# Bovine ampullary and isthmic epithelial spheroids: proteomic profile and physiological features for in vitro studies of gamete-oviduct interactions.

Coline Mahé, Ludivine Marco, Ludivine Laffont, Marie-Véronique Demattei, Karine Reynaud, Emmanuelle Com, Regis Lavigne, Charles Pineau, Marie Saint-Dizier

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## To cite this version:

Coline Mahé, Ludivine Marco, Ludivine Laffont, Marie-Véronique Demattei, Karine Reynaud, et al.. Bovine ampullary and isthmic epithelial spheroids: proteomic profile and physiological features for in vitro studies of gamete-oviduct interactions.. Reproductive biology, 2025, 25 (3), pp.101057. 10.1016/j.repbio.2025.101057. hal-05238375

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Original article

# Bovine ampullary and isthmic epithelial spheroids: proteomic profile and physiological features for *in vitro* studies of gamete-oviduct interactions

<figure><img src="image_3.png" /></figure>

Coline Mahé<sup>a,†</sup>, Ludivine Marco<sup>a,†</sup>, Ludivine Laffont<sup>a</sup>, Marie-Véronique Demattei<sup>a</sup>, Karine Reynaud<sup>a</sup>, Emmanuelle Com<sup>b,c,†</sup>, Régis Lavigne<sup>b,c</sup>, Charles Pineau<sup>b,c</sup>, Marie Saint-Dizier<sup>a,∗</sup>

## ARTICLE INFO

**Keywords:**
Spheroid
Sperm
Isthmus
Ampulla
Fallopian tube

## ABSTRACT

After mating or insemination, spermatozoa reach the first part of the oviduct, known as the isthmus, where a subpopulation binds to the epithelial cilia, forming a reservoir. Then, spermatozoa migrate toward the ampulla, where additional interactions with cilia occur and fertilization takes place. Exploring sperm-oviduct interactions requires physiologically relevant *in vitro* models. Our goal was to characterize isthmic and ampullary epithelial spheroids in terms of cellular, proteomic and sperm binding properties. Spheroids measuring 100–150 µm in diameter, formed after a 3-day culture of pre-ovulatory oviduct mucosa fragments, were selected and analyzed for (i) epithelial, fibroblast and cilia markers by immunostaining, (ii) proteomic content by dia-PASEF nanoLC-MS/MS, and (iii) ability to interact with bull spermatozoa during a 6-h co-culture, assessed by confocal and scanning electron microscopy. Cells within both isthmic and ampullary spheroids showed positive immunostaining for cytokeratin and no staining for vimentin. Ampullary spheroids consistently exhibited a higher proportion of ciliated cells compared to isthmic spheroids (24.5 % vs 20 %; p < 0.05; variation coefficients among replicates of 15 and 11 %, respectively). A total of 6914 proteins were quantified, of which 1010 (14 %) were differentially abundant between isthmic and ampullary spheroids. Following co-culture, 95–97 % of bound spermatozoa were attached by their head to cilia of both spheroid groups. The density of bound spermatozoa was on average 33–45 % higher on ampullary compared to isthmic spheroids (p < 0.001). This study provides a well-differentiated oviduct epithelium model for *in vitro* investigations of gamete and embryo interactions with the oviduct.

## 1. Introduction

During the periconception period, the oviduct lumen serves as the site where male and female gametes migrate and meet, initiating embryo development [1]. The mammalian oviduct is mainly divided into the isthmus, its narrowest lower part, and the ampulla, its upper part where fertilization occurs. Following mating or insemination, gamatozoa first enter the isthmus, where a subpopulation binds to the ciliated epithelial cells, forming a sperm reservoir [2,3]. This close interaction with cilia is thought to help preserve sperm viability until ovulation and to reduce the risk of polyspermy by ensuring a gradual release of spermatozoa toward the oocyte [4]. However, sperm-cilia interactions are not limited to the isthmus, as multiciliated cells are present along the

entire oviductal lumen [5]. *In situ* video recording of sperm behavior in the oviducts of mated mice showed close interactions of spermatozoa with the ampullary epithelium and successive binding and release sequences during migration toward the oocytes [6,7]. Similarly, *in vitro* studies have demonstrated sperm-epithelial interactions in both isthmic and ampullary regions across several mammalian species, including human [8], horse [9], pigs [10,11] and cattle [12,13]. However, the ability of oviduct epithelial cells to bind spermatozoa appears to vary depending on the anatomical region. In cattle, bull spermatozoa were previously shown to interact differentially with ampullary and isthmic secretions during the periconception period [14]. Studies examining sperm binding dynamics and release treatments have reported stronger adhesion of spermatozoa to ampullary than to isthmic cells in mice [6]

<footnote><sup>a</sup> INRAE, CNRS, Université de Tours, PRC, Nouzilly 37380, France</footnote>

<footnote><sup>b</sup> Univ Rennes, Inserm, EHESP, Irset (Institut de recherche en santé, environnement et travail) - UMR_S 1085, F, Rennes 35000, France</footnote>

<footnote><sup>c</sup> Univ Rennes, Biosit - UMS 3480 CNRS, US_S 018 Inserm, Protim Core Facility, F, Rennes 35000, France</footnote>

<footnote>* Correspondence to: INRAE Unité Physiologie de la Reproduction et des Comportements, Nouzilly 37380, France.
E-mail address: marie.saint-dizier@univ-tours.fr (M. Saint-Dizier).</footnote>
and cattle [12]. In addition, a higher binding capacity to bind sperma-
tozoa was reported in ampullary compared to isthmic cells in some [3,
15] but not all [9,11,16] studies in mammals, maybe due to the vari-
ability of the in vitro models used.

Due to the anatomical position of the mammalian oviducts deep
within the abdomen, their narrow diameter (< 3 mm) and the numerous
mucosa folds filling the lumen, the study of gamete-oviduct interactions
requires the development of suitable *in vitro* models. Several models of
oviduct epithelial cells have been reported, but very few have addressed
region-specific cellular characteristics [3,12,17–20]. A commonly used
approach involves culturing oviduct epithelial cells as monolayers on
plastic dishes [3,21]. However, this model is limited by the rapid loss of
columnar, apical-basal polarity, and ciliated characteristics of the
epithelium, enhancing the risk of non-specific interactions [3,21].
Another model consists of oviduct explants or aggregates, used within
hours after tissue collection [3]. This model is hindered by high het-
erogeneity in shape and size, making quantification of sperm binding on
their irregular surfaces difficult. Recently, we developed a standardized,
3D *in vitro* model using bovine isthmic epithelial spheroids for the study
of sperm-oviduct [19,20] and embryo-oviduct [17,18] interactions.
These spheroids spontaneously formed from isthmic mucosa folds after
48–72 h in culture, with the apical surface facing outward, enabling
direct contact between spermatozoa and cilia. To date, explants of
various size and shape from the ampullary part of the oviduct were
previously characterized in cattle [12,22,23], but the characterization of
well standardized spheroids usable for gamete binding assays have not
been reported.

In addition, each part of the oviduct epithelium is known to exhibit distinct transcriptomic signature, as recently demonstrated in mice [24], cattle [25–27] and humans [28]. However, proteomic data remain scarce and have been reported only for the isthmus in cattle [29], the ampulla in rabbits [30], or in mix isthmic/ampullary samples in pigs [31,32].

Therefore, the aim of this study was to characterize and compare
epithelial spheroids from the isthmus and ampulla in terms of epithelial
and ciliary markers, morphometric reproducibility, proteomic content
and ability to interact with spermatozoa.

## 2. Material and methods

2.1. Media and reagents

Unless otherwise specified, all products were obtained from Merck
Life Science. The spheroid culture medium was prepared using M199
(Gibco 31150–022) supplemented with 2.5 % HEPES (Gibco
15630–080), 0.5 % gentamycin (G-1272) and 10 % heat-inactivated
fetal calf serum (Gibco A5256701). For sperm-spheroid co-incubation,
a Tyrode-Lactate-Pyruvate (TLP) medium was used, prepared with
Tyrode's salts (T2145), 1 mM sodium pyruvate (S8636), 27.4 mM so-
dium lactate (L7900), 10 mM HEPES (H0887) and 1 mg/mL polyvinyl
alcohol (P8136). The osmolarity was adjusted to 300 mOsm using NaCl
and the pH was adjusted to 7.4 with NaOH. The TLP medium was then
filtered through a 0.22 µm membrane and stored at 4°C for up to 21
days.

## 2.2. Culture and selection of bovine oviduct epithelial spheroids

Oviduct epithelial spheroids were produced as previously described for the isthmus [19,33] with slight modifications. Pairs of oviducts and ovaries of adult cyclic *Bos taurus* cows (mainly of the Holstein breed) were collected at a commercial slaughterhouse (Elivia, Lion-d'Angers, France), transported on ice to the laboratory and processed within maximum 3 h after animal death. Only females at the pre-ovulatory stage of cycle, i.e. with ovaries presenting a corpus albicans and a pre-ovulatory follicle (11–20 mm in diameter) were used. To obtain a sufficient number of cells, oviducts from three cows were pooled for

each culture. The utero-tubal junction, infundibulum and surrounding
tissue and vessels were isolated from the oviduct using a sterile razor
blade. The oviducts were then laid horizontally on a square box and the
isthmic and ampullary sections were isolated by excising a 3 cm segment
from the central part of the oviduct. Each isthmic and ampullary section
was immersed 5 s in 70 % ethanol then 0.9 % NaCl after which 5 mm
were trimmed from both ends. Mucosa fragments from each section
were expelled into 14 mL of culture medium using sterile forceps, vor-
texed for 1 min, and left to sediment for 10 min in a humidified atmo-
sphere with 5 % CO₂ at 38.5°C. For isthmic cells, the
vortex-sedimentation process was repeated once after discarding the
supernatant. Preliminary experiments indicated that additional me-
chanical dissociation was necessary for successful spheroid formation
from ampullary fragments. Therefore, for ampullary cells only, 600 µL of
the cell pellet was diluted 1:10 in culture medium and dissociated by
drawing the suspension 50 times through a 26-gauge needle using a
1-mL syringe in a 60 mm Petri dish on a heating plate. The dissociated
ampullary cells were then transferred into 14 mL of culture medium,
vortexed for 1 min and left to sediment for 10 min in a humidified at-
mosphere with 5 % CO₂ at 38.5°C. After discarding the supernatant, the
dissociation-vortex-sedimentation process was repeated. Finally, cells
from isthmus and ampulla were diluted 1:100 (600 µL of cells in 5600 µL
of culture medium) and cultured in 60 mm dishes with no surface
treatment for cell attachment (Corning 430589). The dishes were placed
inside square boxes filled with 15 mL of sterile water in a humidified
atmosphere containing 5 % CO₂ at 38.5°C. After 3 days (optimal time to
obtain the greatest number of spheroids), numerous spheroids of varying
shape and size spontaneously formed, with their apical surface and
ciliary beating oriented outward [33]. Spheroids were manually selected
under a stereomicroscope using a mouth drawn Pasteur pipette based on
their morphology: round shape, presence of a cavity, absence of folds,
active ciliary beating, and a diameter ranging from 100 to 150 µm (see
Fig. 1).

2.3. Immunostaining and morphometric analysis of spheroids

Ampullary and isthmic spheroids were used to assess the epithelial
origin of cells (cytokeratin and vimentin immunostaining) and to
quantify the proportion of multiciliated cells (acetylated alpha tubulin
immunostaining). Pools of spheroids from each region were fixed in 4 %
paraformaldehyde in phosphate-buffered saline (PBS) supplemented
with 0.5 % Triton X-100 and 1 % bovine serum albumin (BSA) for
30 min at 37°C. Samples were then washed three times in PBS con-
taining 1 % BSA (PBS-BSA). Then, spheroids were incubated overnight
at room temperature with one of the following primary mouse mono-
clonal antibodies: anti-pan cytokeratin (20 µg/mL; C2931), anti-
vimentin (6.7 µg/mL; V6630) or anti-acetylated tubulin (2 µg/mL;
T7451). Controls were incubated with a mouse IgG1 isotype (M9269) at
the same concentration as the primary antibodies. Spheroids were then
washed three times in PBS-BSA (2 × 5 min then 30 min) under agitation
at room temperature. Spheroids were then incubated for 90 min at room
temperature in PBS-BSA containing a secondary anti-mouse antibody
coupled with Alexa fluor 488 (2 µg/mL; A11001), along with Hœchst
33342 (2 µg/mL; B2261) for nuclei staining. For some samples, Texas-
Red™-X phalloidin was also used for actin staining (2 U/mL; Invitrogen
T7471). Following three washes in PBS-BSA, spheroids were mounted
between a slide and coverslip and observed using a confocal microscope
(Zeiss Confocal LSM 780 Airyscan). For each spheroid, four confocal Z-
stacks pictures were analyzed using the ImageJ software (v1.54d). The
diameter, total number of cells and proportion of multiciliated cells were
manually quantified using Z-stack pictures. Additionally, the height of
10 randomly selected cells along the largest spheroid diameter was
measured. A total of three to four replicates, derived from different pools
of cows and culture days, with at least 15 spheroids per region per
replicate were analyzed.
<figure><img src="image_4.png" /><figcaption>**Fig. 1.** Bright-field microscope pictures of (A) isthmic and (B) ampullary spheroids after selection according to their round shape and diameter. Spheroids spontaneously formed after 3 days of culture of oviduct mucosa fragments, collected from adult slaughterhouse cows at the pre-ovulatory stage of cycle.</figcaption></figure>

## 2.4. Nanoliquid chromatography coupled with tandem mass spectrometry (NanoLC-MS/MS)

Pools of 60 size-selected ampullary and isthmic spheroids were washed from serum 30 min in TLP medium then incubated for 6 h in 1.5 mL new TLP medium in a humidified atmosphere containing 5 % CO₂ at 38.8°C (conditions identical to those used for sperm incubation; see below). Following incubation, spheroids were washed twice with sterile, filtered PBS at 37 °C and snap-frozen in a minimal volume of PBS in 1.5 mL polypropylene tubes, and stored at -70°C until analysis. Cell viability after 6 h in TLP medium was assessed using a LIVE/DEAD™ Viability/Cytotoxicity assay (4 µM calcein and 8 µM ethidium homodimer-1; Invitrogen LIVE/DEAD™ Viability/Cytotoxicity kit L3224) for 30 min at 38.5°C under 5 % CO₂. Confocal microscopy revealed that all cells in spheroids were calcein-positive (live cells) and ethidium homodimer-negative (dead cells) (data not shown).

Pools of spheroids were prepared using the PreOmics iST kit (PreOmics GmbH, Martinsried, Germany) following the manufacturer's instructions with minor adaptations for low-input material. Briefly, proteins were lysed, denatured, reduced and alkylated for 60 min at 95 °C and then Trypsin/LysC digested for 3 h at 37 °C. Salts, contaminants and other detergents were removed using the PreOmics spin cartridge, according to manufacturer's instructions.

Samples were then analyzed by nanoliquid chromatography and tandem mass spectrometry in diaPASEF mode. Two third of each sample were separated on a 75 µm × 250 mm IonOpticks Aurora 3 C18 column (Ion Opticks Pty Ltd., Bundoora, Australia). A gradient of reverse phase buffer (Buffer A: 0.1 % formic acid, 2 % acetonitrile, 97.9 % H₂O; Buffer

B: 0.1 % formic acid, 99.9 % acetonitrile) was run on a nanoElute UHPLC System (Bruker Daltonik GmbH, Bremen, Germany) at a flow rate of 250 nL/min at 50°C controlled by HyStar software (v6.0.30.0, Bruker Daltonik GmbH, Brème, Germany). The liquid chromatography run lasted for 80 min, starting at 2 % buffer B, gradually increasing to 13 % over 42 min, then to 20 % at 65 min, 30 % at 70 min, and 85 % at 75 min, which was maintained for 5 min to wash the column.

The ion transfer capillary temperature was set at 180 °C. Ions were accumulated for 100 ms, and mobility separation was achieved by ramping the entrance potential from -160 V to -20 V within 114 ms. MS and MS/MS mass spectra were acquired with an average resolution of 50000 FWHM full width at half maximum (mass range 100–1700 m/z), respectively. LC-MS/MS data were acquired using a dia-PASEF method. TIMS, MS operation and dia-PASEF window were controlled and synchronized using the control instrument software timsControl 5.1 (Bruker Daltonik GmbH).

The dia-PASEF method was designed using the method editor in PROTIM and a method was manually defined to perform data-independent isolation from several 26 m/z wide precursor windows, also called segments, in a single TIMS separation (0.74 ms) for a total of six segments and twenty-nine boxes. These scans perfectly cover the diagonal area of doubly charged peptides and partially triply charged peptides in the m/z and ion mobility output range. MS and MS/MS data were collected over the m/z range 354.1–1200 and over the mobility range from 1/K₀ = 0.75–1/K₀ = 1.33 Vs cm-2. During each data collection, each TIMS cycle was 0.74 s long and comprised 1 MS and 6 cycles of dia-PASEF MS/MS segments, comprising 4 or 5 boxes, to cover therefore the 29 boxes defined in the acquisition method. The collision energy was increased linearly with mobility from 59 eV at 1/K₀ = 1.6 Vs cm⁻² to 20 eV at 1/K₀ = 0.6 Vs cm⁻². For MS data processing, ion mobility resolved mass spectra, nested ion mobility versus m/z distributions, and fragment ion intensity sums were extracted from the raw data file with DataAnalysis 6.0 (Bruker Daltonik GmbH, Bremen, Germany). The signal-to-noise ratio (S/N) was increased by summing the individual TIMS scans. Mobility peak positions and half-peak widths were determined on the basis of extracted ion mobilograms (EIM, ± 0.05 Da) using the peak detection algorithm implemented in the DataAnalysis software. Feature detection was also performed using DataAnalysis 6.0 software; stored at the raw data level (d).

For the project library, previous DDA raw files from sample acquisition as part of a sample preparation optimization phase, and acquired as previously described [34] were analyzed in Spectronaut software version 18 (Biognosys, Schlieren, Switzerland), using the Pulsar search engine integrated into the Spectronaut software, and a search schema with default settings to generate respective spectral library. Additionally, the DIA files from the individual's samples, based on raw files, were searched in the same way as described above, to generate a combination of DDA and DIA in a so-called "hybrid libraries". The calibration search was dynamic and MS1, MS2 correction factor was 1. Searches were performed against the *Bos taurus* Uniprot database (26,630 entries; July 2024), using trypsin as the protease (up to 2 missed cleavages). To account for post-translational modifications and chemical labelling settings, carbamidomethylation of cysteine residues was defined as a fixed modification, and methionine oxidation as variable modifications. A 1 % false discovery rate (FDR) was applied at precursor, peptide, and protein levels.

For DIA analysis, the raw files from individual samples and acquired in DIA were used. Prior library-based analysis of the DIA data, the raw files were converted into htrms files using the htrms converter (Biognosys). MS1 and MS2 data were centroided during conversion. The other parameters were set to default. The htrms files were then separately analyzed with Spectronaut against the previously generated hybrid libraries and default settings, and allowed quantification of the precursors, peptides and proteins. The results were filtered by a 1 % FDR on precursor, peptide and protein level using a target-decoy approach, which corresponds to a Q value $\le 0.01$ [35]. The abundance of proteins
was estimated by calculating the sum of the abundance of each identified peptide. The abundance of peptides was estimated by calculating the average of the abundance of each identified precursor. Quantification data were then normalized by Spectronaut software to take into account the overall acquisition heterogeneity between samples. Given the number of samples analyzed (less than 500 individuals), the type of data normalization carried out for the whole dataset was a local regression normalization as previously described [36].

All LC-MS data, libraries, and Spectronaut project files are available in the ProteomeXchange Consortium [37] (http://proteomecentral.prot.eomexchange.org) via the PRIDE [38] repository with the dataset identifier PXD065839. The Spectronaut projects can be viewed using the free Spectronaut viewer (www.biognosys.com/technology/spectronaut-viewer).

Proteins identified with at least two unique peptides and log2 abundance > 0 in a minimum of two biological replicates were retained. Contaminants such as human keratins and digestion enzymes (trypsin, LysC) were excluded. Redundant gene names were filtered using BLASTP (≥90% identity), prioritizing UniProt-reviewed entries.

A total of five replicates from different cow pools and culture days were analyzed for both isthmic and ampullary spheroids. Differentially abundant proteins (DAPs) were defined using unpaired Student's t-tests ($p \le 0.05$) and fold-change ratio $\ge 1.3$.

## 2.5. Prediction of protein subcellular localization, secretion and functions

For prediction of subcellular localization and secretion, protein sequences in FASTA format were retrieved using the « ID mapping » tool on Uniprot (https://www.uniprot.org/id-mapping) and used as inputs. Conventional secretion pathways (i.e. signal peptide-mediated secretion) and unconventional secretion were predicted using the OutCyte online tool (http://www.outcyte.com/) [39]. Subcellular localization at the membrane (whether through transmembrane domains or lipid anchorage) was predicted using Deep Loc 2.1 (https://services.healthtech.dtu.dk/services/DeepLoc-2.1/) [40]. Proteins were considered as ciliary if they were previously identified as such in the Human Protein Atlas under the “cilium” category (https://www.proteinatlas.org/). To visualize overlaps between predicted secreted, membrane-associated and ciliary proteins, a Venn diagram was generated using the jvenn interactive tool (https://jvenn.toulouse.inra.fr/app/example.html) [41]. Gene Ontology (GO) enrichment analysis of DAPs was carried out using Metascape (https://metascape.org/gp/index.html#/main/step1) [42] and gene symbols as input. Since the *Bos taurus* database is not yet available in Metascape, human ortholog gene symbols, retrieved via the HUGO Gene Nomenclature Committee (HGNC) database (https://www.genenames.org/), were used as proxies. GO Biological Process (GO BP) and Cellular Component (GO CC) categories were selected for enrichment analyses.

## 2.6. Sperm preparation and incubation with spheroids for binding assays

Bull semen was generously provided by Synetics (Noyal-sur-Vilaine, France) and had been cryopreserved in Optidyl extender (IMV Technologies, L'Aigle, France). To assess sperm binding in a region-specific manner while eliminating individual male effects, a pooled semen sample from three *Bos taurus* bulls (two Holstein, one Normande; one ejaculate per bull) was used for all experiments. For each experiment, three straws (one straw per bull) were thawed for 3 min in a water bath at 38.5°C. Their content was then transferred onto a 4 mL BoviPure-BoviDilute density gradient (40–80%; Nidacon BP-100 BD-100) and centrifuged for 20 min at 700 × g at room temperature. Following centrifugation, the supernatant was discarded and 5 µL of the sperm pellet were diluted in 195 µL distilled water for sperm concentration determination using a Thoma cell counting chamber. The remaining sperm pellet was washed in 1 mL of TLP medium and centrifuged for 10 min at 300 × g at room temperature. Before sperm preparation,

pools of 60 selected spheroids of each region were washed from serum for 30 min in 500 µL TLP medium. During sperm gradient centrifugation, spheroids were transferred in 10 µL of TLP medium to 1.475 mL of new TLP medium in 12-wells plate (Greiner Bio-One 665 180). Then, $1.5 \cdot 10^6$ spermatozoa were added in 15 µL of TLP medium to each well containing the spheroids, to reach a final concentration of $1.10^6$ spermatozoa/mL. Plates were incubated for 6 h at 38.5°C in a humidified atmosphere with 5 % CO₂ in air and sealed with a plastic film to prevent pH fluctuation.

## 2.7. Characterization of sperm-spheroid interaction by confocal microscopy

To quantify bound sperm density per unit surface, spheroids of each region were collected after 1 and 6 h of co-incubation with spermatozoa using a 10-µL pipette. Sperm-spheroids complexes were then rapidly washed three times in 500 µL of TLP medium, and immediately fixed in 2.5 % glutaraldehyde (G5882) diluted in 0.1 M sodium cacodylate (20840) for 30 min at 37°C. After fixation, subgroups of 15 spheroids were washed three times in 500 µL of PBS-BSA, stained with Haecht 33342 (2 µg/mL; B2261) for 15 min and observed under a confocal microscope (Zeiss Confocal LSM 780 Airyscan). For each spheroid, four Z-stacks pictures were analyzed using ImageJ software (v1.54d). The total number of bound spermatozoa was counted and normalized to the total spheroid surface area, yielding sperm density values expressed as spermatozoa per mm². Four biological replicates, each derived from different pools of cows and culture days, were analyzed.

To specifically assess sperm interaction with cilia, spheroids from each region were co-incubated with spermatozoa for 1 h. Subgroups of 15 spheroids were collected for immunostaining of acetylated alpha-tubulin as described above. Confocal microscopy was used to evaluate (i) the proportion of spermatozoa bound to ciliated cells, (ii) the number of bound spermatozoa per ciliated cell and (iii) the proportion of ciliated cells interacting with spermatozoa. Three biological replicates, each derived from different pools of cows and culture days, were analyzed.

## 2.8. Characterization of sperm-spheroid interaction by scanning electron microscopy

After 1 h of co-incubation with spermatozoa, spheroids from each region were fixed in 2.5 % glutaraldehyde diluted in sodium cacodylate solution and stored at 4 °C. Spheroids were subsequently washed in sodium cacodylate and placed onto sterile cover-glasses discs (Marfenfeld, VWR, France). Samples were dehydrated through a graded ethanol series (50–100 %), then subjected to critical point drying under CO₂. Dried samples were mounted onto aluminum scanning electron microscopy stubs (15 mm diameter × 6 mm M4; Micro to Nano, Haarlem, Netherlands) using carbon adhesive discs (Agar Scientific, Oxford Instruments SAS, Gometz-la-Ville, France) and sputter-coated with gold-palladium (Polaron SC7640, Milexia, Verrières-le-Buisson, France) for 220 s at 10 mA. Imaging was performed using a field emission gun scanning electron microscope operating in secondary electron at 2 kV (Hitachi S4500, Milexia, Verrières-le-Buisson, France). Bound spermatozoa were classified into the following categories: (i) bound exclusively by the head or any other part to cilia; (ii) bound exclusively by the head or any other part to microvilli (non-ciliated cells). Additionally, bound spermatozoa were evaluated for acrosomal integrity (presence of an intact acrosome without perforations) and morphological abnormalities (absence or defects of the flagellum or mid-piece) as previously reported in bull semen [43,44]. Two replicates from different pools of cows and culture days were analyzed.

## 2.9. Statistical analysis

Data are represented as means ± SEM. Coefficients of variation (%) among replicates were calculated as the ratio of the standard deviation
to the mean, multiplied by 100. All statistical analyses were performed using Rstudio (v. 2023.06.2) and GraphPad Prism (v6.07) softwares. Data normality was evaluated using the Shapiro-Wilk test. The effect of the anatomical region (ampulla vs. isthmus) on spheroid diameter, total cell number, cell height, bound sperm density (at 1 and 6 h) and number of bound spermatozoa per ciliated cell was analyzed using a linear mixed model in which the region was considered as a fixed effect while replicate and spheroid were treated as random effects. The effect of time of bound sperm density was analyzed using the same model in which the time was considered as a fixed effect. For total cell number and cell height, the lme function (from the nlme package) was used due to normal distribution of data. For sperm densities and number of bound spermatozoa per ciliated cell comparisons, the aovp and nparcomp functions (from the lmPerm and nparcomp packages) were used. Differences in the proportions (%) of ciliated cells, spermatozoa bound to ciliated cells and ciliated cells interacting with spermatozoa between isthmic and ampullary spheroids were analyzed by chi-square tests. A p-value ≤ 0.05 was considered as significant.

# 3. Results

## 3.1. Morphometric and cell characterization of isthmic and ampullary spheroids

The selected isthmic and ampullary spheroids observed under a stereomicroscope displayed very similar morphology, outward ciliary beating and movements (Fig. 1 and Movies 1–2). Confocal microscopy analysis revealed no statistical difference between isthmic and ampullary spheroids in their mean diameter (122.7 ± 4.1 vs. 121.0 ± 2.7 µm, respectively), total cell number (130.9 ± 6.7 vs. 133.2 ± 5.8, respectively) and cell height (12.7 ± 1.0 vs. 12.0 ± 0.4 µm, respectively) (Fig. 2A-B). The coefficients of variation among the four replicates for isthmic and ampullary spheroids were 12.3 % and 8.4 % for mean diameter, 9.2 % and 7.6 % for total cell number, and 15.9 % and 7 % for cell height, respectively.

Supplementary material related to this article can be found online at doi:10.1016/j.repbio.2025.101057.

The epithelial cell marker cytokeratin was immunodetected in all spheroid cells as a cytoplasmic network (Fig. 2D-E), whereas the signal for vimentin, a stromal cell marker, was not detected (Fig. 2F-G). The proportion of ciliated cells, to which spermatozoa are expected to preferentially bind [3], was on average 22.5 % higher in ampullary than in isthmic spheroids (24.5 ± 2.1 % vs. 20.0 ± 1.3 %, respectively, p < 0.0001) with a coefficient of variation of 14.9 % and 11.1 %, respectively among replicates (Fig. 2C).

## 3.2. Proteomic profile of isthmic and ampullary spheroids

Overall, 6914 proteins were identified in the spheroids, of which 6878 (99.5 %) were shared between the two anatomical regions (see list of shared and region-specific proteins in Supplementary Table S1).

Among all proteins, 1829 proteins (26 %) were predicted to be secreted via conventional or unconventional pathways, and 650 (9 %) as membrane proteins. Of the cell membrane proteins, 391 were predicted to be transmembrane while 288 were peripheral or lipid-anchored proteins. Furthermore, 630 proteins (9 % of total proteins) were previously identified in the Human Protein Atlas associated with cilia (Fig. 3 and Supplementary Table S1).

The 15 most abundant proteins in both ampullary and isthmic spheroids included heat shock proteins (HSP90B1, HSPA5, HSPA8), annexins (ANXA1, ANXA2) and cytoskeleton proteins (ACTA1, ACTB, KRT7, KRT8, KRT18, KRT19) (see Supplementary Figure S1). A total of 1010 (14 %) proteins were differentially abundant between anatomical regions (p-value ≤ 0.05; fold change ratio ≥ 1.3). Among those proteins, 517 (including HSP90B1 and HSPA5) were more abundant in isthmic compared to ampullary cells, and 493 showed the opposite pattern of

variation (mean fold-change ratios of 1.53 ± 0.02 and 1.60 ± 0.03, respectively; Fig. 4 for top-20 most abundant DAPs and Supplementary Table S2 for all DAPs). Among these 1010 DAPs, 81 (8 %) were previously identified in human cilia, 175 (17 %) were predicted to be cell membrane proteins and 324 (32 %) to be secreted (Supplementary Table S2).

## 3.3. Gene Ontology enrichment analysis of proteins overabundant in isthmic spheroids

Metascape analysis of proteins overabundant in isthmic compared to ampullary epithelial spheroids revealed highly enriched GO BP terms, including multiple metabolic processes (small molecule biosynthetic process, amide metabolic process, carboxylic acid metabolic process, organic hydroxy compound metabolic process, nucleobase-containing small molecule metabolic process, carbohydrate derivative catabolic process, glycosylation, response to nutrients levels), import into cell, membrane organization, regulation of apoptotic signaling pathway and cellular homeostasis (Fig. 5A and Supplementary Table S3). The most significantly enriched GO CC terms included the lytic vacuole, endoplasmic reticulum lumen, secretory granule lumen, vacuolar lumen, mitochondrial matrix and apical part of cell (Fig. 6A and Supplementary Table 3).

## 3.4. Gene Ontology enrichment analysis of proteins overabundant in ampullary spheroids

Proteins overabundant in ampullary compared to isthmic epithelial spheroids showed enrichment in GO BP terms primarily related to cytoplasm and membrane organization (microtubule cytoskeleton organization, supramolecular fiber organization, epithelial cell differentiation, cell junction organization, regulation of cytoskeleton organization, cell-cell adhesion, regulation of cell projection organization, gland development) as well as DNA repair and some metabolic processes (Fig. 5B and Supplementary Table S3). Consistent with this, the most significantly enriched GO CC terms included focal adhesion, cell-cell junction, centriole, microtubule, protein-DNA complex, desmosome and apical part of cell (Fig. 6B and Supplementary Table S3).

## 3.5. Characterization of sperm-spheroid interaction and morphology of bound spermatozoa

Densities of bound spermatozoa per spheroid surface unit were determined after 1 and 6 h of co-culture. At 1 h, the density of bound spermatozoa was on average 33 % higher on ampullary than on isthmic spheroids (6330 ± 992.5 vs. 4238 ± 566.4 spermatozoa/mm², respectively; p < 0.0001; Fig. 7A). At 6 h, the density of bound spermatozoa significantly decreased in both regions (p < 0.0001) but remained 45 % higher on ampullary than on isthmic spheroids (1110 ± 203.9 vs. 615.8 ± 119.8 spermatozoa/mm²; p < 0.0001; Fig. 7A).

The ability of spheroid cilia to interact with bull spermatozoa was assessed after 1 h of co-culture, followed by washing to remove unbound and loosely bound spermatozoa. In total, 95 % and 97 % of bound spermatozoa were attached to isthmic and ampullary cilia, respectively, and this was confirmed by analysis of scanning electron microscopy pictures (Figs. 7B and 8A-B and Table 1). As evaluated after alpha-tubulin immunostaining and confocal microscopy observation, approximately the same high proportion of multiciliated cells was occupied by sperm heads (89 % vs. 92 % on isthmic and ampullary spheroids, respectively; Fig. 7C and Table 1). However, the multiciliated cells of ampullary spheroids were occupied by a higher number of bound spermatozoa compared to those of isthmic spheroids (means ± SEM of 8.5 ± 0.4 vs. 5.8 ± 1.0, respectively, p < 0.0001, Fig. 7D). Scanning electron microscopy pictures evidenced that a large majority of bound spermatozoa had an intact acrosome and no morphological
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** Epithelial and cellular characteristics of isthmic and ampullary spheroids. Histograms present the means ± standard error of the mean. (A) Cell height (N = 4 replicates using different pools of oviducts), (B) total cell number and (C) proportions of ciliated cells in isthmic and ampullary spheroids (N = 3 replicates using different pools of oviducts). Each dot or square represent one sample of isthmic and ampullary spheroids, respectively. Pictures show (D-E) a positive immunodetection for pan-cytokeratin (in green), an epithelial marker, in the cell cytoplasm and (F-G) a negative detection for vimentin, a fibroblast marker, in both groups of spheroids. Inserts show the isotype controls. The nuclei of cells were stained with Hæchst 33342 (in blue). See Fig. S2 for the positive control of the anti-vimentin antibody.</figcaption></figure>
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** Numbers and subcellular localization of proteins identified in isthmic and ampullary spheroids. A protein was considered as identified with a log₂ abundance > 0 in minimum two biological replicates in each region. Prediction of subcellular localization was performed using the Outcyte [39] and DeepLoc [40] tools online and the protein sequences as inputs. The human protein atlas for "cilium" (https://www.proteinatlas.org/) was used as database. The Venn diagram was performed using the jvenn tool online (https://jvenn.toulouse.inra.fr/app/example.html) [41].</figcaption></figure>

abnormalities (Fig. 8 and Table 1). However, 5 % of spermatozoa bound to isthmic cilia displayed an apparent reacted acrosome, whereas this proportion was 2 % among those bound to ampullary cilia (p < 0.05; Figs. 8E and 8F and Table 1).

## 4. Discussion

Physiological *in vitro* models are needed to study gamete-oviduct interactions during the periconception period in cattle. The main results of this study are that: 1) spheroids derived from both regions of the oviduct contain well-differentiated epithelial cells with distinct proteomic profiles; 2) isthmic and ampullary spheroids support interactions with morphologically normal, acrosome-intact sperm heads on their cilia; 3) ampullary spheroids displayed a higher proportion of ciliated cells and sperm binding capacity than isthmic spheroids.

Cells composing the isthmic and ampullary spheroids exhibited a well-defined cytoplasmic network-like signal for pan-cytokeratin, a broad-spectrum antibody recognizing multiple cytokeratin proteins. Keratins are cytoplasmic intermediate filaments that provide mechanical strength to maintain cell shape and polarity [45]. Proteomic analysis identified 18 keratins in both spheroid groups, with keratins 7, 8, 18 and 19 among the most abundant proteins. Conversely, vimentin, an intermediate filament protein predominantly expressed in mesenchymal cells and associated with epithelial-to-mesenchymal transition [46], was not detected in spheroid cells, in line with our previous reports on isthmic spheroids cultured up to 10 days [17,19]. Positive immunostaining was observed in the stromal connective tissue of bovine oviduct sections, excluding lack of cross-reactivity of the anti-vimentin antibody used (Supplementary Figure S2). However, vimentin was detected by nanoLC-MS/MS in pooled spheroids. The reason for this discrepancy between proteomic and immunohistochemistry is unclear. We hypothesize that some stromal cells present in the mucosal fragments initially cultured persisted within spheroids and were detected by nanoLC-MS/MS in pools of 50 spheroids.

Compared with the monolayer model [3,21], isthmic and ampullary spheroids feature a 3D structure, with well-differentiated epithelial cells (apical-basal polarity and presence of cilia) up to several days in culture, allowing more physiological studies on gamete-oviduct interactions. In addition, a manual selection based on spheroid diameter (100–150 µm) and shape (round with a cavity, without folds), allowed to obtain spheroids with similar and reproducible total cell numbers for both ampullary and isthmic parts of the oviduct. This standardization offered repeatable apical surfaces for sperm interaction among replicates,

making spheroids a more reliable *in vitro* model than oviduct explants of varying shape and size. To better reflect physiology, oviducts used in this study were collected from cows at the pre-ovulatory stage of cycle, when cows are typically inseminated. The average proportion of ciliated cells was 22.5 % higher in ampullary than in isthmic spheroids (24.5 vs. 20 %, respectively). Similarly, scanning electron microscopy observation of bovine oviduct epithelium at the pre-ovulatory phase evidenced 20 % more ciliated cells in the ampulla than in the isthmus [5]. However, one limitation of spheroids is the lower proportions of ciliated cells than the ones reported *in vivo* after scanning electron microscopy (60 % in ampulla and 50 % in isthmus) [5] or after FOXJ1 immunostaining, a marker of ciliated cells (around 40 % in both isthmus and ampulla) [47] of the epithelium prior to ovulation. After 3 days of culture, only a subpopulation of spheroids formed was selected for standardization, possibly explaining the differences in the proportions of ciliated cells found *in vivo*. Another limitation of the spheroids is the absence of oviduct mucosa cells, which probably play a role in the physiology of the epithelial cells. The mean cell height in spheroids was also lower than that reported in oviduct sections from cyclic cows before ovulation (12–12.6 µm here vs. 30–35 µm *in vivo*), with no difference between anatomical region [5]. The organization of cells around a central cavity in spheroids may explain this reduced height. Hormonal stimulation mimicking the pre-ovulatory phase has been shown to increase cell height from 5 to 20 µm in bovine oviduct epithelial cells cultured on polycarbonate membranes in microfluidic systems [48]. Further studies are needed to evaluate the impact of hormonal stimulation on cell morphology and function in oviduct epithelial spheroids.

A total of 6914 proteins were identified in spheroids, representing the most comprehensive proteomic analysis of mammalian oviduct epithelium. Of these proteins, 1010 (14 %) were differentially abundant between ampullary and isthmic epithelial cells. These findings align with transcriptomic data showing region-specific gene expression regulation in bovine oviduct epithelium [25–27]. Around 4000 differentially expressed genes were identified between isthmic and ampullary epithelial cells at day 3 post-estrus [26]. In addition, up to 354 (12 %) proteins in the bovine oviductal fluid differed in abundance between the isthmus and ampulla [49]. Those proteomic differences may be partly due to the higher average proportions of ciliated cells in ampullary spheroids. In accordance, the GO analysis of the proteins overabundant in the ampullary spheroids pointed to the « microtubule cytoskeleton organization » as the most enriched biological process and the « microtubule » term was among the most enriched term for cellular components. However, the difference in the proportions of ciliated cells between ampullary and isthmic spheroids was moderate and the predicted functions of overabundant ampullary proteins evidenced many other biological processes and cellular components, which were not related to the ciliary function. Taken together, these data support the presence of region-specific surface proteins and secretions within the oviduct lumen, supporting specialized interactions with gametes and embryos during the peri-conception period.

Among the proteins overabundant in isthmic spheroids, the heat shock proteins HSP90B1 and HSPA5 were the two most abundant ones (Fig. 4A). The proteomic analysis of the oviduct fluid revealed that several HSPs, including HSP90B1 and HSPA5, were among the most abundant proteins during the peri-ovulatory period of cycle in cows [49]. In addition, HSP90B1 and HSPA5 were previously identified as sperm-interacting proteins in the bovine oviduct fluid [14] and recombinant HSPA5 has also been shown to bind to human sperm [50]. These two proteins were also identified among potential proteins responsible for the hardening of the oocyte zona pellucida in the bovine oviduct fluid [51], suggesting a role in limiting polyspermy during fertilization. Of note, the proteins overabundant in isthmic epithelial cells were mainly localized in the secretory machinery (endoplasmic reticulum, secretory granule lumen, vacuolar lumen, apical part of cells) and involved in small molecule biosynthesis and metabolic processes. Similarly, upregulated genes in isthmic cells from cows at day 3
<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Top 20 differentially abundant proteins between isthmic and ampullary spheroids. (A) Proteins overabundant in isthmic compared to ampullary epithelial cells; (B) Proteins overabundant in ampullary compared to isthmic epithelial cells. Proteins were considered as differentially abundant with a Student's unpaired *t*-test's p-value ≤ 0.05 and a fold-change ratio ≥ 1.3. Each dot or square represent one sample of isthmic and ampullary spheroids, respectively.</figcaption></figure>
<figure><img src="image_9.png" /><figcaption>**Proteins overabundant in isthmic compared to ampullary spheroids**</figcaption></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /><figcaption>**Proteins overabundant in ampullary compared to isthmic spheroids**</figcaption></figure>

<figure><img src="image_12.png" /><figcaption>**Fig. 5. Biological processes overrepresented among differentially abundant proteins between isthmic and ampullary spheroids.** Gene Ontology enrichment analysis of Biological Process was carried out using Metascape [42] and the *Homo sapiens* database.</figcaption></figure>

post-estrus were involved in the synthesis of small molecules like nitrogen compounds, lipids and nucleotides, and their related proteins mainly localized in the Golgi and cytoplasmic vesicles [26]. This suggests that isthmic spheroids provide a suitable environment for the early embryo, in line with their beneficial effect on embryo development and morphology after 8 days of co-culture [17,18]. On the other hand, proteins overabundant in ampullary spheroids were mainly involved in

cytoskeleton remodeling, cell-cell junction and DNA repair. Similarly, bovine ampullary cells analyzed on day 3 post-estrus displayed an overrepresentation of genes involved in cytoskeleton remodeling and DNA damage response [26], suggesting physiological conservation of these functions in ampullary spheroids during culture. Therefore, co-culture with isthmic and ampullary spheroids is expected to differentially influence gamete and embryo physiology, though this requires
<figure><img src="image_13.png" /></figure>

Proteins overabundant in isthmic
compared to ampullary spheroids

<figure><img src="image_14.png" /></figure>

<figure><img src="image_15.png" /></figure>

Proteins overabundant in ampullary
compared to isthmic spheroids

<figure><img src="image_16.png" /><figcaption>**Fig. 6.** Cellular components overrepresented among differentially abundant proteins between isthmic and ampullary spheroids. Gene Ontology enrichment analysis of Cellular Component was carried out using Metascape [42] and the *Homo sapiens* database.</figcaption></figure>

further investigation.

After crossing the utero-tubal junction, a sperm subpopulation binds to isthmic epithelial cells forming a reservoir, while others migrate towards the ampulla, where the cumulus-oocyte complex transits from the infundibulum at ovulation [52,53]. In cattle, fertilization may occur *in vitro* without oviduct components, but *in vivo*, sperm pre-binding to the oviduct epithelium likely precedes fertilization. *In situ* observations in mice suggest that sperm migration implies successive binding

release from isthmic then ampullary epithelial cells [6]. Given the average bovine oviduct length (around 20 cm), proportions of luminal ciliated cells (25–50 %) and the abundance of mucosal folds filling the lumen, it is probable that most spermatozoa interact with cilia from both isthmus and ampulla during their migration toward the oocyte.

Although equal numbers of spermatozoa were used for binding as
says with spheroids, the density of bound spermatozoa was 33–45 %
higher on ampullary compared to isthmic spheroids after 6 h of co-
<figure><img src="image_17.png" /><figcaption>**Fig. 7.** Characteristics of sperm-spheroid interactions analyzed by confocal microscopy. (A) Densities of bound sperm on isthmic and ampullary spheroids after 1 and 6 h of co-incubation (N = 4 replicates using different pools of oviducts); (B) Proportions of spermatozoa bound to ciliated cells in isthmic and ampullary spheroids (N = 3 replicates using different pools of oviducts); (C) Proportions of multiciliated cells interacting with spermatozoa (N = 3 replicates using different pools of oviducts); (D) Number of bound spermatozoa per multiciliated cell in isthmic and ampullary spheroids (N = 3 replicates using different pools of oviducts). The data are presented as means ± SEM of 3 to 4 biological replicates and different letters indicate significant differences (p-value ≤ 0.05). Each dot or square represent one sample of isthmic and ampullary spheroids, respectively. (E-F) Confocal microscopy images of isthmic and ampullary spheroids after 1 h of co-incubation with frozen-thawed washed spermatozoa from a pool of 3 bulls. In green: cilia immunodetected with an acetylated alpha-tubulin antibody; in blue: sperm nuclei stained with Hœchst 33342.</figcaption></figure>
<figure><img src="image_18.png" /><figcaption>**Fig. 8.** Scanning electron microscopy pictures of sperm-spheroid interactions. (A,C,E) Isthmic and (B,D,F) ampullary spheroids. (A,B) Acrosome-intact spermatozoa interacting by their head with cilia; (C-D) Acrosome-intact spermatozoa interacting by their head with microvilli; (E-F) Acrosome-reacted spermatozoa (white arrows) and sperm with an abnormal flagella (black arrow) interacting with microvilli (E) or cilia (F). Scanning electron microscopy pictures were obtained after 1 h of co-incubation with frozen-thawed washed spermatozoa from a pool of 3 bulls.</figcaption></figure>
<table><caption>**Table 1**

**Characterization of sperm-spheroid interactions by scanning electron microscopy.** Statistical differences between isthmic and ampullary spheroids were carried out using chi-square tests (p < 0.05). A total of 239 and 240 spermatozoa bound to 34 isthmic and 39 ampullary spheroids, respectively, were analyzed.</caption><thead><tr><th></th><th>Isthmic<br>spheroids</th><th>Ampullary<br>spheroids</th></tr></thead><tbody><tr><td><strong>Proportions of spermatozoa bound to cilia<br>(%) including:</strong></td><td>94.8 ± 1.2</td><td>96.3 ± 1.7</td></tr><tr><td>- bound exclusively by the head (%)</td><td>94.0 ± 2.0</td><td>96.3 ± 1.7</td></tr><tr><td>- bound by the head and flagella (%)</td><td>0.8 ± 0.8</td><td>0</td></tr><tr><td><strong>Proportions of spermatozoa bound to non-ciliated cells/microvilli (%) including:</strong></td><td>5.2 ± 1.2</td><td>3.7 ± 1.7</td></tr><tr><td>- bound exclusively by the head (%)</td><td>2.3 ± 0.3</td><td>2.0 ± 1.3</td></tr><tr><td>- bound by the head and flagella (%)</td><td>2.9 ± 0.9</td><td>1.8 ± 0.4</td></tr><tr><td><strong>Proportion of bound spermatozoa with a<br>reacted acrosome (%)</strong></td><td>4.6 ± 0.6<sup>a</sup></td><td>1.8 ± 0.4<sup>b</sup></td></tr><tr><td><strong>Proportion of bound spermatozoa with a<br>flagella defect (%)</strong></td><td>0.9 ± 0.2</td><td>1.6 ± 1.6</td></tr></tbody></table>

culture. This higher sperm density likely reflects, in part, the greater proportion of ciliated cells on ampullary than on isthmic spheroids (24.5 % vs. 20 %), and also the higher number of bound spermatozoa per ciliated cell. suggesting a higher density of sperm receptors on ampullary cilia. In addition, some of the 493 proteins more abundant in ampullary compared to isthmic spheroids likely contribute to this differential sperm binding. Annexin A4 (ANXA4), previously identified as a candidate sperm receptor on bovine oviduct epithelium [54], is one such example. Nevertheless, the molecular mechanisms underlying sperm binding to the reservoir in mammals are complex and include glycans and phospholipids in addition to proteins [55,56].

Practically, assessing sperm density on the oviduct epithelium after insemination is challenging due to the inaccessibility of oviducts in live animals and the low sperm number distributed along the long oviduct lumen (thousands of spermatozoa along a 20 cm tube in cows). After insemination, more spermatozoa are expected to bind to the isthmus, as it is the first segment encountered. In six cows slaughtered 2–4 h after insemination, 178 spermatozoa were bound to the isthmus compared to only 3 in the ampulla [3]. Our results suggest that bound spermatozoa progressively detached from cilia regardless of cell origin but remained attached longer to the ampullary compared to isthmic cells. Similarly, in *situ* observations in mice 3 h after mating showed that spermatozoa bound longer to the ampullary than to the isthmic lumen, where frequent detachment and reattachment occurred [6]. In most mammals, sperm binding to isthmic and ampullary epithelial cells was evaluated using oviduct epithelial monolayers [15,57–59], explants [3,11,16, 60–63] or membrane preparations [64] with inconsistent results. In cattle, higher numbers of spermatozoa bound to ampullary compared to isthmic epithelial monolayers during co-culture up to 15 h [3,15]. Using bovine oviduct explants, the density of bound spermatozoa was equivalent between isthmic and ampullary cells when oviducts were collected at the pre-ovulatory phase, but higher on ampullary compared to isthmic cells when oviducts were collected just after ovulation [3]. In horses, at both pre- and post-ovulatory stages, more spermatozoa bound to isthmic than ampullary epithelial explants after 48 h of co-culture [9]. In pigs, one study found equal number of spermatozoa bound to isthmic and ampullary explants [11], while another quantified more spermatozoa bound on isthmic than ampullary explants [61]. Differences in sperm preparation and incubation conditions likely explain interspecies and interstudy discrepancies. Moreover, the number of ciliated cells per replicate, which determines the number of sperm binding sites, was not reported in these studies, possibly contributing to variability. The majority of spermatozoa bound to spheroids from both regions were acrosome intact with no morphological abnormalities in the present study. This is in line with previous reports using oviduct epithelial monolayers [65] and explants [3,16] in cattle. Future studies will test whether spermatozoa selected by sperm binding to spheroids have higher ability

to fertilize oocytes *in vitro*.

In conclusion, this work provides a comprehensive characterization of two physiological models for studying gamete/embryo-oviduct interactions. Isthmic and ampullary epithelial spheroids exhibited similar epithelial and cellular properties, but displayed specific proteomic signatures, ciliary surface and ability to interact with spermatozoa. Further studies are needed to investigate whether binding assays using spheroids could be used to select a superior sperm subpopulation or predict bull fertility in the field.

## Financial support

This research was funded by the French National Research Agency (ANR) under the grant number ANR-23-CE20-0041-01 (SELECT project). Protim Core Facility was supported by structural grants from Infrastructures en Biologie Santé et Agronomie (IBiSA), Conseil Régional de Bretagne (through Biogenouest), European Regional Development Fund, Rennes Métropole and University of Rennes (through Biosit).

## Declaration of Competing Interest

All authors declare that they have no conflict of interest

## Acknowledgments

This work has benefited from the equipment and expertise of the Imaging facility « Plateau d'Imagerie Cellulaire » (PIC) of the PRC Unit. The authors warmly thank Albert Arnould for the collection of oviducts at the slaughterhouse, Olfa Ghazouhani (Synetics, France) for providing bull semen, Thierry Meylheuc and Renaud Fleurot for their expertise in microscopy and Lise Lépinay for technical support.

## Appendix A. Supporting information

Supplementary data associated with this article can be found in the online version at doi:10.1016/j.repbio.2025.101057.

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