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From in vitro development to accessible luminal interface of neonatal bovine-derived intestinal organoids

BMC Veterinary Research volume 21, Article number: 319 (2025) Cite this article

Abstract

Background

Intestinal organoids provide physiologically relevant in vitro models that bridge the gap between conventional cell culture and animal studies. Although these systems have been developed for adult cattle, their use in neonatal calves—who are particularly vulnerable to enteric disease—has not been well established. Neonatal diarrhea remains a major health concern in modern agriculture, yet age-appropriate models for studying its pathogenesis are lacking. Given that host–pathogen interactions vary with developmental stage, there is a need for culture systems that reflect the distinct biology of the neonatal gut. In this study, we developed intestinal organoids and organoid-derived monolayers from 14-day-old dairy calves to enable research on early-life intestinal function and disease.

Results

Organoids were successfully established from five intestinal sections of 14-day-old dairy calves using customized growth media and characterized by immunofluorescence and gene expression analyses. They remained viable for over 300 days of cryopreservation and were serially passaged at least 15 times. Rectal organoid-derived monolayers were further assessed by electron microscopy and barrier function assays, demonstrating stable transepithelial electrical resistance and controlled paracellular permeability.

Conclusions

Optimized methods for adult bovine intestinal organoids and rectal organoid-derived monolayers are applicable to neonatal intestinal epithelial stem cells. Organoids cultured from 14-day-old calves captured key aspects of the multicellularity and functionality of the native epithelium. Future work should focus on adapting monolayer culture methods for additional gut regions, particularly the proximal gastrointestinal tract. Neonatal rectal monolayers represent a promising platform for advancing veterinary research, agricultural innovation, and studies of zoonotic disease.

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Background

The study of intestinal biology has been transformed by the development of three-dimensional (3D) culture techniques of intestinal epithelial cells [1]. In these methods, primary intestinal stem cells from donor species intestinal crypts are cultured within an extracellular matrix (ECM), which supports the self-organization of the multipotent cells into 3D structures known as intestinal organoids. These organoids can be sustained over multiple culture passages without artificially applied immortalization and comprise the various cell types found in the natural intestinal epithelium, including enterocytes, goblet cells, Paneth cells, and enteroendocrine cells [2, 3]. Due to their multicellularity and structural complexity, organoids provide an advanced research platform that bridges traditional cell culture and in vivo gaps, making them particularly valuable for studying intestinal physiology and diseases [4, 5].

Bovine neonatal diarrhea is caused by a variety of enteric pathogens including viruses, bacteria, and protozoa [6]. In 2014, the United States Department of Agriculture reported that over 50% of health events in dairy calves less than one month old are attributed to enteric illnesses, leading to increased mortality, reduced farm efficiency, and negatively impacting the welfare of dairy cattle [7]. Moreover, dairy calves have been linked to outbreaks of human enteric infections caused by Cryptosporidium parvum [8, 9, 10], highlighting the connection of animal health on human health. Investigating these pathogen-host interactions requires robust models to better understand the factors leading to colonization, replication, and disease.

Intestinal organoid technology has been used as a model for bovine associated infectious agents, including bovine Coronavirus [11], Mycobacterium avium subspecies paratuberculosis [12], and gastrointestinal nematodes [13]. These advancements validate the utility of 3D culture techniques in bovine gastrointestinal infectious disease research. However, variability in study design with respect to organoid-donor age, breed, and culture media limits applicability to other research questions, specifically those related to bovine neonatal enteric infections [14, 15]. As cattle age, the maturing intestinal epithelium becomes resistant to infection by pathogens. For example, enterotoxigenic Escherichia coli (ETEC) is a major cause of neonatal diarrhea in calves less than one week old, with calves over one week of age developing resistance to disease [16]. This highlights the importance of refining intestinal organoid culture methods to account for age of cattle, ensuring robust and reliable protocols for researchers. Addressing these variables can improve the accuracy and applicability of organoid-based models to bovine infectious disease studies.

Another major challenge in intestinal organoid technology is accessing the apical luminal surface of the intestinal epithelial cells [17], which is critical for studying pathogen-host interactions [18]. Pathogens typically interact with the apical aspect of these cells, making it essential for researchers to have access to this surface [19]. One method to overcome this challenge is to culture monolayers from dissociated intestinal organoid cells [20, 21, 22]. To advance in vitro models for studying bovine neonatal diarrhea, practical methods are needed to develop and maintain intestinal organoids from neonatal donors and to generate organoid-derived monolayers for detailed functional studies. It remains unclear whether protocols optimized for adult cattle [23] are applicable to neonatal calves.

This study addresses these gaps by presenting methods for culturing intestinal organoids and subsequently generating organoid-derived monolayers from the intestinal epithelial cells of 14-day-old dairy calves. This age group was selected due to its high susceptibility to enteric pathogens such as bovine rotavirus and Cryptosporidium parvum [24], making it ideal for studying intestinal barrier function and host-pathogen interactions. Intestinal organoids were developed using in-house growth media from five sections of intestinal tract, with successful rectal organoid-derived monolayer culture. The organoids and monolayers were characterized with immunofluorescence, gene expression, electron microscopy, and monolayer barrier function assays, including transepithelial electrical resistance measurements and paracellular permeability assays.

Results

Establishment and cryopreservation of neonatal bovine intestinal organoids

Neonatal bovine intestinal stem cells, isolated from the duodenum, jejunum, ileum, colon, and rectum of five donor animals using biopsy-derived fresh tissue and custom-formulated growth media (Table 1), were successfully cultured into 3D intestinal organoids (Fig. 1A). Organoids from each gut section were cryopreserved for over 300 days in a solution containing 10% dimethyl sulfoxide (DMSO) in 90% fetal bovine serum (FBS) before being resuscitated (Fig. 1B). Resuscitation success varied between cryovials, with some yielding a high number of recoverable organoids, while others produced few or none. Post-cryopreservation, organoids from all gut sections were successfully passaged, with at least one line from each section maintained for 15 or more passages (Fig. 1C).

Table 1 Composition of organoid and monolayer culture media reagents
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Fig. 1
figure 1

Phase-contrast images of neonatal bovine intestinal organoid culture. (A) Phase-contrast images of growing organoids embedded in ECM droplets from the initial establishment for five sections of neonatal bovine gut. Organoids from passages 0 or 1 are shown. (B) Phase-contrast images of growing organoids successfully resuscitated after over 300 days of cryopreservation, demonstrating the capacity for long-term storage and revival. (C) Organoids resuscitated from cryopreservation-maintained growth and preserved their morphology over more than 15 serial passages. All images in (A), (B), and (C) were captured between days 4 and 12 of culture. Scale bar = 100 μm

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The culture success for each gut section from the five donor animals is summarized in Table 2. Intestinal organoids were successfully established from all sections of gut for each donor animal. Following cryopreservation, the recovery of at least a single viable organoid was successful in 80% of duodenum samples, 60% of jejunum samples, 80% of ileum samples, 80% of colon samples, and 100% of rectum samples. During serial passaging of resuscitated organoids, duodenal and jejunal organoids exhibited a loss of proliferative capacity, characterized by a dark, granular appearance (S1A Fig). The addition of 10% FBS improved both their morphology and proliferation capacity, enabling continued viable culture. Consequently, FBS supplementation was maintained throughout the culture of duodenal and jejunal organoids (Table 1). Utilizing this adaptation, the success rate of serial passaging after resuscitation was 75% of duodenum samples, and 33% of jejunum samples. In contrast, FBS supplementation was not necessary for ileum, colon, or rectum organoids to maintain their morphology and proliferative capacity, resulting in a 100% success rate for these sections.

Table 2 Summary of viability across stages of neonatal bovine intestinal organoid culture by intestinal segment
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Cellular characterization of neonatal bovine intestinal organoids

Immunofluorescence characterization was performed to confirm the muticellular composition of organoids. The analysis revealed diverse cell populations that self-organize with cellular polarity, self-replicate and differentiate into specific epithelial cell types (Fig. 2). Organoid cells express intercellular epithelial adherens junctions, evidenced by E-cadherin staining colocalizing with F-actin. The cells adopt basal-out polarity and form a central lumen, with enhanced F-actin staining directed toward the intraluminal space and 4’,6-diamidino-2-phenylindole dihydrochloride (DAPI)-stained nuclei positioned peripherally. The epithelial nature of the organoid cells is verified by epithelial cell adhesion molecule (EpCAM) staining. SRY-box transcription factor 9 (SOX9) staining of cells reveals the maintenance of stemness in many, but not all, cells, such as in the rectal organoid, where approximately half the cells are SOX9-positive, demonstrating cellular heterogeneity. Actively replicating cells are identified by 5-ethynyl 2’-deoxyuridine (EdU) uptake. Additionally, fluorescein labelled-Sambucus nigra agglutinin (SNA) staining for mucin is observed intracellularly and within the organoid lumen across all intestinal sections. This suggests goblet cell differentiation, with many directing their secretions toward the lumen, highlighting a mixed cell population within the organoids.

Fig. 2
figure 2

Immunofluorescence staining of neonatal bovine intestinal organoids. (A) Intestinal organoids cultured in organoid expansion media form basolateral epithelial adherens junctions (E-cadherin, green), and exhibit polarity with apical brush borders (F-actin, red) and basal nuclei (DAPI, blue). Intestinal organoids are of diverse cell populations, including epithelial cells (EpCAM, yellow), stem or transit-amplifying cells (SOX9, yellow), actively proliferating cells (EdU, cyan), and mucin-producing goblet cells (SNA, green). Scale bar = 50 μm. (B) To evaluate differences in the cellular makeup of organoids across gut sections, the number of cells positive for SOX9, EdU, and SNA was counted per field of view and normalized to the total number of nuclei. Between four and fifteen independent fields of view were counted from three technical and five biological replicates. Data are shown as mean ± SE for each gut section. Statistical significance was assessed using a one-way ANOVA across gut segments, followed by Tukey’s HSD post-hoc test (* p < 0.05, and ** p < 0.01)

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Quantification of positively stained specialized cells was performed for SOX9, EdU, and SNA to evaluate differences across organoid gut sections. All organoid cells positively stained for EpCAM, and this marker was excluded from quantification. The mean percentage of positively stained cells ± standard error of the mean (SE), and the range across all intestinal organoids were as follows: 62.30 ± 8.58%, range = 39.19–90.98% for SOX9; 32.20 ± 2.70%, range = 19.72–38.74% for EdU; and 15.64 ± 2.75%, range = 1.39–28.68% for SNA (Fig. 2A-C). Analysis of variance revealed a significant effect of gut section on EdU staining (F(4, 144) = 9.19, p < 0.0001). Post-hoc analysis using Tukey’s HSD test indicated statistically significant differences in EdU staining between duodenal organoids and all other gut sections (p = 0.0001–0.016) (Fig. 2B). Similarly, analysis of variance for SNA staining showed a significant effect of gut section (F [4, 58] = 12.21, p < 0.0001). Post-hoc analysis revealed significant differences between duodenal and ileal organoids (p < 0.0001), as well as between ileal and colonic organoids (p < 0.0001) (Fig. 2C). No statistically significant differences were observed in SOX9 staining across gut sections (Fig. 2A).

Gene expression analysis of OEM and ODM-treated organoids

Gene expression analysis was performed to confirm the cellular composition, regional identity, and potential functional characteristics of the organoids under expansion and differentiation culture conditions. Variability in the relative gene expression of target epithelial lineage markers was observed across the different sections of gut. The stem cell marker LGR5 was expressed at lower levels in ODM-treated organoids, with statistically significant differences in ileal (Mann-Whitney U test, W = 0, p < 0.0001) and rectal organoids specifically (Mann-Whitney U test, W = 4, p = 0.0005) (Fig. 3A). The enteroendocrine cell marker CHGA was expressed at higher levels in ODM-treated rectal (Mann-Whitney U test, W = 81, p < 0.0001) organoids. The Paneth cell marker LYZC was generally downregulated in ODM-treated organoids in all sections but no statistical significance noted (Fig. 3C). Interestingly, ODM treatment of colonic organoids tended to increase the expression of MUC2, the goblet cell marker, while differentiation media treatment downregulated MUC2 expression in the other organoids. However, no statistically significant differences in MUC2 expression were observed across ODM-treated organoids (Mann-Whitney U test, ileum: W = 18, p = 0.0503; colon: W = 34, p = 0.6; rectum: W = 22, p = 0.1) (Fig. 3D). Finally, the intestinal epithelial cell marker FABP2 was upregulated in all ODM-treated organoids, with statistically significant increases in ileal (Mann-Whitney U test, W = 66, p = 0.02), colonic (Mann-Whitney U test, W = 67, p < 0.02), and rectal organoids (Mann-Whitney U test, W = 65, p < 0.03) (Fig. 3E).

Fig. 3
figure 3

Relative gene expression of neonatal bovine intestinal organoids cultured in expansion (OEM) or differentiation (ODM) media. To confirm the cellular composition, regional identity, and potential functional characteristics of the organoids under expansion and differentiation culture conditions, gene expression of marker genes for (A) stem cell (LGR5), (B) enteroendocrine cell (CHGA), (C) Paneth cell (LYZC), (D) goblet cell (MUC2), and (E) mature enterocyte (FABP2) was measured via RT-qPCR. Reference genes used for normalization included ACTB, GAPDH, and RPL0. Organoids were cultured for 6 days in OEM, followed by an additional 4 days in either OEM or ODM. Gene expression data were collected from one (duodenum and jejunum) or three (ileum, colon, and rectum) biological replicates, with three technical replicates per biological replicate. Data is summarized as mean ± SE, with individual measurements presented as points. Duodenal and jejunal organoid results are representative only. Statistical significance was assessed using the Mann-Whitney U test or unpaired t-test, as appropriate (* p < 0.05, and ** p < 0.01)

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Rectal-organoid derived monolayer development and barrier integrity assessment

In our study, rectal organoids demonstrated the most stable development compared to organoids from other intestinal segments (Table 2). Based on this observation, we sought to establish sTable 2D monolayers derived from neonatal bovine rectal organoids. Using optimized culture methods previously described for adult bovine rectal monolayers [22, 25], we successfully generated sTable 2D monolayers from neonatal bovine rectal organoids. Cells reached confluence by day 1 of culture (Fig. 4A) and the monolayers were maintained till day 6, both for monolayers cultured in MEM for the entire culture duration and monolayers cultured in MDM after switching the medium on day 3. No morphological differences between MDM and MEM conditions were observed by phase-contrast microscopy at day 6 of culture (Fig. 4B).

Fig. 4
figure 4

Evaluation of stable 2D monolayers derived from neonatal bovine rectal organoids. (A&B) Phase-contrast images of rectal organoid-derived monolayers on cell culture inserts at days 1 (A) and 6 (B) of culture. Monolayers were cultured in monolayer expansion media (MEM) for 3 days, followed by continued culture in MEM or switch to monolayer differentiation media (MDM) for an additional 3 days. No considerable morphologic variation was observed between the conditions. Scale bar = 50 μm. (C) Monolayers were cultured in MEM for 6 days, with transepithelial electrical resistance (TEER) measurement daily, and apparent permeability (Papp) measured on days 1, 3 and 5. Measurements were compared between consecutive days. After 3 days, monolayers demonstrated intact barrier integrity, as indicated by resistance to 4 kDa FITC-dextran diffusion, coinciding with increased TEER. (D) Comparison of TEER measurements between MEM- and MDM-cultured monolayers. MDM-treated monolayers maintained equivocal TEER measurements as MEM-cultured monolayers on days 4 and 6 (p > 0.05). Significant difference was only noted temporarily on day 5 between the group (p < 0.05). Data are shown as mean ± SE. Two technical replicates from three biological replicates were evaluated. Statistical significance was assessed using the Wilcoxon signed-rank test or paired t-test, or the Mann-Whitney U test or unpaired t-test, as appropriate (* p < 0.05, and ** p < 0.001)

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Barrier integrity of the monolayer was confirmed using a combination of transepithelial electrical resistance (TEER) measurements and an apparent permeability (Papp) assay, which assessed the diffusion of a 4 kDa fluorescein isothiocyanate (FITC)-dextran marker across the monolayer. TEER measurements taken daily revealed significant increases from day 1 to day 2 (65.2 ± 26.6 vs. 188.6 ± 34.9 Ω*cm2, t-test [5] = -9.597, p = 0.0002), and from days 2 to 3 (188.6 ± 34.9 vs. 302.9 ± 21.5 Ω*cm2, t-test [5] = -5.044, p = 0.003). The TEER measurements peaked on day 4 (317.1 ± 35.9 Ω*cm2, t-test [5] = -0.326, p = 0.76 vs. day 3) and were maintained through day 5 (290.6 ± 24.2 Ω*cm2, t-test [5] = 1.304, p = 0.25). Additionally, the observed changes in TEER values coincided with increased resistance to the diffusion of 4 kDa FITC-dextran, as indicated by a decrease in Papp value. By day 3 of culture, the monolayer exhibited a significant decrease in Papp value compared with day 1 (5.2 ± 0.7 vs. 1.4 ± 0.3 × 10− 7 cm/s, Mann-Whitney U test, W = 0, p = 0.002) and it was also sustained through day 5 (0.9 ± 0.2 × 10− 7 cm/s, t [5]= -2.525, p = 0.198 vs. day 3) (Fig. 4C).

When comparing barrier integrity of the monolayers cultured in MEM and MDM, TEER measurements taken daily did not reveal statistical difference between the two culture conditions on both day 4 (Wilcoxon signed-rank test, V = 4, p = 0.22) and 6 (t [5] = -0.348, p = 0.74). Significantly lower TEER measurements in MDM-treated monolayers (206.3 ± 11.6 Ω*cm2) compared to MEM monolayers (290.6 ± 24.2 Ω*cm2) were only noted temporarily on day 5 (t [5] = -4.917, p = 0.004) (Fig. 4D).

Cellular characterization of MEM and MDM-treated rectal organoid-derived monolayers

To assess the cellular and molecular characteristics of MEM and MDM-treated rectal organoid-derived monolayers, a series of analyses were conducted. Immunofluorescence revealed that rectal organoid-derived monolayers display a diverse cellular population, including actively proliferating cells (EdU), stem or transit-amplifying cells (SOX9), mucin-producing goblet cells (SNA), and cells expressing epithelial adherens junctions (E-cadherin) and epithelial cell adhesion molecule (EpCAM) (Fig. 5A&B). These findings confirm the presence of similar cell types to those observed in the originating neonatal bovine intestinal organoids (Fig. 2). Z-stack images revealed intracellular localization of SNA staining, apical brush border (F-actin) with basal nuclei (DAPI), and basolateral expression of adherens junctions, further confirming the retention of cellular characteristics seen in the organoids, even within the 2D monolayer structures.

Fig. 5
figure 5

Immunofluorescence and gene expression analysis of 2D rectal monolayers cultured in expansion (MEM) or differentiation (MDM) media. (A) Immunofluorescent staining of 2D rectal monolayers cultured in both MEM and MDM revealed diverse cell populations, including actively proliferating cells (EdU, cyan), stem or transit-amplifying cells (SOX9, yellow), mucin-producing Goblet cells (SNA, green), epithelial adherens junctions (E-cadherin, green), and intestinal epithelial cells (EpCAM, yellow). Phalloidin (red) was used to stain F-actin, highlighting cellular borders where applicable, and DAPI (blue) was used to label nuclei. Top-down and z-stack images are shown. Scale bar = 25 μm. (B) Quantification of cells positive for EdU, SOX9, and SNA per field of view, normalized to the total number of nuclei. Ten independent fields of view were analyzed from three biological replicates. Data are shown as mean ± SE. No statistically significant difference was observed between MEM and MDM-cultured monolayers. (C) Gene expression analysis of marker genes for stem cells (LGR5), enteroendocrine cells (CHGA), Paneth cells (LYZC), goblet cells (MUC2), and intestinal epithelial cells (FABP2) was performed using RT-qPCR. Reference genes (ACTB, GAPDH, and RPL0) were used for normalization. Rectal monolayers were cultured for 3 days in MEM, followed by an additional 3 days in either MEM or MDM. Two technical replicates from three biological replicates were evaluated. Data are shown as mean ± SE. Statistical significance was assessed using the Mann-Whitney U test or unpaired t-test, as appropriate (* p < 0.05, and ** p < 0.01)

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The mean percentage of positively stained cells ± SE across MEM and MDM-treated rectal monolayers were: EdU − 15.1 ± 1.2% (MEM) and 17.8 ± 2.8% (MDM); SOX9–61.0 ± 3.2% (MEM) and 61.7 ± 3.5% (MDM); SNA − 11.0 ± 1.1% (MEM) and 15.0 ± 1.7% (MDM) (Fig. 5B). No statistically significant differences were observed between MEM and MDM-treated rectal monolayers (Mann-Whitney U test, W = 199-258.5, p = 0.12–0.99).

Gene expression analysis revealed downregulation of LGR5 expression and upregulation of epithelial lineage marker genes, namely CHGA, LYZC, MUC2, and FABP2, in MDM-treated monolayers compared to MEM-cultured rectal organoid-derived monolayers (Fig. 5C). Notably, MUC2 expression was significantly upregulated in MDM-treated monolayers (Mann-Whitney U test, W = 31, p = 0.04) relative to MEM-cultured monolayers. However, no statistically significant differences were observed in the expression of LGR5 (t [10] = -2.005, p = 0.07), CHGA (t [10] = 1.240, p = 0.24), LYZC (Mann-Whitney U test, W = 22.5, p = 0.51), or FABP2 (Mann-Whitney U test, W = 23, p = 0.45).

Further cellular detail was assessed using electron microscopy. Scanning electron microscopy (SEM) evaluation of the monolayers revealed the development of microvilli and mucin-like granules, indicative of functional epithelial maturation (Fig. 6A&B). Transmission electron microscopy (TEM) confirmed the formation of intercellular tight junctions, reinforcing the establishment of a robust barrier function (Fig. 6C). Additionally, the presence of well-formed microvilli and glycocalyx suggests an enhanced surface area for nutrient absorption (Fig. 6D). These ultrastructural features highlight the capacity of the rectal organoid-derived monolayers to closely mimic key characteristics of the native intestinal epithelium, further supporting the functional differentiation observed in the cellular and molecular analyses.

Fig. 6
figure 6

Electron microscopic characterization of rectal organoid-derived 2D monolayer. Representative scanning (A & B) and transmission (C & D) electron microscopic images of rectal organoid-derived 2D monolayers at day 6 of culture. Rectal monolayers were cultured for 3 days in MEM, followed by an additional 3 days in MDM, and subsequently fixed with glutaraldehyde (A & B) or glutaraldehyde/paraformaldehyde/alcian blue (C & D). (A) Cells developed diffuse apical microvilli projections. (B) Multifocal dense aggregates with characteristic webbing formed over the apical surface, suggesting mucin-like granules. Both low- and high-power magnification images are shown for (A) and (B). Scale bar = 1 μm. (C) Transmission electron microscopy shows apical microvilli (MV) and a basally located nucleus (N) in the lower magnification image (left). Tight junctions (TJ) between two cells are visible near the apical surface in the higher magnification image (right). Scale bars = 400 nm. (D) Additional emphasis on the apical epithelial morphology, highlighting well-developed microvilli (MV) with an associated glycocalyx (GLX). Scale bar = 400 nm

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Discussion

This study successfully translated optimized culture conditions for adult bovine intestinal organoid development to neonatal bovine intestinal specimens from five distinct intestinal regions [23]. The neonatal organoids self-organized into polarized luminal structures containing diverse epithelial lineages, including progenitor cells and goblet cells, while exhibiting activities such as self-replication and mucin production. Furthermore, the development of rectal monolayers from these organoids addresses a major challenge in accessing the luminal surface, an essential feature for studying host-pathogen interactions [18, 26, 27]. These rectal monolayers displayed cellular polarization, diverse cell populations, and functional characteristics reflective of the organoids from which they were derived. Collectively, these findings highlight the potential of these models to advance in vitro studies of neonatal bovine intestinal biology.

The development of neonatal bovine intestinal organoids from tissue specimens sampled using biopsy forceps and cultured with laboratory-formulated growth media reaffirms previously established methods for adult bovine organoids [23]. This approach provides an alternative to the glass slide method for isolating intestinal crypts [11, 12, 28], which might be somewhat impractical outside of a laboratory setting, such as during field sampling of cattle. Additionally, the use of in-house growth media presents a more cost-efficient alternative to commercially available options, thereby improving accessibility for researchers.

An important aspect of this study was the long-term preservation of established organoids for over 300 days in liquid nitrogen using a freezing medium composed of 10% DMSO and 90% FBS [29]. Although the success rates of resuscitation following cryopreservation were similar to previous reports [21, 23, 29], not all individual cryovials from the same donor were successfully resuscitated, emphasizing the importance of storing multiple samples per donor. Nonetheless, this study demonstrated that at least one cell line from each gut section could be serially passaged for at least 15 passages after resuscitation without altering morphological characteristics or losing proliferative capacity (Fig. 1; Table 2). This finding allows extended time between sampling and experimentation, providing researchers with greater flexibility in scheduling.

During the serial passaging of duodenal and jejunal organoids, proliferation capacity gradually declined, with organoids becoming darkened, granular and eventually disintegrating (S1A Fig). This was likewise encountered during organoid development from the more proximal intestinal tract of adult bovids [23]. It was hypothesized that serial passaging following the thaw process depleted certain growth factors or nutrients that were not accounted for in the organoid growth media. Supplementing the growth media with 10% FBS sustained duodenal and jejunal organoid growth and enabled further serial passaging. This concentration aligns with standard cell culture supplementation rates [30, 31]. While the optimal FBS concentration is unclear, this finding suggests that FBS supplementation may address viability challenges in small intestinal organoids. However, the batch variability of serum components [32] and the potential contamination risks [33] warrant careful consideration, as these factors may impact experimental outcomes [34].

The successful generation of neonatal bovine rectal organoid-derived functional 2D monolayers represents a significant advancement in modeling host-pathogen interactions [35] of neonatal bovids [24]. Barrier integrity of these monolayers was confirmed through stable TEER and Papp values after 3 days of culture (Fig. 4C). TEER measures ion flow across the cellular barrier, while Papp assesses the movement of biomolecules, including polysaccharides [36]. In intestinal monolayers, maturation typically results in increasing TEER values alongside decreases in Papp, reflecting enhanced barrier integrity [37]. The observed rise in TEER values and decline in Papp, indicating resistance to 4 kDa FITC-dextran permeability, confirmed the functionality and integrity of the barrier after 3 days of culture in the presented culture conditions.

In the rectal monolayer, TEER values were slightly higher than those previously reported for adult bovine organoid-derived monolayers in the same laboratory [22], consistent with results from other bovine-origin intestinal monolayers [38, 39], though lower than those observed in certain other bovine-origin monolayers [40]. This variation may be attributed to differences in individual animal donors or laboratory technique. However, in human intestinal organoid research, donor variability in hormone secretion, cell differentiation and metabolic profiling appears to be manageable [41]. The discrepancies observed between neonatal rectal monolayers and other bovine-origin monolayers could be influenced by factors such as variations in media temperature and composition, equipment calibration, and cell passage number [42]. Overall, while inter-laboratory and donor variability may pose challenges for direct comparisons of TEER and Papp values, the simultaneous rise in TEER alongside a decrease in Papp confirms the establishment of stable barrier integrity in the rectal monolayer. This confirmation provides investigators with an accessible physiological intestinal-lumen interface after approximately 3 days of culture.

Both organoids and monolayers contained a variety of cell types identified through immunofluorescence, including goblet cells, stem or transit-amplifying cells, and actively replicating cells. Notably, differences in goblet cell numbers were observed between ileal and duodenal, as well as ileal and colonic organoids. While goblet cells are distributed throughout the intestinal tract, they are more densely concentrated in the large intestine [43]. Therefore, the higher goblet cell density in colonic organoids compared to ileal organoids is expected, though the unexpectedly high number of goblet cells in duodenal organoids remains unexplained. While goblet cell differentiation from intestinal stem cells can be inhibited by infectious agents [44] or promoted by disease [45], neither was observed in the organoid cultures, suggesting other factors may be at play. Evaluating the responses of duodenal organoids to differentiation factors such as Notch and Wnt signaling may provide additional insight [46]. The epithelial cells exhibited adhesion molecules crucial for the formation of adherens junctions, mirroring the characteristics of in vivo epithelial cells [47]. Additionally, ultrastructural analysis of the rectal monolayers confirmed the presence of microvilli, glycocalyx, mucin-like granules, and tight junctions. In vivo, microvilli are critical for nutrient absorption [48], while the glycocalyx and mucin serve as essential mediators, segregating gut microflora from the intestinal epithelial cells [49]. Collectively, these findings suggest the organoids and derived monolayers replicate the complexity of the intestinal epithelium, highlighting the robustness of this in vitro model for investigations in microbial interactions and gut function.

Wnt-3a is a key cytokine that plays an essential role in maintaining intestinal stem cell proliferation and suppressing differentiation [50, 51]. Recombinant murine Wnt-3a has been employed as a practical source of Wnt signaling in previous intestinal organoid cultures [52], despite reports of limited signaling activity [53], particularly in the absence of lipid stabilizers [54]. It is possible that bovine intestinal organoids do not require high levels of active Wnt-3a, or that the presence of conditioned R-spondin and Noggin is sufficient to stabilize and support the minimal Wnt signaling that may occur [55]. The inclusion of FBS in the culture media for duodenal and jejunal bovine neonatal organoids (S1 Fig) may have contributed to a Wnt-stabilizing effect. Additionally, spatiotemporal gradients of Wnt-3a have been shown to contribute to morphological and growth heterogeneity in human intestinal organoids [56]. Such effects, along with interspecies variability, may help explain the thick-walled phenotype of neonatal bovine intestinal organoids observed in OEM (Fig. 1), in contrast to the thinner-walled, cystic morphology reported in human-derived intestinal organoid cultures, which is typically associated with a proliferative stem cell state [57]. Future studies could investigate the responsiveness of bovine intestinal organoids to conditioned Wnt-3a and define optimal inclusion concentrations, which may reveal interspecies differences relative to human or murine models. Nonetheless, the inclusion of recombinant murine Wnt-3a in OEM was effective for culturing adult bovine intestinal organoids [23] and was used in the present study to maintain stem cell proliferative capacity.

In contrast, the removal of Wnt-3a, R-spondin and Noggin in ODM promotes differentiation resulting in reduced stem cell proliferative capacity [1, 58]. To evaluate epithelial cell differentiation towards a more mature state, gene expression analysis was conducted to confirm cellular composition and potential functional characteristics. As expected, ODM-treated neonatal bovine intestinal organoids showed downregulation of the stem cell marker gene, LGR5, across all gut sections, alongside upregulation of the enterocyte marker, FABP2, indicative of differentiation. These results aligned with observations from adult bovine ODM-treated organoids [23].

Paneth cells, which support intestinal stem cell maintenance through Wnt signaling molecules, typically differentiate in the basal crypts where Wnt concentrations are highest [59, 60]. The observed downregulation of the Paneth cell marker LYZC in ODM-treated organoids may reflect insufficient autogenous Wnt production, highlighting the importance of mesenchymal support in vivo. Intriguingly, LYZC gene expression was higher in organoids derived from the large intestine compared to those from the small intestine (Fig. 3). While Paneth cells are the primary source of lysozyme [61], the expected distribution of Paneth cells is typically restricted to the proximal gastrointestinal tract [62], and our findings may reflect a unique feature of the neonatal bovine intestinal epithelium. Alternatively, lysozyme expression may originate from epithelial cells other than Paneth cells. Goblet cells and enterocytes have been reported to produce lysozymes, particularly under inflammatory conditions [63]. This unexpected finding warrants further investigations into potential interspecies variations in intestinal epithelial cell populations.

Other marker genes for differentiated cell types, such as enteroendocrine cells (CHGA), and goblet cells (MUC2), showed less predictable expression patterns. Specifically, MUC2 expression was generally downregulated in all ODM-treated organoids except colonic organoids. This unexpected result may relate to the 4-day ODM exposure exceeding the optimal lifespan of goblet cells, reported as 3 to 7 days in humans [64], potentially reducing RNA yields. Further optimization of ODM exposure duration is necessary to refine differentiation protocols and ensure reliable sampling of ODM-treated bovine organoids.

Several limitations contextualize these findings. First, the absence of donor tissue characterization limits comparisons to in vivo counterparts. Although high fidelity between organoids and donor tissues has been reported [12, 14, 38], discrepancies have also been observed in other studies [11]. Second, direct comparisons of cellular composition were not performed between rectal organoids and their organoid-derived monolayer counterparts, nor across organoids at different passages. Organoids are generally expected to maintain a more stem-cell-rich, proliferative state in their 3D structure compared monolayers [65]. Third, the exclusive use of male donors introduces sex bias, which may affect organoid behavior [66]. Fourth, the in-house culture media, which includes recombinant murine Wnt-3a and nicotinamide, may not be fully optimized for bovine organoids and could influence phenotypic and growth characteristics in ways that differ from organoids of other species [56, 67]. Fifth, variable FBS use across gut sections complicates cross-sectional comparisons. Lastly, reliance on single biological replicates for gene expression in duodenal and jejunal organoids limits statistical robustness, likely due to low RNA yields.

Future directions include evaluating the effects of recombinant versus conditioned Wnt-3a on neonatal bovine intestinal organoids, optimizing media FBS and nicotinamide supplementation, directly comparing organoid cell composition to native donor tissue using histological staining and single-cell RNA sequencing to assess fidelity, and examining how cell composition evolves with serial passaging and in development into organoid-derived monolayers. These efforts would strengthen the utility of neonatal bovine intestinal organoids in broader research applications, including validation of the model to investigate host-pathogen interactions.

Despite these limitations, this study successfully developed and characterized bovine intestinal organoids from 14-day-old dairy calves with key in vivo features, including self-organization, multicellularity, and specialized activity. The successful generation of functional organoid-derived rectal monolayers provides a novel platform for investigating host-pathogen interactions and gut physiology, offering significant potential for advancing neonatal bovine intestinal biology.

Conclusion

This study marks a significant advancement in the development of neonatal bovine intestinal organoids, particularly through the successful generation of rectal organoid-derived monolayers that demonstrate functional barrier properties. By leveraging donor animals at an age vulnerable to bovine neonatal diarrhea, this research introduces a novel and physiologically relevant bovine intestinal model, offering new opportunities for complex studies in intestinal biology. These models replicate the in vivo intestinal epithelium through their multicellularity, self-replication, and differentiation into mature epithelial cell types. This development not only opens new avenues for research in veterinary medicine, agricultural practices and zoonotic disease but also provides a more accurate platform for bridging the gap between in vitro and in vivo research. Moving forward, the integration of these intestinal organoids into neonatal bovine enteric research presents an exciting opportunity to make meaningful contributions to both animal and human health.

Methods

Ethics statement

The use of animals in this study was in accordance with the guidelines and approval of the Washington State University’s (WSU) Institutional Animal Care and Use Committee (IACUC, Protocol Number: 7100).

Donor animals and intestinal tissue sampling

Intestinal tissue samples were collected from five 14-day-old male Holstein dairy calves procured from the WSU-affiliated dairy farm. Details on the animals’ signalment are available in S1 Table. Following euthanasia on-site, sections of the intestine, including the duodenum, jejunum, ileum, colon, and rectum, were identified, dissected, and removed en bloc to a clean work surface. After incising each gut section longitudinally to expose the intestinal mucosa, 15–30 pieces of tissue were sampled using Hildyard Post-Nasal Biopsy Forceps (Med-Plus). This field-adapted method eliminated the need for fragile glass slides commonly used in laboratory settings. Tissue samples were immediately placed in ice-cooled sterile wash medium composed of Dulbecco’s phosphate-buffered saline (PBS, Gibco) supplemented with 1x penicillin/streptomycin (Gibco) and 25 µg/mL gentamicin (Gibco), and transported on ice for further processing in the laboratory.

Isolation of intestinal crypts and establishment of intestinal organoids

Intestinal tissue samples were processed as previously described [23]. Tissue specimens from each section of intestines from all donor animals were washed of mucus and intestinal content with sterile wash medium until the supernatant was visibly clear. Tissue specimens were suspended in 20 mM ethylenediaminetetraacetic acid (EDTA) solution and minced into small fragments using micro-dissecting scissors (Fisher Scientific) in a standard tissue culture dish. Tissue fragments were then collected into a 15 mL conical tube.

Intestinal crypts were isolated from tissue fragments using a tube rotator (Fisher Scientific) set at 10 rpm and 4 °C for variable durations dependent on the intestinal segment: 15 min for the jejunum and ileum, 30 min for the colon, or 60 min for the duodenum and rectum. Following gravitational settling of tissue fragments, suspended intestinal crypts were transferred to a new 15 mL conical tube and centrifuged at 200 × g at 4 °C for 5 min to form a pellet. The supernatant was removed, and the pelleted crypts were resuspended in phenol red-containing, non-growth factor-reduced, LDEV-free extracellular matrix (ECM; Corning® Matrigel® Basement Membrane Matrix, product number 354234, Corning) and placed in 30 µL droplets onto a 24-well tissue culture plate.

After incubating the ECM-embedded intestinal crypts for 10 min at 37 °C to polymerize the ECM, 500 µL of organoid expansion media (OEM) was added to the well. The composition of OEM was previously described [23] and is outlined in Table 1. For the first two to four days of organoid culture, OEM was supplemented with 10 µM rho-associated kinase inhibitor (ROCKi) Y-27,632 (STEMCELL Technologies) and 100 nM glycogen synthase kinase-3 beta (GSK3B) inhibitor CHIR99021 (Sigma-Aldrich). The tissue culture plate was incubated in a humidified incubator set at 37 °C with 5% CO2. Organoid growth was monitored with a phase-contrast microscope (DMi1, Leica) daily, and the culture media was replaced every other day.

Organoid subculture

To passage organoids, the ECM was dissolved by replacing OEM with cold Cell Recovery Solution (Corning) and incubating at 4 °C for 30–60 min. After incubation, the freely suspended organoids were centrifuged at 200 × g at 4 °C for 5 min to form a pellet, and the supernatant removed. Next, TrypLE Express (Gibco) was added and incubated at 37 °C for 1 min (or 40 s for ileal organoids) in a water bath to digest cell-to-cell adhesions. Immediately after incubation, ice-cold basal medium (Table 1) was added to quench the enzymatic reaction, and the mixture was centrifuged at 200 × g at 4 °C for 5 min to pellet the organoids. The supernatant was removed, and the dissociated organoid pellet was resuspended in ice-cold ECM, and pipetted 30–50 times to ensure even suspension in ECM. The organoid-ECM suspension was placed in 30 µL droplets onto a 48-well tissue culture plate, with 300 µL of OEM added after ECM polymerization. The organoid subculture procedure was performed approximately once every 6–10 days.

Organoid cryopreservation, resuscitation and differentiation

For long-term storage of organoids in liquid nitrogen, organoids were recovered from ECM as earlier described. After washing with basal medium, organoids were resuspended in freezing medium composed of 10% dimethyl sulfoxide (DMSO) in 90% fetal bovine serum (FBS) as previously described [23, 29]. The organoid suspension was transferred into cryovials and placed into a cell freezing container (Corning) at -80 °C overnight. Frozen cryovials were then transferred to liquid nitrogen.

To resuscitate frozen organoids, cryovials were thawed in a 37 °C water bath for 1–2 min. The thawed organoid suspension was then diluted with approximately 5 mL of basal medium, transferred to a 15 mL conical tube, and centrifuged at 200 × g at 4 °C for 5 min. The supernatant was removed, and the organoids were washed once with basal medium before being cultured as described above. Organoid growth and proliferation were monitored with a phase-contrast microscope (DMi1, Leica) daily. Lack of observable organoid growth over a period of 10–20 days was considered a non-viable culture.

To evaluate the differentiation of constituent cells, organoids were first cultured in OEM for 6 days, then transferred to organoid differentiation medium (ODM) for an additional 4 days. ODM was prepared similarly to OEM, but omitting Wnt3a, SB202190, and nicotinamide [23, 58].

For duodenal and jejunal organoids, 10% FBS was supplemented to OEM and ODM following resuscitation (Table 1).

Rectal organoid-derived 2D monolayer generation and differentiation

Rectal organoid-derived monolayers were generated from three biological replicates and evaluated as previously described [22, 25]. Briefly, 24-well cell culture inserts with 0.4 μm pores (Falcon) were pre-coated by applying 100 µL of 2% (vol/vol) ECM-supplemented basal medium per insert, followed by incubation at 37 °C with 5% CO2 for 1 h. The extracellular matrix used for insert coating was identical to that used for organoid culture. The coating media was then replaced with monolayer expansion medium (MEM) (Table 1), and incubated at 37 °C with 5% CO2 overnight.

Rectal organoids were recovered from the ECM droplet as described above. The recovered organoids were dissociated into a single-cell suspension using TrypLE Express supplemented with 10 µM of Y-27,632 placed in a 37 °C water bath for 10 min. The enzymatic reaction was quenched with basal medium, and the suspension was immediately filtered through a 70 μm cell strainer (Fisher Scientific). After counting cells with a hemocytometer and adjusting the cell concentration, the cells were seeded into the pre-coated cell culture insert apical chamber at 3 × 105 cells/well in 200 µL of MEM. The basolateral chamber of each well received 500 µL of MEM. The culture medium in both apical and basolateral chambers was replaced every other day.

To induce cellular differentiation, the culture medium in both apical and basolateral chambers was switched to monolayer differentiation medium (MDM) (Table 1) on Day 3 of culture. Monolayer formation and maintenance were monitored and recorded daily over 6 days using a phase-contrast microscope (DMi1, Leica).

Epithelial barrier integrity evaluation

Transepithelial electrical resistance (TEER) measurements and paracellular permeability assays were performed to evaluate the barrier integrity of the monolayers according to previously described protocols [25]. TEER was measured daily using an epithelial Volt-Ohm Meter (Millicell ERS-2, Millipore AG) for both MEM and MDM cultured monolayers. Direct measurements in ohms were converted to Ω*cm2 by subtracting the blank values from the monolayer measurements and multiplying by the cell culture insert surface area (cm2).

Paracellular permeability assays were conducted on days 1, 3 and 5 for MEM cultured monolayers using 0.5 mg/mL 4 kDa fluorescein isothiocyanate (FITC)-dextran (Sigma-Aldrich). The fluorescent intensity of the basolateral medium, reflecting the amount of FITC-dextran that diffused across the monolayer, was measured every 20 min over 60 min using a microplate reader (SpectraMax i3x, Molecular Devices) with excitation and emission wavelengths of 495 and 535 nm, respectively. The apparent permeability coefficient (Papp) (cm/s) was calculated by converting the measured values to concentrations (µg/mL) using a standard curve, and then normalizing based on the cell culture insert surface area (cm2), the initial FITC-dextran concentration in the apical chamber (µg/mL) and the time allowed for diffusion (s).

TEER measurements and permeability assays were performed with at least two technical replicates and three biological replicates. TEER values were compared between MEM and MDM cultured monolayers measured on the same day. Papp values were compared between consecutive time points of the assay.

Immunofluorescence staining

Organoids at Day 6 post-subculture in OEM, and both MEM and MDM cultured monolayers were fixed and stained using previously described methods [23, 25]. At room temperature, cells were fixed with 4% paraformaldehyde (Thermo Scientific) for 15 min, permeabilized with 0.3% Triton X-100 (Thermo Scientific) for 15 min, and blocked with 2% bovine serum albumin (Cytiva) diluted in PBS (BSA-PBS) for 1 h. Primary rabbit antibodies against epithelial cell adhesion molecule (EpCAM) (1:200, Abcam) and SRY-box transcription factor 9 (SOX9) (1:250, Abcam), along with fluorescent markers against E-cadherin (1:200, BD Biosciences) and Sambucus nigra agglutinin (SNA) (1:100, Vector Laboratories) were diluted in 2% BSA-PBS, and applied to the permeabilized organoids. The samples were incubated at 4 °C overnight, protected from light.

After incubation, the samples were rinsed twice with PBS. A secondary antibody (Alexa Fluor 555-conjugated Goat Anti-Rabbit IgG H&L, 1:1000, Abcam) diluted in 2% BSA-PBS was applied at room temperature for 1 hour. Nuclei were stained with 4’,6-diamidino-2-phenylindole dihydrochloride (DAPI) (1:1000, Thermo Scientific) and F-actin was stained with Alexa Fluor 647-conjugated phalloidin (1:400, Invitrogen) according to the manufacturer’s specifications.

For the 5-ethynyl 2’-deoxyuridine (EdU) staining assay (Click-iT EdU Cell Proliferation Kit, Invitrogen), growing organoids were incubated with EdU following the kit instructions for 3 h. Subsequent staining followed the manufacturer’s recommendations.

For all organoid staining protocols, samples were washed with PBS before mounting on glass-bottom plates (Matsunami) with ProLong Gold Antifade Mountant (Invitrogen). The stained monolayer cell culture insert membrane was excised from the insert using a scalpel blade and mounted on glass slides with ProLong Gold Antifade Mountant (Invitrogen). The labelled cells were visualized with a white light point scanning confocal microscope (TCS SP8-X, Leica) and analyzed using Leica Application Suite X (LASX) software (Leica). For organoids, one biological replicate was evaluated for jejunum samples, and three biological replicates for duodenum, ileum, colon and rectum samples. For monolayers, at least two technical replicates using three biological replicates were evaluated.

For EdU, SOX9 and SNA staining, the number of positively stained cells per unit area was quantified and normalized by the total number of nuclei to determine percent positive rates. Ten independent fields of view per sample were evaluated across three biological replicates, with results compared between sections of gut for organoids, and between MEM and MDM cultured monolayers.

Scanning and transmission electron microscopy of 2D monolayers

The ultrastructure of the MDM cultured monolayers was evaluated using scanning electron microscopy (SEM) and transmission electron microscopy (TEM) following previously described protocols [22, 68]. Briefly, the samples were fixed with either 2.5% (vol/vol) glutaraldehyde (Ted Pella) in 0.1 M sodium cacodylate buffer (Ted Pella) (GA), or 2% glutaraldehyde mixed with 2% paraformaldehyde and 1.05% Alcian blue in a 0.1 M sodium cacodylate buffer (GA/PFA/AB) overnight at 4 °C. SEM samples fixed with GA or GA/PFA/AB were post-fixed with either 1% osmium tetroxide (Electron Microscopy Sciences) in 0.1 M sodium cacodylate buffer or 1% osmium tetroxide and 0.5% tannic acid in 0.1 M cacodylate buffer, respectively, for 2 h at room temperature. TEM samples fixed with GA/PFA/AB were post-fixed with 1% osmium tetroxide and 1% ferrocyanide (Braun Knecht Heimann Company) in 0.1 M sodium cacodylate buffer overnight at 4 °C, followed by staining with 2% uranyl acetate (SPI Supplies). All samples were serially dehydrated in 30 to 100% ethanol. SEM samples were finally dried in hexamethyldisilazane (HDMS) (SPI Supplies), mounted on stubs, coated with platinum/palladium, and imaged with a Quanta 200 F SEM (FEI). TEM samples underwent final drying in propylene oxide (Electron Microscopy Sciences), infiltrated with Spurr’s resin (Ted Pella), and polymerized overnight at 60 °C. The samples were sectioned to 80 nm thickness and imaged using a Tecnai G2 20 Twin TEM (FEI).

Gene expression analyses

Gene expression analysis targeted specific intestinal epithelial cell markers including CHGA (enteroendocrine cells), FABP2 (intestinal epithelial cells), LGR5 (stem cells), LYZC (Paneth cells), and MUC2 (goblet cells) [11, 23, 38, 69, 70, 71]. Forward and reverse primers are listed in S2 Table. Following culture in OEM for 6 days, organoids were transferred to ODM for an additional 4 days. Organoids treated with ODM were compared to those cultured for 10 days in OEM, serving as the control group. Gene expression was evaluated between 2D monolayers cultured in MEM and MDM, following a previously described protocol [22].

Following dissolution of ECM as described in the subculturing section, organoids were collected by mechanical dissociation and centrifugation, and total RNA was extracted using the RNeasy Plus Mini Kit (Qiagen) following the manufacturer’s protocol. RNA quality and concentration was assessed with a Nanodrop One spectrophotometer (Thermo Scientific), and complementary DNA (cDNA) was synthesized using the High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems). The cDNA was then standardized for concentration across replicates. Quantitative reverse transcription polymerase chain reaction (RT-qPCR) was conducted using PowerUp SYBR Green Master Mix (Applied Biosystems) on a CFX96 Touch Real-Time PCR Detection System (Bio-Rad). The cycling conditions were: 2 min at 50 °C, 2 min at 95 °C, followed by 40-cycles of 15 s at 95 °C for denaturation and 1 min at 60 °C for annealing and extension. A final melting curve analysis at 95 °C confirmed the purity of the amplified products. ACTB, GAPDH, and RPL0 were used as reference genes to normalize the target gene products [11, 23, 70, 71]. Relative expression was calculated using the standard curve method [72]. Three biological replicates were used for ileal, colonic and rectal organoids, while one biological replicate was used for duodenal and jejunal organoids. For 2D monolayers, two technical replicates per sample were assessed, using three biological replicates.

Statistical analyses

Quantitative data were analyzed using R v.3.4.1 (The R foundation) and plotted using GraphPad Prism 10.1.1 (GraphPad Software). Data normality was assessed using the Shapiro-Wilk test. For immunofluorescence of organoids, comparisons between sections of gut were made using a one-way ANOVA followed by Tukey’s HSD post-hoc test. For immunofluorescence of 2D monolayers, RT-qPCR and TEER, comparisons between expansion (OEM, MEM) and differentiation (ODM, MDM) media conditions were conducted using either the Mann-Whitney U test or unpaired t-test, depending on data normality. Comparisons within expansion (MEM) media conditions were performed using the Wilcoxon signed-rank test or paired t-test for TEER, and the Mann-Whitney U test or unpaired t-test for Papp, based on data normality. Results are presented as the mean ± standard error of the mean (SE), with statistical significance defined as p ≤ 0.05.

Data availability

All relevant data generated or analyzed during this study are included in this published article (and its supplementary information files).

Abbreviations

ECM:

Extracellular matrix

OEM:

Organoid expansion media

ODM:

Organoid differentiation media

MEM:

Monolayer expansion media

MDM:

Monolayer differentiation media

FBS:

Fetal bovine serum

TEER:

Transepithelial electrical resistance

FITC:

Fluorescein isothiocyanate

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Acknowledgements

The authors would like to thank C. Sarinana and R. Griffiths for logistics in donor procurement, I. Nagao, Y. Tachibana, and T. Goyama for technical assistance in specimen collection and organoid development, and G. Weyman, V. Lynch-Holm and B. Wager for their technical expertise in confocal and electron microscopy.

Funding

This study was supported through the Washington State University (WSU) College of Veterinary Medicine Intramural Grant funded by the Robert R. Fast Endowed Food Animal Research and the United States Department of Agriculture National Institute of Food and Agriculture Animal Health and Disease Award to CRB, and the WSU Veterinary Clinical Sciences Student Research Grant to GDD, CSM, and YMA. Additional support was provided in part by the Office of the Director National Institutes of Health (K01OD030515 and R21OD031903) to YMA. The funding sources had no involvement in study design, data collection and analysis, publication decisions, or manuscript preparation.

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Author notes

  1. Gerald D. Dykstra and Minae Kawasaki contributed equally to this work.

Authors and Affiliations

  1. Department of Veterinary Microbiology and Pathology, College of Veterinary Medicine, Washington State University, Pullman, Washington, United States of America

    Gerald D. Dykstra & Claire R. Burbick

  2. Department of Veterinary Clinical Sciences, College of Veterinary Medicine, Washington State University, Pullman, Washington, United States of America

    Minae Kawasaki, Craig S. McConnel & Yoko M. Ambrosini

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  1. Gerald D. Dykstra
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  2. Minae Kawasaki
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  3. Claire R. Burbick
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  5. Yoko M. Ambrosini
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Contributions

GDD and MK generated and analyzed data, wrote and edited text, drafted and edited the figures and tables. CRB, CSM, and YMA facilitated resource allocation, assisted with editing the manuscript and provided technical expertise for the assays.

Corresponding author

Correspondence to Yoko M. Ambrosini.

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Ethics approval and consent to participate

The use of animals in this study was in accordance with the guidelines and approval of the Washington State University’s (WSU) Institutional Animal Care and Use Committee (IACUC, Protocol Number: 7100). Euthanasia was in accordance with the American Veterinary Medical Association’s 2020 Euthanasia Guidelines, with a primary method of captive bolt and secondary method of pithing.

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Dykstra, G.D., Kawasaki, M., Burbick, C.R. et al. From in vitro development to accessible luminal interface of neonatal bovine-derived intestinal organoids. BMC Vet Res 21, 319 (2025). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12917-025-04773-1

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BMC Veterinary Research

ISSN: 1746-6148

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