1 Departments of Obstetrics, Gynecology and Reproductive Sciences, and Physiology, 2 Department of Anatomy and Neurobiology, Center for Studies in Reproduction; University of Maryland School of Medicine, Baltimore, MD 21201 and 3 Department of Physiological Sciences, Eastern Virginia Medical School, Norfolk, VA 23507, USA
4 To whom correspondence should be addressed at: Department of Obstetrics, Gynecology and Reproductive Sciences, University of Maryland School of Medicine, Bressler Research Laboratories 11-019, 655 West Baltimore Street, Baltimore, MD 21201, USA. e-mail ealbrech{at}umaryland.edu
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Abstract |
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Key words: angiogenesis/endometrium/estrogen/glandular/human/
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Introduction |
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Although comparable in vivo studies designed to show a cause and effect relationship between steroid hormones and endometrial angiogenesis have not been conducted for ethical reasons in humans, VEGF mRNA and protein are expressed by human endometrial glandular epithelial and stromal cells (Charnock-Jones et al., 1993; Torry et al., 1996
; Shifren et al., 1996
; Gargett and Rogers, 2001
; Möller et al., 2001
), and estrogen stimulated VEGF expression in cultures of human endometrial cells (Charnock-Jones et al., 1993
; Shifren et al., 1996
; Huang et al., 1998
), through a functional variant estrogen response element (Mueller et al., 2000
). It is well established that VEGF increases angiogenic responses, e.g. cell proliferation, in cultures of purified human microvascular endothelial cells (Ferrara et al., 1992
). However, because microvascular endothelial cells express the estrogen receptor (Critchley et al., 2001
) and estrogen stimulates endothelial cell proliferation (Morales et al., 1995
), estrogen may also promote new blood vessel development by acting directly on microvascular endothelial cells. Thus, it is not clear in the human endometrium whether estrogen promotes angiogenesis directly, and/or indirectly via expression of angiogenic factors by particular endometrial cells. Therefore, in the present study, we developed a co-culture of human endometrial cells and microvascular endothelial cells, as an in vitro model of the morphological and functional interaction that exists between these cell types in vivo, to determine whether the regulatory role shown for estrogen on endometrial VEGF formation and angiogenesis in vivo in the non-human primate would be demonstrable in vitro in the human.
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Materials and methods |
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Endometrial specimens were either immediately frozen in liquid nitrogen for VEGF mRNA analysis by RTPCR, fixed in 10% neutral-buffered formalin for 24 h and embedded in paraffin for VEGF immunocytochemistry, or enzyme-dispersed and separated into enriched glandular epithelial and stromal cell populations by a modification of the method of Osteen et al. (1989). Endometrial tissue was minced into 23 mm pieces in Dulbeccos modified Eagles medium (DMEM)/F12 (Sigma Chemical Co., St Louis, MO) and digested with 5 ml of a solution containing 4 mg/ml collagenase type 4 (Worthington, Biochemical Corp., Lakewood, NJ), 1 mg/ml hyaluronidase (Sigma), 1 mg/ml protease (Sigma), 200 U/ml DNase I type II (Sigma) and 2% chicken serum (Sigma) in magnesium-free Hanks balanced salt solution (HBSS; Gibco, Invitrogen Corp., Grand Island, NY) in an orbital shaking water bath at 37°C for 4560 min. The resultant cell suspension was then washed in calcium- and magnesium-free HBSS and filtered through 85 µm nylon mesh (Small Parts Inc., Miami Lakes, FL). The retentate containing glandular epithelial fragments was washed from the filter with HBSS, resuspended in DMEM/F12 and stored overnight at 4°C. The stromal cell-enriched filtrate was treated with DNase for 10 min, layered over a 66% Percoll (Sigma) gradient to remove erythrocytes, washed in DMEM/F12, DNase-treated and filtered through 20 µm mesh. Cells were then washed in DMEM/F12 and resuspended in DMEM/F12 containing 4% fetal bovine serum (FBS; Hy Clone Laboratories Inc, Logan, UT). Glandular epithelial fragments were digested with collagenasehyaluronidaseproteaseDNase in HBSS at 37°C for 15 min, washed in HBSS, and filtered through 20 µm mesh. The retentate containing gland fragments was digested further for 3045 min, then washed and resuspended in DMEM/F12 containing 4% FBS. Isolated glandular epithelial and stromal endometrial cells (24 x 105 cells/ml) were plated on 15 mm culture wells (Nalge Nunc International, Naperville, IL) in DMEM/F12 supplemented with 4% FBS and incubated at 37°C in a humidified atmosphere of 95% air:5% CO2 for 7 days, with medium change every 2 days.
Culture of human myometrial microvascular endothelial cells
Cryopreserved human uterine myometrial microvascular endothelial cells (HMMECs; UtMVEC-Myo, Clonetics, Walkersville, MD) were obtained at passage 3 and plated on T 75 flasks at 5 x 103 cells/cm2 in a specialized microvascular endothelial cell growth medium (EGM-2-MV, Clonetics) containing 5% FBS, hydrocortisone, ascorbic acid, gentamicin, amphotericin-B, human epidermal growth factor (hEGF), human fibroblast growth factor (hFGF) and human insulin-like growth factor-1 (hIGF-1), but without hVEGF. The attached HMMECs were grown for a pre-confluent period of 57 days at 37°C in a humidified atmosphere of 95% air:5% CO2 with medium change every 2 days, trypsinized and either cryopreserved or subcultured on T 75 flasks. In subsequent experiments, HMMECs from passages 410 were grown for 56 days in EGM-2-MV with medium change to endothelial cell basal medium (EBM, Clonetics) supplemented with 4% FBS for 24 h.
Co-culture of endometrial cells and HMMECs: endothelial cell tube formation
Media from glandular epithelial and stromal cell cultures were replaced after 7 days with 0.5 ml of EBM supplemented with 4% FBS and cells incubated for 24 h in the absence or presence of E2 (106 or 108 mol/l, Sigma). Cell culture inserts (0.2 µm pore size, Anopore, Nalge Nunc International) were coated with 0.15 ml of Matrigel (growth factor reduced without phenol red, BD Biosciences, Bedford, MA) per insert and allowed to polymerize at 37°C for 1 h. HMMECs (25 x 103 cells/0.5 ml) in EBM supplemented with 4% FBS were then added to Matrigel-coated inserts and co-incubated in triplicate in wells with endometrial cells for 24 h at 37°C in the absence or presence of E2 and 50 ng/ml VEGF. Incubation of human endometrial cells with either 106 or 108 mol/l E2 yielded similar results for HMMEC tube formation in our laboratory. Therefore, in the present study with enriched glandular epithelial and stromal cell incubations, 106 mol/l E2 was employed.
HMMEC tube formation was assessed by image analysis in 23 representative fields of view in each culture well. HMMEC images were captured at a final linear magnification of 40x using an Olympus Inverted Research Microscope (Olympus, Tokyo, Japan) coupled to a Polaroid Digital Microscope camera (Polaroid, Cambridge, MA). Images were prepared in Adobe Photoshop 5.0 and exported to an image analysis software package (IP Lab Scientific Image Processing, Scanalytics, Fairfax, VA) for identification of endothelial cell tube-like networks and quantification of total endothelial tube and endothelial cell area. The mean percentage tube formation from each treatment group was calculated as the ratio of the endothelial tube area/total endothelial cell area x 100.
Endothelial cell proliferation assay
The Cell Titer 96 Aqueous One Solution Cell Proliferation Assay (Promega, Madison, WI) was used to confirm the responsiveness of HMMECs to the mitogen VEGF. HMMECs (5 x 103 cells/0.1 ml) were plated in triplicate in 96-well plates in EBM supplemented with 4% FBS and incubated at 37°C in a humidified atmosphere of 95% air:5% CO2 for 24 h. After 24 h, medium was replaced with EBM supplemented with 4% FBS in the absence or presence of increasing concentrations (0.110 ng/ml) of hVEGF (BioSource Int., Camarillo, CA) and/or 1 µg/ml anti-hVEGF antibody (R&D Systems, Inc., Minneapolis, MN). HMMECs were incubated at 37°C in a humidified atmosphere of 95% air:5% CO2 for 4 days with medium change after 2 days. At the end of the 4-day period, medium was replaced with EBM supplemented with 4% FBS and 20 µl of One Solution Reagent and incubated for 2 h at 37°C. Absorbance was measured at 490 nm using a plate reader. The One Solution Reagent contains MTS tetrazolium (3-[4,5-dimethylthiazol-2-yl]-5-[3-carboxymethoxyphenyl]-2-[4-sulfophenyl]-2H-tetrazolium), which is bioreduced by cells into a formazan product. The absorbance of the formazan product is directly proportional to the number of cells.
Enzyme-linked immunoassay (ELISA) of VEGF protein
VEGF protein levels in the conditioned medium at the end of co-culture of glandular epithelial or stromal cells and HMMECs were determined by ELISA using a commercially available kit (R&D Systems), according to the manufacturers instructions. Briefly, 0.20 ml of conditioned medium or recombinant human VEGF standard (01000 pg/ml) were incubated for 2 h at room temperature in wells coated with monoclonal mouse anti-human VEGF capture antibody, washed and incubated for 2 h with 0.2 ml of polyclonal goat anti-human VEGF antibody conjugated to horseradish peroxidase. Wells were then washed and samples incubated for 20 min with 0.2 ml of a 1:1 mixture of colour reagent A (H2O2) and colour reagent B (tetramethylbenzidine) and the reaction stopped by addition of 0.05 ml of 2 mol/l H2SO4. Optical density was then determined at 450 nm (with wavelength correction at 570 nm) using a microplate reader. The limit of sensitivity of the assay was <5 pg/ml, and intra- and inter-assay coefficients of variation were each <10%.
Immunocytochemistry
Endometrium. Immunocytochemistry of VEGF was performed as described previously (Hildebrandt et al., 2001). Paraffin blocks of uterine tissue were serially sectioned (4 µm), deparaffinized and rehydrated in graded alcohols. Tissue sections were boiled in 0.01 mol/l Na citrate for 10 min, incubated in 1% H2O2, and blocked in 10% normal goat serum (Protein Block Serum, Dako Corp, Carpinteria, CA). Tissues were incubated overnight at 4°C with goat anti-human primary antibody to VEGF (AF-293-NA, diluted 1:25 in 5% goat serum, specific for the 121, 165 and 189 splice variants; R&D Systems). Following incubation with biotinylated anti-goat imunoglobulin (Vector Laboratories, Inc., Burlingame, CA), sections were immersed in an avidinbiotin complex solution (Elite Vectastain ABC Kit, Vector Laboratories, Inc.), and incubated with 3,3'-diaminobenzidine (DAB; 0.2 mg/ml, Sigma) to produce a brown reaction product. Negative controls included omission of the primary antibody or pre-absorption of primary antibody with 10-fold excess of human recombinant VEGF peptide (R&D Systems). Sections were counterstained with Harris haematoxylin.
Cell cultures. Glandular epithelial and stromal cells (4 x 105 cells/0.9 ml) were plated on Lab-Tek II chamber slides (Nalge Nunc International) in DMEM/F12 supplemented with 4% FBS and incubated at 37°C in a humidified atmosphere of 95% air:5% CO2 for 7 days. HMMECs (25 x 103 cells/0.9 ml) were plated on chamber slides in EBM supplemented with 4% FBS and incubated for 12 days. Cells were then fixed for 90 min in 10% buffered formalin, rinsed in three changes of 1x phosphate-buffered saline (PBS), incubated in H2O2 for 10 min to inhibit endogenous peroxidase, and blocked with 10% normal goat serum for 1 h. Glandular epithelial cells, stromal cells or HMMECs were incubated overnight at 4°C in a humidified chamber with mono- or polyclonal antibodies to either cytokeratin 7 (Cytokeratin-7-Monoclonal DAKO Clone OV-TL), vimentin (Vimentin-Monoclonal DAKO Clone VIM 3B4) (1:4000 and 1:6000, respectively, DAKO), flt-1 (C-17, 1:500) or KDR/flk-1 (C-1158, 1:750, both from Santa Cruz Biotechnology Inc., Santa Cruz, CA). Cells were incubated for 1 h at room temperature with biotinylated anti-mouse or anti-rabbit immunoglobulins (Vector), followed by avidinbiotinperoxidase complex (ABC Elite, Vector) for 1 h at room temperature using DAB as chromagen. Cells were lightly counterstained with Harris haematoxylin for 30 s, clarified in acid alcohol and blued using lithium carbonate. Slides were dehydrated in graded concentrations of ethanol and cleared in xylene. Negative controls included omission of the primary antibody, substitution of goat IgG for primary antibody, and pre-absorption of primary antibody with 10-fold excess of control peptide for flt-1 (Santa Cruz) or F(ab')2 fragment-specific IgG (Jackson ImmunoResearch Labs Inc., West Grove, PA).
Competitive RTPCR of VEGF mRNA
RNA isolation and oligonucleotide primers. Total RNA was isolated from endometrium by guanidine isothiocyanatecaesium chloride and quantified by UV absorption spectrophotometry to permit normalization of VEGF mRNA levels. Oligonucleotide primers were synthesized by Invitrogen Life Technologies, Inc. and based on the hVEGF cDNA sequence (Tischer et al., 1991), as detailed previously (Hildebrandt et al., 2001
).
Competitive reference standard (CRS). Homologous RNA fragments containing the same primer-binding regions but shortened internal sequence with respect to the target RNA for VEGF were prepared as described previously (Riedy et al., 1995; Babischkin et al., 1997
). Reverse transcription of total RNA (0.53.0 µg) from baboon placenta was performed at 42°C for 60 min in a reaction containing 1 mmol/l each of dNTPs (Invitrogen Life Technologies, Inc.), 200 U of SUPERSCRIPT RNase H-reverse transcriptase (RT) or Molony murine leukaemia virus RT (Invitrogen Life Technologies, Inc.), 1x RT buffer, 40 U of RNAguard (Amersham Pharmacia Biotech, Piscataway, NJ) and 250 ng of random primers (Invitrogen Life Technologies, Inc). A 5 µl aliquot of the RT reaction was added to separate PCRs containing 0.2 mmol/l dNTPs, 1.25 U of cloned Thermus aquaticus DNA polymerase (Amplitaq, Perkin-Elmer Corp/Cetus, Norwalk, CT), 1x PCR buffer and 10 pmol of paired primers to generate cDNA template for VEGF. PCR was performed in a programmable thermal cycler (MJ Research, Inc., Cambridge, MA) for 25 sequential cycles. An aliquot of each reaction was subjected to 2.0% agarose gel electrophoresis, amplified products gel purified, and the CRS synthesized using the MEGAscript T7 transcription kit (Ambion, Inc., Austin, TX). The cDNA templates were extracted with chloroform:isoamyl alcohol, and aliquots of CRS quantitated spectrophotometrically.
RTPCR assay. VEGF was quantified by competitive RTPCR assay (Babischkin et al., 1997; Niklaus et al., 2002
). A constant amount of RNA (10 ng) was added to an RT mixture containing serial dilutions of VEGF-CRS (253 attomol). In all experiments, the presence of possible pseudogene or genomic DNA contamination was checked by control reactions in which either the RT enzyme or RNA was omitted.
A 5 µl aliquot of the RT mixture was added to separate PCR mixtures containing 10 pmol of the primers for VEGF. Endometrial samples were amplified for 34 sequential cycles, and PCR products gel fractionated, visualized with ethidium bromide and photographed using type 665 positive/negative film (Polaroid Crop, Cambridge, MA).
Negatives were scanned using a Gel Doc 1000 imaging system and Multi-Analyst software program (Bio-Rad Laboratories, Hercules, CA). The intensity of amplified target and CRS cDNA products was represented as the relative area under each product band. The logarithm (log) of the ratio of CRS to target area was plotted as a function of the log concentration of VEGF added to each PCR. The concentration of VEGF target mRNA was determined where the ratio of the log of CRS and target area was equal to 0 (i.e. the equivalence point).
Statistical analysis
Data were expressed as the means ± SEM and analysed either by Students t-test or by ANOVA with post hoc comparisons of means by NewmanKeuls multiple comparisons test.
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Results |
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Endometrial VEGF immunocytochemistry
VEGF protein expression was abundant in glandular epithelial cells of the human endometrium (Figure 2A). VEGF protein was also present in the stroma, although the intensity appeared lower than in glandular epithelial cells. Specificity of VEGF immunocytochemistry was evident by the absence of staining when primary antibody was pre-absorbed with recombinant VEGF (Figure 2B).
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Discussion |
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Although E2 alone did not significantly increase myometrial microvessel endothelial cell tube formation in the present study, estrogen receptors are expressed in endothelial cells (Critchley et al., 2001), and estrogen promoted proliferation of and tube formation by human umbilical vein endothelial cells in culture (Morales et al., 1995
). These apparently disparate results may reflect the very different endothelial cell types and cell culture conditions that were employed in the two studies. Under certain conditions, VEGF may be expressed by endothelial cells (Concina et al., 2000
), or by neutrophils focally in association with microvessel endothelial cells (Gargett and Rogers, 2001
). Therefore, it has been proposed that cells within the vasculature are in vivo sources of angiogenic factors for non-sprouting angiogenesis, i.e. intussusception and elongation, within the endometrium (Gargett and Rogers, 2001
).
As shown in the present study and previously by others (Li et al., 1994; Lau et al., 1998
; Charnock-Jones et al., 2000
), VEGF mRNA levels were similar in endometrial tissue collected during the proliferative and secretory phases of the human menstrual cycle, despite cyclical surges in E2. Thus, it has been suggested that estrogen has no role in endometrial VEGF expression or angiogenesis (Smith, 1998
; Gargett and Rogers, 2001
), although the protein levels should be taken into account because VEGF mRNA stability may be altered in response to estrogen. We suggest that the levels of estrogen, although low, immediately preceding and following the late proliferativemidcycle surge in E2, are nevertheless sufficient and necessary to maintain endometrial VEGF expression to progressively promote angiogenesis. Thus, it may only be when estrogen is very low, e.g. after ovariectomy of non-human primates (Niklaus et al., 2002
, 2003; Albrecht et al., 2003
), early in the proliferative phase or after removal of endometrial tissue from in situ for cell culture as in the present study, that VEGF synthesis declines and endometrial cells become responsive to exogenous estrogen. The less marked, but nevertheless significant, increase in HMMEC tube formation elicited by human glandular epithelial cells alone in cultures of the present study may reflect continued VEGF synthesis by endometrial cells previously upregulated by the estrogen environment in situ.
In the present study, we focused on the regulatory role of estrogen, because progesterone was not effective in stimulating endometrial VEGF expression or microvessel permeability in baboons (Albrecht et al., 2003). However, progesterone increased VEGF mRNA expression in vivo in the rat uterus (Cullinan-Bove and Koos, 1993
) and in vitro in human endometrial cells (Shifren et al., 1996
). Thus, additional study with the co-culture of human endometrial and microvascular endothelial cells is needed to determine the potential regulatory role of progesterone in angiogenesis in this system.
In summary, we established a co-culture of human endometrial cells and microvascular endothelial cells to determine whether the regulatory role shown for estrogen on endometrial angiogenesis in vivo in the non-human primate is operable in vitro in the human endometrium. The results of this study show that E2 significantly promoted tube formation by microvascular endothelial cells co-cultured with human glandular epithelial cells. On the basis of the results of the current study and of other in vitro and in vivo studies, we suggest that estrogen, by regulating expression and secretion of angiogenic factors such as VEGF by glandular epithelial cells of the endometrium, regulates endometrial microvascular endothelial cell tube formation and thus angiogenesis.
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Acknowledgements |
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References |
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Submitted on November 7, 2002; resubmitted on March 13, 2003; accepted on June 30, 2003.