Abstract

Understanding of ectopic implantation within the Fallopian tube (FT) is limited. In the human uterus, the putative ‘window of implantation’ in the mid-luteal phase of the menstrual cycle is accompanied by increased endometrial epithelial expression of the integrins α1β1, α4β1 and αvβ3 and its ligand osteopontin. Similar cyclical changes in FT integrin expression have been proposed to contribute to ectopic implantation, but supporting data are limited. In the current study, we present quantitative data on human FT transcription and translation of the integrin subunits α1, α4, αV, β1 and β3 during the follicular and mid-luteal phases of the menstrual cycle, together with a supporting immuocytochemical analysis of their spatial distribution within the FT, and that of osteopontin. In contrast to previous studies, our data indicate that all five integrin receptivity markers are constitutively transcribed and translated in the FT, with no evidence for changes in their expression or distribution during the window of implantation in the mid-luteal phase of the cycle. Furthermore, we could find no evidence for cyclic redistribution of the integrin αvβ3 ligand osteopontin within the FT. Although we do not rule out the involvement of integrin endometrial receptivity markers in the establishment of ectopic pregnancy, our findings do not support their differential expression during a tubal implantation window.

Introduction

An ectopic pregnancy is defined as any pregnancy implanted outside the uterus, with the vast majority (>98%) occurring in the Fallopian tube (FT) (Varma and Gupta, 2009; Sivalingam et al., 2011). The condition has a major clinical and socioeconomic impact worldwide and remains the leading cause of death in the first trimester of pregnancy in the developed world (Farquhar, 2005; Varma and Gupta, 2009). In developing countries, it has been estimated that 10% of women admitted to hospital with a diagnosis of ectopic pregnancy ultimately die from the condition (Leke et al., 2004).

The etiology of tubal implantation is still far from complete, but the bulk of the existing literature supports the hypothesis that it arises from a combination of impaired embryo-tubal transport and changes in the FT environment (Shaw et al., 2010; Brown and Horne, 2011). In the uterus, implantation only occurs when the endometrium is receptive during a putative ‘window of implantation’ in the mid-luteal phase of the menstrual cycle that is associated with marked changes in integrin expression within the endometrial epithelium (Lessey, 1998). The integrins are a family of widely expressed heterodimeric cell surface receptors that mediate cell–cell and cell–extracellular matrix adhesion and, in doing so, regulate many aspects of cell behavior including survival, proliferation, migration and differentiation. Twenty-four different integrin heterodimers are currently recognized in humans, each comprising a pair of non-covalently associated α- and β-subunits (Barczyk et al., 2010). In addition to providing a physical transmembrane link between the extracellular environment and the cytoskeleton, they are capable of transducing bi-directional signals across the cell membrane (Hynes, 2002).

In humans, endometrial transcription of integrins α4, αv, β1 and β3 is significantly higher during the mid-luteal phase of the menstrual cycle, compared with the follicular phase, with the αv and β3 subunits showing the largest increases in expression (Dou et al., 1999). More specifically, the β3 subunit and osteopontin, a ligand for integrin αvβ3, are only expressed at the luminal surface of the endometrium during the receptive period (Lessey et al., 1992; Apparao et al., 2001) supporting the concept that this cycle-dependent protein is involved in implantation in humans (Lessey, 1998).

While endometrial epithelial expression of the integrin heterodimers, α1β1, α4β1 and αvβ3, correlates with receptivity to the presenting embryo in humans (Lessey et al., 1992, 1994a; Lessey, 1998), functional data pertaining to the role of these integrins in implantation is currently limited. Homozygous integrin β1 and α4 null mutations are embryonic lethal (Fassler and Meyer, 1995; Stephens et al., 1995; Yang et al., 1995), while ∼80% of αv null/null mice die in utero, with the remainder dying shortly after birth (Bader et al., 1998). Consequently, there is no data on implantation in female mice that are homozygous null for these three integrins. Homozygous integrins α1 null mice are viable but do not exhibit any reduction in fecundity, suggesting that this integrin does not contribute to implantation in the mouse model (Gardner et al., 1996). The phenotype of integrin β3 knockout mice is more complex, with female homozygous null mice producing ∼50% smaller litters due to a combination of embryo and placental defects (Hodivala-Dilke et al., 1999). No direct analysis of implantation defects has been undertaken in the β3 knockout mice but implantation has been shown to be inhibited in another mouse model by functional blockade of integrin αvβ3 using neutralizing monoclonal antibodies against integrin β3 and integrin αv or the specific αvβ3 disintegrin echistatin (Illera et al., 2000).

A ‘window of implantation’ has also been proposed to occur in the FT, during which time the tubal epithelium is susceptible to ectopic implantation (Sulz et al., 1998). If such a window does occur in the FT, it follows that the integrins α1β1, α4β1 and αvβ3 are likely to be differentially expressed within the FT during the follicular and mid-luteal phases of the menstrual cycle. Semi-quantitative RT–PCR data suggest that the integrin subunits α1, αv and β3 are differentially regulated across the oestrous cycle in the bovine oviduct, with expression levels reaching a minima during the late-luteal phase before, in the case of β3, peaking in the pre-ovulation phase (Gabler et al., 2003). Human studies have also produced data in support of this hypothesis, specifically, reporting increased immunohistochemical labeling intensity for the β3 subunit (Sulz et al., 1998) and more recently the αvβ3 heterodimer (Makrigiannakis et al., 2009) in human tubal epithelium during the mid-luteal phase of the cycle. However, both of these studies were based entirely on immunohistochemistry observations and, given the potential key role for integrin αvβ3 in embryo implantation (Illera et al., 2000), there is a requirement for a quantitative analysis of integrin receptivity marker expression across the menstrual cycle in human FT. The current study was undertaken with the aim of meeting this objective.

Materials and Methods

Tissue collection

Full thickness cross sections of human FT ampulla (follicular phase n = 6, mid-luteal phase n = 6) and Pipelle™ uterine endometrial biopsies (follicular phase n = 2, mid-luteal phase n = 2) were collected from fertile women (Parity ≥2) with regular menstrual cycles (24–35 days) during hysterectomy for benign gynecological conditions (median age = 41; range 27–49 years). Tissues were collected into RNAlater (Applied Biosystems, Warrington, UK) and neutral-buffered formalin, as previously described (Shaw et al., 2011). Menstrual cycle dating was determined by three criteria, each of which had to correlate in order for inclusion in the study: (i) date of last menstrual period (as reported by the patient); (ii) staging by an expert gynecological pathologist of an endometrial biopsy obtained at the time of FT biopsy and (iii) serum estradiol (follicular phase >100 pM) and progesterone (follicular phase <10 nM; mid-luteal >20 nM). Approval for this study was obtained from the Lothian Research Ethics Committee (04/S1103/20, 05/S1103/14, 07/S1103/29), and informed, written consent was obtained from each patient.

Quantitative reverse transcription PCR

RNA was extracted from tissue using TRIzol reagent (Invitrogen, Paisley, UK), treated with DNase and purified using RNeasy kits (Qiagen, Crawley, UK). Two hundred nanograms of RNA were reverse-transcribed into cDNA using random hexamers, according to the recommended method (Applied Biosystems). TaqMan quantitative real-time reverse transcription PCR (qRT-PCR) was used to quantify integrin transcript levels. Specific primers were designed using the Universal Probe Library (UPL) Assay Design Center (www.roche-applied-science.com) and used in conjunction with UPL probes (Roche Applied Science). Details of primer sequences and probes used are given in Table I. Reactions were performed, in triplicate, under standard conditions in an ABI Prism 7900 instrument (Applied Biosystems). Integrin gene expression was normalized to ribosomal 18S expression, using the 2-forumla method, and expressed as relative to a positive standard (a single cDNA sample from follicular FT, included on each reaction plate).

Table I

qRT-PCR primer and probe sequences.

ITGA1 forward 5′-AATTGGCTCTAGTCACCATTGTT-3′ 
ITGA1 reverse 5′-CAAATGAAGCTGCTGACTGGT-3′ 
ITGA1 UPL probe (FAM) 14 
ITGA4 forward 5′-GGAATATCCAGTTTTTACACAAAGG-3′ 
ITGA4 reverse 5′-AGAGAGCCAGTCCAGTAAGATGA-3′ 
ITGA4 UPL probe (FAM) 57 
ITGAV forward 5′-GCCGTGGATTTCTTCGTG-3′ 
ITGAV reverse 5′-GAGGACCTGCCCTCCTTC-3′ 
ITGAV UPL probe (FAM) 64 
ITGB1 forward 5′-CGATGCCATCATGCAAGT-3′ 
ITGB1 reverse 5′-ACACCAGCAGCCGTGTAAC-3′ 
ITGB1 UPL probe (FAM) 65 
ITGB3 forward 5′-GGGCAGTGTCATGTTGGTAG-3′ 
ITGB3 reverse 5′-CAGCCCCAAAGAGGGATAAT-3′ 
ITGB3 UPL probe (FAM) 13 
18S forward 5′-CGGCTACCACATCCAAGGAA-3′ 
18S reverse 5′-GCTGGAATTACCGCGGCT-3′ 
18S probe (VIC) 5′-TGCTGGCACCAGACTTGCCCTC-3′ 
ITGA1 forward 5′-AATTGGCTCTAGTCACCATTGTT-3′ 
ITGA1 reverse 5′-CAAATGAAGCTGCTGACTGGT-3′ 
ITGA1 UPL probe (FAM) 14 
ITGA4 forward 5′-GGAATATCCAGTTTTTACACAAAGG-3′ 
ITGA4 reverse 5′-AGAGAGCCAGTCCAGTAAGATGA-3′ 
ITGA4 UPL probe (FAM) 57 
ITGAV forward 5′-GCCGTGGATTTCTTCGTG-3′ 
ITGAV reverse 5′-GAGGACCTGCCCTCCTTC-3′ 
ITGAV UPL probe (FAM) 64 
ITGB1 forward 5′-CGATGCCATCATGCAAGT-3′ 
ITGB1 reverse 5′-ACACCAGCAGCCGTGTAAC-3′ 
ITGB1 UPL probe (FAM) 65 
ITGB3 forward 5′-GGGCAGTGTCATGTTGGTAG-3′ 
ITGB3 reverse 5′-CAGCCCCAAAGAGGGATAAT-3′ 
ITGB3 UPL probe (FAM) 13 
18S forward 5′-CGGCTACCACATCCAAGGAA-3′ 
18S reverse 5′-GCTGGAATTACCGCGGCT-3′ 
18S probe (VIC) 5′-TGCTGGCACCAGACTTGCCCTC-3′ 

Quantitative dual chemiluminescent western blot

RNAlater stabilized samples of FT were homogenized in pH 8.0 lysis buffer (50 mM Tris–HCl; 150 mM NaCl; 1 mM EDTA; 1% Triton-X100, 1% Na-deoxycholate; EDTA-free complete mini protease inhibitors [Roche Diagnostics, Welwyn Garden City, UK) and 1 mM of the serine protease inhibitor, AEBSF (Sigma, Poole, UK)] using a TissueLyser bead mill (Qiagen). Protein quantification was performed by using the Bradford Assay (Bradford, 1976), adapted for the Cobas Fara centrifugal analyzer (Roche Diagnostics) and samples adjusted to 2 mg/ml total protein in lysis buffer, before further 1:1 dilution in 2× NuPAGE LDS sample buffer (Invitrogen) containing 100 mM dithiothreitol (Sigma). One-dimensional gel electrophoresis was performed in 15-well NuPAGE 4–12% Bis–Tris gels (Invitrogen) using 5–25 µg of total protein/lane alongside SeeBlue® Plus2 prestained molecular weight standards (Invitrogen). A positive control (pooled protein extracts from follicular FT biopsies) was also included in every gel to allow intra-blot comparisons to be made. Gels were equilibrated for 15 min in transfer buffer (50 mM Tris, 40 mM Glycine and 0.05% SDS), before blotting at 20 V (limited to 80 mA/gel) onto polyvinylidene fluoride membrane (Immobilon P: Millipore, Livingston, UK) in the presence of transfer buffer + 10% methanol using a Transblot SD (Bio-Rad Laboratories, Hemel Hempstead, UK).

Blots were blocked for 30 min in TBS-T20 (pH 7.4 Tris-buffered saline containing 0.5% Tween 20) + 2% Marvel (Premier Foods, St Albans, UK) and incubated for 2 h with combinations of mouse and rabbit anti-integrin (Abcam, Cambridge, UK), anti-β actin primary antibodies and/or negative control antibodies (Sigma) diluted appropriately in TBS-T20 + 2% Marvel (Table II). Blots were then washed in TBS-T20 (6 × 3 min) and incubated for 1 h with the appropriate combination of horse-radish peroxidase (HRPO) and alkaline phosphatase (ALKP)-conjugated secondary antibodies (Stratech Scientific, Newmarket, UK) diluted to 20 ng/ml in TBS-T20 + 2% Marvel (Table II). Following washing in TBS-T20 (6 × 3 min), integrin-specific HRPO labeling was visualized using Chemiluminescent Peroxidase Substrate-1 (Sigma). Blots were then washed (3 × 5 min) in TBS-T20 and rinsed in 100 mM Tris–HCl + 100 mM NaCl (pH 9.5), before β-actin-specific ALKP labeling was visualized using CDP-STAR Star (Boehringer-Mannheim, Germany). Images were acquired using a VersaDoc™ Imaging System (Bio-Rad Laboratories). Integrin-specific HRPO chemiluminescent signal and β-actin-specific ALKP chemiluminescent signal were measured using ImageJ software (Rasband, 1997–2011). After normalizing against β-actin, values for integrin-specific labeling were expressed as relative to the positive control. After imaging, blots were stained with Imperial Protein Stain (Fisher Scientific UK, Loughborough, UK) to confirm uniform blotting efficiency.

Table II

Antibodies used in dual chemiluminescent western blots.

Integrin antibody Working conc./dilution Loading control antibody Loading control working conc. Donkey secondary antibodies 
Monoclonal mouse IgG1 anti-integrin α1 0.2 µg/ml Rabbit anti-β actin 0.2 µg/ml Anti-mouse HRPO 
Anti-rabbit ALKP 
Control Mouse IgG1 0.2 µg/ml Rabbit anti-β actin 0.2 µg/ml Anti-mouse HRPO 
Anti-rabbit ALKP 
Monoclonal rabbit anti-integrin α4 1 in 10 000 Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Monoclonal rabbit anti-integrin β1 1 in 10 000 Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Polyclonal rabbit anti-integrin β3 0.2 µg/ml Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Control Rabbit IgG 0.2 µg/ml Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Integrin antibody Working conc./dilution Loading control antibody Loading control working conc. Donkey secondary antibodies 
Monoclonal mouse IgG1 anti-integrin α1 0.2 µg/ml Rabbit anti-β actin 0.2 µg/ml Anti-mouse HRPO 
Anti-rabbit ALKP 
Control Mouse IgG1 0.2 µg/ml Rabbit anti-β actin 0.2 µg/ml Anti-mouse HRPO 
Anti-rabbit ALKP 
Monoclonal rabbit anti-integrin α4 1 in 10 000 Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Monoclonal rabbit anti-integrin β1 1 in 10 000 Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Polyclonal rabbit anti-integrin β3 0.2 µg/ml Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 
Control Rabbit IgG 0.2 µg/ml Mouse anti-β actin 1 in 10 000 Anti-mouse ALKP 
Anti-rabbit HRPO 

Immunocytochemistry

Formal-saline fixed paraffin wax embedded (FFPE) sections were mounted on Snow Coat X-tra™ charged slides (Surgipath Europe, Peterborough, UK), dewaxed in xylene and rehydrated through graded ethanol. Antigen retrieval was performed by pressure cooking, from cold, for 5 min at pressure in 10 mM Tris, 1 mM EDTA and 0.05% T20 (pH 9.0). Once the pressure cooker had returned to ambient pressure, slides were washed in deionized water (DH2O) and endogenous peroxidase activity blocked by a 30-min incubation in TBS-T20 containing 1% H2O2. Slides were washed in TBS-T20 and transferred to a Sequenza immunostaining centre (Thermo Shandon; Runcorn, UK). Non-specific protein binding was blocked by incubating sections for 30 min with staining buffer [TBS-T20 + 10% normal horse serum for integrin labeling or TBS-T20 + 4% bovine serum albumin (BSA) of osteopontin and fibronectin labeling].

For immunohistochemistry (IHC): samples were incubated overnight at 4°C with primary anti-integrin antibodies and controls diluted appropriately in staining buffer (Table III). Slides were subsequently washed in TBS-T20 and incubated for 10 min at room temperature with staining buffer, before being incubated for 30 min with ImmPRESS Universal Antibody (anti-mouse Ig/anti-rabbit Ig, peroxidase) Polymer Detection Kit (Vector Laboratories, Peterborough, UK). Sections were washed in TBS-T20 and incubated for 10 min with 3,3′-diaminobenzidine + Chromogen (Dako UK, Ely, UK), counterstained in Mayer's Haematoxylin and mounted with No. 1.5 glass coverslips using Pertex (Cellpath PLC, Hemel Hempstead, UK).

Table III

Integrin antibody working concentrations for immunocytochemistry. 

Primary antibody Supplier IHC conc. IF conc. 
Monoclonal mouse IgG1 anti-integrin α1 (FB12) Chemicon 0.5 µg/ml — 
Monoclonal mouse IgG1 anti-integrin αvβ3 (BV3) Abcam 2–0.5 µg/ml 2–0.5 µg/ml 
Monoclonal mouse IgG1 anti-integrin αvβ3 (23C6) Chemicon 2–0.5 µg/ml 2–0.5 µg/ml 
Monoclonal mouse IgG1 anti-integrin αvβ3 (LM609) Chemicon 2–0.5 µg/ml 2–0.5 µg/ml 
Monoclonal mouse IgG2a anti-osteopontin (53) Abcam 0.2 µg/ml 0.2 µg/ml 
Monoclonal rabbit IgG anti-integrin α4 (EPR1355Y) Abcam 1 in 3000 — 
Polyclonal rabbit IgG anti-integrin αV Abcam 0.5 µg/ml — 
Monoclonal rabbit IgG anti-integrin β1 (EP1041Y) Abcam 1 in 3000 — 
Monoclonal rabbit IgG anti-fibronectin (F1) Abcam 1 in 5000 1 in 10 000 
Polyclonal rabbit IgG anti-integrin β3 Abcam 0.5 µg/ml 0.2 µg/ml 
Control mouse IgG1 Serotec 2–0.5 µg/ml — 
Control mouse IgG2a Abcam 0.2 µg/ml 0.2 µg/ml 
Control rabbit IgG Vector 1–0.2 µg/ml 0.5–0.2 µg/ml 
Primary antibody Supplier IHC conc. IF conc. 
Monoclonal mouse IgG1 anti-integrin α1 (FB12) Chemicon 0.5 µg/ml — 
Monoclonal mouse IgG1 anti-integrin αvβ3 (BV3) Abcam 2–0.5 µg/ml 2–0.5 µg/ml 
Monoclonal mouse IgG1 anti-integrin αvβ3 (23C6) Chemicon 2–0.5 µg/ml 2–0.5 µg/ml 
Monoclonal mouse IgG1 anti-integrin αvβ3 (LM609) Chemicon 2–0.5 µg/ml 2–0.5 µg/ml 
Monoclonal mouse IgG2a anti-osteopontin (53) Abcam 0.2 µg/ml 0.2 µg/ml 
Monoclonal rabbit IgG anti-integrin α4 (EPR1355Y) Abcam 1 in 3000 — 
Polyclonal rabbit IgG anti-integrin αV Abcam 0.5 µg/ml — 
Monoclonal rabbit IgG anti-integrin β1 (EP1041Y) Abcam 1 in 3000 — 
Monoclonal rabbit IgG anti-fibronectin (F1) Abcam 1 in 5000 1 in 10 000 
Polyclonal rabbit IgG anti-integrin β3 Abcam 0.5 µg/ml 0.2 µg/ml 
Control mouse IgG1 Serotec 2–0.5 µg/ml — 
Control mouse IgG2a Abcam 0.2 µg/ml 0.2 µg/ml 
Control rabbit IgG Vector 1–0.2 µg/ml 0.5–0.2 µg/ml 

For paired immunofluorescence (IF): endogenous biotin was blocked using an avidin/biotin blocking kit (Vector Laboratories). After washing in TBS-T20, samples were incubated for 1 h at room temperature with primary anti-integrin-β3-specific antibody or control rabbit immunoglobulin G (IgG) diluted appropriately in staining buffer (Table III). Sections were then washed and incubated for 1 h at room temperature with staining buffer containing 2 µg/ml of biotinylated goat anti-rabbit IgG (Vector Laboratories). Sections were washed and incubated with R.T.U. ABC Reagent (Vector Laboratories), as per the manufacturer's instructions, before further washing and incubation for 5 min with Tyramide Signal Amplification (TSA) plus FITC (PerkinElmer, Seer Green, UK). To elute existing antibody-peroxidase complexes from the sections, slides were removed from the Sequenza immunostaining center, microwaved at full power in pre-warmed 10 mM sodium citrate (pH 6.0) for 6 min and allowed to cool to room temperature. Slides were then returned to the Sequenza immunostaining center and blocked with staining buffer followed by the avidin/biotin blocking kit. After washing, samples were incubated for 1 h at room temperature with primary anti-osteopontin-specific antibody or control mouse IgG2a diluted appropriately in staining buffer (Table III). Sections were then washed and incubated for 1 h at room temperature with staining buffer containing 2 µg/ml of biotinylated rabbit anti-mouse IgG (Dako). After washing, sections were incubated with R.T.U. ABC Reagent, before further washing and incubation for 5 min with TSA plus Cy3 (PerkinElmer). Finally, sections were washed with TBS and counter stained for 15 min in 1 µM ToPro-3 (Invitrogen) in TBS, before mounting with No. 1.5 glass coverslips using Mowiol (pH 8.5) mounting media (EMD Biosciences, San Diego, CA, USA). Fluorescent images were acquired with a LSM510 META confocal microscope equipped with a ×63 Plan-Apochromat® 1.4 NA objective lens (Carl Zeiss, Welwyn Garden City, UK).

Statistical analysis

All statistical analyses were performed using InStat (GraphPad Software, La Jolla, CA, USA). Differences between groups were analyzed using a two-tailed, Mann–Whitney test and differences were considered significant when P < 0.05.

Results

Quantitative RT–PCR analysis of integrin endometrial receptivity marker gene transcription in follicular and mid-luteal-staged FT biopsies

Messenger RNA transcripts from all five integrin subunit genes studied (ITGA1, ITGA4, ITGAV, ITGB1 and ITGB3) were detected by qRT-PCR in human FT biopsies (Fig. 1). There was little evidence for differences in integrin transcript levels between the follicular and mid-luteal FT groups. Although median ITGB3 transcript levels were higher in the mid-luteal group, the spread of the data and statistical analysis (Mann–Whitney: P = 0.1797) indicate that this observation occurred by chance and that there is no difference in ITGB3 expression between the two groups. With the exception of ITGA4, which appears to be transcribed at lower levels (Fig. 1C), FT (Fig. 1: clear plots) expression of all of the integrins studied here appears to be commensurate with that observed in mid-luteal endometrium (Fig. 1: filled plots).

Figure 1

Quantitative RT–PCR analysis of integrin transcripts in FT (open plots) and endometrial (filled plots) biopsies taken during the follicular and mid-luteal phases of the menstrual cycle. Boxes represent median values ± 1 SD, whiskers denote the full range of the data. Individual panels are presented for: (A) ITGA1; (B) ITGA4; (C) ITGAV; (D) ITGB1 and (E) ITGB3. No significant (P > 0.05) differences in integrin expression were observed between follicular and mid-luteal FT biopsies.

Figure 1

Quantitative RT–PCR analysis of integrin transcripts in FT (open plots) and endometrial (filled plots) biopsies taken during the follicular and mid-luteal phases of the menstrual cycle. Boxes represent median values ± 1 SD, whiskers denote the full range of the data. Individual panels are presented for: (A) ITGA1; (B) ITGA4; (C) ITGAV; (D) ITGB1 and (E) ITGB3. No significant (P > 0.05) differences in integrin expression were observed between follicular and mid-luteal FT biopsies.

Quantitative western blot analysis of integrin endometrial receptivity marker protein levels in follicular and mid-luteal staged FT biopsies

Integrin-α1-, α4-, β1- and β3-specific antibodies reacted with discreet bands in western blots of pooled protein extracts from both follicular and mid-luteal FT biopsies (Fig. 2). No bands were detected with integrin-αv-specific antibodies at total protein loadings of up to 25 µg/lane. Integrin-α1-specific antibodies reacted strongly with a band of ∼190 KDa (expected: 200 KDa), and to a much lesser extent with a band of ∼85 KDa, at a total protein loading of 10 µg/lane. Integrin-α4-specific antibodies reacted with a band of ∼85 KDa (expected: 150 KDa) at a total protein loading of 25 µg/lane. Integrin-β1-specific antibodies reacted strongly with a band of ∼90 KDa (expected size: 88 KDa) at a total protein loading of 5 µg/lane. Integrin-β3-specific antibodies reacted with a band of ∼75 KDa (expected size: 87 KDa), and to a lesser extent ∼45 KDa, at a total protein loading of 25 µg/lane. No bands were observed when integrin-specific antibodies were replaced with equivalent amounts of control mouse IgG1 or control rabbit IgG (data not shown).

Figure 2

Images of dual chemiluminescent western blots for integrins and β-actin in pooled protein extracts from follicular (F) and mid-luteal (ML) FT biopsies. Separate panels are shown for: (A) mouse (IgG1) anti-integrin α1; (B) rabbit anti-integrin α4; (C) rabbit anti-integrin β1 and (D) rabbit anti-integrin β3. Images of β-actin specific labeling are provided in the lower panels.

Figure 2

Images of dual chemiluminescent western blots for integrins and β-actin in pooled protein extracts from follicular (F) and mid-luteal (ML) FT biopsies. Separate panels are shown for: (A) mouse (IgG1) anti-integrin α1; (B) rabbit anti-integrin α4; (C) rabbit anti-integrin β1 and (D) rabbit anti-integrin β3. Images of β-actin specific labeling are provided in the lower panels.

Data derived from quantitative analysis of dual chemiluminescent western blots of individual FT protein extracts are presented in Fig. 3. Integrated density values of integrin α1, α4, β1 and β3 bands were normalized against that of β-actin bands for each lane and the result expressed as a function of the positive control (pooled protein extracts from follicular FT biopsies). There was no evidence for differences in integrin protein levels between the follicular and mid-luteal staged samples.

Figure 3

Quantitative analysis of integrin protein levels in follicular and mid-luteal FT protein extracts. Boxes represent median values ± 1 SD, whiskers denote the full range of the data. Individual panels are presented for: (A) integrin α1; (B) integrin α4; (C) integrin β1 and (D) integrin β3. No significant (P > 0.05) differences in integrin expression were observed between follicular and mid-luteal phases of the menstrual cycle.

Figure 3

Quantitative analysis of integrin protein levels in follicular and mid-luteal FT protein extracts. Boxes represent median values ± 1 SD, whiskers denote the full range of the data. Individual panels are presented for: (A) integrin α1; (B) integrin α4; (C) integrin β1 and (D) integrin β3. No significant (P > 0.05) differences in integrin expression were observed between follicular and mid-luteal phases of the menstrual cycle.

Immunolocalization of integrin endometrial receptivity markers in follicular and mid-luteal-staged FT biopsies

Representative images of follicular (n = 6) and mid-luteal (n = 6) FT tissue sections labeled with integrin-specific antibodies are shown in Fig. 4. Antigen retrieval at pH 9.0 (10 mM Tris, 1 mM EDTA, 0.05% T20) allowed detection of all five integrin subunits under investigation and the integrin αvβ3 ligands, osteopontin and fibronectin, without significant degradation of tissue morphology. Blocking of non-specific protein binding using BSA produced superior results for fibronectin and osteopontin, compared with normal serum, which contains significant levels of both proteins (Mosesson and Amrani, 1980; Scatena et al., 2007). None of the monoclonal antibodies directed against the integrin αvβ3 heterodimer (BV3; 23C6 or LM609) produced convincing specific labeling of FFPE sections, even when highly sensitive TSA  labeling was employed. The epitope recognized by the polyclonal rabbit anti-integrin αv (Fig. 4C) was methanol sensitive, even when quenching of endogenous peroxidase using 0.3% H2O2 in methanol was performed prior to antigen retrieval.

Figure 4

Distribution of integrin subunits and putative integrin αVβ3 ligands within FT biopsies from the follicular and mid-luteal (asterisk) phases of the menstrual cycle. Panels show representative images of tissue sections labeled with: (A) mouse (IgG1) anti-integrin α1; (B) rabbit anti-integrin α4; (C) rabbit anti-integrin αV; (D) rabbit anti-integrin β1; (E) rabbit anti-integrin β3; (F) mouse (IgG2a) anti-osteopontin and (G) rabbit anti-fibronectin. Staining fidelity was confirmed by substituting primary antibodies with equivalent amounts of: (C1) control mouse IgG1 (integrin staining run); (C2) control rabbit IgG (integrin staining run); (C3) control mouse IgG2a (osteopontin staining run); control rabbit IgG (fibronectin staining run). Scale bars represent 100 µm.

Figure 4

Distribution of integrin subunits and putative integrin αVβ3 ligands within FT biopsies from the follicular and mid-luteal (asterisk) phases of the menstrual cycle. Panels show representative images of tissue sections labeled with: (A) mouse (IgG1) anti-integrin α1; (B) rabbit anti-integrin α4; (C) rabbit anti-integrin αV; (D) rabbit anti-integrin β1; (E) rabbit anti-integrin β3; (F) mouse (IgG2a) anti-osteopontin and (G) rabbit anti-fibronectin. Staining fidelity was confirmed by substituting primary antibodies with equivalent amounts of: (C1) control mouse IgG1 (integrin staining run); (C2) control rabbit IgG (integrin staining run); (C3) control mouse IgG2a (osteopontin staining run); control rabbit IgG (fibronectin staining run). Scale bars represent 100 µm.

There was no evidence for changes in integrin labeling intensity, or distribution, between follicular and mid-luteal staged sections of FT (Fig. 4A–E). Similarly, specific labeling of the integrin αvβ3 ligands osteopontin (Fig. 4F) and fibronectin (Fig. 4G) was not influenced by the phase of the cycle in the FT. Integrins α4 (Fig. 4B), αv (Fig. 4C) and β1 (Fig. 4D) specific immunohistochemistry produced intense labeling throughout the FT epithelium, in a pattern consistent with their presence at the plasma membrane, whereas integrin-β3-specific labeling was largely restricted to the luminal plasma membrane of epithelial cells (Fig. 4E). All four of these integrins were detected on leukocytes scattered throughout the FT stromal or, in the case of integrin β3 (Fig. 4E), located near the basement membrane of the epithelium. There was also evidence for the expression of integrins α4, αv and β1 by endothelial cells and, in the case of the αv and β1 subunits, by smooth muscle cells within the FT stroma and surrounding connective tissue. By contrast, integrin-α1-specific labeling was restricted to epithelial cells and exhibited an intense punctuate distribution consistent with it being restricted to the golgi body, with no evidence for it being expressed on the cell surface (Fig. 4A). Osteopontin specific labeling was widely distributed throughout the FT and surrounding smooth muscle and connective tissues but was most prominent within the epithelium compartment of the mucosa, where it appeared to be expressed at high levels by a subset of epithelial cells (Fig. 4F). Although also widely distributed, fibronectin specific labeling was not present with the epithelium of the FT and was largely restricted to endothelial cells and blood pool in the stroma (Fig. 4G). Non-specific labeling was not observed when specific antibodies were substituted with equivalent amounts of appropriate control IgG (Fig. 4C1–C4).

Paired immunofluorescent labeling of integrin β3 and osteopontin in follicular and mid-luteal-staged FT and endometrial biopsies

Further analysis of integrin β3 and osteopontin co-expression was undertaken using paired immunofluorescence. Representative images of follicular and mid-luteal FT (n = 6 per group) and endometrium (n = 2 per group) tissue sections are shown in Fig. 5. There was limited osteopontin- and integrin-β3-specific labeling in the endometrial epithelia during the follicular phase of the menstrual cycle, with intense β3 specific labeling of sporadic cells within the stromal compartment (Fig. 5A), but both molecules were strongly expressed in the luminal epithelium of mid-luteal-staged endometrium (Fig. 5A*). In contrast, FT epithelial cells co-expressed integrin β3 and osteopontin during both the follicular (Fig. 5B) and mid-luteal (Fig. 5B*) phases of the cycle, with no evidence for any cycle-dependent changes in expression level or spatial distribution. Despite being co-expressed in the epithelia of FT and mid-luteal endometria, there was very limited evidence for co-localization (1.4NA objective: ∼200 nm lateral//∼400 nm axial resolution for Cy3) of integrin β3 and osteopontin in either tissue. Integrin β3 also exhibited a different intracellular distribution in the FT and endometrial epithelia, with the vast majority of specific labeling occurring at or near the luminal surface of FT epithelial cells (Fig. 5B and B*). No evidence for non-specific labeling or crosstalk was observed when specific antibodies were substituted with equivalent amounts of control IgG (Fig. 5C1 and C2).

Figure 5

Paired immunofluorescent labeling of integrin β3 and osteopontin within endometrial (A/A*) and FT (B/B*) biopsies from the follicular (A and B) and mid-luteal (A* and B*) phases of the menstrual cycle. (A/A*) and (B/B*) show representative images of tissue sections labeled with rabbit anti-integrin β3 (green), mouse anti-osteopontin (red) and TopPro-3 nuclear counterstain (blue). Staining fidelity was confirmed on sections of mid-luteal FT, substituting mouse anti-osteopontin (C1) or rabbit anti-integrin β3 (C2) with equivalent amounts of control antibodies. Excitation and exposure settings were standardized from each tissue. L indicates the luminal side of the epithelium. Scale bars represent 10 µm.

Figure 5

Paired immunofluorescent labeling of integrin β3 and osteopontin within endometrial (A/A*) and FT (B/B*) biopsies from the follicular (A and B) and mid-luteal (A* and B*) phases of the menstrual cycle. (A/A*) and (B/B*) show representative images of tissue sections labeled with rabbit anti-integrin β3 (green), mouse anti-osteopontin (red) and TopPro-3 nuclear counterstain (blue). Staining fidelity was confirmed on sections of mid-luteal FT, substituting mouse anti-osteopontin (C1) or rabbit anti-integrin β3 (C2) with equivalent amounts of control antibodies. Excitation and exposure settings were standardized from each tissue. L indicates the luminal side of the epithelium. Scale bars represent 10 µm.

Discussion

To date, published data on integrin receptivity marker expression in human FT has been limited to semi-quantitative immunohistochemical studies (Sulz et al., 1998; Makrigiannakis et al., 2009). Here, we present quantitative data on human FT transcription (Fig. 1) and translation (Fig. 3) of the integrin subunits α1, α4, αV, β1 and β3 during the follicular and mid-luteal phases of the menstrual cycle, together with comprehensive supporting immunocytochemistry data (Figs 4 and 5). In contrast to previous studies (Sulz et al., 1998; Makrigiannakis et al., 2009), our data indicate that all five integrin receptivity markers (α1, α4, αV, β1 and β3) are constitutively transcribed and translated in the FT epithelium, with no evidence for changes in their expression or distribution during the putative window of implantation in the mid-luteal phase of the cycle.

Our data are at variance with two previous immunohistochemical studies that reported increased integrin β3 subunit (Sulz et al., 1998) and αvβ3 heterodimer (Makrigiannakis et al., 2009)-specific antibody labeling in the epithelium of the FT during the mid-luteal phase of the menstrual cycle, with little or no labeling of the same area during the follicular phase. While it could be argued that this discrepancy arises from differences in the specificity of the antibodies used, the rabbit polyclonal antibody used in the current study produced results entirely consistent with published data for the temporal/spatial distribution of integrin β3 (Lessey et al., 1992, 1994a) and αvβ3 (Tei et al., 2003) in the endometrium (Fig. 5A and A*). Furthermore, the staining pattern we obtained with rabbit polyclonal anti-integrin β3 on FFPE sections of mid-luteal-staged endometrium (Fig. 5A*) was identical to that previously described for a monoclonal mouse integrin αvβ3 heterodimer-specific antibody (clone LM609) in frozen sections of mid-luteal endometrium (Tei et al., 2003). In the current study, both qRT-PCR (Fig. 1) and western blot (Figs 2 and 3) analysis were undertaken on tissue extracts derived from whole cross-sections of FT and it could be argued that subtle changes in epithelial expression of the integrin subunits could be overlooked against background expression in surrounding tissues. Indeed, the integrin subunits β1 and αV are ‘promiscuous’, forming heterodimmers with multiple partners, and are both expressed by a wide variety of cells, including smooth muscle and stromal cells (Hynes, 2002; Barczyk et al., 2010). However, their partners (α1, α4, and β3) are much more restricted in their distribution, which is predominantly epithelial in the FT and we are confident that any significant changes in epithelial expression of their functional heterodimmers (α1β1, α4β1 and αvβ3) would be detected by the measuring α1, α4, and β3 in whole FT cross-sections. It should be noted that Dou et al. (1999) were able to detect significantly higher transcription of the integrin subunits α4, αv, β1 and β3 during the mid-luteal phase of the menstrual cycle, compared with the follicular phase, using conventional RT–PCR to interrogate cDNA from unfractionated endometrial biopsies.

The absence of cycle-dependent changes in expression of integrin receptivity markers and osteopontin in human FT, reported in the current study, contrasts with the situation in endometrium (Lessey et al., 1992, 1994a; Lessey, 1998; Dou et al., 1999; Apparao et al., 2001). We would argue that this indicates that expression of these receptivity markers, in particular integrin αvβ3 and osteopontin, is likely to be regulated through different mechanisms in the endometrium and the FT. In addition, the intracellular distribution of the α1 and β3 subunits also differ markedly between the epithelia of the FT (Figs 4 and 5) and the endometrium (Lessey et al., 1992), suggesting that these integrins may also fulfill functionally distinct roles in the two tissues. This is, perhaps, unsurprising given the likely evolutionary disadvantage of a FT that is receptive to ectopic implantation at any point during the menstrual cycle, let alone one that increases its receptivity to compete with the endometrium during the window of implantation.

The current study is not the first to report differential regulation of gene expression in the FT and endometrium. We have previously shown that, unlike endometrium, FT constitutively expresses the estrogen receptor isoform ERα throughout the menstrual cycle, with no evidence for a reduction in expression during the mid-luteal phase, when the tissue is exposed to peak levels of circulating progesterone (Horne et al., 2009). These differences in gene regulation are consistent with the distinct embryonic origins of the FT and uterus, and the persistence of differential Hox gene expression in the two organs after birth (Taylor et al., 1997). This spatial Hox axis, more typical of embryonic tissue, is thought play a role in preserving the developmental plasticity of the female reproductive system throughout the menstrual cycle and during pregnancy, allowing tissue remodeling to be differentially regulated in the FT and uterus by steroid hormones (Taylor et al., 1997; Masse et al., 2009).

Defects in integrin β3 expression have been implicated in the pathology of endometriosis, luteal phase defects and polycystic ovarian syndrome (Lessey et al., 1992, 1994b), all of which are associated with infertility or pregnancy loss. Although we do not rule out the involvement of integrin receptivity markers in the establishment of ectopic pregnancy, our data do not support the hypothesis that the temporal/spatial expression profile of integrin receptivity markers (α1β1, α4β1 and αvβ3) or the integrin αvβ3 ligand, osteopontin, reported in endometrium is mirrored in FT. Nor do they support the existence of a window of implantation in the FT. It would, therefore, appear that integrin expression is regulated through different mechanisms in the FT and endometrium, opening up the future possibility of being able to selectively modulate integrin receptivity marker expression in the FT without disturbing normal intrauterine implantation.

Authors’ roles

J.K.B. design, experiments, analysis and interpretation, drafting article and final approval. J.L.V.S. experiments and final approval. H.O.D.C. revising for critical content and final approval. A.W.H. conception and design, interpretation, revising for critical content and final approval.

Funding

This study was supported by the Medical Research Council (MRC G0802808). Funding to pay the Open Access publication charges for this article was provided by the Medical Research Council (MRC G0802808).

Conflict of interest

J.K.B. and J.L.V.S. have nothing to declare. A.W.H. and H.O.D.C. hold a UK patent for a diagnostic biomarker for ectopic pregnancy (#0712801.0). A.W.H. is supported by the UK Medical Research Council (2009–13) and an Albert McKern Bequest (2010–11).

Acknowledgements

We are grateful to Catherine Murray, Sharon McPherson and Helen Dewart for patient recruitment.

References

Apparao
KB
Murray
MJ
Fritz
MA
Meyer
WR
Chambers
AF
Truong
PR
Lessey
BA
Osteopontin and its receptor alphavbeta(3) integrin are coexpressed in the human endometrium during the menstrual cycle but regulated differentially
J Clin Endocrinol Metab
 , 
2001
, vol. 
86
 (pg. 
4991
-
5000
)
Bader
BL
Rayburn
H
Crowley
D
Hynes
RO
Extensive vasculogenesis, angiogenesis, and organogenesis precede lethality in mice lacking all alpha v integrins
Cell
 , 
1998
, vol. 
95
 (pg. 
507
-
519
)
Barczyk
M
Carracedo
S
Gullberg
D
Integrins
Cell Tissue Res
 , 
2010
, vol. 
339
 (pg. 
269
-
280
)
Bradford
MM
A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding
Anal Biochem
 , 
1976
, vol. 
72
 (pg. 
248
-
254
)
Brown
JK
Horne
AW
Laboratory models for studying ectopic pregnancy
Curr Opin Obstet Gynecol
 , 
2011
, vol. 
23
 (pg. 
221
-
226
)
Dou
Q
Williams
RS
Chegini
N
Expression of integrin messenger ribonucleic acid in human endometrium: a quantitative reverse transcription polymerase chain reaction study
Fertil Steril
 , 
1999
, vol. 
71
 (pg. 
347
-
353
)
Farquhar
CM
Ectopic pregnancy
Lancet
 , 
2005
, vol. 
366
 (pg. 
583
-
591
)
Fassler
R
Meyer
M
Consequences of lack of beta 1 integrin gene expression in mice
Genes Dev
 , 
1995
, vol. 
9
 (pg. 
1896
-
1908
)
Gabler
C
Chapman
DA
Killian
GJ
Expression and presence of osteopontin and integrins in the bovine oviduct during the oestrous cycle
Reproduction
 , 
2003
, vol. 
126
 (pg. 
721
-
729
)
Gardner
H
Kreidberg
J
Koteliansky
V
Jaenisch
R
Deletion of integrin alpha 1 by homologous recombination permits normal murine development but gives rise to a specific deficit in cell adhesion
Dev Biol
 , 
1996
, vol. 
175
 (pg. 
301
-
313
)
Hodivala-Dilke
KM
McHugh
KP
Tsakiris
DA
Rayburn
H
Crowley
D
Ullman-Cullere
M
Ross
FP
Coller
BS
Teitelbaum
S
Hynes
RO
Beta3-integrin-deficient mice are a model for Glanzmann thrombasthenia showing placental defects and reduced survival
J Clin Invest
 , 
1999
, vol. 
103
 (pg. 
229
-
238
)
Horne
AW
King
AE
Shaw
E
McDonald
SE
Williams
AR
Saunders
PT
Critchley
HO
Attenuated sex steroid receptor expression in fallopian tube of women with ectopic pregnancy
J Clin Endocrinol Metab
 , 
2009
, vol. 
94
 (pg. 
5146
-
5154
)
Hynes
RO
Integrins: bidirectional, allosteric signaling machines
Cell
 , 
2002
, vol. 
110
 (pg. 
673
-
687
)
Illera
MJ
Cullinan
E
Gui
Y
Yuan
L
Beyler
SA
Lessey
BA
Blockade of the alpha(v)beta(3) integrin adversely affects implantation in the mouse
Biol Reprod
 , 
2000
, vol. 
62
 (pg. 
1285
-
1290
)
Leke
RJ
Goyaux
N
Matsuda
T
Thonneau
PF
Ectopic pregnancy in Africa: a population-based study
Obstet Gynecol
 , 
2004
, vol. 
103
 (pg. 
692
-
697
)
Lessey
BA
Endometrial integrins and the establishment of uterine receptivity
Hum Reprod
 , 
1998
, vol. 
13
 
Suppl 3)
(pg. 
247
-
258
discussion 259–261
Lessey
BA
Damjanovich
L
Coutifaris
C
Castelbaum
A
Albelda
SM
Buck
CA
Integrin adhesion molecules in the human endometrium. Correlation with the normal and abnormal menstrual cycle
J Clin Invest
 , 
1992
, vol. 
90
 (pg. 
188
-
195
)
Lessey
BA
Castelbaum
AJ
Buck
CA
Lei
Y
Yowell
CW
Sun
J
Further characterization of endometrial integrins during the menstrual cycle and in pregnancy
Fertil Steril
 , 
1994
, vol. 
62
 (pg. 
497
-
506
)
Lessey
BA
Castelbaum
AJ
Sawin
SW
Buck
CA
Schinnar
R
Bilker
W
Strom
BL
Aberrant integrin expression in the endometrium of women with endometriosis
J Clin Endocrinol Metab
 , 
1994
, vol. 
79
 (pg. 
643
-
649
)
Makrigiannakis
A
Karamouti
M
Petsas
G
Makris
N
Nikas
G
Antsaklis
A
The expression of receptivity markers in the fallopian tube epithelium
Histochem Cell Biol
 , 
2009
, vol. 
132
 (pg. 
159
-
167
)
Masse
J
Watrin
T
Laurent
A
Deschamps
S
Guerrier
D
Pellerin
I
The developing female genital tract: from genetics to epigenetics
Int J Dev Biol
 , 
2009
, vol. 
53
 (pg. 
411
-
424
)
Mosesson
MW
Amrani
DL
The structure and biologic activities of plasma fibronectin
Blood
 , 
1980
, vol. 
56
 (pg. 
145
-
158
)
Rasband
WS
Image J
 , 
1997
Bethesda, MD, USA
U. S. National Institutes of Health
 
Scatena
M
Liaw
L
Giachelli
CM
Osteopontin: a multifunctional molecule regulating chronic inflammation and vascular disease
Arterioscler Thromb Vasc Biol
 , 
2007
, vol. 
27
 (pg. 
2302
-
2309
)
Shaw
JL
Dey
SK
Critchley
HO
Horne
AW
Current knowledge of the aetiology of human tubal ectopic pregnancy
Hum Reprod Update
 , 
2010
, vol. 
16
 (pg. 
432
-
444
)
Shaw
JL
Wills
GS
Lee
KF
Horner
PJ
McClure
MO
Abrahams
VM
Wheelhouse
N
Jabbour
HN
Critchley
HO
Entrican
G
, et al.  . 
Chlamydia trachomatis Infection Increases Fallopian tube PROKR2 via TLR2 and NFkappaB activation resulting in a microenvironment predisposed to ectopic pregnancy
Am J Pathol
 , 
2011
, vol. 
178
 (pg. 
253
-
260
)
Sivalingam
VN
Duncan
WC
Kirk
E
Shephard
LA
Horne
AW
Diagnosis and management of ectopic pregnancy
J Fam Plann Reprod Health Care
 , 
2011
, vol. 
37
 (pg. 
231
-
240
)
Stephens
LE
Sutherland
AE
Klimanskaya
IV
Andrieux
A
Meneses
J
Pedersen
RA
Damsky
CH
Deletion of beta 1 integrins in mice results in inner cell mass failure and peri-implantation lethality
Genes Dev
 , 
1995
, vol. 
9
 (pg. 
1883
-
1895
)
Sulz
L
Valenzuela
JP
Salvatierra
AM
Ortiz
ME
Croxatto
HB
The expression of alpha(v) and beta3 integrin subunits in the normal human Fallopian tube epithelium suggests the occurrence of a tubal implantation window
Hum Reprod
 , 
1998
, vol. 
13
 (pg. 
2916
-
2920
)
Taylor
HS
Vanden Heuvel
GB
Igarashi
P
A conserved Hox axis in the mouse and human female reproductive system: late establishment and persistent adult expression of the Hoxa cluster genes
Biol Reprod
 , 
1997
, vol. 
57
 (pg. 
1338
-
1345
)
Tei
C
Maruyama
T
Kuji
N
Miyazaki
T
Mikami
M
Yoshimura
Y
Reduced expression of alphavbeta3 integrin in the endometrium of unexplained infertility patients with recurrent IVF-ET failures: improvement by danazol treatment
J Assist Reprod Genet
 , 
2003
, vol. 
20
 (pg. 
13
-
20
)
Varma
R
Gupta
J
Tubal ectopic pregnancy
Clin Evid (Online)
 , 
2009
, vol. 
20
 pg. 
pii: 1406
 
Yang
JT
Rayburn
H
Hynes
RO
Cell adhesion events mediated by alpha 4 integrins are essential in placental and cardiac development
Development
 , 
1995
, vol. 
121
 (pg. 
549
-
560
)

Author notes

2
Present address: Laboratory Medicine and Pathobiology, University of Toronto, 1 King's College Circle, Toronto, Ontario, Canada M5S 1A8.
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