Monash University Department of Obstetrics and Gynecology, Monash Medical Centre, 246 Clayton Road, Clayton, Victoria, 3168 Australia 1 Present address: School of Life Science and Technology, Victoria University of Technology, McKechnie Street, St Albans, Victoria 3021, Australia
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Abstract |
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Key words: angiogenesis/endometrium/endothelial cell/human/VEGF
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Introduction |
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It is clear that vascular endothelial cell growth factor (VEGF) is a major regulator of developmental angiogenesis, as well as of pathological angiogenesis associated with growth of solid tumours and with retinal and synovial disorders (Ferrara and DavisSmyth, 1997). Targeted disruption of even a single VEGF allele resulted in abnormal blood vessel formation and embryonic death in the mouse (Carmeliet et al., 1996
; Ferrara et al., 1996
). VEGF mRNA is upregulated in most malignant tumours (Ferrara and DavisSmyth, 1997
), and tumour growth is inhibited by strategies targeting tumour angiogenesis. Thus, treatment with neutralizing VEGF monoclonal antibodies in vivo, but not in vitro, inhibited tumour cell growth (Kim et al., 1993
), as did tumour cell expression of antisense VEGF (Saleh et al., 1996
). Direct evidence for VEGF as a mediator of normal angiogenesis was recently demonstrated when injection of a truncated, soluble flt-1 receptor prevented corpus luteum angiogenesis, maturation and progesterone release in a rat model, with secondary failure of endometrial development (Ferrara et al., 1998
).
VEGF is expressed in a wide range of cells and tissues (Ferrara and DavisSmyth, 1997), including rodent, primate and human endometrium (Gordon et al., 1995
; Smith, 1995
; Greb et al., 1997
; Torry and Torry, 1997
). There are five isoforms of the VEGF molecule, which are generated by alternative exon splicing of a single VEGF gene (Ferrara and DavisSmyth, 1997
). In the human endometrium, expression of four of these isoforms has been identified, VEGF121, VEGF145, VEGF165 and VEGF189 (Charnock-Jones et al., 1993
; Torry et al., 1996
), although the predominant isoforms expressed are the secreted forms, VEGF121 and VEGF165 (Smith, 1995
). Attempts have been made to relate VEGF expression in the human endometrium to stages of the menstrual cycle and, while a generalized picture is emerging, there is still some disagreement (Torry and Torry, 1997
; Rogers and Gargett, 1999
). From mRNA studies (in-situ hybridization and Northern blots), it appears that VEGF expression in whole endometrium is low during the proliferative phase, increases during the late secretory phase, and reaches a maximum at menses (Charnock-Jones et al., 1993
; Shifren et al., 1996
; Graubert et al., 1997
). However, there is no clear pattern of VEGF protein expression across the menstrual cycle, although expression is greater in glands than stroma (Li et al., 1994
; Shifren et al., 1996
; Lau et al., 1999
). Most studies of VEGF expression in the human endometrium have been descriptive, and only two had large enough sample numbers for meaningful statistical analysis (Graubert et al., 1997
; Lau et al., 1999
). The relative ability of glands and stroma to secrete VEGF in vitro has not been determined. Furthermore, no study has examined the relationship between VEGF expression across the menstrual cycle with indices or markers of endometrial angiogenesis.
The aims of the present study were to: (i) measure in-vitro production of VEGF by human endometrial tissues taken at four stages of the menstrual cycle; (ii) relate VEGF production by cultured endometrial cells to endothelial cell proliferation in the same tissue samples; and (iii) relate VEGF expression as measured by immunohistochemistry to endothelial cell proliferation in the same tissue, using sufficient sample numbers for each stage of the cycle to allow for meaningful statistical analysis.
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Materials and methods |
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A small portion of each biopsy was fixed for 4 h in 10% phosphate-buffered formalin (pH 7.4), for routine paraffin embedding. Serial sections (5 µm) were used for immunohistochemical analysis and staining with haematoxylin and eosin for dating. The remaining biopsy was collected into phenol red-free HEPES-buffered Dulbecco's minimum essential medium Ham's F-12 (DMEM/Ham's F-12) culture medium (Gibco BRL, Gaithersburg, MD, USA) containing 50 µg/ml gentamicin (Gibco BRL) and 0.5 µg/ml fungizone (Gibco BRL).
Cell isolation and culture
Explants (4x10 mg each, weighed after blotting to remove all fluid) of finely chopped tissue were cultured in 0.5 ml phenol red-free DMEM/Ham's F-12 medium containing 0.1% bovine serum albumin (BSA; Gibco BRL) for 24 h at 37°C in a humidified incubator containing 5% CO2 in air for production of conditioned medium (CM).
Glandular epithelial and stromal cells were separated from remaining chopped endometrial tissue using a modified method (Marsh et al., 1994). Briefly, tissue was digested with 45 U/ml collagenase type III (Worthington Biochemical Corporation, Freehold, NJ, USA) and 16 µg/ml deoxyribonuclease type I (Boehringer Mannheim GmbH, Mannheim, Germany) in Ca2+- and Mg2+-free phosphate-buffered saline (PBS; pH 7.4) for 30 min at 37°C and filtered sequentially through 77 µm and 10 µm nylon mesh (Henry Simon & Co., Stockport, UK) filters to collect gland fragments and stromal cells respectively. Erythrocytes were removed from stromal cell preparations by centrifugation over Ficoll-Paque (Pharmacia Biotech, Uppsala, Sweden). Gland fragments were recovered from the filters by backwashing. Washed stromal and gland cell preparations were resuspended in DMEM/Ham's F-12 medium containing 10% charcoal-treated fetal calf serum (FCS; PA Biologicals, Sydney, Australia), gentamicin (50 µg/ml), fungizone (0.5 µg/ml) and 2 mM glutamine (Sigma Chemical Co., St Louis, MO, USA) and seeded in quadruplicate in 24-well plates at 3x105 cells/well and 410x103 glands/well respectively. The purity of the cultures was >95%, confirmed by immunohistochemical staining for cytokeratin using mouse anti-human cytokeratin antibody (Clone MNF116; Dako, Cambridge, UK) and vimentin with mouse anti-vimentin antibody (Clone V9; Zymed, San Francisco, CA, USA), to detect epithelial and stromal cells respectively. Glandular epithelial cells were cultured for 4 days and stromal cells until confluent (26 days) at 37°C in 5% CO2, with medium changes every 48 h.
For collection of CM, cultures were rinsed and the medium changed to 0.5 ml/well serum-free DMEM/Ham's F-12 medium containing 0.1% BSA, antibiotics and glutamine, and cultures were incubated for 24 h. The CM was harvested, centrifuged at 7500 g for 1 min and the supernatant collected and stored at 80°C.
Cell counts
Cells numbers per well were measured after CM collection using the CellTitre96 MTS tetrazolium-based bioassay (Promega, Madison, WI, USA), in which the absorbance of the formazan product is directly proportional to the number of viable cells. Briefly, CM was replaced with 250 µl of standard DMEM/Ham's F-12/10% charcoal-treated FCS medium, 50 µl 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulphophenyl)-2H-tetrazolium (MTS) solution was added, and the cells incubated for 75 min at 37°C. Two 120 µl aliquots were transferred to wells of a 96-well plate, the absorbance measured at 490 nm using a plate reader (Emax, Molecular Devices, Sunnyvale, CA, USA) and analysed with the associated software, SOFTmax, version 2.32. A standard curve was produced for each assay by seeding serial dilutions of passaged endometrial stromal cells (2x104) in 100 µl volumes of DMEM/Ham's F-12/10% charcoal-treated FCS into wells of a 96-well plate. The cells were allowed to attach for 90 min before addition of 20 µl of MTS reagent, incubated for a further 75 min, and the absorbance measured. Gland cell standard curves using primary cultured glandular epithelial cells gave slopes matching those of stromal cells, indicating that the two cell types metabolize the MTS at a similar rate. Values ranged from 7x103 to 67x103/well for glandular epithelial cell cultures, and from 20x103 to 100x103/well for stromal cell cultures, depending on the degree of confluence. In some cultures, cell numbers were estimated using the established relationship between cell number/well and level of confluence.
Enzyme-linked immunosorbent assay (ELISA) for measurement of VEGF in conditioned medium
Purified recombinant mouse VEGF164 (a gift from Dr S.Stacker, Ludwig Institute, Melbourne, Australia) was used to immunize rabbits for production of polyclonal anti-VEGF antibodies by standard methods. The titre and specificity of the anti-VEGF response was evaluated by inhibition of VEGF-mediated proliferation of VEGFR2/EpoR bioassay cells and Western blotting respectively. The IgG fraction of the VEGF antiserum was obtained by affinity chromatography using a protein Aagarose column (Pharmacia Biotech). Biotinylated anti-VEGF IgG was obtained by conjugation with sulpho-biotin-X-NHS (Calbiochem-Novabiochem GmbH, Bad Soden, Germany) for 1 h at 24°C using a 25 molar excess of biotin to IgG. ELISA were performed in Maxisorb 96-well plates (Nunc, Roskilde, Denmark), which were coated with anti-VEGF antibody (2.5 µg/ml) in 50 mmol/l carbonate buffer, pH 9.6 for 20 h at 4°C and blocked with 3% BSA in PBS (0.5 mol/l NaCl, 0.01 mol/l phosphate) overnight at 4°C. Plates were washed in PBS/Tween (0.1% Tween 20), which was also used for subsequent washing steps between addition of reagents. Standards, rhVEGF165 (PeproTech, Rocky Hill, NJ, USA) (0.1610 ng/ml) and CM samples, neat and diluted 1:1, were then added and incubated for 2 h at room temperature with gentle mixing. Bound VEGF was detected with biotinylated anti-VEGF (1.25 µg/ml in 3% BSA/PBS; incubation 2 h) and streptavidinhorseradish peroxidase (HRP) (Zymed, diluted 1/10 000; incubation 30 min) and the substrate was 3,3',5,5'-tetramethylbenzidine dihydrochloride (Sigma) (0.1 mg/ml in 0.05 M phosphate citrate buffer, pH 5.0 containing 0.006% vol/vol H2O2). After a 7 min incubation, the reaction was terminated with 1 M HCl and absorbance measured at 450 nm using a microplate reader. All CM samples from a given patient were analysed on the one plate. The standard curve was fitted by a four-parameter, non-linear regression-fitting program (SOFTMax version 2.32) and was linear between 0.5 and 10 ng/ml VEGF. Intra- and interassay variability were <11% and <15% respectively. The technique was sensitive to 0.30 ng/ml. Serially diluted CM samples produced curves parallel with the standard curve. This ELISA detected both mouse VEGF164 and human VEGF165 and VEGF121, but no VEGF was detected when pre-immune rabbit serum was used to coat the plates.
Immunohistochemistry
Proliferating endothelial cells were detected by a standard double immunohistochemistry protocol using mouse anti-rat antibody to proliferating cell nuclear antigen (PCNA) (clone PC10; Novacastra, Newcastle, UK) and a mouse monoclonal antibody against the cluster determination (CD)34 antigen (clone QBEND/10; Serotec, Oxford, UK), an endothelial cell marker, according to a previously published protocol (Goodger (MacPherson) and Rogers, 1994). All other reagents were from Zymed. A sequential protocol was used with the anti-PCNA as the first primary antibody. Both primary antibodies were applied for 1 h at 37°C, followed by incubation with the same biotinylated secondary antibody (rabbit anti-mouse immunoglobulin). A streptavidinHRP conjugate with AEC chromogen (red) was used for PCNA staining, and a streptavidinalkaline phosphatase conjugate with AP-blue chromogen was used for CD34 staining. Endogenous peroxidase was quenched with 3% H2O2 in 50% methanol. Positive and negative control sections were included in each staining run as described (Goodger (MacPherson) and Rogers, 1994). At least 100 microvessel profiles were counted in sequential fields scanned at x400 magnification from the surface epithelium through the full depth of the section. The number of these vessels containing proliferating endothelial cells was also determined and the percentage of microvessels containing proliferating endothelial cells calculated.
VEGF protein was detected using a rabbit anti-human polyclonal anti-VEGF antibody (Sc-152; Santa Cruz Biotechnology Inc, Santa Cruz, CA, USA) as previously described (Lau et al., 1999). Briefly, sections were boiled for 10 min in 10 mmol/l sodium citrate buffer, pH 6 for antigen retrieval, endogenous peroxidase quenched, and sections blocked with 10% normal goat serum. The primary antibody (2 µg/ml) was applied for 2 h at 37°C, followed by sequential incubations with biotinylated secondary antibody, streptavidinHRP conjugate and AEC chromogen. Positive and negative controls were included in every staining run as described (Lau et al., 1999
). Several images per section were captured using a digital video camera (Fujix, Fuji, Tokyo, Japan) and the same light intensity settings for all sections. For each captured image, the staining intensity of the entire glandular area and the stromal area was measured by Analytical Imaging StationTM software (AIS, Image Research Inc., Ontario, Canada) and corrected for background staining. The average staining intensity of the glandular and stromal regions of several captured images was then calculated for each sample. Similarly, for each captured image, the total area occupied by glands and by stroma was also recorded and the average proportional area for each sample calculated. A staining intensity index (SI) was then obtained for the glandular and stromal compartments of each sample by multiplying the average staining intensity by the average proportion of stained area. Thus, the SI represents the total glandular and total stromal VEGF production for each sample as measured by immunostaining. Interassay variation was <19% (Lau et al., 1999
).
Statistical analysis
Analyses were performed using SPSS, version 6.1.3 (SPSS, Australasia, North Sydney, Australia). The Levene homogeneity test (Snedecor and Cochran, 1989) was used to measure variance and the KolmogorovSmirnovLilliefors/ShapiroWilk test was used to test for normality and to determine whether parametric or non-parametric tests were to be used for further analyses. The KruksalWallis one-way analysis of variance (ANOVA) was used to examine for differences between stages of the menstrual cycle and the MannWhitney U-test was used to confirm a difference between the early proliferative stage and each of the other stages. The Wilcoxon matched-pairs signed-rank test was used for comparing paired data (glands versus stroma), correlations were performed using least squares regression analysis, and Spearman's correlation coefficients (RS) were determined. Some of the data were also transformed into natural logarithms and compared by one-way ANOVA. Results were considered statistically significant when P < 0.05.
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Results |
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Discussion |
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It is well known that VEGF has functions other than inducing endothelial cell proliferation. In the endometrium, VEGF may mediate increases in vascular permeability (Dvorak et al., 1995; Torry et al., 1996
), phenotypic changes observed in microvessel endothelial cells (Roberts et al., 1992
), or provide trophic signals to the implanting embryo (Hornung et al., 1998
).
VEGF may act in a paracrine/autocrine manner and therefore its secretion would be localized. The observation of small, discrete patches of intense VEGF immunoreactivity associated with some blood vessels and also in a population of stromal cells in most of the samples examined supports this hypothesis (Figure 3). These foci of VEGF immunoreactivity were not included in the calculation of the SI, since their quantification requires methods other than described in this study, nor could they be examined in cultured endometrial cells. Localized VEGF immunostaining in perivascular stromal cells and endothelial cells has previously been observed in endometrium (Li et al., 1994
), while focal endothelial cell proliferation has been reported for pathological angiogenesis (Walsh et al., 1998
). The source of focally produced VEGF in human endometrium may be pericytes (Nomura et al., 1995
; Reynolds and Redmer, 1998
), vascular smooth muscle cells (Gu and Adair, 1997
; Abramovitch et al., 1998
) in the spiral arterioles, endothelial cells (Namiki et al., 1995
; Nomura et al., 1995
), or adjacent stromal cells.
It is likely that endometrial angiogenesis is regulated by a complex interaction between angiogenesis promoters and inhibitors (Folkman and Shing, 1992). In this study, a correlation between endothelial cell proliferation and bulk changes in glandular and stromal VEGF was not observed, suggesting that factors other than VEGF may also be involved. Synergism between VEGF and its receptors, VEGFR-1 (flt-1) and VEGFR-2 (KDR), and basic fibroblast growth factor (bFGF), all of which are expressed in human endometrium (Ferriani et al., 1993
), may have a role in endometrial angiogenesis. In addition, changing levels of endogenous angiogenesis inhibitors across the menstrual cycle, such as thrombospondin-1 (IruelaArispe et al., 1996
), may influence endothelial cell proliferation.
VEGF secretion and endometrial angiogenesis detected by PCNA immunoreactivity may occur sequentially due to transient or pulsatile VEGF secretion. VEGF, which acts during G0/G1 may no longer be detectable when endothelial cells become PCNA-positive during late G1 and S phase. Samples shown as outliers or extremes in Figures 1, 2 and 4 may represent transient peaks of VEGF secretion or angiogenic activity. Both hypoxia and oestrogen can rapidly upregulate VEGF within an hour, with return to baseline values within 24 hours of removal of the stimulus (Cullinan-Bove and Koos, 1993
; Gu and Adair, 1997
; Yasuda et al., 1998
).
A second finding in the present study was that in vivo, VEGF was significantly elevated during the early proliferative phase (Figure 2) of the menstrual cycle. However, no relationship was observed between in-vitro VEGF production by cultured endometrial explant, glandular epithelial or stromal cells and the cycle stage when the tissue was collected (Figure 1
). Two other studies, using large sample numbers, have also demonstrated a trend to higher stromal VEGF concentrations during the proliferative stages (Li et al., 1994
; Lau et al., 1999
), although one study of unknown sample number demonstrated increasing VEGF mRNA and protein from early proliferative to late secretory endometrium (Shifren et al., 1996
). Inconsistencies in patterns of VEGF expression in glands and stroma between the various stages of the menstrual cycle may relate to the descriptive nature of some of these studies, as well as the low sample numbers used, considering the degree of between-subject variability. Only two studies were sufficiently large or quantitative to report on variability between individual subjects (Torry et al., 1996
; Lau et al., 1999
). In this study, we have used moderate numbers of samples to allow comparison between four main stages of the menstrual cycle, although menstrual samples were not examined because there was insufficient tissue for culture and immunohistochemical studies. Elevated VEGF mRNA observed in secretory and menstrual stages (Charnock-Jones et al., 1993
; Shifren et al., 1996
; Torry et al., 1996
; Graubert et al., 1997
) probably reflects glandular expression, since considerably more VEGF has been reported in glandular epithelial compared with stromal cells (Table I
) (Li et al., 1994
; Shifren et al., 1996
; Torry et al., 1996
; Lau et al., 1999
). Since the bulk of endometrial VEGF is glandular, and 80% of this VEGF is secreted from the luminal surface (Hornung et al., 1998
), it is unlikely that most of the VEGF produced in the glands has a role in endometrial angiogenesis.
A third observation of the present study was the large degree of variation between subjects for each of the parameters measured. Our laboratory has consistently reported high variability for endothelial cell proliferation indices, and has recently demonstrated that 84% of this variability was attributable to between-sample variation, while only 16% was due to sampling variation within the same uterus (Rogers et al., 1998). Difficulties in obtaining consistency between staining runs may also contribute to the particularly high variability observed in VEGF staining indices, although captured images were always analysed relative to the same positive control section. Nevertheless, immunoreactive VEGF was significantly elevated during the early proliferative phase.
In summary, our results suggest that ovarian steroids, which control the characteristic growth cycle of human endometrium, are not the main regulators of the majority of endometrial VEGF production, or of angiogenesis. While oestrogen and progesterone have been shown to increase VEGF protein secretion in cultured stromal cells, these effects have been modest (Shifren et al., 1996; Huang et al., 1998
). It is uncertain whether the effect of oestrogen in these studies is direct or indirect, since the VEGF gene does not contain oestrogen or progesterone response elements in its promoter region, although there are several half-palindromes for oestrogen (Tischer et al., 1991
; Cullinan-Bove and Koos, 1993
). It is possible that other factors such as hypoxia or locally produced cytokines also play a role in regulating angiogenesis during the menstrual cycle. Our results have demonstrated no obvious correlation between overall VEGF production and endothelial cell proliferation for individual endometrial samples, despite the observed increases in stromal VEGF immunostaining and percentage of proliferating vessels (though not statistically significant) during the early proliferative stage, when angiogenesis undoubtedly occurs. Other approaches are still required to provide further understanding of the regulation of endometrial angiogenesis.
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Acknowledgments |
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Notes |
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References |
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Submitted on December 21, 1998; accepted on April 30, 1999.