Animals ethics statement and experimental conditions
In our study, tissue samples of animals were taken in accordance with the rules of the Regulation on the Working Procedures and Principles of Animal Experiments Ethics Committees of the Ministry of Forestry and Water Affairs (dated 15 February 2014, 28914). This regulation has been prepared on the basis of Animals Protection Law (dated 24 April 2004, 51999) published by Official Gazette dated 1 July 2004 and in accordance with the Universal Declaration of Animal Rights, the European Convention on the Protection of Vertebrate Animals for Experimental and Other Scientific Purposes (Council of Europe ETS 123), and Guide for the Care and Use of Laboratory Animals. In this regulation, it has been reported that the permission of Animal Experiments Local Ethics Committee’ is not required in some cases including the clinical applications for diagnosis and treatment purposes, in procedures with dead animals or their tissues, slaughterhouse materials, aborted fetuses, milk samples, fecal or litter sample collection, and swab sampling, etc. Slaughter of animals in slaughterhouses is carried out under the control of veterinarians following the hygiene rules, at once without frightening, with the least pain. In slaughterhouses, blood drawing process can be done easily with the open draining method. In this method, when the butcher cuts the throat of the animal, the flowing blood is taken directly into the tube with a funnel. For the reasons explained above, no ethics committee approval was obtained before starting our study.
In this study, a total of 30 healthy Holstein cows aged 2-8 years, which were obtained from local abattoirs in Diyarbakır Province, Turkey, were used. Before slaughter, cows were checked for evidence of oestrous behaviors, including mounting or attempting to other cattle, smelling and trailing of other females, vulvar swelling and reddening, clear vaginal mucus discharge, and mucus smeared on the rump (Peralta et al 2005), before they were killed. After the cows had been killed, the entire reproductive tract was removed and macroscopically examined for the presence of disease. Animals without any clinical sign such as purulent uterine discharge, necrotic or hemorrhagic uterine mucosa, cervical or vaginal hyperemia, and edema, malodorous or non-odorous, purulent or mucopurulent vaginal discharge (Millward et al 2019) were included in this study.
Collection of blood samples and measurement of hormone concentrations
To measure serum concentrations of E2 and P4 hormones, the bloods of pre-selected cows were taken into a tube as soon as their throat was cut by the butcher and then, transported to the laboratory immediately after collection. After arriving at the laboratory, the samples were centrifuged (3969g for 5 min at 4 °C). All serum samples were stored at −20 °C until analysis. Serum concentrations of E2 and P4 were measured in a clinical laboratory (PRO-LABORATORY Laboratory Technologies, Istanbul) by enzyme immunoassay (EIA) using commercially available kits (DRG Aurica Elisa Oestradiol Kit (Catalogue no. EIA-2693) and DRG Aurica Elisa Progesterone Kit (Catalogue no. EIA-1561) respectively; DRG International) according to the manufacturer’s protocol.
Collection of tissue samples and histological analysis
After the macroscopic examination of the entire genital tract of the killed cows, small pieces about 2 cm in size depending on the organ were cut out from the ovaries (right and left), uterine horns (right and left), cervix, and vagina, and were immersed in 10% buffered formalin. In the bovine cervix, the cervical mucosa forms three to four annular folds or rings that project into the lumen, as well as numerous smaller longitudinal folds (Breeveld-Dwarkasing 2002). Therefore, the tissue samples used in this study were harvested from all three rings of the cervix, and from the vaginal area adjacent to the vulva. After fixation process, all tissue samples washed in tap water, dehydrated through an ethanol series (70%, 80%, 96%, 100%) and embedded in paraffin. To evaluate the histological changes that occur in the ovary and uterus during the oestrous cycle, the paraffin-embedded tissue samples of the right and left ovaries and the uterine horns of each animal were cut on a microtome into 7 µm-thick sections, and these slides were stained with a modified Mallory’s connective tissue stain (Crossmon 1937). Furthermore, for the histological evaluation of the cervical and vaginal epithelia, the paraffin-embedded cervical and vaginal tissue samples of each animal were cut at the 5-μm thickness, and slides were prepared and stained for mucin with Periodic acid-Schiff (PAS), in view of the bovine cervical and vaginal epithelia containing high levels of carbohydrates, including mucins, during the follicular stage of the oestrous cycle (Wrobel 1971; Wrobel et al 1986; Mullins and Saacke 1989; Miroud and Noakes 1991).
Determination of oestrous cycle phase
The phase of the cycle in the slaughtered cows was determined postmortem. The phase of the oestrous cycle of each cow was determined based on the presence/absence of corpora lutea (CL) or preovulatory follicles in the ovaries, the histological findings detected in the ovaries and uterus, and E2 and P4 concentrations measured in the serum samples (Benbia et al. 2017). The presence of a preovulatory follicle and fully developed CL were assumed as the characteristic features of the follicular and luteal phases of the oestrous cycle, respectively. The mean (±s.d.) serum E2 concentration was higher during the follicular phase (28.55±9.36 pg/mL-1) (ranging from 16.20 to 58.30 pg/mL-1 ; Benbia et al. 2017) compared to the luteal phase (10.73±4.06 pg/mL-1) (range 3.50– 13.20 pg/mL-1 ; Benbia et al. 2017), whereas the mean (±s.d.) P4 concentration was higher during the luteal phase (6.31±0.98 ng/mL-1) (range 4.00–8.20 ng/mL-1 ; Benbia et al. 2017) compared to the follicular phase (0.91±0.33 ng/ mL-1) (ranging from 0.40 to 1.30 ng/mL-1 ; Benbia et al. 2017). Based on these data and literature information (Benbia et al. 2017; Crowe 2016), the cows were divided into two groups, including a follicular phase group (n=13) and a luteal phase group (n=17). Similar to other domestic animals the oestrus cycle in the cow can be divided into four phases: proestrus, oestrus, metoestrus, and dioestrus. Proestrus and oestrus comprise the follicular phase of the ovarian cycle with ovulation taking place 10 to 12 hours after the end of oestrus. Metoestrus and dioestrus constitute the luteal phase of the cycle (Crowe 2016).
Immunohistochemistry
A standard strepavidin-biotin immunoperoxidase technique (Thermo Fisher Scientific Lab Vision Corporation, Fremont) was applied to detect the β-catenin and cadherin proteins. Briefly, the 5-μm paraffin-embedded cervical and vaginal sections were deparaffinised and treated with 3% hydrogen peroxide (H2O2) in methanol for 15 min to block endogenous peroxidase activity. After rinsing thoroughly in phosphate buffer saline (PBS) (pH 7.4), the sections were placed in 0.01 M citrate buffer (pH 6.0), heated in a water bath at 80°C for 30 minutes for antigen retrieval, and cooled for 20 min. Then, the sections were washed in PBS and treated with a blocking solution (Ultra V Block, Thermo Fisher Scientific, LabVision Corporation, Fremont, CA) for 5 min to prevent nonspecific interference of immunoglobulins. Subsequently, the sections were incubated at 4 °C overnight with the following antibodies: anti-pan-cadherin (Ab-4) [RB-1524, Thermo Fisher Scientific Lab Vision Corporation, Fremont, CA, USA, 1:200 dilution], anti-E-cadherin [ab15148, Abcam, 1:50 dilution], anti-P-cadherin [ab-137729, Abcam, 1:200 dilution], anti-N-cadherin [clone 13A9, sc-59987, Santa Cruz Biotechnology, Santa Cruz, CA, USA, 1:100 dilution], and anti-beta-catenin (E-5) [clone E-5, sc-7963, Santa Cruz Biotechnology, Santa Cruz, CA, USA, 1:100 dilution]. Next, the sections were incubated with the secondary antibody and streptavidin peroxidase (Thermo Fisher Scientific Lab Vision Corporation, Fremont, CA), followed by incubation with diaminobenzidine (DAB) substrate for 5 min. Subsequently, the sections were counterstained with Gill’s haematoxylin for 3 min, washed under running tap water, dehydrated through an alcohol series, cleared in xylene and mounted in Entellan (Merck).
As it is known, beta-catenin and cadherin antibodies specific for bovine tissues are not yet commercially available or their commercial production is limited. In the product datasheets of Pan-cadherin and P-cadherin antibodies used in this study have been reported to be positive for bovine tissues. Furthermore, various researchers showed that the N-cadherin antibody used in this study was positive for bovine ovaries (Lee et al. 2019) and beta-catenin antibody for Madin-Darby bovine kidney (MDBK) cells (Fay et al. 2020). We could not perform western blot analysis because the tissue samples examined in this study were prepared in paraffin long before and there was no fresh tissue sample in our laboratory. However, we confirmed in another study that E, P and N-cadherin proteins are expressed in tissue lysates of bovine placenta (unpublished data). Therefore, to determine the distribution of beta-catenin and type I classical cadherins i.e. E-, P-, and N-cadherin, we used polyclonal or monoclonal beta-catenin, E-cadherin, N-cadherin, and P-cadherin antibodies developed for use in several mammalian tissues.
Negative and positive controls were used to control the specificity of the immunostaining of the cadherin and beta-catenin proteins. Archived blocks of the bovine uterus, placenta, and abomasum served as positive controls for E-, P-, and N-cadherin, and beta-catenin. Archived paraffin blocks of the bovine ovary and liver were also stained for N-cadherin. Normal rabbit IgG (Santa Cruz sc-2027) instead of pan-cadherin, and E- and P-cadherin and normal mouse IgG (Santa Cruz sc-2025) instead of anti-N-cadherin and anti-beta-catenin antibodies were used as negative controls. No specific immunostaining was detected in the negative control sections of the cervix and vagina when a normal rabbit or mouse IgG was used instead of primary antibodies (Fig. 2A-D). Whereas, the positive control tissues were immunopositive for cadherins and β-catenin (Fig. 2E-H). These results indicate that the commercial antibodies employed in the present study were suitable for use in bovine tissues.
The semi-quantitative evaluation of immunostainings
Immunostainings for E-, P-, N-cadherin and beta-catenin were evaluated semi-quantitatively using a four-point intensity score (IS) (Detre et al 1995). Positive immunostaining for all cadherins and beta-catenin were determined in high-expression areas by scanning the cervical and vaginal sections at magnifications of X40, X100, X200, and X400. The staining was scored as (−) negative, (+) weak, (++) moderate, or (+++) strong. The subcellular, cellular and tissue localizations of E-, P-, N-cadherin and beta-catenin were evaluated independently for three tissue layers (epithelium, stroma and smooth muscle layer) and blood vessels in the cervix and vagina. The serosa was present in only some cervical sections as it was lost during the fixation and embedding procedures and is, therefore, not included in the results. Furthermore, in the present study, the terminology described by Mullins and Saacke (1989) was used to define the location of the epithelial cells in the cervical mucosa. The epithelium was defined according to its location in the central lumen, primary folds, secondary folds, and grooves, Firstly, the epithelium surrounding the cervical lumen and lining the longitudinal primary folds was described as the central region epithelium, while the epithelium covering the secondary folds was called as the peripheral region epithelium. Secondly, the epithelium was defined according to its location in the grooves. While the term “basal area” was used to identify areas, where epithelial cells were within grooves, the term “apical area” described areas, where epithelial cells were situated between grooves. Accordingly, epithelial immunostaining was evaluated in the epithelia of the central canal, primary and secondary folds, and the apical and basal areas of the grooves.