| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
ARTICLES |
Medical Research Council (C.W., P.T.K.S., G.A.S., H.M.F.), Human Reproductive Sciences Unit, EH3 9ET Edinburgh, United Kingdom; and Regeneron Pharmaceuticals (S.J.W.), Tarrytown, New York 10591
Address all correspondence and requests for reprints to: Dr. C. Wulff, Medical Research Council, Human Reproductive Sciences Unit, 37 Chalmers Street, Edinburgh EH3 9ET, United Kingdom. E-mail: c.wulff{at}hrsu.mrc.ac.uk
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Because one of the most likely factors involved in the regulation of follicular angiogenesis is VEGF, we investigated its physiological role by inhibiting its action in vivo by treating marmosets with a soluble truncated form of the Flt receptor, Flt-1-Fc (VEGF TrapA40). VEGF TrapA40 has a high affinity for all forms of endogenous VEGF, preventing VEGF from binding to its receptors. We examined the effects of VEGF inhibition on thecal vascularization and on the expression of angiogenic factors and their receptors in growing and atretic follicles.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
Flt-1-Fc (VEGF TrapA40)
As described previously (12) the VEGF
TrapA40 used in these experiments comprised a
portion of the extracellular domain of VEGF receptor Flt-1 fused to the
Fc portion of human IgG. This soluble truncated form of the Flt-1
receptor has been demonstrated to neutralize circulating VEGF and
inhibit its action (12). Levels of circulating bound and
unbound VEGF TrapA40 in plasma of marmosets were
measured and by calculation of the ratio of bound to unbound VEGF trap
it was shown that 99% of the circulating VEGF trap was unmodified and
remained available for neutralization of endogenous VEGF
(12).
Animals
Adult female common marmoset monkeys (Callithrix
jacchus) with a body weight of approximately 350 g and
regular ovulatory cycles were housed together with a younger sister or
prepubertal female as described previously (13). Blood
samples were collected three times per week by femoral venipuncture
without anesthesia and plasma was subjected to progesterone assay to
confirm normal ovulatory cycles.
Treatment
Experiments were carried out in accordance with the Animals
(Scientific Procedures) Act, 1986 and approved by the local Ethical
Review Process Committee. To synchronize timing of ovulation, animals
were treated with 1 µg PGF2
analog (Planate,
Coopers Animals Health Ltd., Crewe, UK) im during the mid to late
luteal phase to induce luteolysis. This treatment is normally followed
by ovulation 10 days later (14). Four marmosets were
treated with VEGF trap at a dose of 25 mg/kg, approximately 1 ml
injected at two sc sites over the abdominal region. The same number of
controls were similarly treated with vehicle alone. Treatment commenced
on the day of presumed ovulation (luteal day 0) and was repeated on
days 1 and 2. On day 3 the animals were injected iv with 20 mg BrdU
(Roche Molecular Biochemicals, Essex, UK) in saline. One
hour later, the animals were sedated using 100 µl ketamine
hydrochloride (Parke-Davis Veterinary, Pontypool, Gwent,
UK) and killed with an iv injection of 400 µl Euthetal (sodium
pentobarbitone; Rhone Merieux, Harlow, Essex, UK). After cardiac
exsanguination via a heparinized syringe, ovaries were removed
immediately and fixed in 4% paraformaldehyde. After 24 h, the
ovaries were dehydrated and embedded in paraffin according to standard
procedures.
Immunocytochemistry
Establishment of the thecal vascular network was studied by 1)
quantifying the number of mitotic cells stained for BrdU, 2)
identifying endothelial cells using CD31 staining, and 3) determining
the incidence of colocalization of BrdU and CD31.
Tissue sections (5 µm) were cut onto 3-aminopropyltriethoxy saline Tespa-coated slides (Sigma, St Louis, MO) for immunocytochemistry. Sections were dewaxed in xylene, rehydrated in descending concentrations of ethanol, and washed in distilled water. For BrdU and CD31 immunostaining, antigen retrieval was performed by pressure cooking (Tefal Clypso pressure cooker; Tefal, Essex, UK) sections in 0.01 M citrate buffer, pH 6, for 6 min at high pressure setting 2. Slides were then left for 20 min in hot buffer and washed in Tris-buffered saline (TBS) (0.05 mol/liter Tris, 9 g/liter NaCl). To reduce nonspecific binding sections were blocked in normal rabbit serum (NRS, 1:5 diluted in TBS) for 30 min. Primary antibodies CD31 (mouse antihuman CD31; DAKO Corp., Ely, Cambridgeshire, UK) or BrdU (mouse anti-BrdU; Roche Molecular Biochemicals) were diluted 1:20 or 1:30 in TBS, respectively. Incubation was carried out overnight at 4 C. Slides were washed three times in TBS. Incubation with the secondary antibody (rabbit antimouse Ig, 1:60 diluted in TBS (DAKO Corp.) was performed for 40 min at room temperature followed after 2 washes in TBS by incubation of the APAAP complex (1:100 dilution in TBS, DAKO Corp.) for 40 min at room temperature. Visualization was performed using 500 µl/slide 4-nitro blue tetrazolium chloride solution containing 45 µl NBT substrate (Roche Molecular Biochemicals), 10 ml NBT buffer, 35 µl Xphosphate, and 10 µl levamisole. Sections for BrdU were counterstained with hematoxylin, whereas sections for CD31 were not counterstained so that quantitative image analysis could be performed. For dual labeling, slides were incubated first with CD31 and visualization with fast red (Sigma, St. Louis, MO). The fast red solution contained 1 mg fast red in 1 ml fast red buffer (20 mg naphtol AS-MX phosphate, 2 ml dimethyl formamide, 98 ml 0.1 M Tris, pH 8.2). Staining for CD31 was followed by BrdU, which was visualized with NBT as described above.
Tie-2 immunocytochemistry
Tissue sections were dewaxed, rehydrated, and placed into TBS as
described above. To reduce nonspecific staining, sections were
incubated for 20 min in NRS containing avidin, washed in TBS, and
incubated for another 20 min in NRS biotin (Avidin/Biotin blocking Kit;
Vector Laboratories, Inc., Burlingame, CA). After two
washes in TBS, slides were exposed overnight at 4 C to the Tie-2
antibody (mouse-antirat Tie-2 monoclonal, 1:500 dilution;
Regeneron Pharmaceuticals, Inc., Tarrytown, NY). Slides
were washed two times in TBS followed by incubation for 30 min at room
temperature with the secondary antibody (rabbit-antimouse-biotinylated
antibody, 1:200 dilution; DAKO Corp.). For signal
amplification the ABC (avidin-biotin-complex) method was used.
After incubation with ABC-HPR (horse raddish peroxidase) (DAKO Corp.) for 30 min at room temperature and two washes in
TBS-Tween, signal enhancement was achieved by exposing the slides to
biotinylated tyramide (GenPoint kit; DAKO Corp.) for 20
min at room temperature, followed by incubation with ABC-AP (alkaline
phosphatase) for 30 min. The signal was detected using NBT substrate as
described above.
In situ hybridization
In situ hybridization was performed as described
previously (15) using complementary RNA probes for human
VEGF, Flt, Ang-1, Ang-2, and Tie-2. Sense and antisense probes were
prepared using an RNA transcription kit (Ambion, Inc.,
Austin, TX) and labeled with 35S uridine
5'-triphosphate (NEN Life Science Products, Boston, MA).
Deparaffinized sections were treated with 0.1 N HCl and
then digested in proteinase K (5 µg/ml, Sigma) for 30
min at 37 C. After prehybridization for 2 h at 50 C for VEGF and
55 C for Flt, Ang-1, Ang-2, and Tie-2 subsequent hybridization was
performed in a moist chamber overnight at 50 or 55 C. High stringency
post hybridization washings and RNase treatment were used to remove
excess probe. Slides were then dehydrated, dried, and dipped in Ilford
G5 liquid emulsion (H. A. West, Edinburgh, UK). Exposure
times for VEGF, Flt, Ang-2, and Tie-2 were 2, 4, 7, and 7 weeks,
respectively. Slides were subsequently developed (D19 developer;
Kodak, Rochester, NY) and fixed (Kodak GBX
Fixer/Replenisher). All slides were counterstained with
hematoxylin (Richard-Allan, Richland, MI), dehydrated, and mounted.
Analysis of data
Morphological characterization of ovarian follicles. Stages
of follicular development were defined as follows: primordial (oocytes
surrounded by single flat layer of follicle epithelial cells), primary
(single layer of cuboidal granulosa cells), early secondary (two to
four layers of granulosa cells), late secondary (more than four
granulosa cell layers, no antrum), and tertiary follicles (follicles
containing an antrum). Follicles were classified as healthy if they
contained a normal shaped oocyte surrounded by granulosa cells that
were regularly apposed on an intact basement membrane with normal
appearance of granulosa cell nuclei without signs of pycnosis.
Follicles not fulfilling these criteria were classified as atretic.
Only follicles with a visible oocyte were considered to ensure proper follicular counting and classification. In each ovary a total of 20–30 primordial, 8–10 primary, 5–10 early secondary, 5–8 late secondary, and 4–6 tertiary healthy and atretic follicles were analyzed, respectively.
Quantification of immunocytochemistry and in situ hybridization. Quantitative analysis for BrdU and CD31 immunocytochemistry and in situ hybridization was performed using an image analysis system linked to an Olympus Corp. camera, and the data were processed using Image-Pro plus Version 3.0 for Windows (Microsoft Corp., Reading, Berkshire, UK). In all follicles, the whole cross sections were analyzed. Captured images were thresholded, and the theca and granulosa cell compartment was outlined and analyzed separately.
BrdU labeling. Sections were analyzed under 200x magnification. The image analysis system was set up to measure the number of dark stained nuclei (BrdU positive), and the number of dark and light stained nuclei (total number of cells) in the outlined compartment of interest. A proliferation index (i.e. BrdU-positive cells expressed as a percentage of the total number of cells) was calculated in the theca and granulosa compartment for each follicle. The proliferation index was expressed as a mean value for the number of follicles assessed within each follicular stage and per animal.
CD31 labeling. The endothelial cell area (i.e. CD31-positive cells) was measured at x200 magnification. The captured gray scale image was thresholded and converted to a binary image. The whole area of the theca compartment and the CD31-positive area within the compartment was measured. The CD31-positive area was then calculated per unit area of the theca compartment and expressed as a mean value for the number of follicles assessed within each follicular stage and per animal.
Colocalization of CD31 and BrdU. The number of dual-stained cells (BrdU- and CD31-positive cells) and the total number of proliferating cells (i.e. BrdU-positive cells) were counted in the theca compartment. The proportion of proliferating endothelial cells (dual-stained cells) was expressed as a percentage of the total number of proliferating cells. The mean value was calculated for the number of follicles assessed within each follicular stages and per animal.
In situ hybridization. Slides were analyzed qualitatively under lightfield and quantitatively under darkfield conditions at x 400 magnification. As a value for gene expression the grain density (number of grains/µm2) was measured. The image analyses system was set up to identify mRNA expressing cells, to outline them and measure the grain density within these cells. Tissue background density was measured for each antisense slide and subtracted from the measurements. The mean value was calculated for the number of follicles assessed within each follicular stage and per animal.
Statistical analysis. Data obtained for normal follicular angiogenesis were tested for significant differences using ANOVA followed by Duncans multiple range test. Effects of the treatment as compared with controls, were determined using a two-tailed, unpaired t test. Differences were considered to be significant if P less than 0.05. The tests were performed using SPSS version 6.1 for Macintosh (SPSS, Inc., Chicago, IL). All values are given as mean ± SEM.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
) and
first visible in follicles containing more than four granulosa cell
layers, i.e. the late secondary stage (Fig. 1B). The
endothelial cell area was somewhat increased in tertiary follicles
(Fig. 1C) compared with late secondary follicles; however, this was not
significant. In contrast, the endothelial cell area was markedly
reduced (P < 0.05) by 63–71% in atretic follicles
(Fig. 1D) compared with secondary and tertiary follicles (Fig. 2A).
|
|
and quantified in Fig. 2
. In early
secondary follicles a theca like formation is visible in which single
cells proliferate (Fig. 3A). In the late (Fig. 3B) secondary stage a
significant 15-fold increase of theca proliferation was found compared
with the early secondary follicles (Fig. 2B). No further increase was
observed in tertiary follicles (Figs. 3C and 2B). Proliferation in the
theca was markedly reduced (P < 0.05) by 76% during
follicular atresia (Figs. 3D and 2B).
|
) increased significantly from the primary to early and to late
secondary follicles. The proliferation index of late secondary
follicles was not different compared with the tertiary stage. In
atretic follicles, proliferation of the granulosa was severely reduced
(P < 0.05).
Colocalization of CD31 and BrdU
BrdU-positive cells of the theca in early secondary follicles were
not dual stained with CD31, indicating that cells other than
endothelial cells proliferate in early secondary stages. The earliest
clear evidence of vascular proliferation was found in the theca of late
secondary follicles. In these follicles, 25 ± 5% of
proliferating cells were dual stained. The percentage of double-labeled
proliferating cells in tertiary follicles was 30 ± 5% (Fig. 3C, inset).
In situ hybridization
VEGF mRNA was first detected in late secondary follicles (Fig. 4A), grain density in the granulosa and
theca being similar (0.005 ± 0.001
grains/µm2). A significant increase in VEGF
mRNA expression was observed within the granulosa of follicles in the
tertiary stage (Fig. 5A), whereas in
these follicles VEGF expression in the theca was significantly lower
(0.003 ± 0.001 grains/µm2). During
atresia, the VEGF expression was significantly reduced by 55% in the
granulosa and was no longer detectable in the theca. The Flt receptor
mRNA was expressed in endothelial cells of the theca of late secondary
follicles (Fig. 4B) and increased significantly in the theca of
tertiary follicles (Fig. 5B). During follicular atresia, a significant
50–70% decrease was observed compared with late secondary and
tertiary follicles. Ang-1 mRNA appeared to be expressed at a very low
level (0.003 ± 0.0005 grains/µm2) in the
theca of tertiary follicles. No differences in grain density were found
between tertiary and atretic follicles. No Ang-2 expression was found
in follicles at any stages investigated. Ang-2 mRNA was moderately
expressed only by a subset of blood vessels within the ovarian stroma.
Ang-1 and Ang-2 mRNA were highly expressed in sections of marmoset
placenta included in the run as a positive control (not shown). Tie-2
mRNA expression was found in tertiary follicles (Fig. 4C) mainly
confined to the endothelium of the theca cell compartment. During
follicular atresia, expression of Tie-2 appeared to cease and image
analysis revealed a significant 70% decrease of grain density in the
theca compared with healthy tertiary follicles (Fig. 5C). Unexpectedly,
Tie-2 mRNA expression was found in earliest follicular stages (Fig. 6A) being localized
in the epithelial cells of primordial follicles. The expression of the
Tie-2 receptor was maintained in the granulosa cells of primary and
early secondary follicles (Fig. 6B). The expression of Tie-2 in the
granulosa began to decrease significantly in late secondary follicles
(Fig. 6D) and was barely detectable in the granulosa of tertiary
follicles (Fig. 6, C and D). In a few follicles, Tie-2 expression was
maintained in cumulus cells (Fig. 6C).
|
|
|
, inset),
in the granulosa cells of primary, early, and late secondary follicles
(Fig. 6E) as well as in the endothelium of the theca of tertiary
follicles (Fig. 6F).
Vascular development during folliculogenesis after VEGF trap
treatment
The treatment with VEGF trap effected the endothelial cell area,
cell proliferation, and the expression of angiogenic factors in the
theca of follicles. Furthermore, proliferation in the granulosa was
affected by the treatment.
In the theca of tertiary follicles, a significant 42% reduction of the
endothelial cell area was observed (Figs. 7 and 8A)
after treatment. Proliferation decreased significantly by 87% after
VEGF trap treatment in late secondary and tertiary follicles (Figs. 7 and 8B). Dual staining showed that in controls, 25–30% of
proliferating cells are of endothelial cell origin. The 87% decrease
indicates that in addition to endothelial cell proliferation,
proliferation of other cells are compromised by the treatment. The lack
of double-labeled cells with BrdU and CD31 revealed that
treatment inhibited endothelial cell proliferation (Fig. 7D).
|
|
). After treatment, a significant decrease of 42%
proliferation in the granulosa was observed in late secondary (Fig. 8C)
follicles. In tertiary follicles there was also an apparent decrease
after VEGF trap treatment but, because of a high SEM, this
was not significant.
The Flt receptor expression was severely inhibited by the VEGF
trap treatment (Fig. 9); it was
significantly reduced by 70% in late secondary, 87% in tertiary, and
73% in atretic follicles indicating a switch off of the Flt gene (Fig. 10).
|
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
The fact that proliferation was first seen in the theca in early secondary follicles in which endothelial cell staining was absent shows that the establishment of the theca compartment precedes angiogenesis. As the follicle develops, and more than four granulosa cell layers become established, staining of endothelial cells was detectable demonstrating the onset of theca vascularization. Proliferation in the thecal layer was markedly increased. Double labeling revealed that 25% of all proliferating cells are of endothelial cell origin. In tertiary follicles, theca proliferation continues characterized by further intense endothelial cell proliferation.
With regard to the molecular processes regulating these events, the observation that the expression of VEGF was highest in the granulosa, whereas its receptor Flt was expressed in theca endothelial cells of tertiary follicles suggests VEGF acts mainly in a paracrine manner to stimulate vascularization in the theca. The increase in both VEGF and Flt message in tertiary follicles could be explained by the fact that at that stage follicles undergo rapid growth and antrum formation. To keep pace with this growth, the vasculature has to expand intensively; therefore, VEGF up-regulation is required to induce this vascular development.
Tie-2 expression was localized in the theca endothelial cells of tertiary follicles. It has been shown that Tie-2 activated by its ligand Ang-1 promotes endothelial survival and stabilization of newly formed vessels (10, 16, 17). Thus, it is suggested that Tie-2 could be required to maintain the integrity of theca vessels. It was surprising to discover that the Tie-2 receptor (mRNA and protein), which is believed to be an endothelial-specific tyrosine kinase receptor, was located in the granulosa cells of primordial, primary, and early secondary follicles. The highest levels of Tie-2 mRNA were detected in granulosa cells of early follicular stages. A decrease was observed in late secondary follicles whereas in tertiary follicles the message disappeared in the granulosa except in the granulosa cells of the cumulus cells proximal to the oocyte. Most growth hormones use receptors of the tyrosine kinase family (18, 19, 20, 21). Potentially, this receptor may be used by other ligands than angiopoietins and may be involved in early follicular maturation. However, at this time the function of Tie-2 expression in early follicular stages is unknown.
The relationship between onset of atresia and changes in the vasculature is controversial because of difficulties in establishing the temporal relationship between the two processes. While in the human, a dense capillary network has been reported to remain in atretic follicles (1), decreased vascularity has been observed in atretic follicles of various other mammals (3, 22, 23). In this study, VEGF expression in the granulosa and Flt-1 and Tie-2 mRNA expression in the theca was significantly reduced in atretic follicles. Consistent with this observation, endothelial cell proliferation and the endothelial cell area were significantly reduced in atretic follicle in the marmoset. Thus, the mRNA for a principal angiogenic inducer and its receptor are down-regulated during follicular atresia, which may lead to a decrease in endothelial cell proliferation and survival.
Administration of VEGF antibodies or receptor-based VEGF trap treatment has proven to be a powerful tool by which the actions of VEGF on luteal angiogenesis and function can be investigated (12, 24, 25). This is the first study examining effects of VEGF inhibition by VEGF trap on follicular angiogenesis. After VEGF trap treatment, a reduction of proliferating cells in the theca was found in late secondary and tertiary follicles. The endothelial cell area was also decreased in tertiary follicles. The increased VEGF mRNA in normal tertiary follicles may indicate an increased requirement for VEGF at this stage in development and this, in turn may render them more vulnerable to VEGF inhibition by VEGF trap.
After treatment with VEGF trap, the proliferation in the theca was reduced by up to 87%. Dual staining in controls revealed that only 25–30% of the proliferating cells are of endothelial cell origin. Thus, reduction of cellular proliferation in the theca indicates that the treatment had a secondary inhibitory effect on proliferation of nonendothelial cells in the theca. Furthermore, proliferation of the granulosa of late secondary follicles was reduced after treatment. The likeliest explanation for these findings is that attenuation of the follicular vasculature affected the availability of growth factors, hormones or oxygen to the growing follicle. In addition, as VEGF promotes vascular permeability (26, 27) as well as angiogenesis, permeability may have been altered such that larger molecules like LH cannot penetrate the endothelium and reach the theca cells. In response to reduced exposure to LH their proliferation rate would be reduced. The collateral finding of decreased proliferation of thecal and granulosa cells suggests that this impairment of the development of the thecal vasculature exerts a secondary deleterious effect on follicular development. This hypothesis will be addressed in future studies designed to determine whether prolonged application of VEGF trap early in the cycle impedes the maturation of tertiary follicle and ovulation.
Finally, VEGF trap treatment had severe effects on the Flt receptor mRNA expression. The marked decrease in grain density after the treatment indicates that the Flt gene was switched off due to the lack of functional VEGF, implying that VEGF regulates its own receptor expression in follicles as observed in other tissues (28, 29, 30).
In conclusion, this study provides the first clear experimental support that VEGF, acting in a paracrine manner, is the major inducer of vascular development during follicular growth and that the ability to manipulate its activity using an antagonist such as VEGF TrapA40 could have important clinical applications. It is tempting to speculate that at least two ovarian disorders, i.e. the polycystic ovarian syndrome and the ovarian hyperstimulation syndrome associated with pathological angiogenesis, hypervascularity, and hyperpermeability, may be responsive to an anti-VEGF treatment and VEGF trap may be a possible competent therapeutic candidate.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
Received December 14, 2000.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
This article has been cited by other articles:
 |
|
 |
|
  C. Clapp, S. Thebault, M. C. Jeziorski, and G. Martinez De La Escalera Peptide Hormone Regulation of Angiogenesis Physiol Rev, October 1, 2009; 89(4): 1177 - 1215. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. Abramovich, A. R. Celin, F. Hernandez, M. Tesone, and F. Parborell Spatiotemporal analysis of the protein expression of angiogenic factors and their related receptors during folliculogenesis in rats with and without hormonal treatment Reproduction, February 1, 2009; 137(2): 309 - 320. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A Martelli, N Bernabo, P Berardinelli, V Russo, C Rinaldi, O Di Giacinto, A Mauro, and B Barboni Vascular supply as a discriminating factor for pig preantral follicle selection Reproduction, January 1, 2009; 137(1): 45 - 58. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D Herr, M Rodewald, H M Fraser, G Hack, R Konrad, R Kreienberg, and C Wulff Regulation of endothelial proliferation by the renin-angiotensin system in human umbilical vein endothelial cells Reproduction, July 1, 2008; 136(1): 125 - 130. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 F. Parborell, D. Abramovich, and M. Tesone Intrabursal Administration of the Antiangiopoietin 1 Antibody Produces a Delay in Rat Follicular Development Associated with an Increase in Ovarian Apoptosis Mediated by Changes in the Expression of BCL2 Related Genes Biol Reprod, March 1, 2008; 78(3): 506 - 513. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Rodewald, D. Herr, H.M. Fraser, G. Hack, R. Kreienberg, and C. Wulff Regulation of tight junction proteins occludin and claudin 5 in the primate ovary during the ovulatory cycle and after inhibition of vascular endothelial growth factor Mol. Hum. Reprod., November 1, 2007; 13(11): 781 - 789. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
P.D. Taylor, H. Wilson, S.G. Hillier, S.J. Wiegand, and H.M. Fraser Effects of inhibition of vascular endothelial growth factor at time of selection on follicular angiogenesis, expansion, development and atresia in the marmoset Mol. Hum. Reprod., October 1, 2007; 13(10): 729 - 736. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 W. S.N. Shim, I. A.W. Ho, and P. E.H. Wong Angiopoietin: A TIE(d) Balance in Tumor Angiogenesis Mol. Cancer Res., July 1, 2007; 5(7): 655 - 665. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R.S. Robinson, L.T. Nicklin, A.J. Hammond, D. Schams, M.G. Hunter, and G.E. Mann Fibroblast Growth Factor 2 Is More Dynamic than Vascular Endothelial Growth Factor A During the Follicle-Luteal Transition in the Cow Biol Reprod, July 1, 2007; 77(1): 28 - 36. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. L. Russell and R. L. Robker Molecular mechanisms of ovulation: co-ordination through the cumulus complex Hum. Reprod. Update, May 1, 2007; 13(3): 289 - 312. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. K. Trousdale, S. V. Pollak, J. Klein, L. Lobel, Y. Funahashi, N. Feirt, and J. W. Lustbader Single-Chain Bifunctional Vascular Endothelial Growth Factor (VEGF)-Follicle-Stimulating Hormone (FSH)-C-Terminal Peptide (CTP) Is Superior to the Combination Therapy of Recombinant VEGF plus FSH-CTP in Stimulating Angiogenesis during Ovarian Folliculogenesis Endocrinology, March 1, 2007; 148(3): 1296 - 1305. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. E. Roberts, L. K. Arbogast, C. I. Friedman, D. E. Cohn, P. T. Kaumaya, and D. R. Danforth Neutralization of Endogenous Vascular Endothelial Growth Factor Depletes Primordial Follicles in the Mouse Ovary Biol Reprod, February 1, 2007; 76(2): 218 - 223. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. Abramovich, F. Parborell, and M. Tesone Effect of a Vascular Endothelial Growth Factor (VEGF) Inhibitory Treatment on the Folliculogenesis and Ovarian Apoptosis in Gonadotropin-Treated Prepubertal Rats Biol Reprod, September 1, 2006; 75(3): 434 - 441. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A Martelli, P Berardinelli, V Russo, A Mauro, N Bernabo, L Gioia, M Mattioli, and B Barboni Spatio-temporal analysis of vascular endothelial growth factor expression and blood vessel remodelling in pig ovarian follicles during the periovulatory period J. Mol. Endocrinol., February 1, 2006; 36(1): 107 - 119. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
G. S. Nakhuda, R. C. Zimmermann, P. Bohlen, F. Liao, M. V. Sauer, and J. Kitajewski Inhibition of the Vascular Endothelial Cell (VE)-Specific Adhesion Molecule VE-Cadherin Blocks Gonadotropin-Dependent Folliculogenesis and Corpus Luteum Formation and Angiogenesis Endocrinology, March 1, 2005; 146(3): 1053 - 1059. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 H. M. Fraser, H. Wilson, J. S. Rudge, and S. J. Wiegand Single Injections of Vascular Endothelial Growth Factor Trap Block Ovulation in the Macaque and Produce a Prolonged, Dose-Related Suppression of Ovarian Function J. Clin. Endocrinol. Metab., February 1, 2005; 90(2): 1114 - 1122. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P D Taylor, S G Hillier, and H M Fraser Effects of GnRH antagonist treatment on follicular development and angiogenesis in the primate ovary J. Endocrinol., October 1, 2004; 183(1): 1 - 17. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 Y. F. Zhou, E. Stabile, J. Walker, M. Shou, R. Baffour, Z. Yu, D. Rott, G. D. Yancopoulos, J. S. Rudge, and S. E. Epstein Effects of gene delivery on collateral development in chronic hypoperfusion: Diverse effects of angiopoietin-1 versus vascular endothelial growth factor J. Am. Coll. Cardiol., August 18, 2004; 44(4): 897 - 903. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K.-G. Hayashi, T. J. Acosta, M. Tetsuka, B. Berisha, M. Matsui, D. Schams, M. Ohtani, and A. Miyamoto Involvement of Angiopoietin-Tie System in Bovine Follicular Development and Atresia: Messenger RNA Expression in Theca Interna and Effect on Steroid Secretion Biol Reprod, December 1, 2003; 69(6): 2078 - 2084. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Shimizu, J.-Y. Jiang, K. Iijima, K. Miyabayashi, Y. Ogawa, H. Sasada, and E. Sato Induction of Follicular Development by Direct Single Injection of Vascular Endothelial Growth Factor Gene Fragments into the Ovary of Miniature Gilts Biol Reprod, October 1, 2003; 69(4): 1388 - 1393. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
D. R. Danforth, L. K. Arbogast, S. Ghosh, A. Dickerman, R. Rofagha, and C. I. Friedman Vascular Endothelial Growth Factor Stimulates Preantral Follicle Growth in the Rat Ovary Biol Reprod, May 1, 2003; 68(5): 1736 - 1741. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
T. Shimizu, J.-Y. Jiang, H. Sasada, and E. Sato Changes of Messenger RNA Expression of Angiogenic Factors and Related Receptors During Follicular Development in Gilts Biol Reprod, December 1, 2002; 67(6): 1846 - 1852. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
R. C. Zimmermann, E. Xiao, P. Bohlen, and M. Ferin Administration of Antivascular Endothelial Growth Factor Receptor 2 Antibody in the Early Follicular Phase Delays Follicular Selection and Development in the Rhesus Monkey Endocrinology, July 1, 2002; 143(7): 2496 - 2502. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Wulff, H. Wilson, S. J. Wiegand, J. S. Rudge, and H. M. Fraser Prevention of Thecal Angiogenesis, Antral Follicular Growth, and Ovulation in the Primate by Treatment with Vascular Endothelial Growth Factor Trap R1R2 Endocrinology, July 1, 2002; 143(7): 2797 - 2807. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
N. R. Nayak and R. M. Brenner Vascular Proliferation and Vascular Endothelial Growth Factor Expression in the Rhesus Macaque Endometrium J. Clin. Endocrinol. Metab., April 1, 2002; 87(4): 1845 - 1855. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C. Wulff, H. Wilson, S. E. Dickson, S. J. Wiegand, and H. M. Fraser Hemochorial Placentation in the Primate: Expression of Vascular Endothelial Growth Factor, Angiopoietins, and Their Receptors Throughout Pregnancy Biol Reprod, March 1, 2002; 66(3): 802 - 812. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Endocrinology | Endocrine Reviews | J. Clin. End. & Metab. |
| Molecular Endocrinology | Recent Prog. Horm. Res. | All Endocrine Journals |
