
==== Front
Biol Reprod
Biol Reprod
biolreprod
Biology of Reproduction
0006-3363
1529-7268
Oxford University Press

38847468
10.1093/biolre/ioae090
ioae090
Research Article
AcademicSubjects/MED00773
AcademicSubjects/SCI01070
Induction of in vivo-like ciliation in confluent monolayers of re-differentiated equine oviduct epithelial cells†
Leemans Bart Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands
Department of Internal Medicine, Reproduction, Population Health, Faculty of Veterinary Medicine, Ghent University, Ghent, Belgium

Gadella Bart M Department of Internal Medicine, Reproduction, Population Health, Faculty of Veterinary Medicine, Ghent University, Ghent, Belgium
Biomolecular Health Sciences, Faculty of Veterinary Medicine, Utrecht University, Utrecht,The Netherlands
Population Health Sciences, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands

Marchand Josephine H E A M Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands

Van Soom Ann Department of Internal Medicine, Reproduction, Population Health, Faculty of Veterinary Medicine, Ghent University, Ghent, Belgium

Stout Tom A E Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands

Correspondence: Yalelaan 7, 3584 CM Utrecht, The Netherlands. E-mail: b.m.gadella@uu.nl
9 2024
07 6 2024
07 6 2024
111 3 580599
15 1 2024
09 4 2024
05 6 2024
15 6 2024
© The Author(s) 2024. Published by Oxford University Press on behalf of Society for the Study of Reproduction.
2024
https://creativecommons.org/licenses/by-nc/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (https://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact journals.permissions@oup.com

Abstract

We recently developed re-differentiated equine oviduct epithelial cell (REOEC) monolayers demonstrating various in vivo morphological characteristics, but lacking secondary ciliation. In this study, we evaluated the effects of fetal bovine serum, reproductive steroid hormones, Wnt- and Notch ligands and inhibitors, and different EOEC seeding densities, in both conventional wells and on microporous membranes, on EOEC morphology and, in particular, secondary ciliation. REOEC monolayers were assessed by confocal microscopy after combined staining of nuclei, cilia, and the cytoskeleton. Only Wnt ligands, Notch inhibitors and oviduct explant cell concentration affected EOEC morphology. Undesirable epithelial-mesenchymal transition was observed in REOEC monolayers exposed to Wnt3a containing medium and Wnt ligand CHIR 99021. With respect to secondary ciliation, only the combined effect of oviduct explant cell concentration and Notch inhibition steered REOEC monolayers to in vivo-like ciliation patterns. De-differentiated EOECs, formed 10 days after oviduct explant cell seeding, were reseeded on inserts; only at initial oviduct explant cell concentrations of 1 and 5 × 106 cells per well was the formation of REOEC monolayers with a high rate of diffuse ciliation supported. Within 1 month after air-liquid interface introduction, >40% and >20% of the REOECs showed secondary cilia, respectively. At higher oviduct explant cell seeding densities secondary ciliation was not supported after re-differentiation. Additionally, Notch inhibition helped boost secondary ciliation rates to >60% in REOEC monolayers with diffuse ciliation only. These monolayers demonstrated higher clathrin expression under follicular phase conditions. Overall, the ciliated REOEC monolayers better resemble in vivo oviduct epithelial cells than previous models.

An equine in vitro oviduct epithelium model showing in vivo-like secondary ciliation was established in Transwell inserts using a de-differentiation/re-differentiation protocol.

Graphical Abstract

Graphical Abstract

oviduct
horse
ciliation
in vitro model
Research Foundation Flanders 10.13039/501100003130 12I0517N EU COST Action 16119
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pmcIntroduction

To study oviduct physiology, sperm capacitation, fertilization, and early embryonic development in the horse and other mammalian species, in vivo studies of oviduct function are unrealistic because they would require invasive surgery or laparoscopy to access the female’s oviduct. To examine how the oviduct supports these events it is, therefore, desirable to establish a representative in vitro oviduct model. In man, an immortalized oviduct epithelial cell line was established to study these events because of the restricted availability of primary human oviduct epithelial cells [1]. In the cow, pig, horse, cat, and dog oviductal organoids have been established and characterized [2, 3]. In the horse, previous studies have mainly focused on non-polarized monolayers of de-differentiated oviduct epithelial cells [4, 5] and oviduct explants [6]. Unfortunately, neither of these systems was sufficient to support complete sperm capacitation and fertilization. While the oviduct explant model was sufficient to trigger three critical physiological hallmarks of stallion sperm capacitation, namely increased intracellular pH [7], protein tyrosine phosphorylation in the sperm tail [7–9] and hyperactivated motility [10], it was not able to reliably induce the acrosome reaction and fertilization. Similarly, the percentage of oocytes fertilized by intracytoplasmic sperm injection that developed to blastocysts was not affected by the presence of oviduct explants in the embryo culture medium [11]. In order to improve in vitro oviduct cell culture in terms of phenotype and functionality, our lab and others have developed long term 3D oviduct epithelial cell monolayers for mouse, pig, cow, and horse [12–15]. A monolayer of polarized, differentiated oviduct epithelial cells was established by introducing an air-liquid interface [12, 15]. Moreover, cycle stage-specific hormones stimulated in vivo-like changes in these monolayers [14–17].

Using a direct seeding and a de-differentiation/re-differentiation protocol we recently developed confluent monolayers of polarized, differentiated equine oviduct epithelial cells (EOEC) in Transwell inserts and microfluidic chips [15]. Cultured EOEC monolayers proved to be responsive to cycle stage-specific concentrations of reproductive steroid hormones, in terms of nuclear progesterone receptor expression and sperm binding capacity. Interestingly, using the direct seeding protocol, >60% and 28% of EOECs showed single, non-motile (further referred to as “primary cilia”) and multiple, motile cilia (further referred to as “secondary cilia”) on the apical surface, respectively. Secondary cilia were organized in a random, widely dispersed but evenly distributed (referred to below as “diffuse”) pattern within EOEC monolayers. By contrast, >99% DEOECs/REOECs demonstrated primary cilia, but rarely showed secondary cilia in this culture system. Indeed, over 90% of the monolayers showed <1% secondary ciliated REOECs, while the remaining <10% of the re-differentiated monolayers showed 11–27% EOECs with secondary cilia in a diffuse pattern [15]. Nevertheless, the equine oviduct model exhibited functions of, and close morphological resemblance to, in vivo oviduct epithelium, with the exception of EOEC secondary ciliation rates; neither protocol resulted in secondary ciliation rates similar to the in vivo situation (up to 60% ciliation; [15, 18]).

In an attempt to develop an improved horse oviduct cell culture model, the current study aimed to establish REOEC monolayers, using EOECs originating from the ampulla, with a consistent in vivo-like secondary ciliation pattern. We investigated whether fetal bovine serum (FBS) and reproductive steroid hormones at blood or oviduct tissue levels would support secondary ciliation. Next, modulation of cell differentiation and proliferation by the Wnt- and Notch-pathways in EOEC re-differentiation were assessed by exposing EOEC monolayers to ligands and inhibitors of both. We assessed whether the concentration of harvested oviduct epithelial cells seeded into conventional wells to obtain de-differentiated equine oviduct epithelial cells (DEOECs) and, alternatively, the concentration of DEOECs seeded onto the microporous membrane to establish monolayers of REOECs, affected the subsequent process of re-differentiation to secondary ciliated EOECs. Finally, the effect of cycle stage-specific hormones on the expression of the secretion-associated protein, clathrin, by in vivo-like patchy ciliated REOEC monolayers was assessed.

Materials and methods

Chemicals and reagents

Culture media (DMEM/Ham’s F12 and M199), penicillin/streptomycin, gentamycin, and trypsin were purchased from Gibco by Life technologies (Thermo Fisher Scientific, Waltham, Massachusetts, USA). Wnt-inhibitor N-(6-Methyl-2-benzothiazolyl)-2-[(3,4,6,7-tetrahydro-4-oxo-3-phenylthieno [3,2-d]pyrimidin-2-yl)thio]-acetamide (IWP-2) and gamma-secretase or Notch-inhibitor N-[(1S)-2-[[(7S)-6,7-Dihydro-5-methyl-6-oxo-5H-dibenz[b,d]azepin-7-yl]amino]-1-methyl-2-oxoethyl]-3,5-difluoro-benzeneacetamide (DBZ) were purchased from Sigma Aldrich (I0536; 565 789). Jagged-1 protein (JAG1), a Notch-ligand, was purchased from Genscript (RP20331, Piscataway, New Jersey, USA). Wnt-agonist 6-[[2-[[4-(2,4-Dichlorophenyl)-5-(5-methyl-1H-imidazol-2-yl)-2-pyrimidinyl]amino]ethyl]amino]-3-pyridinecarbonitrile (CHIR 99021) was purchased from Selleckchem.com (S2934, Munich, Germany). All other chemicals were purchased from Merck and Sigma-Aldrich (Darmstadt, Germany) unless otherwise indicated.

Animals

Equine oviducts were collected at a local slaughterhouse no more than 30 minutes after the slaughter of healthy mares aged between 4 and 25 years (average 16 years), and without any visible reproductive tract pathologies. Oviducts from mares that had one or more large follicles and no corpus luteum on the ovaries, together with uterine edema, were considered to have been harvested during the follicular phase; oviducts collected from mares with a corpus luteum and without uterine edema were considered to come from the luteal phase. The oviducts of each mare were processed separately. In total, 136 oviducts were collected from 68 mares to perform the experiments described.

Harvesting and culturing EOECs

EOECs were harvested as described by Leemans et al. [15]. Briefly, after collection, oviducts were transported on ice to the laboratory in tubes filled with Dulbecco’s Phosphate Buffered Saline (PBS; 0.9 mM CaCl2, 2.68 mM KCl, 1.47 mM KH2PO4, 0.5 mM MgCl2, 137 mM NaCl, 8.1 mM Na2HPO4; pH: 7.30, 300 ± 10 mOsmol/kg; Gibco BRL, Paisley, UK) supplemented with penicillin/streptomycin (both 100 μg/mL) and gentamycin (50 μg/mL), arriving within 2 hours. The oviducts were dissected free from surrounding connective tissue, straightened as much as possible, and washed three times in the antibiotic-supplemented PBS. In a flow cabinet, the ampulla was opened longitudinally with scissors and the inner lining of the oviduct, including the mucosa, was scraped using the blunt side of a scalpel blade. The harvested EOECs were transferred into a petri dish containing M199 medium supplemented with 20% FBS, penicillin and streptomycin (both 100 μg/mL) and gentamycin (50 μg/mL). The suspension was centrifuged for 5 minutes at 200 g at room temperature. The cell pellet was resuspended in DMEM/F12 medium containing 10% FBS, 10 ng/mL epidermal growth factor (EGF), 5 μg/mL insulin, 5 μg/mL apo-transferrin, 100 μg/mL penicillin/streptomycin, 50 μg/mL gentamycin and 1 μg/mL amphotericin B (adapted from previous publications [4, 19, 20]). The cells were then cultured for 24 hours in 6-well plates (Greiner Bio-one, Alphen aan den Rijn, The Netherlands). During this 24 hour culture, the cells arranged themselves into floating vesicles or “oviduct explants” with outward facing, actively beating cilia, while contaminating mesodermal fibroblast cells attached to the bottom of the polystyrene 6-well plates. The oviduct epithelial vesicles were collected and centrifuged at 200 g for 5 minutes at 25°C and resuspended in DMEM containing 10% FBS culture medium. After thorough mixing of the pellet by pipetting, an aliquot of the cell suspension was removed to assess cell concentration and viability. Cell concentration was determined using a hemocytometer chamber. For viability assessment, the cell suspension was incubated with trypan blue solution (0.4% w/v in 0.85% w/v NaCl in water; Gibco BRL, Paisley, UK) at a ratio of 1:1 (v:v) for 4 minutes at room temperature. At least 100 oviduct explant cells per donor were assessed. Blue stained cells were considered to be dead, whereas white to grey cells were considered alive. Ten million live oviduct explant cells were seeded per well; the wells had a surface area of 9.6 cm2.

For this study, the de-differentiation/re-differentiation protocol was applied (a schematic overview of this protocol is shown in Figure 1B of Leemans et al. [15]). The oviduct explants were incubated for 10 days, to allow attachment of the EOECs to the bottom of a 6-well plate and the formation of DEOEC (flattened and de-ciliated) monolayers. After this incubation period, the DEOEC monolayers were detached using 0.25% w/v Trypsin/0.1% w/v EDTA (trypsin/ethylenediaminetetraacetic acid) in PBS. After a 10–15 minutes incubation at 38°C in an incubator with a rocking platform, trypsin activity on the DEOECs was blocked by adding M199 medium containing 20% FBS. The detached DEOECs were collected and subsequently washed by centrifugation-resuspension in the same medium (200 g, 5 minutes). These DEOECs were then seeded onto the microporous membrane inside hanging Transwell inserts in DMEM culture medium containing 10% FBS at 0.5 × 105 EOECs per 0.33 cm2. Each insert was seeded with EOECs from a single mare. The volume of culture medium in the Transwell was 800 μL in the basal section and 200 μL in the apical part. The oviduct culture medium was refreshed completely twice a week. After 3 days, an air-liquid interface was introduced to allow re-differentiation of the EOEC monolayers. Any culture medium leaking from the basolateral to the apical compartment was removed daily until membrane confluency was reached 8–10 days after cell seeding (5–7 days after air-liquid interface introduction). Cells were cultured in a humidified atmosphere of 5% CO2-in-air at 38.5°C for up to 60 days after introduction of the air-liquid interface.

Figure 1 (a) Effect of cycle stage on the presence of secondary cilia in oviducts retrieved from mares in the luteal and follicular phases. Approximately 60% of the equine oviduct epithelial cells (EOECs) lining the ampullary oviductal lumen demonstrated the presence of secondary cilia regardless of whether oviducts were retrieved from mares in the luteal or follicular phase. Data are mean (± SD) percentage of EOECs displaying secondary cilia (n = 3 mares per cycle stage; 5 oviduct tissue sections per mare). (b) Representative immunohistochemistry pictures from oviduct sections in the (A) luteal and (B) follicular phase showing that more than 50% of cells of the in vivo oviduct epithelium are ciliated (secondary cilia are stained in red with the as visualized with the AEC chromogen) (scale bar: 20 μm).

Immunohistochemistry of oviduct tissue sections to assess secondary ciliation rate

Reference samples for the phenotype that the cultured REOEC monolayers should resemble were collected from three oviducts in the luteal and three in the follicular phase of the estrous cycle. The ampullae of these oviducts were dissected from any other tissue and fixed in 4% w/v formaldehyde (Klinipath, 4186, Olen, Belgium) for 3 days. Next, the ampulla tissue was embedded in paraffin, and 3 μm sections were subsequently cut and mounted on Superfrost plus glass slides. Tissue sections were successively deparaffinized with xylene (3 times 5 minutes), decreasing concentrations of ethanol (first 100%, then 50%; 2 times 10 minutes per concentration) and demi water. A final washing step was performed with PBS. Subsequently, samples were probed without antigen retrieval for the presence of cilia and counterstained with hematoxylin–eosin.

A DAKO pen (Dako, S2002, Heverlee, Belgium) was used to create a hydrophobic barrier to keep all liquids on the slide. Oviduct sections were treated for 5 minutes with endogenous peroxidase activity blocking solution (S2023; Dako, Heverlee, Belgium). Slides were then washed twice with PBST for 5 minutes and blocked with normal goat serum for 15 minutes. Next, the monoclonal mouse anti-acetylated alpha tubulin antibody (sc-23 950, Santa Cruz Biotechnology; 1:200 in normal antibody diluent, 33 593; Immunologic, Duiven, The Netherlands) was added for 1 h at room temperature. After three washing steps, the polyclonal goat anti-mouse/anti-rabbit radish peroxidase-conjugated secondary antibody (code: VWRKDPVO110HRP; Immunologic, Duiven, The Netherlands) was added to the slides at the concentration indicated by the manufacturer and incubated for 30–60 minutes at room temperature. The slides were washed again in PBST before 2 drops of the chromogen AEC (K3464; Dako, Heverlee, Belgium) were added. After a 20-min incubation, the slides were rinsed with demineralized water for 5 minutes. Oviduct sections were subsequently counterstained with hematoxylin–eosin (HE; J.T.Baker, 3870, Landsmeer, The Netherlands). The sections were initially stained with hematoxylin for 4 minutes and rinsed with water for 5 minutes. Next, the sections were incubated successively for 1 minute in 70% and 96% ethanol. The sections were then stained with eosin Y for 1 minute before dehydration by immersion in 96% ethanol (twice for 1 minute) and xylene (twice for 1 minute). The slides were rinsed with tap water for 5 minutes and sealed under a cover slip with water soluble glue (Aquatex; Merck, Amsterdam, The Netherlands).

Brightfield images were taken using an Olympus BX60 microscope at magnifications of 200x, 400x and 1000x. The images were analyzed using Fiji software [21]. At three different locations on secondary folds of the reference tissue, 100 cells were assessed for the presence of secondary cilia. The average value of these three locations was considered representative for an individual mare. For comparison, 100 REOECs per monolayer were assessed and the percentage of ciliated cells was determined (see below).

Immunofluorescent staining of oviduct tissue cryosections to assess clathrin expression

Clathrin was used as a marker for secretory activity in the equine oviduct [22–24] via immunofluorescent staining. Three dissected oviducts per cycle stage from mares in the luteal and follicular phase were cut into pieces of approximately 1.5 cm in length and frozen in Neg-50 frozen section medium (VWR International B.V., Amsterdam, The Netherlands) in liquid nitrogen. Next, five to ten μm thick sections were cut at −20°C on a cryostat (Model CM 3050S, Leica, Nussloch, Germany), and mounted on Superfrost glass slides (VWR International B.V., Amsterdam, The Netherlands). The tissue was fixed in acetone at 4°C for 10 minutes before permeabilization and blocking using PBS containing 0.5% Triton X-100 and 5% donkey serum for 30 minutes. The cryosections were then incubated with polyclonal goat anti-clathrin primary antibodies (sc-6579, Santa Cruz Biotechnology; 1:100 in normal antibody diluent (33 593; Immunologic, Duiven, The Netherlands)) for 1 hour followed by a polyclonal donkey anti goat-Alexa fluor 488 secondary antibody (sc-45 091, 1:100 dilution with PBS, Santa Cruz Biotechnology, Immunologic, Duiven, The Netherlands) for 2 hours at room temperature. The nuclei and F-actin were stained for 1 hour with 5 μg/ml Hoechst 33342 and phalloidin-Alexa Fluor 568 (1:100 dilution), respectively. The sections were mounted in a 1:1 solution of Vectashield (Vector Laboratories, Burlingame, USA) and PBS, covered with a coverslip and sealed with nail polish. Sections were imaged using a fluorescence microscope (BX60, Olympus, Zoeterwoude, The Netherlands) equipped with a CCD camera (Leica DFC425C) connected to LAS-AF software (Leica Microsystems GmbH, Wetzlar, Germany) at 100–400x magnification. The light source was an UV lamp (EL-6000, Leica) and filters were used for blue (wavelengths for filters were, for excitation: 340–380/20 nm, dichroic mirror: 400 nm, emission: >425 nm), green (excitation: 445–535 nm, dichroic mirror: 510 nm, emission: >515 nm), and red fluorescence (excitation: 515–560 nm, dichroic mirror: 580 nm, emission: >590 nm). During imaging of samples, exposure time, gain, and gamma values for the green fluorescence (clathrin labeling) were kept constant between stained samples and the corresponding negative controls, while settings for blue and red fluorescence were adjusted to optimize the balance of the color in the overlay images.

Detection of confluency in REOEC monolayers by trans-epithelial electrical resistance

To quantitatively assess the confluency of the epithelial monolayer, transepithelial electrical resistance (TEER) measurements were performed as described by Chen et al. [25]. Electrodes (World Precision Instruments GmbH, Friedberg, Germany) were equilibrated and sterilized for 10 minutes in 70% ethanol and connected to a digital volt-ohm meter (Millicell, Billerica, Massachusetts, USA) according to the manufacturer’s recommendations. To eliminate the influence of temperature and atmospheric conditions, measurements were performed immediately after changing the DMEM +10% FBS culture medium with antibiotic supplemented M199 + 20% FBS on a heated plate at 38°C. Using this protocol, no reading drift was observed. Electrodes were inserted into both the apical and the basolateral compartments. After 1 minute of stabilization, the electrical resistance was recorded. The electrical resistance of a blank insert (without cells) was measured in parallel. To obtain the TEER (in Ω*cm2), the blank value was subtracted from the total resistance of the sample, and the final unit area resistance (Ω*cm2) was calculated by multiplying the sample resistance by the effective area of microporous membrane onto which the EOECs were attached (insert: 0.33 cm2).

Cell morphology including secondary cilia formation in REOEC monolayers

At designated times after introduction of the air–liquid interface, REOEC monolayers cultured in Transwell inserts were assessed for cell morphology including secondary cilia formation. Three Transwell inserts were used per animal (n = 3) per experiment, and each experimental condition was tested three times. REOEC monolayers containing a confluent monolayer of re-differentiated polarized EOECs were fixed in 4% w/v paraformaldehyde in PBS for 15 minutes at room temperature and rinsed twice with PBS for 5 minutes. Subsequently, REOEC monolayers were permeabilized and incubated with PBS containing 0.5% Triton X-100 and 5% normal goat serum for 30 minutes at room temperature to block non-specific binding. Next, the monolayers were washed twice in PBS containing 0.5% Triton X-100 for 5 minutes, before overnight incubation at 4°C with a monoclonal mouse anti-acetylated α-tubulin primary antibody (sc-23 950, 1:100 dilution, Santa Cruz Biotechnology, Immunologic, Duiven, the Netherlands). The next morning the REOEC monolayers were washed three times in PBS containing 0.5% Triton X-100 (5 minutes per wash) and incubated with a polyclonal Alexa 488 conjugated goat-anti-mouse antibody (sc-362 257, 1:100 dilution with PBS, Santa Cruz Biotechnology, Immunologic, Duiven, The Netherlands) at room temperature for two hours. REOEC monolayers were again washed three times with PBS containing 0.5% Triton X-100 for 5 minutes. Subsequently, cell nuclei and actin filaments were stained using, respectively, 5 μg/ml Hoechst 33342 and Alexa 568-conjugated phalloidin (1:100 dilution) in PBS containing 0.5% Triton X-100. After 1 hour incubation, the REOEC monolayers were washed twice with PBS for 5 minutes and excised from the hanging insert. Excised REOEC monolayers were mounted on glass slides (Marienfeld, Germany) using Vectashield (1/3 dilution, Vector Laboratories, Burlingame, USA) as antifade, and sealed with nail polish. Negative controls were performed by omitting the primary antibody. Imaging was performed using a TCS SPE-II laser scanning confocal microscopy system (Leica Microsystems GmbH, Wetzlar, Germany) attached to an inverted semi-automated DMI4000 microscope (Leica) with a 10x objective with a numerical aperture (NA) of 0.25 and 40x objective with a numerical aperture (NA) of 1.25. Z-stacks with 0.2 μm step size and 1.5x digital zoom were obtained. The 3D constructs of the monolayer EOECs were performed using ImageJ software (National Institutes of Health, Bethesda, MD, USA) to demonstrate and assess different aspects of cell morphology including ciliation patterns (diffuse or patchy), cellular length and nuclear diameter; and cell density per mm2. In addition, two image stacks per timepoint, donor, cycle stage or treatment condition were obtained using the 40x lens to quantify the percentage of ciliated REOECs. A mean of 166 REOECs (minimum: 105, maximum: 209) per stack were evaluated. Five randomly selected fields of view in the center of the membrane were imaged for each monolayer and, at least, 800 REOECs per membrane were classified. The images were imported into Imaris 8.2 software (Bitplane AG, Zurich, Switzerland) to determine the percentage of ciliated REOECs on each membrane. The ratio of nuclei without overlying cilia to the total number of nuclei was assessed.

Mimicking cycle stage by exposing REOEC monolayers to steroid hormones

Reproductive steroid hormone concentrations resembling the concentrations found in blood and oviduct tissue during the follicular and luteal phases [26–28] were used to mimic an estrous cycle. From the time of air–liquid interface introduction, REOEC monolayers were cultured in DMEM containing 10% FBS culture medium supplemented with estradiol 17-β (E2; P8783–16, Sigma, Zwijndrecht, The Netherlands) and progesterone (P4; E2758–16, Sigma, Zwijndrecht, The Netherlands) in concentrations chosen to mimic the luteal and follicular stages of the mare’s estrous cycle. To this end, the medium in the basal compartment was supplemented for 14 days with 20 pg/mL E2 and 10 ng/mL P4 (blood concentration); or 10 ng/mL E2 and 1000 ng/mL P4 (oviduct concentration) to mimic the luteal phase. To subsequently mimic the follicular phase, hormone concentrations were changed to 40 pg/mL E2 and 0 ng/mL P4 (blood concentration); or 80 ng/mL E2 and 40 ng/mL P4 (oviduct concentration) for seven consecutive days. Ethanol was used as the solvent for hormone supplementation with a final ethanol concentration of 1%. For the negative controls, 1% ethanol was added to the culture medium. Culture medium was refreshed every other day.

Clathrin expression in steroid hormone-exposed REOEC monolayers

To assess the effect of steroid hormones in clathrin expression in patchy ciliated REOEC monolayers, a modification of the staining protocol described above for cell morphology was performed. Firstly, the secondary antibody used to label cilia was a goat anti-mouse anti-acetylated α-tubulin antibody-Alexa Fluor 647 conjugate (1:100 dilution with PBS; Santa Cruz Biotechnology, Immunologic, Duiven, the Netherlands). Secondly, an additional indirect immunofluorescent staining step was included to label clathrin. After cilia labeling, the monolayers were washed twice in PBS containing 0.5% Triton X-100 for 5 minutes, and subsequently blocked for 30 minutes with PBS containing 0.5% Triton X-100 and 5% donkey serum. Next, the monolayers were again washed twice in PBS containing 0.5% Triton X-100 for 5 minutes, before overnight incubation at 4°C with a polyclonal goat anti-clathrin primary antibody (sc-6579, Santa Cruz Biotechnology; 1:100 dilution, 33 593; Immunologic, Duiven, The Netherlands). The next morning, the REOEC monolayers were washed three times in PBS containing 0.5% Triton X-100 (5 minutes per wash) and incubated with a polyclonal Alexa 488 conjugated donkey-anti-goat antibody (sc-45 091, 1:100 dilution with PBS, Santa Cruz Biotechnology, Immunologic, Duiven, the Netherlands) at room temperature for two hours. Phalloidin and Hoechst 33342 were again used to stain the cytoskeleton and the nuclei; and imaging was identical to above. For quantification of clathrin expression in patchy ciliated REOEC monolayers, five images per monolayer were analyzed by evaluating fluorescence intensity for clathrin and DNA in each area using ImageJ software (National Institutes of Health, Bethesda, MD, USA). After maximum projection reconstruction of Z-stacks, the fluorescence intensity of each channel was measured. The average intensity of fluorescence for clathrin was then normalized by dividing the clathrin intensity by Hoechst 33342 fluorescence to normalize for DNA content.

Experiments to steer secondary cilia formation in an EOEC de-differentiation/re-differentiation protocol

Various protocols were tested to support complete in vivo-like re-differentiation of EOEC monolayers cultured in Transwell inserts, i.e., in vivo-like morphology including the formation of secondary cilia:

(1) In a first approach, the effect of FBS concentration was examined 1 and 2 months after air-liquid interface introduction. From the start of air-liquid interface introduction, oviduct culture medium was enriched with 2.5, 5, 10 or 20% FBS.

(2) Next, the effect of cycle stage-specific hormones was assessed. After confluency was obtained, 21 days after air-liquid interface introduction, oviduct culture medium was supplemented with estradiol and progesterone at blood and oviduct tissue concentrations mimicking the luteal and follicular phases sequentially (see above). More specifically, REOEC monolayers were initially exposed for 14 days to luteal and then 7 days to follicular phase progesterone/estradiol concentrations. The results were compared to EOEC secondary ciliation rates in oviduct tissue sections from mares in the luteal and follicular phases.

(3) We next examined whether EOEC re-differentiation was supported by stimulation and/or inhibition of the Wnt and Notch pathways. For this, confluent REOEC monolayers were exposed for a week to either Wnt or Notch ligands/inhibitors, starting 3 and 7 weeks after air liquid-interface introduction. These triggers were tested in specific concentration ranges [29]. Initially, a Wnt inhibitor (IWP 2: 0, 2 or 20 μM) and ligand (CHIR 99021: 0, 1, 3, 5, or 10 μM) and Notch inhibitor (DBZ - γ secretase inhibitor: 0, 1, or 10 μM) and ligand (JAG-1: 0, 1, or 10 μM) were added individually to the cell culture medium via the basolateral compartment of the Transwell insert. Additionally, the effect of Wnt3a containing medium was assessed. This medium contains Wnt3a secreted by a genetically modified cell line (mouse L cells - American Type Culture Collection (ATCC) CRL-2648) into DMEM culture medium [30, 31]. For experimental use, oviduct cell culture medium was enriched with 0, 30, 50, 70, or 100% Wnt3a containing medium.

(4) The effect of experiment, EOEC donor and cycle stage of the EOEC donor were verified because REOEC monolayers with a diffuse ciliation pattern (>10% REOECs with secondary cilia) were only obtained occasionally.

(5) After the observation that secondary cilia formation in a diffuse pattern occurred only in specific experiments and in areas of the REOEC monolayer with increased cell density, the effect of the seeding concentration of trypsinized DEOECs on membrane inserts was assessed. On a membrane surface of 0.33 cm2, 0.1 × 105, 0.5 × 105, 1 × 105, 2.5 × 105, 5 × 105 and 10 × 105 trypsinized DEOECs were seeded.

(6) In addition, the effect of initial oviduct explant cell concentration used to obtain DEOEC monolayers in conventional wells, on the subsequent formation of spontaneously REOEC monolayers was evaluated. After one day preculture, oviduct explants were seeded at different concentrations, namely 1 × 106, 5 × 106, 10 × 106 and 30 × 106 cells in 4 mL culture medium in wells with a surface area of 9.6 cm2.

(7) Since seeding 1 and 5 × 106 oviduct explant cells per well resulted in diffuse ciliated EOEC monolayers after re-differentiation, the effect of cycle stage-specific hormones and Wnt and Notch ligands/inhibitors were re-assessed. Identical experimental conditions were applied as tested above on REOEC monolayers with a low ciliation rate (less than 2%).

(8) The combined effect of oviduct explant cell concentration during de-differentiation and Notch inhibition during re-differentiation was examined. REOEC monolayers exhibiting an in vivo-like patchy ciliation pattern were obtained by the following protocol: (1) oviduct explant cell seeding concentration of up to 5 × 106 cells per 9.6 cm2 well to first establish a monolayer of DEOECs in 10 days, (2) the trypsinized and reseeded DEOECs were re-differentiated for 3 weeks by air-liquid interface introduction and (3) exposed to 10 μM DBZ for one more week.

(9) The ciliation status of in vivo-like patchy ciliated REOEC monolayers was assessed 0, 7, and 14 days after the end of DBZ treatment.

(10) Finally, the expression of the secretory marker clathrin was assessed in patchy ciliated REOEC monolayers after exposure to cycle stage-specific hormones at oviduct tissue level concentrations. Similar hormone concentrations were used to those tested above on REOEC monolayers with a low ciliation rate.

Statistical analysis

All experiments were performed at least three times, and EOECs for a given comparison were derived from three different Warmblood mares. Before analysis, normality of the variables was checked using the Shapiro–Wilk and Kolmogorov–Smirnov tests (P < 0.05). The effect of different de- or re-differentiation conditions, i.e., FBS concentration, cycle stage-specific hormone supplementation, Wnt/Notch agonists and inhibitors, oviduct explant cell concentration to obtain DEOECs or DEOEC seeding concentration to obtain REOEC monolayers, and clathrin expression levels, were assessed by analysis of variance (ANOVA). Overall differences were identified using repeated measures ANOVA with Greenhouse–Geisser and Bonferroni corrections, as implemented in the general linear model. Scheffé post-hoc tests were performed for pairwise comparisons. Differences were considered significant if P < 0.05. All analyses were performed using SPSS version 20 for Windows (SPSS IBM, Brussels, Belgium).

Results

About 60% of the horse oviduct ampullary epithelial cells show secondary ciliation, independent of cycle stage

In vivo reference tissue sections obtained from luteal and follicular phase oviducts demonstrated that the epithelial lining contains 55.0 ± 8.6% and 58.9 ± 7.7% secondary ciliated EOECs, respectively (Figures 1a and 1b). These in vivo ciliation rates were subsequently used as the control reference level of ciliation for all in vitro experiments with REOEC monolayers.

Different FBS concentrations and estrous cycle mimicking hormone regimes did not affect secondary cilia formation

Exposing EOEC monolayers to culture medium supplemented with 2.5, 5, 10, or 20% FBS during re-differentiation did not affect secondary cilia formation after incubation for 1 or 2 months. Overall, less than 2% of the EOECs from re-differentiated monolayers demonstrated secondary cilia (further referred as “poorly ciliated REOEC monolayers”), independent of FBS concentration (Figure 2a).

Figure 2 (a) Effect of different concentrations of FBS in culture medium on the formation of secondary cilia in re-differentiating equine oviduct epithelial cells (EOECs) assessed one and two months after air-liquid interface (ALI) introduction. Secondary ciliation rates after 1 and 2 months of culture were low for all FBS concentrations tested. Data are mean (± SD) percentage of EOECs displaying secondary cilia (n = 3 mares; 3 inserts per mare per FBS concentration). (b) Effect of estradiol and progesterone to mimic estrous cycle phase on secondary cilia formation in re-differentiating EOECs, after 14 days of luteal (blood concentration: 20 pg/mL E2 and 10 ng/mL P4; oviduct concentration: 10 ng/mL E2 and 1000 ng/mL P4) and 7 days of subsequent follicular phase hormone exposure (blood concentration: 40 pg/mL E2 and 0 ng/mL P4; oviduct concentration: 80 ng/mL E2 and 40 ng/mL P4). Ethanol was used as the solvent for hormone supplementation at a final ethanol concentration of 1%. Tested hormone conditions did not have an effect on secondary ciliation rates and did not differ from the control treatment (culture without hormones). Data are mean (± SD) percentage of secondary ciliated EOECs (n = 3 mares; 3 inserts per mare per experimental condition).

Similarly, addition of reproductive steroid hormones to EOEC culture medium at concentrations designed to mimic blood and oviduct tissue concentrations at different stages of the estrous cycle had no effect on secondary cilia formation. Oviduct tissue hormone concentrations mimicking the luteal and follicular phases did not result in more than 2% of REOECs developing secondary cilia 14 days after luteal (1.2 ± 0.4% ciliated EOECs) and 7 days after subsequent follicular phase-specific hormone exposure (0.6 ± 0.3% ciliated EOECs) (Figure 2b). Similar results were obtained for the percentage of secondary ciliated REOECs in monolayers exposed to culture medium containing hormones at blood concentrations (luteal phase: 1.0 ± 0.4% ciliated EOECs; follicular phase: 0.7 ± 0.2% ciliated EOECs) (Figure 2b). The control medium containing 1% ethanol (1.0 ± 0.3% ciliated EOECs) and medium without hormone or ethanol supplementation (0.9 ± 0.4% of ciliated EOECs) also had low levels of secondary cilia formation. These percentages of ciliated cells are in marked contrast to those observed in in vivo reference tissue sections (Figure 1).

Wnt stimulation induced a concentration-dependent epithelial-mesenchymal transition in REOEC monolayers, associated with decreased monolayer confluency

The Wnt and Notch pathways are well-known for their roles in cell differentiation and proliferation [32–34]. No effect on cytoskeleton or nucleus morphology was observed using 0 or 30% Wnt3a containing medium for 1 month (Figures 3aA and 3aB). Moreover, TEER values (0% Wnt3a containing medium: 843 ± 53 Ω/cm2 and 30% Wnt3a containing medium: 787 ± 73 Ω/cm2; Figure 3b) remained above the confluency threshold, i.e., 462 Ω/cm2 (as determined by Leemans et al. [25]). Neither did any of the tested concentrations of Wnt inhibitor (IWP-2), Notch agonist (JAG-1) or Notch inhibitor (DBZ) have any adverse effect on cytoskeleton and nucleus morphology. Moreover, TEER measurements after 1 week exposure to these triggers did not drop below the confluency threshold (Figure 3c).

Figure 3 (a) Representative images of re-differentiated equine oviduct epithelial cell (REOEC) monolayers cultured for 1 month in (A) 0%, (B) 30%, (C) 50%, (D) 70% and (E) 100% Wnt3a containing medium or for 7 days in (F) 0 μM, (G) 1 μM, (H) 3 μM, (I) 5 μM, and (J) 10 μM Wnt agonist CHIR 99021. No effect on cell or nucleus morphology was observed after 1 month culture in 30% Wnt3a containing medium. Epithelial-mesenchymal transition was observed within 1 month in re-differentiated EOEC monolayers cultured in 50 and 70% Wnt3a containing medium, with cells becoming elongated with enlarged round to oval nuclei, growing in multiple cell layers, and lacking primary cilia. A similar effect was observed after 1 week exposure to 1, 3, and 5 μM Wnt agonist CHIR 99021. The EOECs detached from the polycarbonate membrane in 100% Wnt3a conditioned medium and 10 μM Wnt agonist CHIR 99021. Moreover, none of the tested conditions supported cilia formation [green (anti-acetylated α-tubulin antibodies-AlexaFluor 488 secondary antibody): primary (single green dots at each EOEC) and secondary cilia (many green dots on some EOECs), red (phalloidin conjugated to Alexa Fluor 568): cytoskeleton, blue (Hoechst 33342): nuclei] (original magnification, 400x – scale bar: 25 μm). (b) The effect on transepithelial electrical resistance (TEER) of confluent re-differentiated EOEC monolayers cultured in 0%, 30%, 50%, 70%, and 100% Wnt3a containing medium after 1, 2, 3, and 4 weeks of incubation is shown. At 3 weeks, TEER values dropped below the confluency threshold (indicated by the horizontal line) in 50% and 70% Wnt3a containing medium; a similar observation was observed after 1 week in 100% Wnt3a containing medium (n = 3 mares; 1 insert per mare per concentration at each time point). Values that differ significantly between TEER values are indicated by small letters. (c) The effect on TEER of confluent re-differentiated EOEC monolayers exposed to CHIR 99021 (Wnt ligand; 0, 1, 3, 5, and 10 μM), IWP-2 (Wnt inhibitor; 0, 2, and 20 μM), JAG-1 (Notch ligand; 0, 1, and 10 μM) and DBZ (Notch inhibitor; 0, 1, and 10 μM) was assessed after 1 week. TEER values dropped below the confluency threshold (indicated by the horizontal line) under 1, 3, 5, and 10 μM CHIR 99021 conditions (n = 3 mares; 1 insert per mare for each concentration). Values that differ significantly between TEER values are indicated by small letters.

Using 50% and 70% Wnt3a containing medium, TEER values did drop below the confluency threshold within 3 weeks of the start of culture (for 50% Wnt3a containing medium TEER values dropped from 752 ± 174 Ω/cm2 to 291 ± 49 Ω/cm2; for 70% Wnt3a containing medium from 825 ± 145 Ω/cm2 to 190 ± 40 Ω/cm2; Figure 3b). This was associated with unfavorable concentration-dependent EOEC epithelial-mesenchymal transition. Cytoskeleton rearrangements and nuclear morphology changes observed were related to de-differentiation of EOECs from columnar to flat and included elongation of EOECs (mean EOEC length almost doubled from 11 ± 3 μm in 0% to 19 ± 4 μm in 50% Wnt3a conditioned medium; Table 1) and enlarged round to oval nuclei (mean nuclear diameter increased almost 1.5 times from 8 ± 1 μm in 0% to 12 ± 3 μm in 50% Wnt3a conditioned medium; Table 1), multiple cell layer growth and a lack of primary cilia (the latter is considered to be an EOEC polarization marker [12, 20]; Figures 3aC and 3aD). Similar observations were made after 1 week exposure to 1, 3, or 5 μM CHIR 99021 (Wnt agonist; Figures 3aG, 3aH, and 3aI). TEER values dropped below the confluency threshold (Figure 3c) with concomitant cytoskeleton and nuclear alterations (Table 1).

Table 1 Quantification of cellular length and nuclear diameter within re-differentiated equine oviduct epithelial cell (EOEC) monolayers to assess the effect of Wnt3a containing medium and Wnt agonist CHIR 99021 on cytoskeleton and nuclear morphology

Wnt agonist	Concentration	Mean EOEC length	Mean nuclear diameter	
Wnt3a containing medium	0%	11 ± 3 μma	8 ± 1 μma	
	30%	10 ± 2 μma	8 ± 1 μma	
	50%	19 ± 4 μmb	12 ± 3 μmb	
	70%	25 ± 6 μmb	15 ± 4 μmb	
	100%	–	–	
CHIR 99021	0 μM	11 ± 2 μma	7 ± 2 μma	
	1 μM	14 ± 3 μmb	8 ± 3 μma	
	3 μM	22 ± 5 μmb	11 ± 4 μmb	
	5 μM	27 ± 6 μmb	13 ± 3 μmb	
	10 μM	–	–	
a,bCellular length and nuclear diameter were compared to their reference condition (0% Wnt3a containing medium and 0 μM CHIR 99021). Overall, EOEC length and nuclear diameter was significantly higher in REOECs cultured for 1 month in 50 and 70% Wnt3a containing medium. Similar observations were observed when REOEC monolayers were cultured for 7 days with 1, 3, 5 μM Wnt agonist CHIR 99021. As an exception, nuclear diameter in EOEC monolayers exposed to 1 μM CHIR 99021 did not differ from the control (0 μM CHIR 99021). No results could be obtained from REOEC monolayers exposed to either 100% Wnt3a containing medium or 10 μM Wnt agonist CHIR 99021 because EOECs detached from the microporous membrane in these conditions.

After the onset of culture for REOEC monolayers in 100% Wnt3a containing medium, TEER values dropped steeply within a week to below the confluency threshold (from 812 ± 56 Ω/cm2 to 128 ± 26 Ω/cm2; Figure 3b). Consequently, unfavorable EOEC epithelial-mesenchymal transition was observed within 2 weeks, followed shortly by complete detachment of the REOEC monolayer from the microporous membrane (Figure 3aE). Interestingly, similar observations were made after 1 week of exposure to 10 μM CHIR 99021 (with TEER values dropping from 733 ± 221 Ω/cm2 to 28 ± 11 Ω/cm2; Figure 3C). In addition, REOECs showed cytoskeleton rearrangements, nucleus morphology changes and subsequent cell detachment within 7 days after the start of exposure (Figure 3aJ).

Secondary cilia formation in REOEC monolayers remained low after incubation in Wnt3a containing medium, or exposure to ligands or inhibitors from the Wnt and notch pathways

Culturing poorly ciliated (<2–3% secondary ciliated cells) REOEC monolayers in 30% Wnt3a containing medium did not induce higher rates of secondary cilia formation, even after long term exposure. After a 1 month culture, 1.1 ± 0.8% of the REOECs showed secondary cilia formation (Figure 3aB) and, after a further month, an average of 2.6 ± 1.6% of the re-differentiated monolayer EOECs demonstrated secondary cilia. Similarly, control medium without Wnt3a medium, and 50 and 70% Wnt3a containing medium failed to support secondary cilia formation (Figures 3aA, 3aC, and 3aD, respectively). REOEC monolayers exposed to 100% Wnt3a containing medium could not be assessed because of EOEC detachment from the membrane (see above; Figure 3aE).

Besides the fact that Wnt activation induced adverse morphological changes, no effect on secondary ciliation rates was observed after one week of exposure to 0, 1, 3, or 5 μM CHIR 99021 starting 3 (Figures 3aF, 3aG, 3aH, and 3aI) or 7 weeks after air-liquid interface introduction. Similar to 100% Wnt3a containing medium, REOEC monolayers exposed to 10 μM CHIR 99021 could not be assessed because epithelial-mesenchymal transition resulted in EOEC detachment from the membrane (see above; Figure 3aJ). Inhibition of the Wnt pathway using 2 or 20 μM IWP-2, and activation or inhibition of the Notch pathway using 1 or 10 μM JAG-1 or DBZ respectively, also failed to support secondary ciliation.

A diffuse pattern of secondary ciliation in REOEC monolayers is related to an experimental variable independent of cell donor (mare), donor age, and donor cycle stage

In three out of 11 experiments, diffuse patterns of secondary cilia were observed in spontaneously REOEC monolayers cultured for 1 month. Secondary ciliation rates in these 3 experiments varied from 10.5 to 24.4% and were consistent for all three replicates, but independent from cell donor (mare), donor age, and donor cycle stage (Table 2). In the other eight experiments, only poorly ciliated REOEC monolayers were obtained (<2% secondary ciliated EOECs per monolayer) (Table 2). Overall, these results suggest an unknown experimental variable with a marked effect on secondary ciliation.

Table 2 Relationship between experiment, cell donor (mare), donor age, cycle stage and secondary ciliation rates 1 month after air-liquid interface introduction in re-differentiated equine oviduct epithelial cell (EOEC) monolayers cultured in hanging inserts

Experiment	Mare	Age	Cycle stage	Insert 1	Insert 2	Insert 3	Mean % secondary cilia per mare	Mean % secondary cilia per experiment	Mean % secondary cilia per cycle stage	
1	1	4	LUT	1	1	1.5	1.2	0.5		
2	19	FOL	0.4	0.0.3	0.1	0.3		
3	23	FOL	0.2	0.1	0.1	0.1		
2	4	7	FOL	0.3	0.7	0.5	0.5	1		
5	9	LUT	1.0	1.6	1.4	1.3		
6	22	LUT	1.9	0.9	0.8	1.2		
3	7	5	FOL	0.7	0.4	0.6	0.6	0.4		
8	6	LUT	0.3	0.1	0.3	0.2		
9	25	LUT	0.8	0.2	0.6	0.5		
4	10	25	LUT	0.7	0.5	0.6	0.6	0.8		
11	16	FOL	0.8	0.4	0.9	0.7	LUT	
12	18	LUT	1.2	1.1	0.8	1	5.4 ± 7.1	
5	13	14	FOL	15.8	16.4	15.6	15.9	16.2		
14	24	LUT	10.8	14.8	12.8	12.8		
15	6	LUT	20.4	22.5	24.4	22.4		
16	15	FOL	12.8	15.8	12.9	13.8		
6	17	22	LUT	1.1	0.8	1.3	1.1	1		
18	20	FOL	0.6	0.3	0.6	0.5		
19	21	FOL	1.5	1.3	1.2	1.3		
7	20	9	LUT	0.3	0.8	0.5	0.5	1		
21	19	FOL	1.5	1.1	1.4	1.3		
22	23	LUT	1.0	1.6	1.1	1.2	FOL	
8	23	21	FOL	0.1	0.5	0.8	0.5	1.1	3.9 ± 5.8	
24	18	FOL	1.2	1.7	1.9	1.6		
9	25	14	FOL	10.5	12.9	10.8	11.4	13.3		
26	19	LUT	14.9	13.9	17.5	15.4		
27	17	LUT	14.5	11.8	12.8	13		
10	28	9	FOL	0.8	0.2	0.4	0.5	0.6		
29	24	LUT	0.4	0.8	1.1	0.8		
30	7	FOL	0.3	0.5	0.5	0.4		
11	31	9	FOL	13.5	15.1	12.1	13.6	12.7		
32	22	LUT	11.0	12.3	11.7	11.7		
Twenty seven of 96 (28.1%) EOEC monolayer cultures on inserts were classified as “diffusely ciliated” whereas 69 (71.9%) were “poorly ciliated”. Mean (± s.d.) secondary ciliation rate in poorly ciliated EOEC monolayers was 0.8 ± 0.5% whereas in diffusely ciliated EOEC monolayers it was 14.4 ± 6.3%. Interestingly, an experiment-related effect was observed where all EOEC monolayers in 3 out of 11 experiments demonstrated diffusely ciliated EOECs. However, no effect of donor, donor age or cycle stage on secondary cilia formation was observed (P > 0.05) (LUT: luteal cycle stage; FOL: follicular cycle stage).

Higher seeding concentrations of trypsinized DEOECs did not support a diffuse secondary ciliation pattern in spontaneously REOEC monolayers

In the 3 experiments where spontaneously REOEC monolayers showed >20% secondary ciliated EOECs (Table 2; Figure 4a), we observed areas with an increased cell density of 40 ± 12 EOECs per mm2, compared to 15 ± 7 EOECs per mm2 in more sparsely ciliated areas (Figure 4b; see also Leemans et al., 2022). In a follow-up experiment, the effect on ciliation of the reseeding density of trypsinized DEOECs onto the insert membranes was assessed in EOEC monolayers after spontaneous re-differentiation. Increasing cell concentration from 0.1 to 10 × 105 trypsinized DEOECs per 0.33 cm2 insert did not result in increased monolayer EOEC density and ciliation rate (<1% of the EOECs showed secondary ciliation, Figure 4c).

Figure 4 (a) Representative overview image of a spontaneously re-differentiated equine oviduct epithelial cell (REOEC) monolayer 21 days after air-liquid interface introduction [green (anti-acetylated α-tubulin antibodies-AlexaFluor 488 secondary antibody): primary (single green dots at each EOEC) and secondary cilia (many green dots on some EOECs), red (phalloidin conjugated to Alexa Fluor 568): cytoskeleton, blue (Hoechst 33342): nuclei] (original magnification, 100x – scale bar: 100 μm). (b) Two images of the same area of a REOEC monolayer, showing that (A) secondary cilia were present mainly in spontaneously REOEC monolayer areas with (B) a higher cell density (left of white line) (original magnification, 400x – scale bar: 25 μm). (b) The effect of varying seeding density of trypsinized de-differentiated EOECs from 0.1 to 10 × 105 cells per 0.33 cm2 insert membrane was assessed 1 month after air-liquid interface introduction. No relationship was observed between EOEC seeding density and secondary ciliation rates in spontaneously re-differentiated EOEC monolayers. Data are mean (± SD) percentage of secondary ciliated cells (n = 3 mares; 3 inserts per mare per seeding concentration).

Oviduct explant cell concentration influences the ability of trypsinized DEOECs to spontaneously re-differentiate and form secondary cilia in a diffuse pattern

DEOEC density after 10 days of oviduct explant cell de-differentiation was significantly lower after seeding 1 (32 ± 5 EOECs per mm2) or 5 × 106 EOECs per 9.6 cm2 well (38 ± 4 EOECs per mm2) compared to 10 (61 ± 5 EOECs per mm2) or 30 × 106 EOECs per 9.6 cm2 well (73 ± 5 EOECs per mm2). This observation can also be appreciated in Figure 5a.

Figure 5 Representative (a) bright field (differential interference contrast) light images of de-differentiated equine oviduct epithelial cell (EOEC) monolayers 10 days after oviduct explant seeding in conventional wells (magnification 100x – scale bar: 100 μm) and (b) fluorescent images of diffusely ciliated re-differentiated EOEC monolayers 1 month after air-liquid interface introduction in hanging inserts. If (A) 1 × 106 or (B) 5 × 106 oviduct explant cells were initially seeded in conventional wells to obtain de-differentiated EOEC monolayers, diffuse ciliated EOEC monolayers were obtained after re-differentiation. In contrast, seeding (C) 10 × 106 or (D) 30 × 106 oviduct explant cells resulted in monolayers with <2% ciliated EOEC after re-differentiation [green (anti-acetylated α-tubulin antibodies-AlexaFluor 488 secondary antibody): primary (single green dots at each EOEC) and secondary cilia (many green dots on some EOECs), red (phalloidin conjugated to Alexa Fluor 568): cytoskeleton, blue (Hoechst 33342): nuclei] (original magnification, 400x – scale bar: 25 μm). (c) The effect on secondary cilia formation of varying oviduct explant cell seeding concentration from 1 × 106 to 30 × 106 cells per 9.6 cm2 well to establish de-differentiated EOEC monolayers, prior to re-differentiation with an air-liquid interface for 1 month. For comparison, in vivo ±60% of EOECs exhibit secondary cilia. Using a lower oviduct explant cell concentration to obtain de-differentiated EOEC monolayers was critical to obtaining spontaneously re-differentiated EOEC monolayers with secondary cilia in a diffuse pattern. Data are mean (± SD) percentages of secondary ciliated cells (n = 3 mares; 3 inserts per mare per seeding concentration). Values that differ significantly between EOEC concentration are indicated by small letters.

Interestingly, DEOECs harvested from trypsinized de-differentiated monolayers 10 days after oviduct explant cell seeding at a concentration of 1 or 5 × 106 EOECs per 9.6 cm2 well, showed diffuse ciliated REOEC monolayers within 1 month after air-liquid interface introduction (Figure 5b). On average, 47.5 ± 4.8% of the EOECs showed secondary cilia in a diffuse pattern after initial seeding at 1 × 106 oviduct explant cells (Figures 5bA and 5C) per well whereas 28.9 ± 4.4% secondary ciliated EOECs were observed using an oviduct explant cell seeding concentration of 5 × 106 EOECs per well (Figures 5bB and 5C). Seeding concentrations higher than 5 × 106 EOECs per well did not result in secondary ciliation rates above 1% (10 × 106 oviduct explant cells: 0.7 ± 0.2%, Figures 5bC and 5C; 30 × 106 oviduct explant cells: 0.6 ± 0.3%, Figures 5bD and 5C). These results demonstrate that oviduct explant cell seeding concentration is a crucial factor for trypsinized DEOECs to retain their capacity to re-differentiate into secondary ciliated EOECs. However, in vivo ciliation levels (about 60%, Figure 1) were not reached under any tested oviduct explant cell seeding concentration.

Notch inhibition enhances secondary cilia formation in diffuse ciliated REOEC monolayers to an in vivo-like patchy ciliation pattern

Exposure of diffusely ciliated spontaneously REOEC monolayers to cycle stage-specific hormones at blood or oviduct tissue concentrations mimicking the luteal and follicular phases did not affect secondary cilia formation (Figure 6a). Overall, diffuse secondary ciliation was maintained during exposure to cycle phase mimicking hormone regimes.

Figure 6 Effect of (a) estrous cycle phase mimicking hormone concentrations on diffusely ciliated re-differentiated equine oviduct epithelial cell (REOEC) monolayers, after 14 days of luteal (blood concentration: 20 pg/mL E2 and 10 ng/mL P4; oviduct concentration: 10 ng/mL E2 and 1000 ng/mL P4) and 7 days of subsequent follicular phase hormone exposure (blood concentration: 40 pg/mL E2 and 0 ng/mL P4; oviduct concentration: 80 ng/mL E2 and 40 ng/mL P4). Control treatment was REOEC monolayer culture without added hormones. Tested hormone conditions did not have an effect on secondary ciliation rates and did not differ from the control treatment (culture without hormones). Data are mean (± SD) percentage of secondary ciliated EOECs (n = 3 mares; 3 inserts per mare per experimental condition). Effect of (b) Wnt (0–1-5 μM CHIR 99021 and 0–2-20 μM IWP-2) and Notch (0–1-10 μM JAG-1 and 0–1-10 μM DBZ) ligands / inhibitors on diffusely ciliated REOEC monolayers. The Notch inhibitor DBZ enhanced secondary ciliation rates up to in vivo-like levels. This stimulatory effect on secondary ciliation was not observed after exposure to Wnt ligand or inhibitor, or the Notch agonist. Data are mean (± SD) percentage of secondary ciliated cells (n = 3 mares; 3 inserts per mare per experimental condition). Values that differ significantly between conditions are indicated by small letters.

None of Wnt or Notch ligands (CHIR 99021 and JAG-1) or Wnt inhibitor (IWP 2) had any effect on secondary ciliation patterns of the diffusely ciliated REOEC monolayers (Figure 6b). Consequently, a diffuse ciliation pattern was maintained after one week under these conditions. Although a trend towards a concentration-dependent decrease in secondary ciliation rates could be observed after one week of Wnt activation, apparent differences between different CHIR 99021 concentrations were not significant (P > 0.05 for all comparisons; Figure 6b). In contrast, the Notch inhibitor DBZ increased secondary ciliation in diffuse ciliated REOEC monolayers to achieve a patchy ciliation pattern (Figure 6b) similar to the in vivo situation (± 60% secondary ciliated EOECs; Figure 1). Ciliation rates above 60% were obtained after 1 week of exposure to 1 and 10 μM DBZ (1 μM DBZ: 63.7 ± 6.2%, 10 μM DBZ: 65.9 ± 5.5%; Figure 6b).

To confirm the importance of the two effective experimental factors, an experiment was performed in which oviduct explant cell seeding concentration and Notch inhibition were combined to generate REOEC monolayers with in vivo-like patchy ciliation. Monolayers with >60% secondary ciliated REOECs were established if (1) oviduct explant cell concentrations up to 5 × 106 cells per 9.6 cm2 well were seeded to first establish a monolayer of DEOECs within 10 days. (2) Subsequently, trypsinized reseeded DEOECs were re-differentiated for 3 weeks by air-liquid interface introduction and (3) further exposed for 1 week to 10 μM DBZ (1 × 106 oviduct explant cells, 10 μM DBZ: 64.7 ± 6.9%; 5 × 106 oviduct explant cells, 10 μM DBZ: 62.3 ± 7.0%; Figures 7a and 7bC). Diffusely ciliated REOEC monolayers were obtained if EOEC monolayers were cultured under similar conditions, but in the absence of DBZ (1 × 106 oviduct explant cells, 0 μM DBZ: 40.8 ± 5.9%; 5 × 106 oviduct explant cells, 0 μM DBZ: 23.0 ± 6.2%; Figures 7a and 7bA). Higher starting oviduct explant cell seeding concentrations (10 and 30 × 106 cells) did not support secondary cilia formation (poorly ciliated EOEC monolayers: <2%) after re-differentiation and exposure to 0 or 10 μM DBZ (Figures 7a, 7bB, and 7bD). A videoclip is added to demonstrate cilia beating in a patchy ciliated REOEC monolayer (supplemental video clip 1).

Figure 7 (a) The combined effect of oviduct explant cell seeding concentration and Notch inhibition on secondary cilia formation. De-differentiated equine oviduct epithelial cells (EOECs) were obtained 10 days after seeding oviduct explant cells at concentrations varying from 1 to 30 × 106 cells per 9.6 cm2 well. Subsequently, secondary ciliation rates in re-differentiated EOEC monolayers were assessed 1 month after air-liquid interface introduction. During the last week of culture, EOEC monolayers were incubated in the absence or presence of 10 μM DBZ. A lower oviduct explant cell concentration to obtain de-differentiated EOEC monolayers supported a diffuse ciliation pattern after re-differentiation. Subsequent exposure to DBZ enhanced ciliation rates up to a patchy ciliation pattern showing ciliation rates similar to the in vivo situation (± 60% secondary ciliated EOECs; Figure 1). In contrast, initially seeding 10 or 30 × 106 oviduct explant cells resulted in <2% secondary ciliated EOECs after re-differentiation, irrespective of DBZ. Data are mean (± SD) percentage of secondary ciliated EOECs (n = 3 mares; 3 inserts per mare per seeding concentration). Values that differ between oviduct explant cell concentration are indicated by small letters. (b) Representative fluorescent images of re-differentiated EOEC monolayers 1 month after air-liquid interface introduction. To obtain (A) diffuse ciliated EOEC monolayers, a maximum seeding concentration of 5 × 106 oviduct explant cells was required. Subsequent exposure to DBZ during re-differentiation supported the development of (C) in vivo-like patchy ciliated EOEC monolayers. (B, D), Higher oviduct explant cell seeding concentration did not support secondary ciliation, regardless of whether re-differentiated EOEC monolayers were exposed to DBZ [green (anti-acetylated α-tubulin antibodies-AlexaFluor 488 secondary antibody): primary (single green dots at each EOEC) and secondary cilia (many green dots on some EOECs), red (phalloidin conjugated to Alexa Fluor 568): cytoskeleton, blue (Hoechst 33342): nuclei] (original magnification, 400x – scale bar: 25 μm).

Ultimately, we observed that the in vivo-like patchy ciliation status was maintained in REOEC monolayers for up to 14 days after the last day of 10 μM DBZ treatment (0 days after the end of DBZ treatment: 64.5 ± 5.7% ciliated EOECs; 7 days after the end of DBZ treatment: 60.7 ± 4.9% ciliated EOECs; 14 days after the end of DBZ treatment: 58.9 ± 5.2% ciliated EOECs). These data suggest that in vivo-like ciliated REOEC monolayers can potentially be used after stopping DBZ exposure, to study events related to capacitation, fertilization, and early embryonic development consecutively.

Clathrin expression is higher in in vivo-like patchy ciliated REOEC monolayers during the follicular phase

Staining for clathrin revealed the presence of secretory vesicles in in vivo oviduct tissue cryosections and patchy ciliated REOEC monolayers in a cycle stage-dependent density (Figure 8). After 14 days exposure to luteal phase oviduct tissue hormone conditions, clathrin fluorescence intensity per cell in REOECs (0.21 ± 0.08) was significantly lower than after 7 days exposure to follicular phase oviduct tissue hormone conditions (0.53 ± 0.76) (P < 00.1) (Figure 8). This observation indicates cycle stage-dependent secretory responsiveness of patchy ciliated REOEC monolayers. Interestingly, clathrin expression was present in both ciliated and non-ciliated EOECs after exposure to either luteal or follicular oviduct tissue hormone conditions.

Figure 8 (a) Representative fluorescent images showing the secretory marker clathrin (green) in (A, F) in vivo oviduct tissue sections from mares in the luteal and follicular phase, respectively, and in re-differentiated EOEC monolayers after (B, C, D, E) 14 days of luteal (oviduct concentration: 10 ng/mL E2 and 1000 ng/mL P4) and (G, H, I, J) 7 days of subsequent follicular phase hormone exposure (oviduct concentration: 80 ng/mL E2 and 40 ng/mL P4). Higher clathrin expression was observed for re-differentiated EOEC monolayers exposed to follicular than to luteal phase hormone conditions; [pink (anti-acetylated α-tubulin antibodies-Alexa Fluor 647 secondary antibody): primary (single pink dots at each EOEC) and secondary cilia (many pink dots on some EOECs), green (anti-clathrin antibodies- Alexa Fluor 488 secondary antibody) red (phalloidin conjugated to Alexa Fluor 568): cytoskeleton, blue (Hoechst 33342): nuclei] (original magnification, 400x; A, F - scale bar: 50 μm; B, C, D, E, G, H, I, J – scale bar: 25 μm). (b) Cross sections through re-differentiated EOEC monolayers confirms higher clathrin expression after exposure to (B) follicular compared to (A) luteal phase hormonal conditions. Moreover, clathrin expression was observed in ciliated and non-ciliated EOECs. (C) Effect of estradiol and progesterone to mimic estrous cycle phase on mean clathrin fluorescence intensity per cell in re-differentiating EOECs, after 14 days of luteal and 7 days of subsequent follicular phase hormone exposure. Significantly higher clathrin expression in re-differentiated EOECs was observed after follicular phase hormone exposure. Data are mean (± SD) clathrin fluorescence intensity per cell (n = 3 mares; 3 inserts per mare per hormonal condition). Values that differ between clathrin fluorescence intensity per cell are indicated by small letters.

Discussion

In mammalian species such as the horse, the oviductal ampulla is the site at which an oocyte is fertilized by a spermatozoon and where early embryonic development begins. To date, equine ex vivo oviduct models have failed to fully support or permit sperm capacitation and fertilization. In the current study, an oviduct model was optimized to more closely recapitulate the oviduct luminal environment. We obtained confluent monolayers of REOECs by first allowing collected oviduct epithelial explants to form DEOEC monolayers and subsequently stimulating re-differentiation by introducing an air-liquid interface. Initial seeding at a maximum of 5 × 106 oviduct explant cells per well supported the formation of diffusely ciliated EOEC monolayers after de- and re-differentiation. Notch inhibition further enhanced secondary ciliation to in vivo-like patchy ciliation patterns. This improved strategy significantly enhanced the capacity of REOECs to form secondary cilia. Moreover, patchy ciliated REOEC monolayers showed higher clathrin expression in follicular compared to luteal phase conditions.

The development of primary or solitary, non-motile cilia at the apical side of OECs is driven by cell polarization. This type of cilia does not contain a microtubule core. In contrast, secondary or motile cilia do contain a microtubule core which is arranged in a 9 + 2 pattern, and within which ATP-dynein-dependent sliding of microtubule doublets generates cilia movement. Various factors such as adrenergic and cholinergic stimulation, cycle stage-dependent hormones, prostaglandins, platelet activating factors, angiotensin II receptors, and adrenomedullin have been shown to regulate the beat cross frequency of secondary cilia in the mammalian oviduct to fine tune gamete and embryo transport (reviewed by Ezzati et al. [35]). Ciliation is also pivotal to developing a more physiological in vitro oviduct model because both the primary and secondary cilia play key roles in monitoring and modulating the extracellular environment, by processing mechanosensory signals, and by generating fluid flow [36]. In addition, cilia act as secretory organelles and transduce information by releasing small vesicles, termed ectosomes, that play a role in cell–cell communication, intracellular signaling and cell cycle-related processes [37, 38]. In fact, spermatozoa have been shown to interact specifically with ciliated epithelial cells in the oviduct, and this appears to be important for in vivo sperm capacitation since sperm cells released from binding to ciliated oviduct epithelial cells show hyperactivated motility and the ability to fertilize an oocyte [7, 10, 39, 40]. The current study shows that it is possible to culture REOEC monolayers with several in vivo-like morphological characteristics thought to be instrumental to physiological function.

Unfortunately, the mechanism responsible for inducing de novo formation of secondary cilia in EOECs is not yet fully understood. In this study, we show that reproductive steroid hormones in cycle stage-specific concentrations do not affect secondary ciliation levels regardless of whether they were added to poorly ciliated (<2%) or diffusely ciliated REOEC monolayers. In general, OEC ciliation has been proposed to be triggered by hormonal cues, e.g., the injection of estrogens promotes ciliogenesis in the OECs of newborn rats [41]. Similarly, in the pig, a clear effect of cycle-stage specific hormones, i.e., estradiol and progesterone, on OEC cell height, ciliation rates, and secretory markers, has been reported [16]. Clear evidence of EOEC functionality was obtained by exposing REOEC monolayers to cycle stage-specific hormones [15]. Although cycle stage-specific hormones do not support secondary cilia formation in REOEC monolayers, the affinity for sperm binding by cilia increased under follicular phase culture conditions [15]. Moreover, exposing REOECs to progesterone under luteal phase conditions downregulated progesterone receptor abundance in REOEC monolayers [15]. In this study, we additionally observed that in vivo-like patchy ciliated REOEC monolayers show higher clathrin expression when cultured under follicular compared to luteal stage hormonal conditions. Clathrin, a protein that is known to coat the membrane of cellular transport vesicles, is an indirect secretory marker in that it regulates cell transport, endo- or exocytosis [22–24]. Besides cycle stage-dependent expression, a clear clathrin signal was present in nearly all cells of the REOEC monolayers, irrespective of ciliation status. This may indicate that secretory activity is not restricted to non-ciliated OECs. This contrasts with the concept that oviduct epithelium consists of secretory, non-ciliated and non-secretory, ciliated OECs [42]. Moreover, an electron microscopic study of human oviduct tissue showed that occasionally OECs have features of both ciliated and secretory OECs [43]. In the horse, Desantis et al. [44] obtained similar results, confirming the presence of secretory granules in ciliated EOECs. Interestingly, Ghosh et al. [45] suggested that secretory OECs are proliferative and act as a precursor for ciliated OECs.

Wnt and Notch pathways are key signaling pathways during development (proliferation) and adult tissue homeostasis (differentiation) [32–34, 46]. A key factor of the Wnt signaling pathway is the stability and localization of the soluble pool of β-catenin [47–50]. In the absence of a Wnt ligand, free cytosolic β-catenin interacts with a destruction complex containing glycogen synthase kinase 3β which supports β-catenin phosphorylation, to target degradation. Binding of Wnt ligands to the cell surface receptors Frizzled and LRP will initiate the destruction process and prevents phosphorylation of β-catenin. The accumulation of β-catenin in the cytoplasm and, subsequently, the nucleus will induce expression of target genes [47–50]. On the other hand, Notch signaling is triggered by the interaction between Notch receptors and DSL (Delta, Serrate, Lag2) ligands on adjacent cells. This leads to the proteolytic cleavage of Notch to release the intracellular domain. This domain will translocate to the nucleus to induce expression of target genes [51]. Wnt and Notch activation appear to act antagonistically as either proliferation or differentiation triggers in different tissues [52]. In a given tissue, both pathways are often active and have opposing effects on cell fate specification [52]. Therefore, strict regulation of both pathways within a given tissue is essential to prevent conflict and to support status, i.e., Wnt-on/Notch-off or Notch-on/Wnt-off. The crosstalk between these pathways generates diversity in many cell types. It is hypothesized that Notch signaling reduces Wnt activation (Notch-on/Wnt-off state) by limiting the transcriptional activity of β-catenin [53].

In our study, Wnt signaling caused an unfavorable EOEC epithelial-mesenchymal transition, instead of maintaining REOEC monolayers. Cytoskeleton reorganization resulted in the transformation of columnar REOECs into flat fibroblast-like, spindle-shaped cells. This was accompanied by increased cellular length and nuclear diameter, and a reduced monolayer confluency, likely due to reduced cell–cell coherence via a loss of stable tight-junctions [25, 54]. Interestingly, a significant proportion of the transformed EOECs were pushed out of the original EOEC monolayer. Subsequently, these EOECs tended to grow over or on top of each other, or lost contact with other EOECs completely. We hypothesize that, as a result of multiple layer growth, relatively few EOECs are able to polarize and they subsequently lose their capacity to develop primary cilia. This phenomenon was previously reported for other tissues by Lee and Nelson [55]. According to the latter, typical features of epithelial-mesenchymal transition include a spindle-shaped phenotype, absence of stable tight junctions, increased mobility and absence of primary cilia [55]. The apparent epithelial-mesenchymal transition is presumably related to activation of the Wnt/β-catenin signaling pathway, which is known to activate genes responsible for epithelial-mesenchymal transition. This has been reported to result in a loss of cadherin-mediated cell–cell adhesion [56, 57]. Similarly, Notch stimulation is reported to be a key regulator of epithelial-mesenchymal transition induction [58, 59], although this effect was not observed for the Notch activator concentration range tested in the current study. Conversely, the phenomenon mesenchymal-epithelial transition was not observed in the adult mouse oviduct and uterus [60].

Secondary cilia formation in REOEC monolayers is considered to be an important morphological feature of epithelial cell differentiation. In the ampulla of the horse oviduct, around 60% of the EOECs have secondary cilia, independent of cycle stage [this study; 15, 44]. Interestingly, inhibiting Notch signaling in vitro decreases cell proliferation, and enhances the formation of secondary cilia, as previously demonstrated in human oviduct epithelial cell organoids [61] and skin tissue from Xenopus embryos [62]. In the present study, Notch inhibition using DBZ only increased secondary ciliation if there was already a “threshold” proportion of ciliated EOECs in a diffuse pattern. This contrasts to previous findings of Kessler et al. [61]. As suggested above, it appears that the effects of Wnt- and Notch-signaling differ between species and tissues. For example, it was recently reported that Notch inhibition turns proliferative cells in murine intestinal crypts and adenomas into goblet (secretory) cells [63]. On the other hand, stimulation of Wnt-signaling in a murine tracheal tube resulted in an increase in ciliated epithelial cells and a decrease in Clara-like (secretory) cells [64], whereas Wnt-signaling in mouse OECs was essential for both self-renewal and the differentiation into secretory cells [45]. Although these findings show marked species and cell-type variability in the effects of Wnt and Notch ligands and inhibitors, in our study only inhibition of Notch signaling supported secondary ciliation in EOECs.

The likelihood of obtaining secondary ciliated EOECs in re-differentiated monolayers proved to be related to cell density in the EOEC monolayer. We found a similar correlation in a previous study [15]. Interestingly, increasing the seeding density of trypsinized DEOECs on insert membranes did not result in higher cell density or secondary ciliation rates. There is evidence that cell density influences cell behavior, including migration, proliferation, and differentiation [65–67]. Using mouse bone marrow cells, cell proliferation rates decreased and expression of osteoclastogenesis-related genes increased as initial plating density increased [68]. Cardiomyocyte differentiation is also affected greatly by cell seeding density [69, 70]. Differentiation of mouse endothelial cells from embryonic stem cells was also related to cell density and seeding concentration [71]. Moreover, a high cell seeding density correlated with up-regulation of several genes including cell adhesion molecules from the notch (NOTCH1 and NOTCH4) and cadherin (CDH5) families related to vascular development/differentiation [71]. In alignment with our study, the density of reseeded EOECs was important for EOEC differentiation. The reason why increased seeding concentration did not result in increased cell density might be that seeding higher concentrations of DEOECs resulted in increased medium acidosis and cell death. Under these conditions, a certain percentage of trypsinized DEOECs might undergo cell deterioration before proper membrane attachment. Similar observations were made in human induced pluripotent stem cell cultures induced to undergo neural differentiation [72].

The seeding concentration of oviduct explant cells proved to be a crucial factor for EOECs to retain their capacity to form secondary cilia after de-differentiation and subsequent re-differentiation. As discussed by Leemans et al. [15], DEOECs have “transit-amplifying cell” (TAC) characteristics, i.e., a cell population that becomes differentiated after several rounds of cell division [73, 74]. Under the experimental conditions described, after adherence of oviduct explants to the floor of the culture well, EOECs flattened, formed a DEOEC monolayer and acquired mitotic competence. As a result, monolayer cell density increased progressively during 10 days of incubation. Interestingly, after initially seeding of 10 or 30 × 106 oviduct explant cells per well, DEOECs lost their capacity to form cilia after re-differentiation whereas they did not at lower seeding densities (1 or 5 × 106 oviduct explant cells per well). We hypothesize that the experimental variability observed in Table 2 is related to this phenomenon. Slightly lower or higher seeding concentrations (between 5 and 10 × 106 oviduct explant cells) during the 10 day de-differentiation step may explain differences in ciliation rates between experiments. It seems reasonable to propose that the maximal cell density per well would be achieved more quickly after seeding >5 × 106 oviduct explant cells per well, since fewer mitotic cycles would be necessary to reach this stage. Maximal cell density has been proposed to be associated with a phenomenon called “cell contact inhibition”, a physiological process through which cells cease dividing and proliferating when in close contact with one another [75]. Incubation time might also play a crucial role in this respect. Seeding ≤5 × 106 oviduct explant cells per well will likely not achieve maximal cell density or result in contact inhibition within 10 days. It can consequently be hypothesized that in conditions where >5 × 106 oviduct explant cells are seeded, a shorter de-differentiation period (<10 days) before maximal cell density is obtained will allow the DEOECs to retain the capacity to form secondary cilia after re-differentiation.

In conclusion, a reproducible way was developed for generating equine oviduct epithelial cell monolayers on hanging inserts that support EOEC polarization and morphological differentiation, including secondary cilia formation. Re-differentiated EOECs have shown functionality by expressing proteins associated with cell ciliation, sperm binding characteristics [25], dynamic changes in hormone receptor expression [15] and clathrin-dependent secretory activity. The level of EOEC differentiation achieved is an important step in the development of an in vitro model recreating an in vivo-like oviductal environment. Future research should concentrate on validating full biological functionality by comparing the (epi)transcriptome and proteome of re-differentiated EOEC monolayers with in vivo oviductal tissue. Ultimately, this oviduct model will provide a system to better study sperm capacitation, fertilization and early embryonic development.

Supplementary Material

Supplemental_video_clip_1-cilia_beating_ioae090

Acknowledgment

Images were acquired in the Center for Cellular Imaging (CCI) at the Faculty of Veterinary Medicine Utrecht, and we thank Dr R. Wubbolts and E. van ‘t Veld for their excellent help and technical advice. Wnt3a containing medium was kindly provided by Dr B. Spee (Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, The Netherlands). We would also like to thank Heiko Henning (Department of Clinical Sciences, Faculty of Veterinary Medicine, Utrecht University, The Netherlands) for his experimental support.

Footnotes

Conflict of interest

The authors have declared that no conflict of interest exists.

Data availability

The data underlying this article will be shared on reasonable request to the corresponding author.

† Grant Support: This study was supported by the Research Foundation—Flanders (FWO-Flanders; grant number 12I0517N) and EU COST Action 16119 CellFit.
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