FASEB J.
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Published as doi: 10.1096/fj.05-5137fje.
This Article
Right arrow Abstract Freely available
Right arrow Summary
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
fj.05-5137fjev1
20/9/1495    most recent
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Li, B.
Right arrow Articles by Young, P. P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Li, B.
Right arrow Articles by Young, P. P.
(The FASEB Journal. 2006;20:1495-1497.)
© 2006 FASEB

VEGF and PlGF promote adult vasculogenesis by enhancing EPC recruitment and vessel formation at the site of tumor neovascularization

Bin Li*, Emerson E. Sharpe*, Amanda B. Maupin*, Amylynn A. Teleron*, Amy L. Pyle*, Peter Carmeliet{ddagger} and Pampee P. Young*,{dagger},1

Departments of
* Pathology and

{dagger} Internal Medicine, Vanderbilt University Medical Center, Nashville, Tennessee, USA; and

{ddagger} Center for Transgene Technology and Gene Therapy, Flanders Interuniversity Institute for Biotechnology, University of Leuven, Leuven, Belgium

1Correspondence: Vanderbilt University School of Medicine, Department of Pathology, 1161 21st Ave. South, C2217 MCN, Nashville, TN 37232. E-mail: pampee.young{at}vanderbilt.edu

ABSTRACT

There are growing data to suggest that tissue hypoxia represents a critical force that drives adult vasculogenesis. Vascular endothelial growth factor (VEGF) expression is dramatically up-regulated by hypoxia and results in enhanced neovascularization. Although the role of VEGF in angiogenesis has been well characterized, its role in adult vasculogenesis remains poorly understood. We used two distinct murine bone marrow transplantation (BMT) models to demonstrate that increased VEGF levels at the site of tumor growth promoted vasculogenesis in vivo. This effect of VEGF was downstream of its effect to enhance either mobilization or survival of circulating endothelial progenitor cells (EPCs). Both VEGFR1 (flt1) and VEGFR2 (flk1) are expressed on culture expanded human EPCs. Previous studies suggest that the effect of VEGF on endothelial cell migration is primarily mediated via VEGFR2; however, VEGF-induced EPC migration in vitro was mediated by both receptors, suggesting that VEGF-VEGFR1 interactions in EPCs are distinct from differentiated endothelial cells. We used specific blocking antibodies to these receptors to demonstrate that VEGFR1 plays an important role in human EPC recruitment to tumors. These findings were further supported by our finding that tumor-associated placental growth factor (PlGF), a VEGFR1-specific agonist, increased tumor vasculogenesis in a murine BMT model. We further showed that both VEGF receptors were necessary for the formation of functional vessels derived from exogenously administered human ex vivo expanded EPCs. Our data suggest local VEGF and/or PlGF expression promote vasculogenesis; VEGF plays a role in EPC recruitment and subsequent formation of functional vessels.—Li, B., Sharpe, E. E., Maupin, A., Teleron, A. A., Pyle, A., Carmeliet, P., Young, P. P. VEGF and PlGF promote adult vasculogenesis by enhancing EPC recruitment and vessel formation at the site of tumor neovascularization.


Key Words: VEGF receptors • BM transplant models • angiogenesis • placental growth factor

THE ROLE of vasculogenesis during postnatal neovascularization in both pathological and physiological conditions is under intense study (1 2 3) . Multiple, recent studies have demonstrated that ex vivo isolated, culture-expanded endothelial progenitor cells (CE-EPCs) from human peripheral blood (hPB) are preferentially recruited to sites of ischemia and tumor formation, incorporate into functional vasculature, and promote vascularization (4 5 6 7 8) . However, the mechanisms by which circulating EPCs are recruited to sites of neovascularization remain incompletely understood (2 , 3 , 9) .

Recently, it has been proposed that postnatal vasculogenesis is driven primarily by local tissue ischemia, and this is likely to involve the local release of soluble factors that promote EPC recruitment (10 11 12) . The chemokine, stromal cell-derived factor-1 (SDF-1), has been well characterized as a mediator of hematopoietic stem cell recruitment and homing (13) . More recently, it has been shown to be involved in hypoxia-mediated recruitment of CE-EPCs in mouse models of tissue ischemia (14 15 16 17 18) . Other cytokines are also known to be up-regulated by hypoxia (19) . Perhaps the most widely studied of these is the endothelial cell (EC) mitogen, vascular endothelial growth factor (VEGF; also known as VEGF-A; refs. 20 , 21 ). The role of VEGF in angiogenesis is well established by both in vitro and in vivo studies (21) . VEGF has also been shown to be important in recruiting myeloid cells to tumor site (22) . However, its role in postnatal vasculogenesis is not well understood (1 , 3 , 21 , 23) . Thus far, multiple groups have shown that recombinant VEGF promotes mobilization of immature hematopoietic progenitors into peripheral circulation as well as the numbers of CE-EPCs/ml of peripheral blood (PB) in mice (24 , 25) . Consistent with that observation, blocking VEGFR2 signaling resulted in poor mobilization of EPCs (26) . The presence of tissue ischemia, which resulted in increased circulating VEGF levels, also increased circulating progenitors and CE-EPCs in mice and humans (11 , 27 , 28) .

In our previous work, we showed that coadministration of recombinant VEGF with donor bone marrow (BM) cells to neonatal mice promoted engraftment of donor-derived ECs into recipient liver and heart, suggesting that the effect of VEGF may be downstream of EPC mobilization (29) . This finding was supported by another study in which VEGF164 gene transfer augmented proliferative activity, adhesion, and incorporation of CE-EPCs to EC monolayers in vitro and in vivo engraftment to ischemic limbs (30) .

VEGF regulates blood vessel growth primarily by interacting with two cell surface tyrosine kinase receptors, VEGF receptor-1 (VEGFR1 or flt1) and VEGF receptor-2 (VEGFR2 or flk1; ref 21 ) and with membrane protein neuropilin-1, which does not contain a tyrosine kinase domain (31) . VEGFR1 and VEGFR2 have distinct functions during embryonic vasculogenesis (32 , 33) . Whereas absence of VEGFR2 results in complete absence of both ECs and hematopoietic cells (32) , ablation of VEGFR1 results in abundant ECs but failure to form functional vessels (33) . The role of these receptors in postnatal vasculogenesis has not been well studied. There has been growing interest in placenta growth factor (PlGF), a member of the VEGF family that binds specifically to VEGFR1, and its role in tumor vascularity. PlGF has been found to promote tumorigenesis in mice, and very recent studies report PlGF overexpression in human cancers (34 35 36 37) . In this study, we provide data to support the hypothesis that VEGF promotes tumor vasculogenesis by promoting EPC chemotaxis, recruitment (through VEGFR1), and vessel formation. We also investigate the role of PlGF in tumor vasculogenesis.

MATERIALS AND METHODS

Mice and cell lines
The NOD/LtSz-scid (NOD/SCID) mouse with a deficiency in ß-glucuronidase (GUSB) activity (NOD/SCID/GUSB–/–) was recently generated (38) and bred in a colony maintained by P. P. Young at Vanderbilt University School of Medicine. Offspring homozygous for the GUSB mutation (NOD/SCID/ß-glucuronidase–/–) were identified by a specific polymerase chain reaction (PCR) assay (29 ,38) . The GUSB–/– tumor line arose spontaneously in a homozygous NOD/SCID mouse and was passaged in culture, and cells from passage 6–20 were used for all experiments. Tumor cell proliferation in vitro was not affected by rVEGF (10 uM). To measure tumor vasculogenesis, GUSB–/– mice were transplanted with BM from syngeneic GUSB+/+ donors resulting in GUSB/BMT. After hematopoietic engraftment, mice were implanted with tumors that were analyzed by quantitative enzyme (39) , flow cytometric (29) , and immunofluorescent GUSB assays (29) to determine BM engraftment and EPC recruitment.

For CE-EPC recruitment studies, after tumor growth {approx}0.5 cm3 (10 days after 1x106 cells were implanted in untransplanted GUSB–/– mice), 5 x 105 hPB-derived CE-EPCs were preincubated for 1 h at 4°C with 8 µg of neutralizing antibody to either VEGFR1 (clone 6.12, IgG1, ImClone Systems) or VEGR2 (clone 89106, IgG1, R&D Systems) or control mouse IgG without serum. Cells were washed in PBS after incubation and subsequently injected intravenously by tail vein into tumor-bearing recipients. Mice were killed 12 h later and tumor tissue, and lung and skeletal muscles were removed for quantitative GUSB analysis. Six mice were used for each condition (n=18) over three independent experiments. To study in situ vessel formation, hPB-derived CE-EPCs were preincubated with anti-VEGFR antibodies (as noted above) or control mouse IgG, washed, and then admixed with tumor cells before subcutaneous implantation in untransplanted GUSB–/– mice. After 8 days, the mice were killed and the tumor was removed for histochemical and immunofluorescent analysis to identify GUSB-positive, human vessels using human specific anti-von Willebrand factor (vWF; VW1–2, Takara). Each animal received all three conditions, and three animals were used for this experiment.

Flk-1/LacZ mice in C57Bl/6 background (Jackson Laboratory) represent mice heterozygous for the VEGFR2 null mutation in which the target gene was replaced with a promoterless lacZ gene from Escherichia coli. In the heterozygous mouse (flk/lacZ), expression of lacZ is subject to regulation by an EC-specific promoter (32) . Wild-type littermates transplanted with BM from flk/lacZ transgenic donors, generating flk/lacZ/BMTs. BMT mice were injected subcutaneous in one flank with 1 x 106 B16F10 mouse melanoma tumor cells [American Type Culture Collection (ATCC)] that had been stably transfected with VEGF cDNA (n=7) or mouse PlGF2 cDNA (n=6) or with vector control in the contralateral flank (34) . Two to four clones of each type were combined before injection. Animals were killed 14 days later, and tumors were resected and measured as described above. Donor BM-derived vessels (lacZ-positive) were distinguished by histochemistry, immunofluorescence, and quantitative ß-galactosidase (ß-gal) assay.

For BMT, unfractionated BM from syngeneic donor mice was obtained as described (29) . We injected 5 x 106 nucleated cells through the lateral tail vein from same sex donors into recipients that had been preconditioned with either 300 (NOD/SCID) or 800 (C57Bl/6) rads of {gamma}-radiation from a 137Cs source. This concentration of radiation resulted in >95% engraftment in all animals studied (data not shown). Studies post-BMT were performed 2 months after transplant. All protocols were approved by the Vanderbilt Institutional Animal Care and Use Committee.

CE-EPC culture
Human peripheral blood (hPB) leukocytes were isolated from human blood donor leukocyte reduction filters (LeukotrapRC, Pall Corporation) that were obtained from the American Red Cross (Nashville, TN) as described previously (40) ; three to four filters were pooled per prep to reduce donor variability. Mononuclear cells (MNCs) were isolated from blood leukocytes by density centrifugation and were directly plated at 1 x 108 cells/cm2 on 100 mM culture dishes or on 22 mm2 glass coverslips (placed in 6-well plates) coated with human fibronectin (Sigma) diluted 1:50 in HBSS and maintained in EBM-2 (Clonetics) with supplements as described previously (40 , 41) . CE-EPCs were used after 7 days by detaching with PBS and 1 mM EDTA.

Cell lines and generation of stable transfectants
Human umbilical vein endothelial cells (HUVECs; ATCC) and human microvascular endothelial cell (HMEC) (CDC, Atlanta) were maintained in EBM-2 with supplements. B16F10 murine melanoma cells (ATCC) were grown in Dulbecco’s modified Eagle’s medium media with 10% FCS, at 37°C with 5% CO2. Stable transfections with pcDNA3 plasmid encoding VEGF164 or PIGF-2 cDNA driven by the cytomeglovirus promoter were carried out with Lipofectamine 2000 (Invitrogen Life Technologies). After selection for 14 days with 1500 µg/ml G418 (Sigma), individual colonies were picked into 24-well plates cultured with medium containing 500 µg/ml G418. Mouse VEGF and PlGF-2 secreted into culture medium were measured as described by ELISA Quantikine kits (R&D Systems).

Quantitative enzyme biochemistry
Tissues were homogenized (150 mM NaCl, 10 mM Tris, pH7.5, 0.2% Triton X-100, and 1 mM DTT), and samples were centrifuged at 14,000 g for 15 min, 4°C. To quantify ß-gal activity, 50 µl of homogenized tumor tissue from flk/lacZ/BMT was tested in triplicate following the manufacturer protocol in the All-in-One ß-gal assay reagent (Pierce), PBS alone, as well as samples obtained from wild-type mice, was used as negative control. Tumors from flk/lacZ mice served as positive control.

For GUSB-specific biochemical assay, a portion of tissue homogenates were incubated at 37°C for 1 h in 100 µl 5 mM 4-MU-ß-D-glucuronide substrate (Sigma). Reactions were stopped with 0.1 M sodium carbonate and then assayed fluorometrically as described previously (42) . Specific activity (1 U=1 nmol substrate cleaved/h/mg protein) was calculated for lung and liver; total tumor GUSB activity was calculated by multiplying specific activity by total tumor mass. The protein concentrations were measured by using the Coomassie dye binding assay (Pierce).

Migration assay
CE-EPCs, HUVECs, or HMECs (8x105 for all) were isolated by EDTA treatment and incubated in the presence of anti-VEGFR1 (clone 6.12, Imclone Systems), -VEGFR2 (Clone 89106, R&D Systems), or -CXCR4 (clone 44716, R&D Systems) or control mouse IgG for 1 h on ice and then added to the upper chamber. Migration was assessed after 5 h using a modified Boyden chamber assay as described previously (43) . Migrating cells adhering to the undersurface of the filters were fixed and stained with crystal violet and counted. Results are indicative of four independent experiments performed in duplicate. Quantitative (q)RT-PCR to examine VEGF receptors were performed as described previously (44) . Primer sequences are available on request.

Histochemistry, immunofluorescence, immunohistochemistry, and morphometry
Tissues were collected and processed as described previously (29) . For immunofluorescence, sections were fixed for 20 min at 4°C in 100% acetone and then incubated with MOM block [5% (wt/wt) papain/2 uM EDTA, pH 7.4, and 20 mM L-cysteine] at dilution of 1:20, a PBS-blocking buffer (0.01 g/ml BSA/2 µg powdered milk/ml, and 3 µl/ml Triton X-100) and 10% v/v goat serum for 1–2 h as described previously (29) . Fluorescent staining of adherent cells on glass cover slips was conducted after 7 days in culture (40) . The images were visualized using a Ziess Axioplan 2 microscope equipped with a 20x/0.75 objective lens (Carl Ziess MicroImaging). Images were photographed with a CoolSNAP Hq CCD camera (Photometrics) and acquired with Metamorph v5.0 software (Universal Imaging Corporation). For GUSB-specific histochemical assay, samples were frozen in OCT embedding medium (Sakura, Torrance, CA) for histochemical analysis; 10 µm sections were stained for active GUSB as described previously by using naphthol-AS-BI-ß-D-glucuronide (Sigma) as a substrate and then counterstained with methyl green (42) . For ß-gal staining, frozen sections were prepared and stained as described previously (45) . In parallel, paraffin embedded, formalin fixed tissue slides were also stained with a polyclonal antibody to ß-gal (1:1000, Novus) and analyzed using histochemistry (46) . Histochemistry for GUSB was performed as described previously (46) . To determine whether CE-EPCs formed functional vessels, animals were injected intravenously with biotinylated BS-1 lectin (200 µl of 500 µg/ml; Vector Labs) 30 min before death. Tumors were resected, embedded in OCT, and processed as described before staining (29) . Coimmunofluorescent staining was performed with human-specific vWF and lectin was detected by rhodamine streptavidin.

Flow cytometry
Tumor tissues from GUSB/BMT were finely minced, washed to remove tumor-associated leukocytes, and treated with collagenase type IV (1 mg/ml, Worthington) for 30 min. Large debris was allowed to settle, and the suspension was red cell lysed and analyzed as described previously by using a fluorogenic GUSB substrate (Molecular Probes; ref 29 ) and VE-cadherin/PE (BV9, Cell Sciences). Tumor suspensions obtained from GUSB-negative and GUSB-positive animals were used as negative and positive controls, respectively, and used to set the GUSB gates. Isotype control antibodies were used as control for the VE-cadherin Ab. Nonviable cells, identified by 7-aminoactinomycin D (7-AAD; Molecular Probes) staining, were excluded. Quantitative analyses were performed on a FACScan flow cytometer and subsequently analyzed using Cellquest software (Becton-Dickinson).

Antibodies used for cell/tissue staining
Active caspase 3 (Promega), polyclonal hVEGFR2 (R&D Systems), hVEGFR1 (89106, R&D Systems), polyclonal hTie-1 (Santa Cruz), polyclonal Tie-2 (Santa Cruz), hVE-Cadherin (BV9, Cell Sciences), human-specific vWF (VW1–2, Takara), vWF (Dako), and polyclonal goat anti-human GUSB (provided by M. S. Sands). Appropriate secondary antibodies were purchased and used from Jackson ImmunoResearch and Molecular Probes.

Statistical analysis
Two-tailed paired Student’s t tests were performed to compare different data sets in which appropriate. All data are mean ± SE; P < 0.05 denotes statistical significance.

RESULTS

Recombinant VEGF admixed with tumor cells increased BM-derived ECs in tumors
We used two different mouse models of tumor growth to quantify the effect of VEGF on vasculogenesis. The first model used the GUSB-deficient mouse engrafted with GUSB-positive (wild-type) BM, resulting in GUSB/BMT (Fig. 1 E). This BMT model has been previously used to study the in vivo role of EPCs (29) . GUSB is a lysosomal enzyme that is expressed in virtually all cell types, including BM-derived cells (47) . On transplantation into a GUSB-deficient host, cells from normal mouse or human donors can be identified by virtue of their GUSB expression with several sensitive biochemical and histochemical assays (42) . A spontaneously occurring tumor was isolated from a GUSB-deficient mouse. Histological analysis of H&E stained sections from formalin-fixed, paraffin-embedded tumor tissue suggested that this was a malignant high-grade sarcoma (Fig. 1A -B). The tumor was negative for lysozyme sarcoplasmic actin, CD31, keratin, LCA, factor 13, vimentin, NE filament, ubiquitin, vWF, desmin, and HHF 35. The tumor was positive for CD117 (ckit), actin, and reticulin (Young and Vogler, data not shown). These findings excluded the diagnosis of angiosarcoma. Absence of GUSB enzyme activity was confirmed by two methods. First PCR analysis was performed on the tumor to demonstrate GUSB-specific mutation (Fig. 1D ). Second, histochemical stain for GUSB performed on a frozen section of the primary tumor tissue demonstrated the absence of GUSB enzyme activity (Fig 1C ). To determine tumor growth kinetics, 8- to 10-wk-old NOD/SCID mice both homozygous or wild type for the GUSB mutation were injected subcutaneously with 5 x 105 GUSB–/– tumor cells admixed with VEGF (16 ng/ml) and heparin in one flank and without VEGF in the contralateral flank. Mice were killed, and tumor volume was determined using the formula 0.52 x width2 x length (Fig. 1F ; ref. 43 ). Because there were no statistical differences in tumor growth among animals homozygous for GUSB mutation from wild type, the data were combined. Faster growth in VEGF admixed tumor was evident by day 5 (Fig. 1F ). We confirmed no direct effect of VEGF on tumor proliferation (data not shown). To determine whether BM-derived cells were contributing to tumor vasculature, we examined tumors obtained from GUSB/BMT after confirming >90% donor engraftment (Fig. 1E depicts the experimental design; ref 29 ). Each transplanted recipient received subcutaneous implantation of 2 x 106 GUSB–/– tumor cells with and without admixed VEGF in opposing flanks. The mice were killed either 5 (n=15) or 25 (n=6) days later, and tumors were analyzed for GUSB-positive (BM-derived) vessels. Donor-derived, GUSB-positive vascular structures that also costained with BS1 lectin (to mark blood vessels) were evident in tumors from both time points (Fig. 2 A shows only day 25 data) and clustered around tumor periphery. To determine whether VEGF affected the numbers of BM-derived cells within the tumor, we quantified GUSB-enzyme activity of tumor homogenates obtained at both time points. Tumor tissue admixed with VEGF that were obtained 5 days after implantation contained a fourfold greater GUSB specific activity than contralateral tumors without VEGF (Fig. 2B ). No statistical difference was evident between the VEGF vs. control samples for day 25 samples (Fig. 2B ). Since GUSB enzyme activity was present in all BM-derived cells and did not selectively quantify BM-derived ECs, we used flow cytometric analysis of collagenase-treated tumor suspensions to quantify the numbers of GUSB-positive (BM-derived) cells that coexpressed VE-cadherin, a marker specific for ECs (Fig. 2 C -D shows the control samples used to establish the gates and representative analysis of tumor suspensions obtained after day 5 of implantation). VEGF-containing tumors contained significantly greater percentage of BM-derived ECs (GUSB/VE-cadherin dual positive), 30.8 ± 8.2 vs. 10 ± 4.8%, P < 0.05, but only in tumors obtained 5 days after implantation (Fig. 2E -G). This effect of VEGF was not statistically different from control in tumors obtained 25 days after implantation. The lack of effect of VEGF at day 25 may be due to the short bioavailability of the admixed VEGF. The analysis of tumor homogenates after 5 days of implantation supported our hypothesis that locally increased VEGF levels promoted tumor vasculogenesis.


Figure 1
View larger version (92K):
[in this window]
[in a new window]
  		Figure 1. Tumor-associated VEGF promotes tumor growth of GUSB–/– tumor. H&E of primary GUSB–/– tumor that was generated spontaneously in a NOD/SCID/GUSB–/– animal. Shown are representative formalin fixed paraffin-embedded sections that demonstrate tumor tissue (T) adjacent to normal (Nl) stroma (A) or infiltrating normal muscle (NlMu; B). C) Staining of a tumor cryosection histochemically for GUSB (red) and counterstaining with methyl green shows absence of GUSB activity. Inset shows wild-type skeletal muscle stained in parallel as positive control. D) PCR analysis was performed to detect specific GUSB mutation. First lane: marker; lanes 2 and 3: GUSB–/– and wild-type tail DNA samples, respectively; lane 4: GUSB–/– tumor DNA. GUSB mutation results in loss of restriction cleavage size and a higher molecular mass fragment. E) Schematic of GUSB/BMT tumor model. Engrafted animals were implanted subcutaneously in left flank with tumor cells admixed with rVEGF and in right flank tumor cells with vehicle. Tumors were analyzed at day 5 or 25 after implantation by immunofluorescence, flow cytometry, or GUSB biochemistry. F) Tumor volume of GUSB–/–sarcoma in the absence and presence of VEGF was measured at day 5 after subcutaneous implantation. *P < 0.05., statistically significant.


Figure 2
View larger version (52K):
[in this window]
[in a new window]
  		Figure 2. Local VEGF promotes tumor vasculogenesis in the GUSB/BMT model. A) Representative photomicrographs of cryosections obtained from VEGF-containing tumors implanted into GUSB/BMT. Top left panel) Tumor vasculature as detected by BS1 lectin staining (arrows). Top right panel) Donor BM-derived vascular structures detected by indirect immunofluorescence with anti-GUSB antibody (arrows). Bottom left panel) Merge in which a subset of tumor vessels costained with donor marker; 4',6'-diamidino-2-phenylidole (DAPI) staining of same slide shows location of the nuclei. Bottom right panel) Histochemical stain for GUSB (red) of a tumor cryosection demonstrating donor-derived vascular structure (arrow). B) GUSB specific activity (nmol substrate cleaved/h/mg total protein) to quantify donor BM-derived cells was measured in tumor homogenates obtained at defined time points grown with and without rVEGF. C–F) Flow cytometric analysis of single cell GUSB/BMT tumor suspension after collagenase treatment were used to detect BM-derived (GUSB+), ECs (VE-cadherin+). VE-cadherin gates were set by isotype control and GUSB gates were set using tumors implanted in wild-type (C) or GUSB–/– animals (D). Representative dot plots of tumor cell suspensions obtained from a GUSB/BMT animal inoculated with tumor cells admixed with VEGF (E) or vehicle (F) in contralateral flanks. GUSB+/VE-cadherin+ dual-positive cells represent BM-derived ECs. The flow cytometry data from 3 independent experiments shown graphically in G. *Statistical significance.

Flk/lacZ/BMT model showed that increased local VEGF-promoted tumor vasculogenesis
We confirmed these findings with a second, independent mouse BMT model using transgenic mice that constitutively expressed ß-gal (lacZ) transcriptionally regulated by an EC-specific promoter, VEGFR2 (flk1; ref 32 , 48 ). They will be noted as flk/lacZ/BMT. The absence of a single VEGFR-2 allele on the EPCs may result in underestimation of vasculogenesis in this model by reducing donor-derived EPC recruitment or assembly into vessels. The use of this model, however, enabled clear distinction of vasculogenesis from angiogenesis, because the expression of lacZ in tissues was restricted to BM-derived cells of the EC lineage (32 , 48) . B16F10 melanoma clones stably expressing murine VEGF164 cDNA or stably transfected with control plasmid expressed 27.4 ± 6.4 vs. 12.7 ± 6.3 pg VEGF/µg total protein, respectively (P=0.01). Clones overexpressing VEGF were also compared with control in a proliferation assay to exclude direct effects of VEGF on tumor proliferation (data not shown). VEGF-overexpressing clones were combined and implanted subcutaneously into the left flank of flk/lacZ/BMT mice. The right flank was implanted with equal numbers of B16F10 cells that had been stably transfected with control plasmid. Animals were sacrificed, and tumor growth and vasculogenesis were analyzed after 14 days (schematic in Fig. 3 A). As expected, the tumor volume generated by VEGF-overexpressing melanoma cells was significantly larger, >16-fold (Fig. 3B ), than the contralateral nonvascular VEGF overexpressing tumor in each mouse (n=7), although there was wide variation in the size of tumors generated from mouse to mouse. Immunohistochemistry using anti-ß-gal antibody (Fig. 3C ) of tumors from flk/lacZ (untransplanted) animals confirmed ubiquitous lacZ-positive vascular structures, whereas tumors from wild-type mice not expressing the lacZ transgene were devoid of positive staining (shown in Fig. 3A inset as negative control). Immunohistochemistry was also performed on tumors obtained from flk/lacZ/BMT to identify donor-derived (lacZ+/vasculogenic) vessels. Few (<5%) lacZ-positive vessels were evident in control tumors, and almost all were located in the tumor periphery (Fig. 3D ). A greater number of lacZ-positive cells, again located primarily in the periphery, were identified in most sections of tumors obtained from VEGF-overexpressing tumors (Fig. 3E -F). To quantify tumor vasculogenesis, we measured ß-gal enzyme activity in tumor homogenates obtained from half of each resected tumor mass. This avoided sampling error caused by quantifying immunoreactive areas on multiple histological sections. The average ß-gal activity in control tumors vs. VEGF-overexpressing tumors was 0.65 ± 0.33 and 1.6 ± 0.77 optical density (OD)/mg total protein P < 0.05, respectively. The average ratio of ß-gal activity between VEGF and control tumor pairs isolated from each mouse is shown (Fig. 3G ). This difference was supported by our qualitative analysis (data not shown). Hence, tumor cells that over-expressed VEGF exhibited an {approx}2.5-fold increase in vasculogenesis.


Figure 3
View larger version (63K):
[in this window]
[in a new window]
  		Figure 3. Enhanced tumor VEGF expression promotes tumor vasculogenesis in the flk/LacZ/BMT model. A) Schematic demonstrates flk/lacZ/BMT model. After engraftment, animals were implanted s.c. with tumor cells stably transfected with control vector or vector encoding VEGF in opposing flanks. Tumors were analyzed at d 14 after implantation and assessed by immunohistochemistry and quantitative lacZ assay. B) Average of the fold change in tumor volumes generated by VEGF overexpressing B16F10 tumor cells versus control obtained from the same mouse 2 wk after implantation (n=7). C) Tumors implanted into flk/lacZ mice (positive controls) were analyzed by lacZ immunohistochemistry (arrows point to immunoreactive ECs of both large and small vessels). Parallel analysis of negative controls in inset. D) Representative histology from Bl6F10vector tumor obtained from flk/lacZ/BMT demonstrates a longitudinal section through a LacZ positive small vessel containing red blood cells (arrow). EûF) Representative histology from Bl6F10VEGF tumor obtained from flk/lacZ/BMT demonstrates numerous LacZ-positive vessels (arrows). While most donor BM-derived vessels were small capillaries (as in F), a larger vessel containing numerous red blood cells are evident in panel E. G) The fold change in ?-gal enzyme activity, as a marker for donor (BM-derived) vessels, was measured in VEGF-overexpressing and control B16F10 tumor homogenates residing within the same mouse to quantify the effect of VEGF on vasculogenesis.

VEGFR1 mediates EPC recruitment to site of tumorigenesis
Both in vivo models suggested that local VEGF expression increased vasculogenesis. Several studies have shown that CE-EPCs, generated after ex vivo expansion and differentiation of hPBMNCs, home to the site of neovascularization and incorporate into new vessels and promote vascularization (4 , 5) . Phenotypic characterization of hPB-derived CE-EPCs confirmed the expression of various EC marker proteins such as VEGFR1, VEGFR2, VE-cadherin, tie-1, tie-2, vWF, and UEA-1 lectin and uptake of DiI-acLDL (data not shown; ref 40 ). The role of VEGFRs in EPC recruitment in vivo was investigated by preincubation of CE-EPCs with receptor-specific antibodies that blocked both ligand binding and activation (49) . After tumors were evident, mice were injected intravenously with 5 x 105 CE-EPCs preincubated with either anti-VEGFR1-, -VEGFR2-blocking antibodies or control IgG (Fig. 4 A depicts experimental design). Since homing of stem and progenitor cells is believed to occur within 6–12 h (50 , 51) , the mice were killed 12 h after administration. The tumors were homogenized and assayed for GUSB enzyme activity as a quantitative marker for administered CE-EPCs. To control for experimental variation in total number of EPCs administered, the total tumor GUSB activity from each animal was normalized to lung GUSB specific activity before calculating fold change between experimental groups. GUSB specific activity (1 U=nmol/mg protein/h) in lung (as an example of normal tissue) was very low and similar between treated and control animals, 140 ± 20 vs. 110 ± 29 U, respectively. By contrast, GUSB specific activity in tumor tissue was {approx}10-fold (1320±80 U) higher than that in lung in control animals (n=3). Preincubation with VEGFR1 antibody resulted in a 5-fold decrease in total GUSB activity in tumors as compared with tumors from mice receiving EPCs preincubated with control IgG (Fig. 4B ; P=0.001). Preincubation of EPCs with anti-VEGFR2 antibody also diminished EPC homing to tumors, as determined by measuring fold change in GUSB activity, but this difference was not statistically significant (Fig. 4B ; P=0.13).


Figure 4
View larger version (51K):
[in this window]
[in a new window]
  		Figure 4. VEGFR1 mediates EPC recruitment to site of tumor growth and VEGFR1 agonist promotes tumor vasculogenesis. A) Experimental schematic used to study role of VEGFR1 vs. VEGFR2 in CE-EPC recruitment to tumor. Briefly, after tumor growth, GUSB–/– mice were injected with GUSB+/+ EPCs preincubated with either VEGFR1- or VEGFR2-blocking antibodies. GUSB activity at tumor site was measured after 12 h as a surrogate measure of EPC recruitment. B) Fold change in total GUSB enzyme activity in tumor homogenates as a quantitative measurement for CE-EPC homing to tumor site after preincubation with control IgG, or antibodies to VEGFR1 or VEGFR2. C) Schematic shows flk/lacZ/BMT model used to study role of PlGF (a specific ligand for VEGFR1) in tumor vasculogenesis. Tumor cells stably transfected with vector were implanted subcutaneously in right flank; left flank received tumor cells stably transfected with vector encoding PlGF. Tumors were analyzed after day 14 postimplantation. D) Average of the fold change in tumor volumes generated by PlGF overexpressing B16F10 tumor cells vs. control obtained from the same mouse 2 wk after implantation (n=6). E) Bl6F10PlGF-derived tumors obtained from flk/lacZ/BMT demonstrated several red blood cell containing, LacZ-positive vessels (arrow). F) Fold change in ß-gal enzyme activity, as a marker for donor (BM-derived) vessels, was measured in PlGF-overexpressing and control B16F10 tumor homogenates residing within the same mouse to quantify the effect of PlGF on vasculogenesis.

Tumor-associated PlGF increases tumor vasculogenesis
Our unexpected finding that VEGFR1 affects CE-EPC recruitment to tumor site led us to hypothesize that the VEGFR1-specific agonist PlGF also promotes vasculogenesis. To test this hypothesis, we generated stable transfectants of B16F10 melanoma cells overexpressing PlGF cDNA or control plasmid. Pooled clones expressed a 2.3-fold higher PlGF2 protein than pooled control transfectants (1112 vs. 492 pg/ml medium P<0.001, data not shown). A cell proliferation assay was used to exclude direct effects of PlGF on tumor proliferation (data not shown). PlGF-overexpressing clones were implanted subcutaneously into the left flank of flk/lacZ/BMT mice and equal numbers of control B16F10 cells into the right flank (Fig. 4C shows schematic). Tumors were analyzed after 14 days. Local overexpression of PlGF enhanced tumor size over control by {approx}4-fold (n=6; Fig. 4D ), consistent with published reports (34) . Immunohistochemistry using anti-ß-gal antibody of tumors obtained from flk/lacZ/BMT to identify donor-derived (lacZ+/vasculogenic) vessels of control tumors showed very few positive cells (representative image in Fig. 3D ). A greater number of lacZ-positive cells occurring in clusters were identified in several sections of tumors obtained from PlGF-overexpressing tumors (Fig. 4E ). To quantify tumor vasculogenesis between PlGF and control tumors, we measured ß-gal enzyme activity in tumor homogenates obtained from half of both control- and PlGF-expressing tumors. The average ß-gal activity in control tumors vs. PlGF-overexpressing tumors was 0.33 ± 0.14 and 0.64 ± 0.09 OD/mg total protein, P < 0.05, respectively, {approx}2-fold difference (Fig. 4F ). These data suggest that tumor-derived PlGF also promotes formation of BM-derived blood vessels.

VEGFR-blocking antibodies prevent CE-EPC-mediated vessel formation in tumors
VEGFR2 is known to be critical for EC survival and hence likely to be important for CE-EPCs (21) . To distinguish CE-EPC survival and vessel formation from recruitment effects of receptor-blocking antibodies, we preincubated 5 x 105 CE-EPCs in the presence of anti-VEGFR2 or -VEGFR1 antibodies or IgG control for 60 min before admixing them with 2 x 106 tumor cells and implanting them subcutaneously. Each mouse was implanted with all three conditions, thereby generating 3 foci of solid tumors. Tumors were removed after 8 days for analysis of functional vessel formation by GUSB-positive CE-EPCs. Control tumors contained numerous GUSB-positive cells, many that had assembled into vascular-like structures (Fig. 5 A). Abundant GUSB-positive cells were also evident in tumors to which CE-EPCs preincubated with VEGFR1 antibodies had been added (Fig. 5B ); however, no cord-like structures were evident. Unlike VEGFR1-blocking antibodies, the presence of VEGFR2-blocking antibodies significantly diminished the numbers of intact CE-EPCs in tumors as determined by counting intact numbers of GUSB-positive cells in 5 representative fields each from three different tumors (Fig. 5C, J ). VEGFR2 has been shown to be important for EC survival (21) ; hence, we examined the section for evidence of increased CE-EPC apoptosis using an antibody recognizing active caspase 3. Few intact, isolated GUSB-positive cells were detected in tumors containing CE-EPCs pretreated with VEGFR2 (Fig. 5G ); several of the immunoreactive cells costained for active caspase 3 (Fig. 5H , merge in inset). No caspase-3/GUSB+ dual positive CE-EPCs were identified in samples preincubated with VEGFR1 blocking antibodies (Fig. 5I ). CE-EPC-derived functional vessels (incubated with control IgG) were identified in tumors by colocalizing cells positive for human-specific anti-vWF antibody with BS1-lectin positive structures, which marked functional tumor vessels (Fig. 5D -F). However, preincubation of the CE-EPCs with either anti-VEGFR1 or -R2 disrupted vessel formation, as no vWF-positive vessels were detected within tumors (Fig 5B -C, insets). Thus, either VEGFR1 or VEGFR2 blocking antibodies prevented CE-EPC assembly into vessels within the tumor mass.


Figure 5
View larger version (94K):
[in this window]
[in a new window]
  		Figure 5. VEGFR blocking antibodies prevent CE-EPC-mediated vessel formation in tumors. Representative cryosections (A–F) from tumors removed after 8 days after implantation were processed with a histochemical stain (A–C) to detect GUSB-positive cells (human CE-EPCs, red) in tumor tissue. CE-EPCs were pretreated with IgG (A, D–F, negative control) or anti-VEGFR1 (B) or anti-VEGFR2 (C) before coimplantation with tumor cells. CE-EPCs treated with control antibody (IgG) formed vascular-structures (A, arrow) that were positive for human-specific vWF by indirect immunofluorescence (D), and the same section was also stained to detect BS-1 lectin (functional vessels), injected before death (E). F) Merge; DAPI staining of same slide shows location of nuclei. Similar analysis of cryosections obtained from anti- VEGFR1 or -R2-treated CE-EPCs showed no human-specific vWF staining (inset in b and c; DAPI-stained nuclei evident only). A tumor cryosection containing CE-EPCs preincubated with anti-VEGFR2 was costained with GUSB antibody (G) and active caspase 3 antibody (H) with merge shown as inset in H. I) Similar analysis of representative tumor section containing CE-EPCs preincubated with anti-VEGFR1 showed numerous GUSB-positive cells (green) but no caspase 3 staining of the same section (inset). J) Bar graph of average number of GUSB-positive CE-EPC cells/x 20 field in tumors containing pretreated CE-EPCs. Five random fields were quantified from 2 independent experiments. K) After 7 days in culture, anti VEGFR1, VEGFR2 or control IgG were added to culture medium of CE-EPCs for 48 h. Cells were trypsynized and viability evaluated by trypan blue exclusion. Graph shows percentage of live cells per 100 cells counted. Data represent 3 independent experiments, each performed in duplicate.

VEGF/VEGFR1 interactions play a unique role in VEGF-mediated CE-EPC migration in vitro
Published studies suggest that VEGF-induced EC migration, survival, and proliferation are mediated primarily by VEGFR2 (21 , 52) . Hence, we were surprised to identify VEGFR1 as an important mediator of CE-EPC recruitment to the tumor site. Using a modified Boyden chamber migration assay, we examined the role of VEGFR1 and VEGFR2 in mediating in vitro CE-EPC chemotaxis in response to VEGF as compared withECs (e.g., HUVECs and HMECs; Fig. 6 A–D). CE-EPCs migrated in a dose-dependent manner to both VEGF and the VEGFR1-specific agonist, PlGF (Fig. 6B ). To examine receptor specificity, CE-EPCs were preincubated in receptor-blocking anti-VEGFR1 or -VEGFR2 for 60 min before migration assay (Fig. 6C ). Cell viability of nonmigrated cells at the end of the assay was >90% (data not shown). Migration of CE-EPCs was equally responsive to PlGF, a VEGFR1-specific agonist (53) , as VEGF (Fig. 6B -C). Concentration of inhibition by VEGFR1 blocking antibody was commensurate with blocking anti-VEGFR2. By contrast, in vitro migration in response to VEGF of HMECs (Fig. 6D ) and HUVECs (data not shown) was mediated primarily by VEGFR2, consistent with published studies (21 , 52) . By (q)RT-PCR, we found that relative VEGFR2 transcript levels in cultured EPCs obtained from independent preparations were 75-fold (P=0.004) lower than ECs (i.e., HMECs), whereas VEGFR1 transcript levels were consistently {approx}3-fold higher (P=0.008) than in ECs, and neuropilin transcript levels were not statistically different between ECs and EPCs. Hence, unlike ECs, CE-EPCs express relatively higher levels of VEGFR1 transcripts, and their migration in vitro was mediated by both receptors.


Figure 6
View larger version (36K):
[in this window]
[in a new window]
  		Figure 6. VEGFR1 agonist has unique effects on VEGF-mediated CE-EPC chemotaxis. A) Representative photomicrographs of cells stained with crystal violet that have migrated toward the VEGFR1-selective agonist, PlGF in the absence (left panel) and presence (right panel) of a VEGFR1 blocking Ab. B) Quantification of basal, dose-escalating VEGF- and PlGF-induced migration of CE-EPCs. C) Relative migration of CE-EPCs toward VEGF or PlGF stimuli with or without preincubation of anti-human VEGFR1 or VEGFR2 Ab. D) Relative migration of HMECs toward VEGF in the absence or presence of specific VEGFR antibodies compared with migration in response to PlGF stimuli. E) Relative migration of CE-EPCs in response to VEGF, SDF-1, or combination in the absence and presence of specific receptor blocking antibodies. *P < 0.05., statistically significant.

No crosstalk between SDF-1- and VEGF-mediated migratory signals
Recently, it has been shown that the rise in plasma SDF-1 after insult is correlated with an increase in numbers of CE-EPCs that can be obtained from culturing human periferal blood MNCs (14 15 16 17 18) . Tissue SDF-1 is also increased during ischemia and has been shown to recruit CE-EPCs. We were interested in determining if there was crosstalk between the SDF-1 and VEGF pathway in mediating CE-EPC migration in vitro. Migration of CE-EPCs to rVEGF (17 nM) was similar to migration in response to 13 nM SDF-1 in a modified Boyden chamber assay (Fig. 6E ). The combination of VEGF and SDF-1 resulted in an additive increase in CE-EPC migration. Preincubation of CE-EPCs with neutralizing antibodies against CXCR4 did not diminish VEGF-induced migration but did abolish migration in response to SDF-1. Conversely, neutralizing antibody against VEGFR1 and VEGFR2 abolished migration in response to VEGF but failed to affect migration in response to SDF-1 (Fig. 6E ). These data suggest that VEGF and SDF-1 mediated CE-EPC migration is mediated by independent mechanistic pathways.

DISCUSSION

The proangiogenic functions of VEGF have been extensively studied (21 , 54) . Less well understood is its precise role in vasculogenesis (32 , 33 , 55 56 57 58 59) . There are emerging data that VEGF is also important in postnatal vasculogenesis, specifically in EPC mobilization into the circulation (18 , 24 25 26 , 30) . Coadministration with BM resulted in increased vasculogenesis in a neonatal model (29) . Finally, VEGF gene transfer into CE-EPCs promoted survival and adhesion in vitro and subsequently resulted in greater engraftment in vivo (24 , 30) . In this study, we further defined how VEGF may promote vasculogenesis. We show that VEGF overabundance at the site of tumor growth results in increased vasculogenesis using two different murine BMT models. In the first model, rVEGF admixed with tumor cells elicited 3-fold greater numbers of donor BM-derived ECs that was evident at 5 days of growth, but not after 25 days, a likely result of admixing recombinant VEGF (with limited half-life) at time of implantation. The finding that the enhanced vasculogenesis observed at early time points (when VEGF was available) did not persist at the later time point supports the notion that vasculogenic vessels regress over time. This is also supported by our observation that majority of the donor-derived vessels were located selectively in the periphery of the tumor. Aspects such as stability, remodeling, or expansion of postnatal vessels derived from vasculogenesis have not been well studied.

The second BMT model used tumor cells that were stably transfected with VEGF. In control tumors, the number of donor-derived, lacZ positive cells was rare in most histological fields. At 14 days of tumor growth in this model, vasculogenesis was 2.5-fold greater in VEGF-overexpressing tumors by a quantitative measurement of lacZ (expressed in only BM-derived vessels). Since the VEGF and control sites were within the same animal in both models, the effect of VEGF was local and not secondary to EPC mobilization or direct effects of VEGF on survival and adhesion of the circulating vascular stem cell population. Our data support a role of VEGF downstream of EPC mobilization or general effects on the circulating EPC population.

We further show that the VEGFR1 agonist PlGF, expressed by tumor cells, also led to an almost twofold increase in vasculogenesis over control. Interestingly, PlGF has been shown to enhance tumor growth and vascularity in mice (34) , and more recently, its overexpression has been associated with worse prognosis in human tumors (35 36 37) . To our knowledge, the role of PlGF in vasculogenesis has not been reported. Together, the VEGF and PlGF data support the hypothesis that the local microenvironment (i.e., local VEGF and/or PlGF levels) plays an important role in vasculogenesis. The present findings help to address the controversy resulting from the wide variation in the degree of tumor vasculogenesis observed in different model systems (22 , 60 61 62) . Some studies have demonstrated substantial contribution of EPCs to tumor endothelium. Absence of EPC-derived vessels in other studies may be the result of different models systems (e.g., use of different reporter constructs, such as SCL or different tumor types) or in method of tissue sampling. For example, we found substantially greater levels of vasculogenesis in the GUSB–/– tumors than in the B16F10-derived tumors. Although we did not rigorously quantify the number of vasculogenic vessels relative to angiogenic vessels, lacZ-positive vessels comprised <5% in control B16F10 tumors, consistent with published reports (60 61 62) . Interestingly, we have found that donor BM-derived vessels were not located uniformly through the tumor but occurred in clusters around the tumor periphery, suggesting that local cues, in addition to VEGF/PlGF overexpression, modulate this process. Ex vivo expanded EPCs (CE-EPCs) were used to further study the role of VEGF and VEGFRs in postnatal vasculogenesis as used in other studies (4 , 41 , 63 , 64) . VEGFR1 immunoneutralization resulted in consistent, statistically significant reduction in homing of exogenously administered EPCs, whereas VEGFR2 blockade had variable effects. VEGFR1 transcript expression was modestly higher in cultured EPCs than in ECs whereas VEGFR2 transcripts were much higher in ECs. The relatively lower VEGFR2 expression on CE-EPCs may explain the greater role played by VEGFR1 in CE-EPC migration in vitro and may be a partial explanation for the variable inhibition of homing observed with the VEGFR2-blocking antibodies. Consistent with the predominant role of VEGFR1 in cultured EPC recruitment in vivo, the VEGFR1 agonist PlGF had a significantly more robust effect on EPC chemotaxis as compared with ECs. It is well established that VEGF is a potent monocyte chemoattractant, and this effect is mediated by VEGFR1 (22 , 65 , 66) . VEGF-induced EPC migration in vivo was mediated primarily by VEGFR1. The data suggest that VEGF-VEGFR1 interactions in EPCs are distinct from differentiated ECs and that receptor specificity for the migration response may be lineage dependent.

Recent work has implicated ischemia-associated SDF-1 expression as important for CE-EPC recruitment in murine wound models (14 , 17) . Our data suggest that VEGF and SDF-1 likely act independently on EPCs to promote vasculogenesis.

In addition to EPC recruitment to the site of tumor vasculogenesis, our data also support a role for VEGF in the formation of functional vessels by exogenous CE-EPCs. Pretreatment of CE-EPCs with blocking antibodies to either VEGFR blocked assembly into functional vessels. In addition, blocking VEGFR2, but not VEGFR1, resulted in reduced viability of CE-EPCs. This suggests that VEGF/VEGFR2 signaling may provide necessary survival cues. These studies do not exclude the possibility that VEGF also has direct effects on the vascular endothelium to up-regulate adhesion molecules that promote leukostasis or its potential effects to increase vessel permeability and resultant extravasation of serum proteins leading to enhanced blood vessel formation (21 , 67) . Studies are underway to more completely define VEGF’s role in postnatal vasculogenesis to determine how modulating VEGF signaling may affect this process during tumor and ischemic neovascularization.

ACKNOWLEDGMENTS

The anti-VEGFR1 blocking antibody was a gift from Dr. Daniel J. Hicklin at ImClone Systems. GUSB antibody was a kind gift from Dr. M. Sands (Washington University, St. Louis, MO). The authors thank Drs. Jeffrey Davidson and Samuel A. Santoro for critical reading of the manuscript. Research described in this article was supported in part by Philip Morris USA Inc., and Philip Morris International (P. Carmeliet). This work was also supported by the Vanderbilt Physician Development Award, Pfizer Atorvastatin Research Award, Veterans Affairs, and KO8HL84020 from the National Institutes of Health, Bethesda, MD (P.P. Young).

Received for publication October 3, 2005. Accepted for publication February 21, 2006.

REFERENCES

  1. Raffi, S., Lyden, D. (2003) Therapeutic stem and progenitor cell transplantation for organ vascularization and regeneration. Nature Med. 9,702-712[CrossRef][Medline]
  2. Rabelink, T. J., de Boer, H. C., de Koning, E. J. P., van Zonneveld, A. (2004) Endothelial progenitor cells: more than an inflammatory response. Arteroscler. Thromb. Vasc. Biol. 24,1-6[Free Full Text]
  3. Urbich, C., Dimmeler, S. (2004) Endothelial progenitor cells characterization and role in vascular biology. Circ. Res. 95,343-353[Abstract/Free Full Text]
  4. Kalka, C., Masuda, H., Tomono, T., Kalka-moll, W. M., Silver, M., Kearney, M., Li, T., Isner, J. M., Asahara, T. (1999) Transplantation of ex vivo expanded endothelial progenitor cells for therapeutic neovascularizaition. Proc. Natl. Acad. Sci. U. S. A. 97,3422-3427
  5. Schatteman, G., Hanlon, H. D., Jiao, C., Dodds, S. G., Christy, B. A. (2000) Blood-derived angioblasts accelerate blood-flow restoration in diabetic mice. J. Clin. Invest. 106,571-578[Medline]
  6. Urbich, C., Heeschen, C., Aicher, A., Sasaki, K., Bruhl, T., Farhadi, M. R., Vajkoczy, P., Hofmann, W. K., Peters, C., Pennacchio, L. A., et al (2005) Cathepsin L is required for endothelial progenitor cell-induced neovascularization. Nat. Med. 11,206-213[CrossRef][Medline]
  7. Jiang, S., Walker, L., Afentoulis, M., Anderson, D. A., Jauron-Mills, L., Corless, C. L., Fleming, W. H. (2004) Transplanted human bone marrow contributes to vascular endothelium. Proc. Natl. Acad. Sci. U. S. A. 101,16891-16896[Abstract/Free Full Text]
  8. Hristov, M., Weber, C. (2004) Endothelial progenitor cells: characterization, pathophysiology, and possible clinical relevance. J. Cell Mol. Med. 8,498-508[Medline]
  9. Rumpold, H., Dominik, W., Koeck, R., Gunsilius, E. (2004) Endothelial progenitor cells: A source for therapeutic vasculogenesis. J. Cell Mol. Med. 8,509-518[Medline]
  10. Tepper, O. M., Capla, J. M., Galiano, R. D., Ceradini, D. J., Callaghan, M. J., Kleinman, M. E., Gurtner, G. C. (2005) Adult vasculogenesis occurs through in situ recruitment, proliferation and tubilization of circulating bone marrow-derived cells. Blood 105,1068-1077[Abstract/Free Full Text]
  11. Takahashi, T., Kalka, C., Haruchika, M., Chen, D., Silver, M., Klearny, M., Magner, M., Isner, J. M., Asahara, T. (1999) Ischemia- and cytokine-induced mobilization of bone marrow-derived endothelial progenitor cells for neovascularization. Nature Med. 5,434-438[CrossRef][Medline]
  12. Ii, M., Nishimura, H., Iwakura, A., Wecker, A., Eaton, E., Asahara, T., Losordo, D. W. (2005) Endothelial progenitor cells are rapidly recruited to myocardium and mediate protective effect of ischemic preconditioning via "imported" nitric oxide synthase activity. Circulation 111,1114-11120[Abstract/Free Full Text]
  13. Thomas, J., Liu, F., Link, D. C. (2002) Mechanisms of mobilization of hematopoietic progenitors with granulocyte-colony stimulating factor. Curr. Opin. Hematol. 9,183-189[CrossRef][Medline]
  14. De Falco, E., Porcelli, D., Torella, A. R., Straino, S., Lachininoto, M. G., Orlandi, A., Truffa, S., Biglioli, P., Napolitano, M., Capogrossi, M. C., Pesce, M. (2004) SDF-1 involvement in endothelial phenotype and ischemia-induced recruitment of bone marrow progenitor cells. Blood 104,3472-3482[Abstract/Free Full Text]
  15. Yamaguchi, J., Kusano, K. F., Masuo, O., Kawamoto, A., Silver, M., Murasawa, S., Bosch-Marce, M., Masuda, H., Losordo, D. W., Isner, J. M., Asahara, T. (2003) Stromal cell-derived factor-1 effects on ex vivo expanded endothelial progenitor cell recruitment for ischemic neovascularization. Circulation 107,1322-1328[Abstract/Free Full Text]
  16. Raffi, S., Heissig, B., Hattori, K. (2002) Efficient mobilization and recruitment of marrow-derived endothelial and hematopoietic stem cells by adenoviral vectors expressing angiogenic factors. Gene Ther. 2002,631-641
  17. Ceradini, D. J., Kulkarni, A. R., Callaghan, M. J., Tepper, O. M., Basitidas, N., Kleinman, M. E., Capla, J. M., Galiano, R. D., Levine, J. P., Gurtner, G. C. (2004) Progenitor cell trafficking is regulated by hypoxic gradients through HIP-1 induction of SDF-1. Nat. Med. 10,858-864[CrossRef][Medline]
  18. Moore, M. A., Hattori, K., Heissig, B., Shieh, J. H., Dias, S., Crystal, R. G., Rafii, S. (2001) Mobilization of endothelial and hematopoietic stem and progenitor cells by adenovector-mediated elevation of serum levels of SDF-1, VEGF and angiopoietin-1. Ann. N. Y. Acad. Sci. 938,45-47
  19. Murdoch, C., Giannoudis, A., Lewis, C. E. (2004) Mechanisms regulating the recruitment of macrophages into hypoxic areas of tumors and other ischemic tissues. Blood 104,2224-2234[Abstract/Free Full Text]
  20. Xie, K., Wei, D., Shi, Q., Huang, S. (2004) Constitutive and inducible expression and regulation of vascular endothelial growth factor. Cytokine Growth Factor Rev. 15,297-324[CrossRef][Medline]
  21. Ferrara, N. (2001) Role of vascular endothleial growth factor in regulation of physiological angiogenesis. Am. J. Physiol. Cell Physiol. 280,C1358-C1366[Abstract/Free Full Text]
  22. Lyden, D., Hattori, K., Dias, S., Costa, C., Blaikie, P., Butros, L., Chadburn, A., Heissig, B., Marks, W., Witte, L., et al (2001) Impaired recruitment of bone-marrow-derived endothelial and hematopoietic precursor cells blocks tumor angiogenesis and growth. Nature Med. 7,1194-1201[CrossRef][Medline]
  23. Poole, T. J., Finkelstein, E. B., Cox, C. M. (2001) The role of FGF and VEGF in angioblast induction and migration during vascular development. Dev. Dyn. 220,1-17[CrossRef][Medline]
  24. Kalka, C., Masuda, H., Takahashi, T., Gordon, R., Tepper, O. M., Gravereaux, E., Pieczek, A., Iw, H., Hayashi, S., Isner, J. M., Asahara, T. (2000) Vascular endothelial growth factor165 gene transfer augments circulating endothelial progenitor cells in human subjects. Circ. Res. 86,1198-1202[Abstract/Free Full Text]
  25. Asahara, T., Takahashi, T., Masuda, H., Kalka, C., D, C., Iwaguro, H. (1999) VEGF contributes to postnatal neovascularization by mobilizing bone marrow-derived endothelial progenitor cells. EMBO J. 18,3964-3972[CrossRef][Medline]
  26. Hattori, K., Dias, S., Beate, H., Hackett, N. R., Lyden, D., Tateno, M., Hicklin, D. J., Zhu, Z., Witte, L., Crystal, R. G., et al (2001) Vascular endothelial growth factor and angiopoietin-1 stimulate postnatal hematopoiesis by recruitment of vasculogenic and hematopoietic stem cells. J. Exp. Med. 193,1005-1014[Abstract/Free Full Text]
  27. Shintani, S., Murohara, T., Ikeda, H., Ueno, T., Honma, T., Katoh, A., Sasaki, K., Shimada, T., Oike, Y., Imaizumi, T. (2001) Mobilization of endothelial progenitor cells in patients with acute myocardial infarction. Circulation 103,2776-2779[Abstract/Free Full Text]
  28. Gill, M., Dias, S., Hattori, K., Rivera, M. L., Hicklin, D. J., Witte, L., Giradi, L., Yurt, R., Himel, H., Rafii, S. (2001) Vascular trauma induces rapid but transient mobilization of VEGFR2(+)AC133(+) endothelial precursor cells. Circ. Res. 88,167-174[Abstract/Free Full Text]
  29. Young, P. P., Hofling, A. A., Sands, M. S. (2002) VEGF increases engraftment of bone marrow-derived endothelial progenitor cells (EPCs) into vasculature of newborn murine recipients. Proc. Natl. Acad. Sci. U. S. A. 99,11951-11956[Abstract/Free Full Text]
  30. Iwaguro, H., Yamaguchi, J.-I., Kalka, C., Murasawa, S., Masuda, H., Hayashi, S., Silver, M., Li, T., Isner, J. M., Asahara, T. (2002) Endothelial progenitor cell vascular endothelial growth factor gene transfer for vascular regeneration. Circulation 12,732-738
  31. Soker, S., Takashima, S., Miao, H. Q., Neufeld, G., Klagsburn, M. (1998) Neuropilin-1 is expressed by endothelial and tumor cells as an isoform-specific receptor for vascular endothelial growth factor. Cell 92,735-745[CrossRef][Medline]
  32. Shalaby, F., Rossant, J., Yamaguchi, T. P., Gertenstein, M., Wu, X. F., Breitman, M., Schuh, A. C. (1995) Failure of blood island formation and vasculogenesis in Flk-1 deficient mice. Nature 376,62-66[CrossRef][Medline]
  33. Fong, G. H., Rossant, J., Gertenstein, M., Breitman, M. (1995) Role of Flt-1 receptor tyrosine kinase in regulation of assembly of vascular endothelium. Nature 376,66-67[CrossRef][Medline]
  34. Luttun, A., Tjwa, M., Moons, L., Wu, Y., Angelillo-Scherrer, A., Liao, F., Nagy, J. A., Hooper, A., Priller, J., De Klerck, B., et al (2002) Revascularization of ischemic tissues by PIGF treatment, and inhibition of tumor angiogenesis, arthritis and atherosclerosis by anti-Flt-1. Nature Med. 8,831-840[CrossRef][Medline]
  35. Parr, C., Watkins, G., Boulton, M., Cai, J., Jiang, W. G. (2005) Placenta growth factor is over-expressed and has prognostic value in human breast cancer. Eur. J. Cancer 41,2819-2827[CrossRef][Medline]
  36. Fragoso, R., Pereira, T., Wu, Y., Zhu, Z., Cabecadas, J., Dias, S. (2005) VEGFR-1 (FLT-1) activation modulates acute lymphoblastic leukemia localization and survival within the bone marrow, determining the onset of extramedullary disease. Blood 107,1608-1616[Medline]
  37. Zhang, L., Chen, J., Ke, Y., Mansel, R. E., Jiang, W. G. (2005) Expression of placental growth factor (PlGF) in non-small cell lung cancer (NSCLC) and the clinical and prognostic significance. World J. Surg. Oncol. 3,68[CrossRef][Medline]
  38. Hofling, A. A., Vogler, C., Creer, M. H., Sands, M. S. (2002) Engraftment of human CD 34+ cells leads to widespread distribution of donor derived cells and correction of tissue pathology in a novel murine xenotransplantation model of lysosomal storage disease. Blood 101,2054-2063[Medline]
  39. Freeman, B. J., Roberts, M. S., Vogler, C., Nicholes, A., Hofling, A. A., Sands, M. S. (1999) Behavior and therapeutic efficacy of beta-glucuronidase-positive mononuclear phagocytes in a murine model of mucopolysaccharidosis type VII. Blood 94,2142-2150[Abstract/Free Full Text]
  40. Teleron, A. A., Carlson, B., Young, P. P. (2004) Blood donor WBC reduction filters as a source of human peripheral blood-derived endothelial progenitor cells (EPCs). Transfusion 45,21-25[CrossRef]
  41. Hill, J. M., Zalos, G., Halcox, J. P. J., Schenke, W. H., Waclawiw, M. A., Quyyumi, A. A., Finkel, T. (2003) Circulating endlthelial progenitor cells vascular function, and cardiovascular risk. N. Engl. J. Med. 248,593-600
  42. Glaser, J. H., Sly, W. S. (1973) beta-glucuronidase deficiency mucopolysaccharidosis: methods for enzymatic diagnosis. J. Lab. Clin. Med. 82,969-977[Medline]
  43. Brantley, D. M., Cheng, N., Thompson, E. J., Qing, L., Brekken, R. A., Thorpe, P. E., Muraoka, R. S., Cerretti, D. P., Pozzi, A., Jackson, D., Lin, C., Chen, J. (2002) Soluble Eph A receptors inhibit tumor angiogenesis and progression in vivo. Oncogene 21,7011-7026[CrossRef][Medline]
  44. Young, P. P., Modur, V., Teleron, A. A., Ladenson, J. (2005) Enrichment of genes in the aortic initma whic are associated with stratified epithelium: implications of underlying biomechanical and barrier properties of the arterial intima. Circulation 111,2382-2390[Abstract/Free Full Text]
  45. Hofling, A. A., Devine, S., Vogler, C., Sands, M. S. (2004) Human CD34+ hematopoietic progenitor cell-directed lentiviral-mediated gene therapy in a xenotransplantation model of lysosomal storage disease. Mol. Therapy 9,856-865
  46. Young, P. P., Vogler, C., Hofling, A. A., Sands, M. S. (2003) Biodistribution and efficacy of donor lymphocytes in a murine model of lysosomal storage disease. Mol. Therapy 7,52-61
  47. Fishman, W. H., Goldman, S. S., Delellis, R. (1967) Dual localization of beta-glucuronidase in endoplasmic reticulum and in lysosomes. Nature 213,457-460[CrossRef][Medline]
  48. Asahara, T., Masuda, H., Takahashi, T., Kalka, C., Pastore, C., Silver, M., Kearne, M., Magner, M., Isner, J. M. (1999) Bone marrow origin of endothelial progenitor cells responsible for postnatal vasculogenesis in physiological and pathological neovascularization. Circ. Res. 85,221-228[Abstract/Free Full Text]
  49. Vincent, L., Jin, D. K., Karajannis, M. A., Shido, K., Hooper, A. T., Rashbaum, W. K., Pytowski, B., Wu, Y., Hicklin, D. J., Zhu, Z., et al (2005) Fetal stromal-dependent pracrine and intracrine vascular endothelial growth factor-A/vascular endothelil growth factor receoptor-1 signaling promotes proliferation and motility of human primary myeloma cells. Cancer Res. 65,3185-3192[Abstract/Free Full Text]
  50. Asaumi, N., Omoto, E., Mahmut, N., Katayama, Y., Takeda, K., Shinagawa, K., Harada, M. (2001) Very late antigen-5 and leukocyte function-associated antigen-1 are critical for early stage hematopoietic progenitor cell homing. Annals Hematol. 80,387-392
  51. Solanilla, A., Crosset, C., Duchez, P., Legembre, P., Pitard, V., Dupouy, M., Belloc, F., Viallard, J.-F., Reiffers, J., Boiron, J.-M., et al (2003) Flt3-lingand induces adhesion of haematopoietic progenitor cells via a very late antigen (VLA)-4- and VLA-5-dependent mechanism. Brit. J. Hematol. 120,782-786[CrossRef][Medline]
  52. Kanno, S., Oda, N., Abe, M., Tera, Y., Ho, M., Shitara, K., Tabayashi, K., Shibuya, M., Sata, Y. (2000) Roles of two VEGF receptors, Flt-1 and KDR, in the signal transduction of VEGF effects in human vascular endothelial cells. Oncogene 19,2138-2146[CrossRef][Medline]
  53. Autiero, M., Luttun, A., Tiwa, M., Carmeliet, G. (2003) Placental growth factor and its receptor, vascular endothelial growth factor receptor-1: novel targets for stimulation of ischemic tissue revascularization and inhibition of angiogenic and inflammatory disorders. J. Throm. Haem. 1,1356-1370
  54. Ferrara, N., Alitalo, K. (1999) Clinical applications of angiogenic growth factors and their inhibitors. Nature Med. 5,1359-1364[CrossRef][Medline]
  55. Argraves, W. S., Larue, A. C., Fleming, P. A., Drake, C. J. (2002) VEGF signaling is required for the assembly but not the maintenance of embryonic blood vessels. Dev. Dyn. 225,298-304[CrossRef][Medline]
  56. Bautch, V. L., Redick, S. D., Scalia, A., Harmaty, M., Carmeliet, P., Rapoport, R. (2000) Characterization of the vasculogenic block in the absence of vascular endothelial growth factor-A. Blood 95,1979-1987[Abstract/Free Full Text]
  57. Drake, C. J., Little, C. D. (1995) Exogenous vascular endothelial growth factor induces malformed and hyperfused vessels during embryonic neovascularization. Proc. Natl. Acad. Sci., U. S. A. 92,7657-7661[Abstract/Free Full Text]
  58. Drake, C. J., LaRue, A., Ferrara, N., Little, C. D. (2000) VEGF regulates cell behavior during vasculogenesis. Dev. Biol. 224,178-188[CrossRef][Medline]
  59. Risau, W., Flamme, I. (1995) Vasculogenesis. Annu. Rev. Cell Dev. Biol. 11,73-91[CrossRef][Medline]
  60. Conejo-Garcia, J. R., Buckanovich, R. J., Benencia, F., Courreges, M., Rubin, S. C., Carroll, R. G., Coukos, G. (2005) Vascular leukocytes contribute to tumor vascularization. Blood 105,679-681<