Nagao and Ambrosini ﻿
BMC Molecular and Cell Biology(2024) 25:14
https://doi.org/10.1186/s12860-024-00512-w

BMC Molecular andCell Biology

Open Access

RESEARCH

High‑fat diet enhances cell proliferationand compromises intestinal permeability
in a translational canine intestinal organoid
model
Itsuma Nagao1,2 and Yoko M. Ambrosini1*

Abstract
Background Emerging evidence underscores the responsiveness of the mammalian intestine to dietary cues, notably through the involvement of LGR5 + intestinal stem cells in orchestrating responses to diet-driven signals. However,
the effects of high-fat diet (HFD) on these cellular dynamics and their impact on gut integrity remain insufficiently
understood. Our study aims to assess the multifaceted interactions between palmitic acid (PA), cell proliferation,and the intestinal epithelial barrier using a canine colonoid model. Canine models, due to their relevance in simulating human intestinal diseases, offer a unique platform to explore the molecular mechanisms underlying HFD derived
intestinal dysfunction.
Results Canine colonoids were subjected to PA exposure, a surrogate for the effects of HFD. This intervention
revealed a remarkable augmentation of cell proliferative activity. Furthermore, we observed a parallel reduction
in transepithelial electrical resistance (TEER), indicating altered epithelium barrier integrity. While E-cadherin exhibited
consistency, ZO-1 displayed a noteworthy reduction in fluorescence intensity within the PA-exposed group.
Conclusions By employing canine intestinal organoid systems, we provide compelling insights into the impactof PA on intestinal physiology. These findings underscore the importance of considering both cell proliferative activity and epithelial integrity in comprehending the repercussions of HFDs on intestinal health. Our study contributes
to a deeper understanding of the consequences of HFD on intestinal homeostasis, utilizing valuable translationalin vitro models derived from dogs.
Keywords Canine, Intestinal organoids, High-fat diet, Palmitic acid

*Correspondence:
Yoko M. Ambrosini
yoko.ambrosini@wsu.edu
1
Department of Veterinary Clinical Sciences, College of Veterinary
Medicine, Washington State University, Pullman, WA, USA
2
Department of Veterinary Internal Medicine, Graduate Schoolof Agricultural and Life Sciences, The University of Tokyo, Tokyo, Japan

Background
The mammalian intestine exhibits responsiveness todietary cues [1–6]. Notably, LGR5 + intestinal stem cells
play a crucial role in reshaping intestinal composition in
response to diet-driven signals [1, 7, 8]. These cells regulate the production of daughter stem cells and progenitor cells, which subsequently differentiate into various
cell types within the intestine [9, 10]. The rise in obesity
has been closely linked to elevated levels of plasmatic freefatty acids (FFAs), which in turn lead to insulin resistance [11, 12] and tissue inflammation [13, 14]. However,© The Author(s) 2024. Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which
permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to theoriginal author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images orother third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line
to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory
regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of thislicence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

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little is known about how the adaptation of stem andprogenitor cells to pro-obesity high-fat diet (HFD) alters
the potential of these cells and leads to intestinal tumor
formation. Recent studies have highlighted the adverse
impact of fatty acid from HFD on gut homeostasis andintestinal epithelial function [1, 7]. Of particular interest is palmitic acid (PA), a prevalent saturated fatty acidin HFD made from animal fat [15, 16], renowned for its
lipotoxic effects across various organs [17–20]. Notably,
PA has been associated with compromised gut insulin
sensitivity [21] and gut barrier function [22]. Additionally, prolonged exposure to PA has been shown to hindercell differentiation in different tissues [7, 23, 24].
Despite growing recognition of the relationshipbetween metabolic disorders and intestinal cell dysfunction, the underlying molecular mechanisms remain
poorly elucidated [25, 26]. This knowledge gap can be
attributed to the dearth of translational animal andin vitro models specifically tailored to study obesity’s
impact on intestinal physiology. In this context, canine
models have gained prominence due to their relevanceas a spontaneous, large animal model for various humanintestinal diseases [27–33].
In canines subjected to a HFD, biopsies revealed anotable augmentation in stem cell characteristics [34].
Intriguingly, employing PA as a surrogate for mimicking the effects of a HFD yielded similar results, withelevated stemness characteristics coinciding with heightened expression of the peroxisome proliferator‐activatedreceptor‐γ, a pivotal nuclear receptor transcription factor[34]. Additionally, consumption of a HFD was reported
to decrease the gut barrier function in dogs, which isconsistent with findings in humans and mice [35]. Hence,
Investigation of the effects of HFD on intestinal function
in dogs can be expected to serve as a model for HFD inhumans to bridge the knowledge gap.
This study aims to address this gap by utilizing canine
intestinal organoid systems. These systems have beendeveloped to directly assess how nutritional factors, particularly palmitate-induced effect, impact both the development of stem cell populations and the integrity of theintestinal epithelial barrier. Through this proof-of-concept investigation, we endeavor to shed light on the intricate interplay between palmitate-induced effect, stem cell
dynamics, and epithelial barrier function in the context
of HFD-indcuced intestinal dysfunction.

effect of PA on the proliferative activity of intestinal organoids following PA exposure. Canine colonic organoids,
also known as colonoids, were cultured in organoid
medium. This culturing process was maintained fromthe point of passaging until day 4, allowing the organoids
to reach a suitable level of maturity. Subsequently, themedium was substituted with PA-containing medium,and the organoids were incubated for a duration of 24 h.
Figure 1A and B demonstrate the alteration in organoid dimensions following exposure to PA. At concentrations of 30 µM and 60 µM, PA elicited a notable impact
on organoid size. Specifically, the organoid dimensions
increased by 2.66-fold at 30 µM (p < 0.01) and 2.22-fold at
60 µM (p = 0.02) in comparison to the control group, suggesting increased cell proliferation in colonoid exposed
to PA.
In order to explore the underlying cause of the alteredorganoid size, we conducted an investigation into
whether increased stem cell pupulation played a role. The
gene expression of LGR5, a prominent stem cell marker,
using quantitative reverse transcription polymerase
chain reaction (qPCR) (Fig. 1C) was performed. Notably, the expression level of LGR5 was found to be significantly increased by exposure to 30 µM PA (1.61 times:
p < 0.01), although such an increase was not observed at
60 µM (p = 0.26). Building upon these observations, weestablished 30 µM as the dosage that manifests a representative effect of PA. This aligns with earlier in vivo
and in vitro investigations involving mice and dogs, asevidenced by previous studies [7, 34]. Consequently, we
opted to employ 30 µM PA for subsequent experiments.

Results
Palmitic acid increases organoid size and LGR5 expression
in canine colonoids

In this study, PA, a well-established main component of
the HFD [7, 26], was used to model the effect of HFD
in vitro. First, 3D organoids were used to examine theInfluence of palmitic acid on the proliferative activityof canine colonoids

To demonstrate increased cell proliferative activity ofcolonoids, we employed 5-ethynyl-2-deoxyuridine (EdU)
assay to assess the proliferative activity of colonoids. Following a 24-h exposure to 30 µM PA, an EdU assay was
conducted. The rate of EdU-positive cells exhibited a significant increase upon PA exposure (2.22 times: p < 0.01),
providing conclusive evidence of PA-induced heightened
proliferative activity (Fig. 2A and B).
Palmitic acid exposure and its impact on intestinal barrier
integrity

Subsequently, we examined the influence of PA on intestinal barrier function. To achieve this, a colonoid-derived
monolayer was established by seeding single cells enzymatically isolated from Matrigel-grown colonoids ontoa cell culture insert, a method previously documented[36]. These organoid-derived monolayers were cultured
in organoid medium until day 5, a point at which thetransepithelial electrical resistance (TEER) value reachedNagao and Ambrosini ﻿BMC Molecular and Cell Biology(2024) 25:14

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Fig. 1 Revealing enlarged dimensions and enhanced LGR5 expression in palmitic acid-exposed 3D canine colonoids. A Representative phase
contrast brightfield microscopy depicting 3D colonoids cultured in Matrigel following to 24 h of exposure to varying concentrations of palmitic acid(3, 30, 60 µM). For 0 µM, a carrier control consisting of 0.1% dimethyl sulfoxide (DMSO) in the media was used. Scale bar = 100 µm. B The organoid
area (µm.2) was quantified from five randomly selected fields for each condition, encompassing a minimum of ten organoids, as assessed using
ImageJ. This assessment was carried out across three biological replicates, each with three technical replicates. The error bars depict the standard
error of the mean. ** p < 0.01, * p < 0.05. C The gene expression level of the stem cell marker LGR5 in colonoids exposed to varying concentrationsof palmitic acid was quantified using qPCR. Each dataset was compiled from three biological replicates, each with three technical replicates. The
error bars represent the standard error of the mean. ** p < 0.01

Fig. 2 Assessment of proliferative activity in 3D canine colonoids exposed to palmitic acid. A Representative confocal microscopy images
displaying 3D colonoids cultivated within Matrigel following a 24-h exposure to 30 µM of palmitic acid. Actively proliferating cells were marked
by EdU staining (Cyan Blue), accompanied by nuclear counterstaining (DAPI; Blue). The left panel presents the image following 24 h of exposure
to the carrier control (DMSO), while the right panel depicts the image after 24 h of exposure to 30 µM of palmitic acid. Scale bar = 25 µm. B
The percentage of EdU-positive cells per nucleated cells in each condition was quantified using ImageJ. Each dataset was compiled from threebiological replicates, each with three technical replicates. For each condition, five randomly chosen fields of view at 40 × magnification wereselected from each sample, evaluating a minimum of ten organoids. The error bars represent the standard error of the mean. ** p < 0.01

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a stable state (Supplemental Fig. 1) [36]. On day 5, theapical medium was substituted with a PA-containing
medium, and the assessment of epithelial barrier integrity was conducted 24 h subsequent to this medium
alteration.
While no overt disparity in cellular morphology was
discernible between the Control and PA groups, as
observed through phase contrast microscopy (Fig. 3A), asignificant reduction in TEER value was evident after a
24-h PA exposure (0.79 times: p = 0.02) (Fig. 3B), indicating gut barrier integrity was decreased by addition of PA.

causes a decrease in gut barrier integrity as indicated by
TEER. To further examine changes in tight junction protein expression, we evaluated gene expression levels ofOccludin and Junctional adhesion molecule A (JAM-A),
which have been reported to be expressed in the canine
intestine other than ZO-1 and E-cadherin [39, 40]. However, no significant differences between the two groups
were detected in both Occludin and JAM-A (Supplemental Fig. 2).

Palmitic acid exposure leads to downregulation of tightjunction protein expression

To elucidate the underlying cause behind the observed
decline in epithelial barrier integrity, substantiated by
TEER measurements, the expression levels of E-cadherin and Zonula occludens-1 (ZO-1) were evaluatedby immunocytochemistry. These are proteins that play a
crucial role in maintaining the integrity and function ofcell–cell junctions, particularly in epithelial tissues [37,38].
In the control group, both ZO-1 and E-cadherin displayed distinct staining patterns at cell boundaries. However, within the PA-exposed group, E-cadherin exhibited
staining comparable to the control, whereas ZO-1 demonstrated a conspicuous reduction in fluorescence intensity (Fig. 4A).
The quantification of these fluorescence intensities
using Image J, revealed no significant variance in fluorescence intensity concerning E-cadherin. However, amarked reduction in fluorescence intensity was detected
for ZO-1 within the PA group (0.78 times: p < 0.01)(Fig. 4B), implying that decreased expression of ZO-1

Discussion
The presented results shed light on the impact of PA
exposure on canine colonoids as well as on canine colonoid-derived monolayers, extending our understanding of how dietary components can influence intestinal
homeostasis.
The enhanced cell proliferative activity observed in
response to PA exposure prompts discussions on thepotential molecular pathways and signalling mechanisms that mediate these effects. In studies using mice
HFD model, there is evidence suggesting that PA exposure can influence stemness in the intestinal epithelium[7]. This suggests a necessity for additional experimentation on canine organoids, particularly throughcolony formation assays, to more accurately evaluate
and quantify any enhancement in stem cell function
and the population of stem cells with self-renewal abilities following PA treatment. Elevated levels of PA havebeen correlated with alterations in the intestinal stem
cell niche and increased proliferation of stem cells [7].
This may have implications for tissue repair and regeneration but could also contribute to hyperproliferative
disorders if dysregulated. In parallel, canine investigations have revealed a similar trend where PA exposure

Fig. 3 Evaluation of epithelial barrier integrity in canine colonoid-derived monolayers. A Representative phase contrast brightfield microscopy
images of colonoid-derived monolayers cultured on cell culture inserts following a 24-h exposure to palmitic acid. The left panel presents the image
following 24 h of exposure to the carrier control (DMSO), while the right panel depicts the image after 24 h of exposure to 30 µM of palmitic acid.
Scale bar = 100 µm. B The intestinal epithelial barrier integrity was assessed through the measurement of TEER values. TEER values were recorded
after treating with media containing the carrier control (DMSO) or 30 µM palmitic acid for 24 h. This assessment encompassed three biologicalreplicates, each with three technical replicates. The error bars represent the standard error of the mean. * p < 0.05

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Fig. 4 Tight junction proteins in canine colonoid-derived monolayers. A Representative immunocytochemistry image depicting tight junction
proteins (ZO-1: Yellow, E-cadherin: Green) within a colonoid-derived monolayer treated with media containing the carrier control (DMSO)
or 30 µM palmitic acid. The left panel represents the control group, and the right panel depicts the palmitic acid-treated group. Scale bar = 25 µm.
B Fluorescence intensity in the entire field of view was quantified using ImageJ and subsequently normalized by dividing each intensity
by the average of the intensity of the control group. The upper panel displays the normalized fluorescence intensity of ZO-1, while the lower panel
showcases that of E-cadherin. This assessment was conducted across three biological replicates, each with three technical replicates. For eachcondition, five randomly selected fields of view at 63 × magnification were evaluated in each sample. The error bars represent the standard errorof the mean. ** p < 0.01

is linked to an amplification of stemness in intestinal
cells [34]. Notably, studies employing canine enteroids
have disclosed that PA exposure elicits an elevationin the expression of stem cell markers such as LGR5,
concurrent with heightened expression of peroxisome
proliferator-activated receptor-γ, a pivotal nuclear
receptor transcription factor [34]. Studies on mice havedemonstrated an upregulation of peroxisome proliferator-activated receptor-δ (PPAR-δ) in the intestinal
stem cells of those on a high-fat diet (HFD) [1]. Furthermore, when mouse intestinal organoids are treatedwith PPAR-δ agonists, an enhancement in stemness
is observed, mirroring that seen in organoids derived
from HFD-fed mice [1]. By integrating these findings
with observations from dogs on a HFD, we proposethat a similar upregulation of peroxisome proliferatoractivated receptors might contribute to the increased
proliferative activity observed in canine colonoids. Thissuggests a potential molecular pathway through which
PA exposure may influence stemness and intestinal

barrier integrity, highlighting the relevance of PPAR-δ
pathways in the observed phenotypic changes.
The compromised integrity of the intestinal barrier,
resulting from exposure to PA in canine colonoidderived monolayers, underscores the critical role oftight junction proteins in upholding epithelial integrity.
Among dietary components, PA has garnered significant attention within the scientific community, primarily due to its well-established lipotoxic effects on
various human organs. Notably, our study aligns withprevious research, revealing the growing understanding of PA’s impact on intestinal barrier function [41].
This underlines the complexity of its effects and lends
credence to our current findings showcasing compromised intestinal barrier integrity following PA exposure. The consistent outcomes further accentuate thepotential of utilizing dogs as a valuable in vitro model
to explore human intestinal homeostasis. However,the cause of the reduction of gut barrier integrity by
HFD remains to be elucidated. Some reports indicate

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that HFD-induced oxidative stress causes a decrease intight junction proteins in the intestinal tract [15, 42],while other reports indicate that tight junction proteinsare decreased in HFD models even though oxidative
stress is not induced [41]. Exploring the cause whichtight junction protein levels decrease in a human-relevant canine colonoid model could uncover the factors behind the decline in gut barrier integrity linked to
HFD in humans.
The congruence between our observations in canine
colonoids and the findings from in vivo human [15,43] and dog studies [34, 35] bolsters the translationalsignificance of our research. The consistent outcomes
observed across diverse models emphasize the potential relevance of our findings to human health, aligning with the principles of the One Health initiative. In
humans, feeding a HFD is known to impair intestinal
function or to be involved in tumor formation, but themolecular mechanisms of the effects of a HFD on theintestinal epithelium have not yet been fully elucidated
due to the lack of appropriate in vitro models. Although
HFD has been studied in vitro in mice using intestinal organoids [7, 26], mice and humans differ greatly
in their living environment, diet, and intestinal microflora composition, and there is a large gap as an animalmodel [44]. Therefore, we examined the direct interaction between PA and intestinal epithelial cells using
canine intestinal organoids, whose living environmentis close to that of humans and whose intestinal microbiota composition is relatively similar to that of humans[44, 45]. Hence, the canine in vitro HFD model in thisstudy is considered to be an important tool for elucidating the mechanism of intestinal dysfunction related
to HFD not only in dogs but also in humans.
While our canine colonoids and colonoid-derived
monolayers offer a valuable platform for nutritional
assays aimed at investigating early events in the intestinal alterations induced by HFD consumption, it isimportant to acknowledge the limitations inherent
in our models. Notably, our in vitro models comprisevarious cell types, yet they only consist of intestinal
epithelial cells. The intricate maintenance of intestinal
homeostasis involves a complex interplay among multiple key players, including the intestinal microbiome,
local immune cells, and intestinal epithelial cells [46].
It is imperative to recognize that PA, though animportant dietary component, represents just one facetof the diverse array of fatty acids present in HFD. Our
study’s focus on PA offers insight into specific mechanisms, but future research should explore the effects
of other fatty acids commonly found in HFD [15].
This expanded investigation could provide a morecomprehensive understanding of the multifaceted
impact of dietary fat on intestinal function.
Moreover, the exploration of more advanced in vitro
models, such as Gut-on-a-Chip systems [47, 48], presents a new avenue for research. These systems can better
mimic the complexity of the intestinal environment byincorporating aspects like fluid flow, mechanical forces,
and cellular interactions [48]. Integrating these elements
could enhance our understanding of how dietary factors
interact with the gut milieu.

Conclusion
In conclusion, this study demonstrated that exposure
to PA has diverse effects on canine colonoids as well ascolonoid-derived monolayers, mirroring similar findings
observed in mice colonoids and in vivo studies with dogs
[7, 34, 35]. It increases proliferative activity and impairs
intestinal barrier integrity by downregulating tight junction protein expression. These consistent findings acrossdifferent experimental settings contribute to our understanding of how PA, a prevalent component of HFD,
can potentially disrupt gut homeostasis and function in
direct way to the gut epithelium, without the involvement of gut microbiota.
Materials and methods
Generation and culture of canine colonoids

Intestinal biopsies were obtained by colonoscopy fromthree clinically healthy dogs presented to Washington
State University Veterinary Teaching Hospital Community Service for a dental procedure. These dogs were initially screened based on physical exam and blood work,no history of chronic disease (heart, kidney, liver, andintestinal) and be between 1 and 12 years old. If the dog
is clear of these aspects and deem appropriate to seek
the elective anesthetic procedure at Washington State
University Veterinary Teaching Hospital Community
Service, the dogs were considered clinically healthy. All
animal procedure were approved by Washington State
University Institutional Animal Care and Use Committee(IACUC approval: ASAF#6993). The signalment of these
healthy dogs is listed in Supplemental Table 1. Colonicstem cells were collected and cultured as previously
reported [49]. Briefly, colonic biopsies were washed using
ice-cold Dulbecco’s phosphate-buffered saline (PBS,
Gibco) containing 1 × Penicillin/Streptomycin (Gibco)for five times, cut into small pieces, and incubated in
30 mM EDTA solution (Invitrogen) for 60 min at 4 °C torelease crypts containing intestinal stem cells. Collected
intestinal crypts were suspended in Matrigel (Corning) and seeded to a 48-well plate (Thermo Scientific) in
30 µl per well. After the Matrigel dome was solidified in
37 °C incubator, 500 µl of organoid culture medium wasNagao and Ambrosini ﻿BMC Molecular and Cell Biology(2024) 25:14

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added to each well. For composition of organoid culture
medium, we followed the previous study; DMEM/F12(Gibco) supplemented with 2 mM GultaMAX (Gibco),
10 mM HEPES (Gibco), 1 × Penicillin/Streptomycin(Gibco), 10% (vol/vol) conditioned medium of Noggin[50], 20% (vol/vol) conditioned medium of R-spondin,
100 ng/ml recombinant murine Wnt-3a (PeproTech),
50 ng/ml murine Epidermal Growth Factor (EGF) (PeproTech), 10 nM Gastrin (Sigma-Aldrich), 500 nM A-83–
01 (Sigma-Aldrich), 10 µM SB202190 (Sigma-Aldrich),
1 mM N-Acetyl-L-Cysteine (MP Biomedicals), 10 mM
Nicotinamide (Sigma-Aldrich), 1 × B27 supplement(Gibco), 1 × N2 MAX media supplement (R&D Systems), 100 µg/ml Pimocin (Invitrogen). 10 µM Y-27632(Stem Cell Technologies) and 2.5 µM CHIR 99021 (StemCell Technologies) were added to the organoid culture
medium for the first two days after the crypts were isolated. The culture medium was changed every other day,and colonoids were passaged once a week by breaking
down the colonoids using TrypLE Express (Gibco), spinning down them (200 g, 5 min, 4 °C), resuspending themin Matrigel, and seeding them in a 48-well plate.

colonoids were then subjected to be exposed to organoidculture medium containing varying concentrations of
PA (3, 30, 60 µM) for 24 h, duration already reported as
an in vitro model of short-term HFD exposure [26, 41].
The concentrations of PA were chosen based on existing literature, where 3 µM represents a minimal effective
dose [9], 30 µM approximates physiological responses tohigh-fat diets [1], and 60 µM assesses potential toxicity
thresholds in intestinal organoid viability [10]. For theassessment of colonoid gut barrier function, a colonoidderived monolayer culture spanning 5 days was utilized.
After this period, only the apical medium was replacedwith organoid culture medium containing 30 µM of PA.
As a control, a carrier control consisting of 0.1% dimethyl
sulfoxide (DMSO) in the media was used. Each experiment was conducted with three biological replicates andthree technical replicates.

Establishment of canine colonoid‑derived monolayer
cultureThe 3D colonoids were harvested from Matrigel after
being cultured for 4–6 days. This was achieved by dissolving the Matrigel dome using Cell Recovery Solution(Corning), following the manufacturer’s instructions. The
collected organoids, along with the solution, were thentreated by incubating them in TrypLE Express containing
10 µM of Y-27632 for 10 min at 37 °C. Subsequently, thedisrupted organoids were filtered through a 70 µm cellstrainer (Fisher Scientific) and centrifuged at 200 g, 4 °C
for 5 min.
For the preparation of Falcon cell culture inserts(0.4 µm pores, Falcon), a coating process was carriedout using 100 µg/ml Matrigel and 30 mg/ml collagen I(Gibco) in DMEM/F12 supplemented with 2 mM GlutaMAX, 10 mM HEPES, and 1 × Penicillin/Streptomycin.
This coating process took place at 37 °C for 1 h. Dissociated single cells obtained from the colonoids were thenquantified using a Hemocytometer and seeded at a concentration of 1
­ 06 cells/ml within the pre-coated cell culture insert. The culture medium was changed every otherday.
Palmitic acid exposure in canine colonoids

In this study, we employed PA (Cayman chemical), a wellestablished key component of the HFD [7, 26], to simulate
the effects of a HFD in an in vitro setting. To evaluate thecell proliferation of colonoids, we utilized 3D colonoids
that were cultured in Matrigel domes for 4 days. These

Analysis of canine colonoids growth

After a 24-h exposure to PA, images of colonoids were
captured under a phase contrast microscope (DMi1,
Leica) at 10 × magnification to assess their growth rate.
The surface area of the colonoids was measured using
ImageJ (FIJI, http://​fiji.​sc/​Fiji) [51]. For each condition,
five fields of view were randomly selected at 10 × magnification within each well, and a minimum of ten organoids
were analyzed.
Total RNA extraction and quantitative reverse transcription
polymerase chain reaction analysis

Colonoids exposed to PA were harvested by dissociating
the Matrigel dome using Cell Recovery Solution. Subsequently, total RNA was extracted from the colonoids
utilizing the RNeasy mini kit (Qiagen) and subjected toreverse transcription using the High-Capacity cDNA
Reverse Transcription Kit (Applied Biosystems). qPCR
was conducted by PowerUp SYBR Green Master Mix(Applied Biosystems) and the CFX96 Touch Real-time
PCR Detection System (Bio-Rad) using 20 ng of cDNA toevaluate the gene expression levels of LGR5, a significant
stem cell marker, Occludin and JAM-A, tight junction
proteins. For cDNA normalization, internal control genes
were employed, namely, Succinate dehydrogenase complex subunit A (SDHA), Hydroxymethyl-bilane synthase(HMBS), and Hypoxanthine phosphoribosyl-transferase
1 (HPRT1), as previously reported in colon tissue [52].
Details of the primers used in this study can be found in
Supplemental Table 2.
Assessment of proliferative activity

To evaluate the proliferative activity of colonoids exposed
to PA, we conducted a EdU assay using the Click-iT
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as we have previously reported [53]. Briefly, colonoids
exposed to PA for 24 h were preincubated with a 10 µM
EdU solution for 3 h, followed by fixation with 4% PFA(Thermo Scientific) for 15 min. Subsequently, these organoids were permeabilized using 0.3% Triton-X (Thermo
Scientific) for 10 min and stained with the EdU staining
solution and 4’,6-diamidino-2-phenylindole dihydrochloride (DAPI) (Thermo Fisher Scientific) for 30 min.
Then, stained colonoids were mounted by Prolong Gold
Antifade reagent (Thermo Fisher Scientific) onto glassbottom dish (Matsunami). EdU-positive cells were captured using a confocal microscope (SP8-X, Leica) withexcitation laser sources of 405 nm, 499 nm, 553 nm,
and 653 nm and high efficiency Leica HyD detector, theacquired images were further processed using LAS X
software (Leica) for analysis, and the percentage of EdUpositive cells was quantified using ImageJ software. For
each condition, we selected five randomly chosen fieldsof view at 40 × magnification from each sample, evaluating a minimum of ten organoids.

respectively. The monolayer was then mounted with Prolong Gold Antifade reagent (Thermo Fisher Scientific)
and imaged using a confocal microscope (SP8-X, Leica)
using 63 × magnification with excitation laser sourcesof 405 nm, 499 nm, 553 nm, and 653 nm and high efficiency Leica HyD detector, the acquired images were further processed using LAS X software (Leica) for analysis.
For each condition, five randomly selected fields of viewat 63 × magnification were evaluated in each sample,
and fluorescence intensity in the entire field of view wasquantified using ImageJ software. Subsequently, the normalized fluorescence intensity was calculated by dividing
the fluorescent intensity of each sample by the average offluorescence intensity of control wells.

Assessment of intestinal barrier integrityin colonoid‑derived monolayer

The intestinal barrier integrity of the colonoid-derived
monolayer was assessed by measuring the TEER value
following exposure to PA. Electrical resistance (Ωt) was
quantified using Ag/AgCl electrodes connected to a VoltOhm meter (Millicell ERS-2, Millipore), and this value
was then transformed into a TEER measurement using
the following equation: TEER = (Ωt—Ωblank) x A. Here,Ωblank represents the resistance of the blank well in ohms,
and A denotes the surface area of the culture insert in­cm2. For each condition, measurements were made with
3 biological replicates with 3 technical replicates.
Immunocytochemistry of colonoid‑derived monolayer

After exposure to PA, the colonoid-derived monolayer
was fixed using 4% PFA at room temperature for 15 min.
Following permeabilization with 0.3% Triton-X for
15 min at room temperature and blocking with 2% bovine
serum albumin (Cytiva) for 30 min at room temperature,primary antibodies were applied to the colonoid-derived
monolayer overnight at 4 °C. For visualization of E-cadherin and ZO-1, monoclonal anti-mouse E-cadherin
antibody (36/E-cadherin, dilution 1:200, BD Biosciences)and polyclonal anti-rabbit ZO-1 antibody (61–7300, dilution 1:50, Invitrogen) were utilized as primary antibodies. Subsequently, the colonoid-derived monolayers were
washed with PBS and incubated with secondary antibodies (Anti-Rabbit IgG H&L labeled with Alexa Fluor 555,
dilution 1:1000, Abcam). After another round of washing, nuclei and F-actin were stained using DAPI and
Alexa Fluor 647 Phalloidin (Thermo Fisher Scientific),

Statistical analysis

Statistical analysis was conducted using R Studiov1.4.1717 (RStudio) and figures were generated using
GraphPad Prism 10.0.2.232 (Dotmatics). To confirm
the normality of each dataset, the Shapiro–Wilk’s test
was performed. The Kruskal–Wallis and Dunnett tests
were employed for the comparison of organoid sizeand LGR5 gene expression among different conditions.
The Wilcoxon test was utilized to compare the rates of
EdU-positive cells and TEER values across conditions.
Additionally, Student t-tests were applied to compare
the fluorescence intensity of tight junction proteins. All
results were presented as mean ± standard error of themean. A significance level of p < 0.05 was considered statistically significant.
Abbreviations
EdU	5-Ethynyl-2-deoxyuridine
FFAs	Free fatty acids
HFD	High-fat diet
HMBS	Hydroxymethyl-bilane synthase
HPRT1	Hypoxanthine phosphoribosyl-transferase 1
PA	Palmitic acid
PBS	Phosphate-buffered salineqPCR	Quantitative reverse transcription polymerase chain reaction
SDHA	Succinate dehydrogenase complex subunit A
TEER	Transepithelial electrical resistanceZO-1	Zonula occludens-1

Supplementary Information
The online version contains supplementary material available at https://​doi.​
org/​10.​1186/​s12860-​024-​00512-w.
Supplementary Material 1.
Acknowledgements
Authors would like to thank WSU Small Animal Internal Medicine service (Dr.
Jillian Haines, Dr. Sarah Guess, Shelley Ensign LVT, Sybil Fiedler VTA), WSU Community Practices service (Dr. Jessica Bell, Dr. Cassidy Cordon, Dr. Matt Mason,
Dr. Jenna Waltzek, Mrs. Melody Gerber, Ms. Maggie deSouza, Ms. Becky Brodie),and WSU VTH Clinical Studies Coordinator Valorie Wiss for their support in caserecruitment and sample collection from citizen scientists (patient donors). We

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also would like to thank Dr. Minae Kawasaki, Dr. Takashi Goyama, Dr. Yurika
Tachibana, and Dr. Meg Nakazawa at Ambrosini lab for their help in sample
collection, organoid generation, and organoid maintenance.

9.

Authors’ contributions
I.N. and Y.M.A. designed the experiments and I.N. performed the experiments.
I.N. and Y.M.A. wrote and edited the manuscript, tables and figures.

11.

Authors’ information
Not applicable.

12.

Funding
This work was supported in part by the Office Of The Director, National
Institutes Of Health (K01OD030515 and R21OD031903 to Y.M.A.) and Japan
Society for the Promotion of Science Overseas Challenge Program for Young
Researchers (202280196 to I.N.). The content (i.e., the design, analysis andreporting of the study) is solely the responsibility of the authors and does notnecessarily represent the official views of the funding bodies.
Availability of data and materials
All relevant data are within the paper and its Supporting Information files.

10.

13.

14.

15.
16.

Declarations
Ethics approval and consent to participate
All procedures on client-owned dogs in this study were approved by the
Washington State University Institutional Animal Care and Use Committee(IACUC protocol: ASAF #6993). All clients who owned the dogs participated inthis study were informed and signed the informed consent forms approvedby the IACUC. All the experiment in this study were conducted in accordance
to the guideline and regulations for experiments on samples derived fromclient-owned animals (IBC protocol: BAF #00814).

17.

Consent for publication
Not applicable.

20.

Competing interests
The authors declare no competing interests.

21.

Received: 14 February 2024 Accepted: 26 April 2024

18.
19.

22.

23.
References
1. Xie Y, Ding F, Di W, Lv Y, Xia F, Sheng Y, et al. Impact of a high-fat diet on
intestinal stem cells and epithelial barrier function in middle-aged female
mice. Mol Med Rep. 2020;21:1133–44.
2. Cheng H, Zhou J, Sun Y, Zhan Q, Zhang D. High fructose diet: a risk factor
for immune system dysregulation. Hum Immunol. 2022;83:538–46.
3. Qu D, Wang G, Yu L, Tian F, Chen W, Zhai Q. The effects of diet and gut
microbiota on the regulation of intestinal mucin glycosylation. Carbohydr
Polym. 2021;258:117651.
4. Calibasi-Kocal G, Mashinchian O, Basbinar Y, Ellidokuz E, Cheng C-W,
Yilmaz ÖH. Nutritional control of intestinal stem cells in homeostasis andtumorigenesis. Trends Endocrinol Metab. 2021;32:20–35.
5. Zimmermann C, Wagner AE. Impact of food-derived bioactive compounds on intestinal immunity. Biomolecules. 2021;11:1901.
6. Tomé D. The roles of dietary glutamate in the intestine. Ann Nutr Metab.
2018;73(Suppl 5):15–20.
7. Beyaz S, Mihaylova MM, Lamming DW, Dogum R, Guo G, Yuan G, et al.
High fat diet enhances stemness and tumorigenicity of intestinal progenitors. Nature. 2016;531:53–8.
8. Choi J, Zhang X, Li W, Houston M, Peregrina K, Dubin R, et al. Dynamic
intestinal stem cell plasticity and lineage remodeling by a nutritional
environment relevant to human risk for tumorigenesis. Mol Cancer Res.
2023;21:808–24.

24.

25.

26.

27.
28.
29.
30.

Leushacke M, Barker N. Lgr5 and Lgr6 as markers to study adult stem cellroles in self-renewal and cancer. Oncogene. 2012;31:3009–22.
Leung C, Tan SH, Barker N. Recent advances in Lgr5+ stem cell research.
Trends Cell Biol. 2018;28:380–91.
Groop LC, Saloranta C, Shank M, Bonadonna RC, Ferrannini E, DeFronzo
RA. The role of free fatty acid metabolism in the pathogenesis of insulin
resistance in obesity and noninsulin-dependent diabetes mellitus. J Clin
Endocrinol Metab. 1991;72:96–107.
Weidinger C, Ziegler JF, Letizia M, Schmidt F, Siegmund B. Adipokines andtheir role in intestinal inflammation. Front Immunol. 2018;9:1974.
Villaret A, Galitzky J, Decaunes P, Estève D, Marques M-A, Sengenès C,
et al. Adipose tissue endothelial cells from obese human subjects: differences among depots in angiogenic, metabolic, and inflammatory gene
expression and cellular senescence. Diabetes. 2010;59:2755–63.
Mao Y, Luo W, Zhang L, Wu W, Yuan L, Xu H, et al. STING-IRF3 triggers
endothelial inflammation in response to free fatty acid-Induced mitochondrial damage in diet-Induced obesity. Arterioscler Thromb Vasc Biol.
2017;37:920–9.
Rohr MW, Narasimhulu CA, Rudeski-Rohr TA, Parthasarathy S. Negative
effects of a high-fat diet on intestinal permeability: a review. Adv Nutr.
2020;11:77–91.
Buettner R, Parhofer KG, Woenckhaus M, Wrede CE, Kunz-Schughart
LA, Schölmerich J, et al. Defining high-fat-diet rat models: metabolic and molecular effects of different fat types. J Mol Endocrinol.
2006;36:485–501.
Vesga-Jiménez DJ, Martin C, Barreto GE, Aristizábal-Pachón AF, Pinzón A,
González J. Fatty acids: An insight into the pathogenesis of neurodegenerative diseases and therapeutic potential. Int J Mol Sci. 2022;23:2577.
Marra F, Svegliati-Baroni G. Lipotoxicity and the gut-liver axis in NASH
pathogenesis. J Hepatol. 2018;68:280–95.
Garcia-Martinez I, Alen R, Pereira L, Povo-Retana A, Astudillo AM, Hitos AB,
et al. Saturated fatty acid-enriched small extracellular vesicles mediate acrosstalk inducing liver inflammation and hepatocyte insulin resistance.
JHEP reports Innov Hepatol. 2023;5:100756.
Yang Y, Yu Q, Li B, Yang Z, Zhang S, Yuan F. Palmitate lipotoxicity is closely
associated with the fatty acid-albumin complexes in BV-2 microglia. PLoS
One. 2023;18:e0281189.
Tran TTT, Postal BG, Demignot S, Ribeiro A, Osinski C, de Pais Barros J-P,
et al. Short term palmitate supply impairs intestinal insulin signaling via
ceramide production. J Biol Chem. 2016;291:16328–38.
Ghezzal S, Postal BG, Quevrain E, Brot L, Seksik P, Leturque A, et al.
Palmitic acid damages gut epithelium integrity and initiates inflammatory cytokine production. Biochim Biophys Acta Mol Cell Biol Lipids.
2020;1865:158530.
Bernard JN, Chinnaiyan V, Andl T, Le Bras GF, Qureshi MN, Altomare
DA, et al. Augmented CPT1A expression is associated with proliferation and colony formation during Barrett’s tumorigenesis. Int J Mol Sci.
2022;23:11745.
Ghareghani M, Zibara K, Azari H, Hejr H, Sadri F, Jannesar R, et al. Safflower
seed oil, containing oleic scid and palmitic acid, enhances the stemnessof cultured embryonic neural stem cells through Notch1 and induces
neuronal differentiation. Front Neurosci. 2017;11:446.
Horcas-Nieto JM, Versloot CJ, Langelaar-Makkinje M, Gerding A, Blokzijl
T, Koster MH, et al. Organoids as a model to study intestinal and liver
dysfunction in severe malnutrition. Biochim Biophys Acta Mol Basis Dis.
2023;1869:166635.
Filippello A, Di Mauro S, Scamporrino A, Torrisi SA, Leggio GM, Di PinoA, et al. Molecular effects of chronic exposure to palmitate in intestinalorganoids: a new model to study obesity and diabetes. Int J Mol Sci.
2022;23(14):7751.
German AJ. The growing problem of obesity in dogs and cats. J Nutr.
2006;136(7 Suppl):1940S–1946S.
Patronek GJ, Waters DJ, Glickman LT. Comparative longevity of pet dogs
and humans: Implications for gerontology research. Journals Gerontol Ser A Biol Sci Med Sci. 1997;52:B171.
Lutz TA. Mammalian models of diabetes mellitus, with a focus on type2 diabetes mellitus. Nat Rev Endocrinol. 2023. https://​doi.​org/​10.​1038/​
s41574-​023-​00818-3
Allenspach K, Culverwell C, Chan D. Long-term outcome in dogs withchronic enteropathies: 203 cases. Vet Rec. 2016;178:368.

Nagao and Ambrosini ﻿BMC Molecular and Cell Biology(2024) 25:14

Page 10 of 10

31. Patnaik AK, Hurvitz AI, Johnson GF. Canine gastrointestinal neoplasms. Vet
Pathol. 1977;14:547–55.
32. Saito T, Chambers JK, Nakashima K, Nibe K, Ohno K, Tsujimoto H, et al.
Immunohistochemical analysis of beta-catenin, e-cadherin and p53 incanine gastrointestinal epithelial tumors. J Vet Med Sci. 2020;82:1277–86.
33. Kopper JJ, Iennarella-Servantez C, Jergens AE, Sahoo DK, Guillot E,
Bourgois-Mochel A, et al. Harnessing the biology of canine intestinalorganoids to heighten understanding of inflammatory bowel disease
pathogenesis and accelerate drug discovery: a One Health approach.
Front Toxicol. 2021;3:1–13.
34. Allenspach K, Borcherding DC, Iennarella-Servantez CA, Mosichuk AP,
Atherly T, Sahoo DK, et al. Ketogenic diets in healthy dogs induce gut andserum metabolome changes suggestive of anti-tumourigenic effects: amodel for human ketotherapy trials. Clin Transl Med. 2022;12:1–8.
35. Moinard A, Payen C, Ouguerram K, André A, Hernandez J, Drut A, et al.
Effects of high-fat diet at two energetic levels on fecal microbiota, colonicbarrier, and metabolic parameters in dogs. Front Vet Sci. 2020;7:566282.
36. Ambrosini YM, Park Y, Jergens AE, Shin W, Min S, Atherly T, et al. Recapitulation of the accessible interface of biopsy-derived canine intestinal organoids to study epithelial-luminal interactions. PLoS One. 2020;15:1–17.
37. Barbara G, Barbaro MR, Fuschi D, Palombo M, Falangone F, Cremon C,
et al. Inflammatory and microbiota-related regulation of the intestinal
epithelial barrier. Front Nutr. 2021;8:718356.
38. Suzuki T. Regulation of intestinal epithelial permeability by tight junctions. Cell Mol Life Sci. 2013;70:631–59.
39. Ahn C, Shin D-H, Lee D, Kang S-M, Seok J-H, Kang HY, et al. Expression ofclaudins, occludin, junction adhesion molecule a and zona occludens 1
in canine organs. Mol Med Rep. 2016;14:3697–703.
40. Ogawa M, Osada H, Hasegawa A, Ohno H, Yanuma N, Sasaki K, et al. Effectof interleukin-1β on occludin mRNA expression in the duodenal andcolonic mucosa of dogs with inflammatory bowel disease. J Vet Intern
Med. 2018;32:1019–25.
41. Gori M, Altomare A, Cocca S, Solida E, Ribolsi M, Carotti S, et al. Palmitic
acid affects intestinal epithelial barrier integrity and permeability in vitro.
Antioxidants (Basel). 2020;9:417.
42. Rao RK, Basuroy S, Rao VU, Karnaky KJ, Gupta A. Tyrosine phosphorylationand dissociation of occludin-ZO-1 and E-cadherin-β-catenin complexes
from the cytoskeleton by oxidative stress. Biochem J. 2002;368:471–81.
43. Pourvali K, Monji H. Obesity and intestinal stem cell susceptibility tocarcinogenesis. Nutr Metab (Lond). 2021;18:37.
44. Kol A, Arzi B, Athanasiou KA, Farmer DL, Nolta JA, Rebhun RB, et al. Companion animals: translational scientist’s new best friends. Sci Transl Med.
2015;7:308ps21.
45. Ambrosini YM, Borcherding D, Kanthasamy A, Kim HJ, Willette AA, Jergens
A, et al. The gut-brain axis in neurodegenerative diseases and relevanceof the canine model: a review. Front Aging Neurosci. 2019;11:1–14.
46. Okumura R, Takeda K. Maintenance of gut homeostasis by the mucosal
immune system. Proc Jpn Acad Ser B Phys Biol Sci. 2016;92:423–35.
47. Min S, Than N, Shin YC, Hu G, Shin W, Ambrosini YM, et al. Live probiotic
bacteria administered in a pathomimetic Leaky Gut Chip ameliorate impaired epithelial barrier and mucosal inflammation. Sci Rep.
2022;12:22641.
48. Shin W, Kim HJ. 3D in vitro morphogenesis of human intestinal epithelium in a gut-on-a-chip or a hybrid chip with a cell culture insert. Nat
Protoc. 2022;17:910–39.
49. Chandra L, Borcherding DC, Kingsbury D, Atherly T, Ambrosini YM,
Bourgois-Mochel A, et al. Derivation of adult canine intestinal organoids
for translational research in gastroenterology. BMC Biol. 2019;17:1–21.
50. Heijmans J, van Lidth de Jeude JF, Koo B-K, Rosekrans SL, Wielenga MCB,
van de Wetering M, et al. ER stress causes rapid loss of intestinal epithelial
stemness through activation of the unfolded protein response. Cell Rep.
2013;3:1128–39.
51. Schindelin J, Arganda-Carreras I, Frise E, Kaynig V, Longair M, Pietzsch
T, et al. Fiji: an open-source platform for biological-image analysis. Nat
Methods. 2012;9:676–82.
52. Peters IR, Peeters D, Helps CR, Day MJ. Development and applicationof multiple internal reference (housekeeper) gene assays for accurate
normalisation of canine gene expression studies. Vet Immunol Immunopathol. 2007;117:55–66.

53. Nagao I, Ambrosini YM. Ion channel function in translational bovine
gallbladder cholangiocyte organoids: establishment and characterizationof a novel model system. Front Vet Sci. 2023;10:1–7.

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