin carcinogenesis model, Hagendoorn et al. (40) revealed alterations in blood vasculature, architectural changes, and impairment of lymphatic vessels along with increased IFP in premalignant tissues and thus hypothesized that blood and lymphatic vasculature loose their functional interdependence during premalignant progression. Our data, however, indicate that, during squamous carcinogenesis in HPV16 mice, the two vessel systems retain a functional interrelationship in premalignant tissue and in well-differentiated carcinomas as evidenced by increased blood vessel leakage accompanied by retained ability to clear fluid, albeit in tissue with higher tissue fluid content than normal.

Lymphatic vessel functionality. We observed that intratumoral lymphatic vessel functionality inversely correlates with SCC grade. In the published literature, there are contradicting reports on the existence of intratumoral lymphatics in human (9, 10, 32, 33, 51) and experimental rodent (37, 38, 40, 49, 52) tumors, where only peripheral lymphatic structures are hypothesized to retain functionality, whereas intratumoral lymphatic structures were rendered nonfunctional due to compression by rapidly proliferating tumor cells (52). Similarly, we found functional lymphatic vessels at tumor peripheries but also observed open lumen/functional lymphatic vessels in centers of low-grade SCCs. Our interpretation of these observations is that functional properties of higher-grade, less-differentiated carcinomas correlate with, and is effected by, limited lymphatic vessel functionality that, in turn, likely regulates tumor cell physiology and/or behavior.

In summary, by examining proliferation, diameter, density, and functionality of blood versus lymphatic vasculature during de novo cancer development, we have revealed unique features of each vasculature that illuminates their interdependency during premalignant progression as opposed to their independence in higher-grade carcinomas. Maintenance of the interdependence between blood and lymphatic systems during premalignancy and in low-grade carcinomas enables efficient fluid transport from leaky blood vasculature via interstitia into draining, open lumen lymphatic vessels, which would result in minimizing IFP. The closed nature of lymphatic vessels in higher-grade carcinomas would exclude this and support elevated IFPs common to less-differentiated carcinomas. The implication of these findings supports the notion that delivery of anticancer therapeutics would best be realized in premalignant tissue, in situ carcinomas, or carcinomas displaying well-differentiated characteristics, where tissue hemodynamics better support drug delivery.


   Acknowledgments
 
Grant support: Serono Foundation for the Advancement of Medical Science (A. Eichten) and NIH grants CA72006, CA94168, and CA098075, Sandler Program in Basic Sciences, National Technology Center for Networks and Pathways grant U54 RR020843, and Department of Defense Breast Cancer Research Program Era of Hope Scholar Award BC051640 (L.M. Coussens).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Drs. K. de Visser, D. Soto, and N.E. Sounni for critical comments and suggestions; Lidiya Korets, Aleksandr Rudik, and Nadia Milshteyn for technical assistance in animal husbandry; Eva Soliven and the UCSF Comprehensive Cancer Center (CCC) Mouse Pathology Shared Resource for tissue sectioning; members of the UCSF CCC Laboratory for Cell Analysis for microscopic and cytometric assistance; and members of the Coussens laboratory for encouragement and critical discussion.


   Footnotes
 
4 J. Nagy, personal communication. Back

Received 12/20/06. Revised 3/30/07. Accepted 4/ 9/07.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

  1. Folkman J. Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat Med 1995;1:27–31.[CrossRef][Medline]
  2. Braverman IM, Sibley J. Role of the microcirculation in the treatment and pathogenesis of psoriasis. J Invest Dermatol 1982;78:12–7.[CrossRef][Medline]
  3. Detmar M, Brown LF, Claffey KP, et al. Overexpression of vascular permeability factor/vascular endothelial growth factor and its receptors in psoriasis. J Exp Med 1994;180:1141–6.[Abstract/Free Full Text]
  4. Folkman J, Watson K, Ingber D, Hanahan D. Induction of angiogenesis during the transition from hyperplasia to neoplasia. Nature 1989;339:58–61.[CrossRef][Medline]
  5. Bergers G, Benjamin LE. Angiogenesis: tumorigenesis and the angiogenic switch. Nat Rev Cancer 2003;3:401–10.[CrossRef][Medline]
  6. Swartz MA. The physiology of the lymphatic system. Adv Drug Deliv Rev 2001;50:3–20.[CrossRef][Medline]
  7. Heldin CH, Rubin K, Pietras K, Ostman A. High interstitial fluid pressure—an obstacle in cancer therapy. Nat Rev Cancer 2004;4:806–13.[CrossRef][Medline]
  8. Beasley NJ, Prevo R, Banerji S, et al. Intratumoral lymphangiogenesis and lymph node metastasis in head and neck cancer. Cancer Res 2002;62:1315–20.[Abstract/Free Full Text]
  9. Franchi A, Gallo O, Massi D, Baroni G, Santucci M. Tumor lymphangiogenesis in head and neck squamous cell carcinoma: a morphometric study with clinical correlations. Cancer 2004;101:973–8.[CrossRef][Medline]
  10. Giorgadze TA, Zhang PJ, Pasha T, et al. Lymphatic vessel density is significantly increased in melanoma. J Cutan Pathol 2004;31:672–7.[CrossRef][Medline]
  11. Boucher Y, Brekken C, Netti PA, Baxter LT, Jain RK. Intratumoral infusion of fluid: estimation of hydraulic conductivity and implications for the delivery of therapeutic agents. Br J Cancer 1998;78:1442–8.[Medline]
  12. Padera T, Stoll B, Tooredman J, Capen D, di Tomaso E, Jain R. Cancer cells compress intratumour vessels. Nature 2004;427:695.[CrossRef][Medline]
  13. Coussens LM, Hanahan D, Arbeit JM. Genetic predisposition and parameters of malignant progression in K14-HPV16 transgenic mice. Am J Pathol 1996;149:1899–917.[Abstract]
  14. Smith-McCune K, Zhu Y-H, Hanahan D, Arbeit J. Angiogenesis and VEGF mRNA upregulation in progressive squamous carcinogenesis: cross-species comparison of human cervix and K14-HPV16 transgenic mice. Cancer Res 1997;57:1294–300.[Abstract/Free Full Text]
  15. van Kempen LCL, Rhee JS, Dehne K, Lee J, Edwards DR, Coussens LM. Epithelial carcinogenesis: dynamic interplay between neoplastic cells and their microenvironment. Differentiation 2002;70:610–23.[CrossRef][Medline]
  16. Coussens LM, Raymond WW, Bergers G, et al. Inflammatory mast cells up-regulate angiogenesis during squamous epithelial carcinogenesis. Genes Dev 1999;13:1382–97.[Abstract/Free Full Text]
  17. Coussens LM, Tinkle CL, Hanahan D, Werb Z. MMP-9 supplied by bone marrow-derived cells contributes to skin carcinogenesis. Cell 2000;103:481–90.[CrossRef][Medline]
  18. Junankar S, Eichten A, Kramer A, de Visser KE, Coussens LM. Analysis of immune cell infiltrates during squamous carcinoma development. J Invest Dermatol 2006;126:36–43.[Medline]
  19. Arbeit JM, Munger K, Howley PM, Hanahan D. Progressive squamous epithelial neoplasia in K14-human papillomavirus type 16 transgenic mice. J Virol 1994;68:4358–68.[Abstract/Free Full Text]
  20. Daniel D, Meyer-Morse N, Bergsland EK, Dehne K, Coussens LM, Hanahan D. Immune enhancement of skin carcinogenesis by CD4+ T cells. J Exp Med 2003;197:1017–28.[Abstract/Free Full Text]
  21. de Visser KE, Korets LV, Coussens LM. Early neoplastic progression is complement independent. Neoplasia 2004;6:768–76.[CrossRef][Medline]
  22. Thurston G, Baluk P, Hirata A, McDonald DM. Permeability-related changes revealed at endothelial cell borders in inflamed venules by lectin binding. Am J Physiol 1996;271:2547–62.
  23. Eichten A, Shen H-CJ, Coussens LM. Three-dimensional visualization of blood and lymphatic vasculature in tissue whole mounts using confocal microscopy. In: Robinson JP, editor. Current protocols in cytometry. New Jersey: John Wiley & Sons, Inc.; 2005. p. 12.5.1–12.5.11.
  24. Nagy JA, Vasile E, Feng D, et al. Vascular permeability factor/vascular endothelial growth factor induces lymphangiogenesis as well as angiogenesis. J Exp Med 2002;196:1497–506.[Abstract/Free Full Text]
  25. DeLisser HM, Newman PJ, Albelda SM. Molecular and functional aspects of PECAM-1/CD31. Immunol Today 1994;15:490–5.[CrossRef][Medline]
  26. Cursiefen C, Chen L, Borges LP, et al. VEGF-A stimulates lymphangiogenesis and hemangiogenesis in inflammatory neovascularization via macrophage recruitment. J Clin Invest 2004;113:1040–50.[CrossRef][Medline]
  27. Miles AA, Miles EM. Vascular reactions to histamine, histamine-liberator and leukotaxine in the skin of guinea pigs. J Physiol 1952;118:228–57.[Free Full Text]
  28. Nathanson SD, Nelson L. Interstitial fluid pressure in breast cancer, benign breast conditions, and breast parenchyma. Ann Surg Oncol 1994;1:333–8.[Medline]
  29. Curti BD, Urba WJ, Alvord WG, et al. Interstitial pressure of subcutaneous nodules in melanoma and lymphoma patients: changes during treatment. Cancer Res 1993;53:2204–7.[Abstract/Free Full Text]
  30. Gutmann R, Leunig M, Feyh J, et al. Interstitial hypertension in head and neck tumors in patients: correlation with tumor size. Cancer Res 1992;52:1993–5.[Abstract/Free Full Text]
  31. Hirst DG, Denekamp J, Hobson B. Proliferation studies of the endothelial and smooth muscle cells of the mouse mesentery after irradiation. Cell Tissue Kinet 1980;13:91–104.[Medline]
  32. Sipos B, Klapper W, Kruse ML, Kalthoff H, Kerjaschki D, Kloppel G. Expression of lymphangiogenic factors and evidence of intratumoral lymphangiogenesis in pancreatic endocrine tumors. Am J Pathol 2004;165:1187–97.[Abstract/Free Full Text]
  33. Koukourakis MI, Giatromanolaki A, Sivridis E, et al. LYVE-1 immunohistochemical assessment of lymphangiogenesis in endometrial and lung cancer. J Clin Pathol 2005;58:202–6.[Abstract/Free Full Text]
  34. Karpanen T, Alitalo K. Lymphatic vessels as targets of tumor therapy? J Exp Med 2001;194:F37–42.[Free Full Text]
  35. Pepper MS. Lymphangiogenesis and tumor metastasis: myth or reality? Clin Cancer Res 2001;7:462–8.[Abstract/Free Full Text]
  36. Achen MG, McColl BK, Stacker SA. Focus on lymphangiogenesis in tumor metastasis. Cancer Cell 2005;7:121–7.[CrossRef][Medline]
  37. Skobe M, Hawighorst T, Jackson DG, et al. Induction of tumor lymphangiogenesis by VEGF-C promotes breast cancer metastasis. Nat Med 2001;7:192–8.[CrossRef][Medline]
  38. Karpanen T, Egeblad M, Karkkainen MJ, et al. Vascular endothelial growth factor C promotes tumor lymphangiogenesis and intralymphatic tumor growth. Cancer Res 2001;61:1786–90.[Abstract/Free Full Text]
  39. Hirakawa S, Kodama S, Kunstfeld R, Kajiya K, Brown LF, Detmar M. VEGF-A induces tumor and sentinel lymph node lymphangiogenesis and promotes lymphatic metastasis. J Exp Med 2005;201:1089–99.[Abstract/Free Full Text]
  40. Hagendoorn J, Tong R, Fukumura D, et al. Onset of abnormal blood and lymphatic vessel function and interstitial hypertension in early stages of carcinogenesis. Cancer Res 2006;66:3360–4.[Abstract/Free Full Text]
  41. Smith-McCune K, Zhu YH, Hanahan D, Arbeit J. Cross-species comparison of angiogenesis during the premalignant stages of squamous carcinogenesis in the human cervix and K14-HPV16 transgenic mice. Cancer Res 1997;57:1294–300.[Abstract/Free Full Text]
  42. de Visser KE, Korets LV, Coussens LM. De novo carcinogenesis promoted by chronic inflammation is B lymphocyte dependent. Cancer Cell 2005;7:411–23.[CrossRef][Medline]
  43. Leung DW, Cachianes G, Kuang WJ, Goeddel DV, Ferrara N. Vascular endothelial growth factor is a secreted angiogenic mitogen. Science 1989;246:1306–9.[Abstract/Free Full Text]
  44. Oh SJ, Jeltsch MM, Birkenhager R, et al. VEGF and VEGF-C: specific induction of angiogenesis and lymphangiogenesis in the differentiated avian chorioallantoic membrane. Dev Biol 1997;188:96–109.[CrossRef][Medline]
  45. Paavonen K, Puolakkainen P, Jussila L, Jahkola T, Alitalo K. Vascular endothelial growth factor receptor-3 in lymphangiogenesis in wound healing. Am J Pathol 2000;156:1499–504.[Abstract/Free Full Text]
  46. Jeltsch M, Kaipainen A, Joukov V, et al. Hyperplasia of lymphatic vessels in VEGF-C transgenic mice. Science 1997;276:1423–5.[Abstract/Free Full Text]
  47. Mandriota SJ, Jussila L, Jeltsch M, et al. Vascular endothelial growth factor-C-mediated lymphangiogenesis promotes tumour metastasis. EMBO J 2001;20:672–82.[CrossRef][Medline]
  48. Stacker SA, Caesar C, Baldwin ME, et al. VEGF-D promotes the metastatic spread of tumor cells via the lymphatics. Nat Med 2001;7:186–91.[CrossRef][Medline]
  49. Skobe M, Hamberg LM, Hawighorst T, et al. Concurrent induction of lymphangiogenesis, angiogenesis, and macrophage recruitment by vascular endothelial growth factor-C in melanoma. Am J Pathol 2001;159:893–903.[Abstract/Free Full Text]
  50. He Y, Kozaki K, Karpanen T, et al. Suppression of tumor lymphangiogenesis and lymph node metastasis by blocking vascular endothelial growth factor receptor 3 signaling. J Natl Cancer Inst 2002;94:819–25.[Abstract/Free Full Text]
  51. Williams CS, Leek RD, Robson AM, et al. Absence of lymphangiogenesis and intratumoural lymph vessels in human metastatic breast cancer. J Pathol 2003;200:195–206.[CrossRef][Medline]
  52. Padera TP, Kadambi A, di Tomaso E, et al. Lymphatic metastasis in the absence of functional intratumor lymphatics. Science 2002;296:1883–6.[Abstract/Free Full Text]



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