VEGF-C gene therapy, Lymphangiogensis and Lymphedema

Lymphangiogenic Gene Therapy, yellow nail syndrome, lymphatic vascular development, Intratumoral lymphatics, peritumoral lymphatics, stem cell research, Angiopoietins, VEGF, PIGF, FOXC1, FOXC2, Lymphatic Insufficiency. SOX18, lymphatic hyperplasia, Molecular lymphangiogenesis, PROX1, FLT3, Telan­giectasia, Lymphatic endothelial cells, adult vasculogenesis, LYVE1

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VEGF-C gene therapy, Lymphangiogensis and Lymphedema

Postby patoco » Wed Sep 20, 2006 1:17 pm

** Article from 2003, however good foundation for the understand of the
VEGF-C gene**

Clin Invest. 2003 March 1; 111(5): 717–725.
doi: 10.1172/JCI200315830.
Copyright © 2003, American Society for Clinical Investigation

VEGF-C gene therapy augments postnatal lymphangiogenesis and ameliorates secondary lymphedema

Young-sup Yoon,1 Toshinori Murayama,1 Edwin Gravereaux,1 Tengiz
Tkebuchava,1 Marcy Silver,1 Cynthia Curry,1 Andrea Wecker,1 Rudolf
Kirchmair,1 Chun Song Hu,1 Marianne Kearney,1 Alan Ashare,2 David G.
Jackson,3 Hajime Kubo,4 Jeffrey M. Isner,1 and Douglas W. Losordo1

1Department of Vascular Medicine and Department of Cardiovascular
Research, and2 Department of Nuclear Medicine, St. Elizabeth’s
Medical Center, Tufts University School of Medicine, Boston,
Massachusetts, USA3 Molecular Immunology Group, Institute of Molecular
Medicine, University of Oxford, Oxford, United Kingdom4
Molecular/Cancer Biology Laboratory, Haartman Institute, University of
Helsinki, Helsinki, Finland

Address correspondence to: Douglas W. Losordo, Cardiovascular Research, St. Elizabeth’s Medical Center, 736 Cambridge Street, Boston,
Massachusetts 02135, USA. Phone: (617) 789-3346; Fax: (617) 779-6362;

Received April 30, 2002; Accepted January 7, 2003


Although lymphedema is a common clinical condition, treatment for this
disabling condition remains limited and largely ineffective. Recently,
it has been reported that overexpression of VEGF-C correlates with
increased lymphatic vessel growth (lymphangiogenesis). However, the
effect of VEGF-C–induced lymphangiogenesis on lymphedema has yet to
be demonstrated. Here we investigated the impact of local transfer of
naked plasmid DNA encoding human VEGF-C (phVEGF-C) on two animal models of lymphedema: one in the rabbit ear and the other in the mouse tail.

In a rabbit model, following local phVEGF-C gene transfer, VEGFR-3
expression was significantly increased. This gene transfer led to a
decrease in thickness and volume of lymphedema, improvement of
lymphatic function demonstrated by serial lymphoscintigraphy, and
finally, attenuation of the fibrofatty changes of the skin, the final
consequences of lymphedema. The favorable effect of phVEGF-C on
lymphedema was reconfirmed in a mouse tail model. Immunohistochemical analysis using lymphatic-specific markers: VEGFR-3, lymphatic endothelial hyaluronan receptor-1, together with the proliferation marker Ki-67 Ab revealed that phVEGF-C transfection potently induced new lymphatic vessel growth. This study, we believe for the first time, documents that gene transfer of phVEGF-C resolves lymphedema through direct augmentation of lymphangiogenesis. This novel therapeutic strategy may merit clinical investigation in patients with lymphedema.


Lymphedema is defined as the progressive accumulation of protein-rich
fluid in the interstitial spaces that results from an anatomic or
functional obstruction in the lymphatic system (1). While primary
lymphedema occurs infrequently on a hereditary or idiopathic basis,
secondary lymphedema is common worldwide, primarily due to the increase in radical surgery and radiotherapy for cancer in developed countries and infectious disease (filariasis) in developing countries (2, 3).
Despite substantial advances in both surgical and conservative
techniques, therapeutic options for management of lymphedema are
limited (3, 4). Pathophysiologically, restoration of the
lymph-transporting capacity would appear to represent the optimal
treatment for lymphedema. However, no means for accomplishing new
lymphatic channel development currently exists. Growth of new lymphatic
vessels (lymphangiogenesis) in healthy animals is rapid. The best
example of the natural recovery of lymphatic drainage in animals is the
complete restoration of lymphatic flow after limb reimplantation
(5–9). Therefore the primary difficulty found in lymphedema animal
models is to develop a method to sustain lymphedema long enough to
allow evaluation of therapies.

Recent molecular studies have begun to elucidate the basis for
lymphangiogenesis that can be stimulated by various cytokines,
including VEGF-C (VEGF-2) (10, 11). VEGF-C, the first ligand to be
discovered for VEGFR-3 (Flt4), is a member of the VEGF family of
polypeptide growth factors. VEGF-C binds to endothelial cell receptors
VEGFR-2 (Flk1) and VEGFR-3 (12–15). Although VEGFR-3 plays a critical
role for both vascular and lymphatic endothelial cell development, its
expression becomes limited to the lymphatic endothelium beginning in
the late stages of development (16–18). Overexpression of VEGF-C cDNA
in the skin of transgenic mice induced lymphatic endothelial cell
proliferation and hyperplasia of the lymphatic vasculature, and
recombinant VEGF-C specifically stimulated lymphangiogenesis in
chorioallantoic membrane (11, 19). Recently, direct evidence of the
link between VEGFR-3 and lymphedema has been found: it has been
reported that human hereditary lymphedema is associated with
heterozygous missense mutation of the Flt4 gene, which leads to
insufficient VEGFR-3 signaling (20, 21). Recently, it was demonstrated
that subcutaneous injection of adenovirus or adeno-associated virus
encoding VEGF-C could generate lymphatic vessels in the skin of normal
mice (22) and in a mouse model (chy mouse) of primary lymphedema (23).

Although these studies showed that VEGF-C could induce
lymphangiogenesis in vivo, they failed to show that this
VEGF-C–induced lymphangiogenesis could improve overall lymphatic
vascular dysfunction and prevent chronic changes accompanied by
lymphedema, which are the key determinants of whether VEGF-C can be
used as a therapeutic option to treat human lymphedema.

Accordingly, first we sought to establish reliable animal models of
secondary lymphedema. The two animal models used here provided
complementary measurements: the rabbit ear had the advantage of size,
which is conducive to direct measurements and lymphoscintigraphy,
whereas the mouse tail had advantages for immunohistochemistry owing to the availability of lymphatic vessel–specific Ab’s. Next, using
these animal models, we investigated whether local transfer of naked
plasmid DNA encoding human VEGF-C (phVEGF-C) could promote
lymphangiogenesis and improve physical, functional, and pathologic
aspects of lymphedema.


All animal protocols were approved by the Institutional Animal Care and
Use Committee of St. Elizabeth’s Medical Center. Investigators for
the follow-up examinations were blinded to the identity of the
treatment given.

Rabbit ear model of lymphedema. We modified several previous rabbit ear models to overcome the shortcomings of rapid lymphatic regeneration and to provide the bed for new lymphatic vessel growth (5, 24, 25). To meet those requirements, we used old (3–4 years of age) New Zealand White rabbits and created a skin bridge. Before the operation, the lymphatic vessels were identified by intradermal injection of 0.2 ml of 1% Evans blue dye at the dorsal tip of the right ear. A strip of skin,
subcutaneous tissues, and perichondrium 3 cm wide was circumferentially
excised from the base of the ear, except for the central portion (1 cm
in width) of the dorsal skin, i.e., a “skin bridge” underneath
which runs the neurovascular bundle (Figure 1b). After the distal edge
of the skin bridge was incised, lymphatic channels were dissected and
the lymphatic stumps were resected under a dissecting microscope. Other
edges of skin were inversely sutured to the perichondrium to prevent
reapproximation of skin edges and recanalization of the lymphatic
vessels. This created a strip of bare cartilage, leaving only the skin
bridge for lymphatic growth (Figure 1a).

Preparation of phVEGF-C and gene transfer protocol in a rabbit model. A
total of 54 rabbits was randomized into two groups in a blinded fashion
for treatment with phVEGF-C or control (saline). In the VEGF-C group,
500 μg of phVEGF-C in 0.5 ml volume was injected intradermally and
subcutaneously at the skin bridge using a 27-gauge needle on days 1, 6,
and 11 after lymphedema surgery.

Measurement of ear thickness and volume. The thickness of the rabbit
ears was measured 1 cm medial and distal to the medial border of the
skin bridge with a vernier caliper. The ear was put in a 50-ml cylinder
filled with water. After removing the ear, the volume of water
displaced by the ear was measured (25). The thickness and volume of all
ears was measured before surgery and every week for 6 weeks, and
thereafter every 2 weeks until the 12-week point (n = 12 in each

Measurement of skin thickness in histologic sections. Thickness of the
ear skin at 8 weeks after surgery was measured in a cross section of
the skin just distal to the skin bridge in paraffin-embedded histologic
specimens after elastic-tissue trichrome staining as described
previously (26) n = 5 in each group).

Lymphoscintigraphy and quantitative analysis. Tc-99m–filtered sulfur
colloid was injected intradermally into the dorsal tip of rabbit ears
at a dose of 50 μCi. Imaging was performed using a large-field-of-view
Genesys γ camera (ADAC Laboratories, Milpitas, California, USA).

To quantitatively compare lymphatic drainage of the injected
radiotracers, radioactivity within the ears was counted. The γ counts
at injection sites were similar in both ears of the saline and VEGF-C
groups (P = 0.93). For standardization, the ratio of radioactivity of
the operated ear to that of the normal (contralateral) ear, designated
the radioactivity index, was used to compare lymphatic drainage at 4,
8, and 12 weeks (see Figure 2, g–i). The validity of the
radioactivity index was verified by repeated examination of normal ears
(n = 7) for intraindividual variation, which was 6% (coefficient of
variation [cv %]), and by comparison of the day 1 postoperative
lymphoscintigrams for interindividual variation (n = 20), which was 10%
(cv %).

Western analysis of VEGF-C transgene expression in tissue. Samples
harvested from the skin bridge and from tissue proximal and distal to
the skin bridge of the operated ears, and from the bridge site of the
contralateral ears were snap frozen in liquid nitrogen 7 days after the
second injection of phVEGF-C (postoperative day 13) (n = 5 in each
group). Western analysis was performed as described (27).

Molecular cloning of partial rabbit VEGFR-3 cDNA. Because the rabbit
VEGFR-3 DNA sequence has not been identified, we sequenced part of the
VEGFR-3 cDNA using degenerate oligonucleotides. Degenerate
oligonucleotides were designed from conserved amino acid sequences
NVSDSLEM and WEFPRER, located at the transmembrane domain of human and mouse VEGFR-3 (28, 29). The deduced oligonucleotide sequences were 5′-AACGTGAG (CT)GACTC (GC) (CT)T (AGCT)GA (AG)ATG-3′ and 5′-CC (GT)YTC (CT)C (GT)GGG (AG)AA (CT)TCCCA-3′, respectively. A single PCR product of 470 bp was obtained from all the tissues (see Figure 3d).

Semiquantitative RT-PCR analysis of VEGFR-3. Using samples harvested
from the bridge site of both ears at postoperative day 13 (n = 5 in
each group), total RNA was isolated and RT-PCR was performed as
described above. The primer pair used, designed on the basis of the
sequenced cDNA’s for rabbit VEGFR-3, was
5′-ACAGGTATTCACATTGCTCCT-3′ (antisense). To quantify the VEGFR-3
mRNA, we used the “competimer” quantitative PCR technique (Ambion
Inc., Austin, Texas, USA) according to the manufacturer’s
instructions. To the VEGFR-3 PCR mix, we added a mix of 18S primer
pairs and 18S 3′-end modified primers (competimers) at a ratio of
1:9, yielding a 488-bp product. PCR was performed as follows: 94°C, 2
min (once); 94°C, 15 s; 50°C, 30 s; 72°C, 1 min (40 cycles); 72°C,
10 min (once). PCR products were separated on 1.5% agarose gel and
quantified by integrated density analysis software (EagleSight Software
3.2; Stratagene, La Jolla, California, USA).

Mouse tail model of lymphedema. Male nude (nu/nu) mice (Harlan,
Indianapolis, Indiana, USA) 12 weeks of age were used. A mouse
lymphedema model was created by modifying a previous model (30).

Gene transfer protocol in a mouse tail model. In total, 115 mice were
randomized into five groups: no operation, VEGF-C, VEGF165, LacZ, and
saline (n = 23 in each group). The unoperated group served as negative
control. The other groups underwent the operation as described. In the
VEGF-C group, 100 μg of phVEGF-C was given in 100 μl volume on days
1, 6, and 11 after the operation, respectively. The phVEGF165 plasmid
(18, 31), pGSV-nlsLacZ (32) (a nuclear targeted LacZ gene plasmid
encoding the protein β-galactosidase), and saline were injected in an
identical fashion in the VEGF165, LacZ, and saline groups,

Immunoprecipitation of receptor phosphorylation. To investigate the
effect of VEGF-C overexpression on phosphorylation of VEGFR-2 and
VEGFR-3, immunoprecipitation and Western blot analysis was performed in the mouse tail model. Lysis of tissues, immunoprecipitation, and
Western blot analysis were performed as described (31, 33). Aliquots of
protein extracts (1 mg) were incubated for 2 hours at 4°C with 3 μg
of mAb against phosphotyrosine (Upstate Biotechnology Inc., Lake
Placid, New York, USA), followed by incubation with 40 μl of protein
G–agarose beads (Roche Diagnostics GmbH, Mannheim, Germany) overnight at 4°C. Immunoprecipitates of tyrosine-phosphorylated proteins were separated by 7.5% SDS-PAGE and electrotransferred onto PVDF membranes.

The membranes were immunoblotted overnight at 4°C with a rabbit
polyclonal Ab against VEGFR-3 (1:500; Santa Cruz Biotechnology Inc.,
Santa Cruz, California, USA) or VEGFR-2 (1:500; Santa Cruz
Biotechnology Inc.).

Immunohistochemistry and morphometric analysis. The skin from the
bridge area was harvested 3 weeks after plasmid/saline injections. Skin
sections were stained using a rat mAb against mouse VEGFR-3 (34) and a
rabbit polyclonal Ab against the lymphatic marker lymphatic endothelial
hyaluronan receptor-1 (LYVE-1), a receptor for hyaluronan and a
homologue to the CD44 glycoprotein (35).

In double fluorescent immunohistochemistry of LYVE-1 and Ki-67, LYVE-1
staining was performed with the use of Texas red–streptavidin (NEN
Life Science Products Inc., Boston, Massachusetts, USA), and Ki-67
staining was performed with rabbit polyclonal Ab against Ki-67
(Novocastra Laboratories Ltd., Newcastle, United Kingdom) and
Cy2-conjugated goat anti-rabbit IgG (Jackson ImmunoResearch
Laboratories Inc., West Grove, Pennsylvania, USA). Endothelial cells
were identified by immunohistochemical staining for platelet
endothelial cell adhesion molecule-1 (PECAM-1 or CD31) with a rat mAb
against mouse CD31 (BD Biosciences, San Diego, California, USA) (36) in
mouse tissues and with a mouse mAb against human CD31 in rabbit

Statistical analysis. All results were expressed as mean ± SEM.
Statistical analysis was performed with an unpaired Student t test for
comparisons between two groups and ANOVA followed by Scheffe’s
procedure for more than two groups. P values < 0.05 were considered to
denote statistical significance.


VEGF-C gene therapy induces remission of lymphedema. Seeking to
establish an appropriate animal model, initial experiments using young
(6–8 months old) rabbits showed expedited regression of lymphedema
such that we could not properly evaluate the effect of gene transfer.
In the older rabbits (3–4 years old), a substantial degree of
lymphedema developed and was sustained for more than 12 weeks.

To investigate the effect of phVEGF-C gene transfer on this lymphedema
model, we measured ear thickness and volume over a 12-week period. Ear thickness was consistently smaller in the VEGF-C group than in the
saline group. The difference between the groups was statistically
significant beginning at 2 weeks and was maintained for the duration of
the study (2 weeks: 4.5 ± 0.3 vs. 5.4 ± 0.2 mm, P < 0.05; 3 weeks:
3.9 ± 0.2 vs. 4.6 ± 0.3 mm, P < 0.05; 8 weeks: 2.8 ± 0.2 vs. 3.6 ±
0.3 mm, P < 0.05; 10 weeks: 2.6 ± 0.2 vs. 3.5 ± 0.3 mm, P < 0.05; 12
weeks: 2.4 ± 0.2 vs. 3.3 ± 0.2 mm, P < 0.01) (Figure 1c). Similarly,
ear volume was consistently smaller in the VEGF-C group than in the
saline group for the duration of the study (2 weeks: 33.1 ± 2.2 vs.
38.2 ± 1.5 ml, P < 0.05; 3 weeks: 29.4 ± 1.5 vs. 34.1 ± 1.0 ml, P <
0.05; 4 weeks: 26.5 ± 1.8 vs. 31.7 ± 1.7 ml, P < 0.05; 8 weeks: 18.3
± 2.3 vs. 26.3 ± 1.9 ml, P < 0.05; 10 weeks: 17.1 ± 2.1 vs. 25.5 ±
1.8 ml, P < 0.01; 12 weeks: 14.6 ± 2.3 vs. 24.5 ± 1.3 ml, P < 0.01)
(Figure 1d). The VEGF-C group had significantly thinner skin than the
saline group at 8 weeks (2.8 ± 0.2 vs. 3.8 ± 0.2 mm, P < 0.05).

Lymphoscintigraphy demonstrates enhanced lymphatic drainage after
VEGF-C gene transfer. In normal ears, lymphatic flow assumes a linear
pattern, and the draining LNs are clearly visible at the base of the
skull. (Figure 2b). Imaging performed at day 1 after surgery showed
successful surgical blockade of lymphatic egress in all animals (Figure
2, c and e). Follow-up lymphoscintigraphy at 4, 8, and 12 weeks showed
dynamic changes of radiotracer clearance from the operated ears that
was more efficient in the phVEGF-C–transfected ear than in the
saline-injected ear. Images at 12 weeks revealed that the
saline-injected ear still showed a dermal backflow pattern with faint
visualization of LNs, while the phVEGF-C–transfected ear shows a
linear pattern of lymphatic drainage and clear visualization of LNs
(Figure 2, d and f).

Quantification of lymphatic drainage (Figure 2, g and h) over the study
period revealed consistently lower retention of radioactivity in the
VEGF-C group than in the saline group; this achieved statistical
significance at 8 weeks (radioactivity index, 3.8 ± 0.4 vs. 5.0 ±
0.5, P < 0.05) and 12 weeks (radioactivity index, 2.2 ± 0.3 vs. 4.2 ±
0.4, P < 0.05) (Figure 2i).

Transgene expression of phVEGF-C in a rabbit ear model. To assess
transgene expression of injected phVEGF-C in ear skin, we performed
Western blotting of VEGF-C protein. In our experiments, two different
bands were detected using two anti–VEGF-C Ab’s. A 58-kDa band
corresponds to the earliest processed form, while a 31-kDa band
represents the major secreted form of VEGF-C polypeptides.
Densitometric analysis of multiple experiments revealed that expression
of the 58-kDa VEGF-C isoform in the phVEGF-C–transfected bridge was
2.6 times and 3.2 times higher than that in the saline-injected and the
normal skin, respectively (P < 0.01) (Figure 3a). Expression of the
31-kDa VEGF-C isoform in the phVEGF-C–transfected bridge was 3.6
times and 3.9 times higher than that in the saline-injected and the
normal skin, respectively (P < 0.01) (Figure 3b).

Gene transfer of phVEGF-C increases VEGFR-3 expression. A partial
470-bp rabbit VEGFR-3 cDNA was cloned by RT-PCR using degenerate
oligonucleotide primers (GenBank accession number AF453570). The amino acid sequence displayed 92.9%, 93.6%, and 94.3% identity with human, bovine, and mouse VEGFR-3 (Figure 3c). We investigated VEGFR-3 expression using RT-PCR, revealing a nearly 1.7-fold induction of
VEGFR-3 mRNA levels by VEGF-C compared with saline control (P < 0.01,
Figure 3g).

Gene transfer of phVEGF-C increases phosphorylation of VEGFR-3 in a
mouse tail model. We investigated the effect of phVEGF-C gene transfer
on the tyrosyl phosphorylation of VEGFR-3 and VEGFR-2 by
immunoprecipitation with anti-phosphotyrosine Ab followed by Western
blot analysis with anti–VEGFR-3 and anti–VEGFR-2 Ab’s,
respectively. Phosphorylated VEGFR-3 (195 kDa) in the
phVEGF-C–transfected samples was 1.6 times and 1.8 times higher than
in the samples from the saline and LacZ groups, respectively (P < 0.05)
(Figure 3h). Gene transfer of phVEGF165 did not increase phosphorylated
VEGFR-3 compared with the controls. Phosphorylated VEGFR-2 in the
phVEGF165-transfected sample was 2.0, 1.8, and 1.6 times higher
(235-kDa band) than in the samples of the saline, LacZ, and VEGF-C
groups, respectively (P < 0.05) (Figure 3i). Phosphorylated VEGFR-2 was
slightly higher in the phVEGF-C–transfected samples than in the
control groups (saline and LacZ) but was not statistically significant.

Gene transfer of phVEGF-C improves lymphedema in a mouse tail model.

To determine whether the effect of phVEGF-C could be reproduced in another lymphedema model, similar experiments were performed in a mouse tail model (30) (Figure 4a). In the VEGF-C group, compared with the saline, LacZ, and VEGF165 groups, the tail thickness was consistently smaller beginning at 3 weeks (3 weeks: 4.05 ± 0.08 vs. 4.37 ± 0.07, 4.32 ±
1.08, and 4.30 ± 0.07 mm, P < 0.05; 4 weeks: 4.01 ± 0.09 mm vs. 4.42
± 0.08, 4.31 ± 0.08, and 4.28 ± 0.07 mm, P < 0.05; 5 weeks: 4.00 ±
0.07 mm vs. 4.35 ± 0.08, 4.28 ± 0.08, and 4.28 ± 0.08 mm, P < 0.05),

Gene transfer of phVEGF-C promotes lymphatic vessel growth in a mouse
tail model. The VEGF-C group showed higher density of LYVE-1–positive
lymphatic vessels than the other groups (VEGF-C, 85 ± 7 per mm2;
saline, 38 ± 4 per mm2; LacZ, 42 ± 5 per mm2; VEGF165, 46 ± 5 per
mm2, P < 0.01) (Figure 4, b–f and l). Skin sections stained with
VEGFR-3 Ab showed similar results (VEGF-C, 81 ± 8 per mm2; saline, 37
± 4 per mm2; LacZ, 43 ± 5 per mm2; VEGF165, 44 ± 5 per mm2, P <
0.01) (Figure 4, g–k and m). After phVEGF-C transfection, lymphatic
vessels appeared hyperplastic (Figure 4, e and j). In sections at
3-week follow-up, the number of lymphatic vessels containing Ki-67+
nuclei was 2.5 times higher in the VEGF-C group than in the saline or
LacZ groups (VEGF-C, 58 ± 7 per mm2; saline, 23 ± 3 per mm2; LacZ, 26 ± 4 per mm2, P < 0.01) (Figure 5, a–j).

Blood capillary density analysis. Rabbit ear and mouse tail skins were
stained for an endothelial cell marker, CD31 (36). In the rabbit
lymphedema model, capillary density was not significantly different
among the saline (193 ± 18 per mm2), LacZ (198 ± 22 per mm2), or
VEGF-C (201 ± 20 per mm2) groups (Figure 5, k–o). Similar findings
were observed in the capillary density of operated mouse tail groups
(saline, 172 ± 18 per mm2; LacZ, 181 ± 19 per mm2; VEGF-C, 189 ± 20
per mm2, P value not significant) (Figure 5b, F–K). However, the
VEGF165 group (302 ± 27 per mm2) showed significantly higher capillary
density than the saline, LacZ, or VEGF-C groups (P < 0.01).


Chronic lymphedema is a disabling condition characterized by thickening
of the skin due to fibrofatty deposition in underlying tissues as well
as disfiguring swelling of affected limbs. In most cases of secondary
lymphedema in humans, depletion of lymphatic vessels is the culprit in
its pathogenesis (1–4). Here we show that phVEGF-C gene therapy, by
promoting lymphangiogenesis, favorably modulates all the phenotypic
changes associated with secondary lymphedema. We believe the present
study is the first to document improvement in the clinical and
pathologic features of lymphedema resulting from enhancement of
lymphatic drainage by phVEGF-C gene therapy.

In two animal models, we demonstrate significant attenuation of
lymphedema by phVEGF-C gene transfer. The effect prevailed over the
chronic phase as well as the acute phase of lymphedema. This
improvement was also confirmed in histologic sections, which reflect
chronic fibrofatty changes more accurately. Prevention or reduction of
fibrotic change is one of the most important goals of therapy for
lymphedema, since this secondary change can drive lymphedema into a
vicious cycle by increasing interstitial solid pressure (by fibrofatty
deposition) and thus collapsing already reduced or impaired lymphatic
vessels (3–5, 24, 25). That the improvement in the physical indices
of lymphedema was the actual result of improved lymphatic drainage was
documented by quantitative lymphoscintigraphy. Although VEGF-C plasmid transgene expression is usually limited to less than 30 days (27), our results indicate that once the lymphatic connection is reestablished,
the recovery of drainage function can be maintained.

VEGF-C protein expression was documented in situ after phVEGF-C gene
transfer. We detected the partially processed form (58 kDa) and major
secreted form (31 kDa) of VEGF-C (37, 38) after local gene delivery.
VEGF-C is produced as a 61-kDa prepropeptide form which undergoes
multistep proteolytic maturation. The secreted 31-kDa form
predominantly activates VEGFR-3 (37). VEGFR-3 mRNA expression was very low in normal skin, slightly higher in the saline-injected experimental
ears, and strongly upregulated following phVEGF-C gene transfer. We
also directly measured the number of lymphatic vessels using
lymphatic-specific markers. LYVE-1 or VEGFR-3 staining confirmed
augmentation of lymphangiogenesis in phVEGF-C–transfected mouse
tails. The hyperplastic nature of proliferating lymphatic vessels was
consistent with previous reports (19, 22). Ki-67 staining documented
that proliferating lymphatic endothelial cells exist in more than half
the lymphatic vessels in phVEGF-C–transfected skin, suggesting a
potent lymphangiogenic effect of phVEGF-C.

As VEGF-C is also known to activate VEGFR-2 and thus to induce
angiogenesis in vitro and in ischemic tissues (15, 32), we evaluated
capillary density from both animal models and found it was slightly
higher in the VEGF-C group but not significantly different from that of
the saline or LacZ control groups. The apparent absence of fully
processed 21-kDa product of VEGF-C, which has potent angiogenic
activity, and thus the weak activation of VEGFR-2 phosphorylation,
could explain the lack of obvious angiogenesis in these animal models.
These findings are compatible with previous reports that claimed no
discernible angiogenesis in transgenic mice overexpressing VEGF-C in
the skin (19) and in normal mouse skin that was transfected with
adeno–VEGF-C (22). Physiologic function of any ligand is dependent on
the temporal and spatial expression of its specific receptors. In the
case of VEGF-A–induced angiogenesis, the absence of ischemia-induced
regional upregulation of VEGFR-2 has been shown to result in
nullification of the angiogenic effect of transient overexpression of
VEGF-A (39, 40). To further address the concern that increased
angiogenesis might improve lymphedema, we investigated the effect of
VEGF-A (phVEGF165) plasmid gene transfer in the mouse tail model and
found that augmenting angiogenesis but not lymphangiogenesis did not
improve lymphedema.

The potential clinical relevance and limitations of our study derive
from certain features of the design and the findings. First, the models
are pathophysiologically similar to the secondary forms of human
lymphedema. The models we used do not represent the entire spectrum of lymphedema found in humans, especially the primary form of lymphedema, which is an inherited developmental disorder of the lymphatic system.

However, pathophysiologically our models are approximations of
secondary lymphedema, which comprises most cases of lymphedema and
results primarily from surgical removal of lymphatic vessels and lymph
nodes in industrialized countries (1–4). Additionally, in our models,
we used gene therapy in an acute/subacute stage of lymphedema.
Therefore, whether this gene therapy can be effective in chronic cases
is uncertain. As there are concerns about the potential enhancement of
tumor growth and metastasis by VEGF-C in tumor models (41), we need to consider the potential advantages and dangers of using local VEGF-C
therapy in patients with lymphedema caused by cancer treatment. This
issue can be resolved after performing experiments adopting tumor
implantation and treatment such as a combination of surgery,
chemotherapy or radiation, and local VEGF-C gene transfer. Third, the
approach of gene therapy using naked plasmid DNA (phVEGF-C) has been
used in early clinical trials and has an accumulating record of safety
(42). Finally, to the best of our knowledge, this study represents the
first experimental proof of a beneficial effect of VEGF-C gene therapy
on lymphedema per se. Our findings clearly indicate a favorable effect
of phVEGF-C–induced lymphangiogenesis on lymphedema and thus
represent a novel therapeutic paradigm for the treatment of this
otherwise difficult-to-manage condition.


This paper is dedicated to Jeffrey M. Isner, who passed away on October
31, 2001. We would like to gratefully acknowledge his inspirational
leadership. We gratefully acknowledge M. Neely, I. Johnson, and T.
Shiojima for their excellent secretarial assistance. This study was
supported in part by NIH grants HL-53354, HL-60911, HL-63414, HL-63695, and HL-66957, and by the Shaughnessy Center for Clinical Genetics, Boston, Massachusetts, USA. Young-sup Yoon is the recipient of a fellowship from the American Heart Association, New England Affiliate.


Conflict of interest:

The authors have declared that no conflict of
interest exists.

Nonstandard abbreviations used:

naked plasmid DNA encoding human VEGF-C (phVEGF-C); coefficient of variation (cv); lymphatic endothelial
hyaluronan receptor–1 (LYVE-1); platelet endothelial cell adhesion
molecule–1 (PECAM-1).


Browse NL. The diagnosis and management of primary lymphedema. J. Vasc.
Surg. 1986;3:181–184. [PubMed] Almeida AB, Freedman DO. Epidemiology and immunopathology of
bancroftian filariasis. Microbes Infect. 1999;1:1015–1022. [PubMed]
3.Szuba A, Rockson SG. Lymphedema: classification, diagnosis and
therapy. Vasc. Med. 1998;3:145–156. [PubMed]
4.Ko DS, Lerner R, Klose G, Cosimi AB. Effective treatment of
lymphedema of the extremities. Arch. Surg. 1998;133:452–458. [PubMed]
5.Slavin SA, Upton J, Kaplan WD, Van den Abbeele AD. An investigation
of lymphatic function following free-tissue transfer. Plast. Reconstr.
Surg. 1997;99:730–741; discussion 742–743. [PubMed]
6.Drinker CK, Field MJ, Homans J. The experimental production of edema
and elephantiasis as a result of lymphatic obstruction. Am. J. Physiol.
7.Clodius L. Experimental lymphedema and therapeutic concepts. Acta
Chir. Plast. 1976;18:113–116. [PubMed]
8.Casley-Smith JR, Clodius L, Foldi M. Experimental blood vascular and
lymphatic occlusion in the rabbit ear and the effect of benzopyrones.
Arzneimittelforschung. 1977;27:379–382. [PubMed]
9.Lee-Donaldson L, et al. Refinement of a rodent model of peripheral
lymphedema. Lymphology. 1999;32:111–117. [PubMed]
10.Leak LV, Jones M. Lymphangiogenesis in vitro: formation of lymphatic
capillary-like channels from confluent monolayers of lymphatic
endothelial cells. In Vitro Cell. Dev. Biol. Anim. 1994;30A:512–518.
11.Oh SJ, et al. VEGF and VEGF-C: specific induction of angiogenesis
and lymphangiogenesis in the differentiated avian chorioallantoic
membrane. Dev. Biol. 1997;188:96–109. [PubMed]
12.Aprelikova O, et al. FLT4, a novel class III receptor tyrosine
kinase in chromosome 5q33-qter. Cancer Res. 1992;52:746–748. [PubMed]
13.Galland F, et al. Chromosomal localization of FLT4, a novel
receptor-type tyrosine kinase gene. Genomics. 1992;13:475–478.
14.Lee J, et al. Vascular endothelial growth factor-related protein: a
ligand and specific activator of the tyrosine kinase receptor Flt4.
Proc. Natl. Acad. Sci. U. S. A. 1996;93:1988–1992. [PubMed]
15.Joukov V, et al. A novel vascular endothelial growth factor, VEGF-C,
is a ligand for the Flt4 (VEGFR-3) and KDR (VEGFR-2) receptor tyrosine
kinases. EMBO J. 1996;15:290–298. [PubMed]
16.Kaipainen A, et al. Expression of the fms-like tyrosine kinase 4
gene becomes restricted to lymphatic endothelium during development.
Proc. Natl. Acad. Sci. U. S. A. 1995;92:3566–3570. [PubMed]
17.Kukk E, et al. VEGF-C receptor binding and pattern of expression
with VEGFR-3 suggests a role in lymphatic vascular development.
Development. 1996;122:3829–3837. [PubMed]
18.Baumgartner I, et al. Constitutive expression of phVEGF165 after
intramuscular gene transfer promotes collateral vessel development in
patients with critical limb ischemia. Circulation. 1998;97:1114–1123.
19.Jeltsch M, et al. Hyperplasia of lymphatic vessels in VEGF-C
transgenic mice. Science. 1997;276:1423–1425. [PubMed]
20.Ferrell RE, et al. Hereditary lymphedema: evidence for linkage and
genetic heterogeneity. Hum. Mol. Genet. 1998;7:2073–2078. [PubMed]
21.Karkkainen MJ, et al. Missense mutations interfere with VEGFR-3
signalling in primary lymphoedema. Nat. Genet. 2000;25:153–159.
22.Enholm B, et al. Adenoviral expression of vascular endothelial
growth factor-C induces lymphangiogenesis in the skin. Circ. Res.
2001;88:623–629. [PubMed]
23.Karkkainen MJ, et al. A model for gene therapy of human hereditary
lymphedema. Proc. Natl. Acad. Sci. U. S. A. 2001;98:12677–12682.
24.Piller NB, Clodius L. Lymphoedema of the rabbit ear following
partial and complete lymphatic blockade; its effects on fibrotic
development, enzyme types and their activity levels. Br. J. Exp.
Pathol. 1978;59:319–326. [PubMed]
25.Fu K, Izquierdo R, Vandevender D, Warpeha RL, Fareed J.
Transplantation of lymph node fragments in a rabbit ear lymphedema
model: a new method for restoring the lymphatic pathway. Plast.
Reconstr. Surg. 1998;101:134–141. [PubMed]
26.Tsurumi Y, et al. Reciprocal relation between VEGF and NO in the
regulation of endothelial integrity. Nat. Med. 1997;3:879–886.
27.Schratzberger P, et al. Reversal of experimental diabetic neuropathy
by VEGF gene transfer. J. Clin. Invest. 2001;107:1083–1092. [PubMed]
28.Finnerty H, et al. Molecular cloning of murine FLT and FLT4.
Oncogene. 1993;8:2293–2298. [PubMed]
29.Galland F, et al. The FLT4 gene encodes a transmembrane tyrosine
kinase related to the vascular endothelial growth factor receptor.
Oncogene. 1993;8:1233–1240. [PubMed]
30.Slavin SA, Van den Abbeele AD, Losken A, Swartz MA, Jain RK. Return
of lymphatic function after flap transfer for acute lymphedema. Ann.
Surg. 1999;229:421–427. [PubMed]
31.Schratzberger P, et al. Favorable effect of VEGF gene transfer on
ischemic peripheral neuropathy. Nat. Med. 2000;6:405–413. [PubMed]
32.Witzenbichler B, et al. Vascular endothelial growth factor-C
(VEGF-C/VEGF-2) promotes angiogenesis in the setting of tissue
ischemia. Am. J. Pathol. 1998;153:381–394. [PubMed]
33.Brogi E, et al. Hypoxia-induced paracrine regulation of vascular
endothelial growth factor receptor expression. J. Clin. Invest.
1996;97:469–476. [PubMed]
34.Kubo H, et al. Involvement of vascular endothelial growth factor
receptor-3 in maintenance of integrity of endothelial cell lining
during tumor angiogenesis. Blood. 2000;96:546–553. [PubMed]
35.Banerji S, et al. LYVE-1, a new homologue of the CD44 glycoprotein,
is a lymph-specific receptor for hyaluronan. J. Cell Biol.
1999;144:789–801. [PubMed]
36.Rivard A, et al. Age-dependent impairment of angiogenesis.
Circulation. 1999;99:111–120. [PubMed]
37.Joukov V, et al. Proteolytic processing regulates receptor
specificity and activity of VEGF-C. EMBO J. 1997;16:3898–3911.
38.Pepper MS, Mandriota SJ, Jeltsch M, Kumar V, Alitalo K. Vascular
endothelial growth factor (VEGF)-C synergizes with basic fibroblast
growth factor and VEGF in the induction of angiogenesis in vitro and
alters endothelial cell extracellular proteolytic activity. J. Cell.
Physiol. 1998;177:439–452. [PubMed]
39.Takeshita S, et al. Therapeutic angiogenesis. A single intraarterial
bolus of vascular endothelial growth factor augments revascularization
in a rabbit ischemic hind limb model. J. Clin. Invest.
1994;93:662–670. [PubMed]
40.Isner JM, et al. Arterial gene transfer for therapeutic angiogenesis
in patients with peripheral artery disease. Hum. Gene Ther.
1996;7:959–988. [PubMed]
41.Skobe M, et al. Induction of tumor lymphangiogenesis by VEGF-C
promotes breast cancer metastasis. Nat. Med. 2001;7:192–198. [PubMed]
42.Isner JM, Vale PR, Symes JF, Losordo DW. Assessment of risks
associated with cardiovascular gene therapy in human subjects. Circ.
Res. 2001;89:389–400. [PubMed] ... ed&pubme...


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