Abstract
In vitro research on host–microbe interactions in the human gut has
been challenging due to the differing oxygen requirements of mammalian
cells and intestinal microbiota. Few models of this environment have
been developed, and those available are complex, limiting the
extraction of important information during experiments. Here we report
an in vitro model that by simple means creates an anaerobic environment
for microbiota growing on living, cultured human epithelium under
physiological flow. This model enables long-term co-culture of
intestinal epithelial cells with obligate anaerobic bacteria,
exemplified here by Clostridioides difficile and Bacteroides fragilis.
Anaerobic conditions are maintained through the integration of an
anaerobization unit, developed to facilitate online deoxygenation of
media via liquid-to-liquid gas diffusion, eliminating the need for
encapsulation in complex gas chambers. We show that stable oxygen
levels of less than 1% can be maintained in the model for several days
without compromising the viability of the intestinal epithelium.
Furthermore, we demonstrate the performance of the model by simulating
prolonged colonization with C. difficile and B. fragilis, as well as
the clinically relevant persistence of C. difficile following treatment
with vancomycin.
Subject terms: Clinical microbiology, Pathogens
Introduction
The human colon is a highly complex biological niche hosting trillions
of bacteria that exist in symbiosis with the colon epithelium. This
bacterial community, known as the microbiota, includes at least
500–1,000 different species, predominantly obligate
anaerobes^[40]1–[41]4. The microbiota plays a vital role in maintaining
various aspects of human health^[42]5, including serving as a barrier
to gastrointestinal pathogens^[43]6. Disruption or dysbiosis of the
microbiota is a major risk factor for acquiring intestinal infections,
such as Clostridioides difficile infection (CDI). Understanding the
complex interaction between host, microbiota, and pathogen is critical
for the development of effective treatment strategies. Until recently,
researchers have primarily relied on animals as surrogate models of
human intestinal infection^[44]7 and static microtiter plate cell
culture models^[45]8,[46]9, both of which have significant limitations.
For example, traditional in vitro cell culture systems, such as cell
culture microtiter plate inserts^[47]8,[48]9, fail to mimic the complex
conditions of the colon as they lack key in vivo-like features, such as
the three-dimensional (3D) architecture of the intestinal epithelium,
its anaerobic environment, and shear stress. Furthermore, these static
systems can only be used for short-term studies. In contrast, animal
models provide valuable insights into gut physiology, but they do not
fully replicate the human gut environment or human susceptibility to
infection. As a result, critical aspects of the pathogenesis of
anaerobes remain poorly understood, complicating the development of
treatment strategies. This is particularly the case for C. difficile
infection, where the lack of qualified surrogate models recently led
researchers to suggest developing a C. difficile infection model in
humans^[49]10.
In recent years, intestine-on-a-chip systems have emerged as a
promising platform for studying long-term co-cultures of bacteria and
intestinal cells. The gold standard approach to manufacturing these
chips is by soft lithography with polydimethylsiloxane (PDMS), a highly
flexible, biocompatible material which is typically coated with
extracellular matrix (ECM) for cell adhesion^[50]11. Indeed, PDMS chips
have revolutionized the field of organ-on-chips, including models
simulating the human intestine^[51]12. However, PDMS and other rubber
materials compatible with soft lithography are also highly gas
permeable, which compromises the chip’s compatibility with cultures
that require strict separation and control of oxygen. Engineers have
solved these problems in pilot setups by placing the chips in nitrogen
containers^[52]13 or by shielding against oxygen leakage by various
means^[53]14,[54]15. A common characteristic of these solutions is that
the systems have become complex. Combined with the expert knowledge
required to produce and handle soft lithography chips, this has limited
the dissemination of in vitro models to frontline clinical and industry
researchers that develop solutions for infections and dysbiosis caused
by anaerobic microorganisms^[55]16. The Caco-2 cell line, derived from
a human colorectal tumor, is commonly used in these systems due to its
ease of cultivation and proven barrier properties^[56]17,[57]18. To
mimic epithelial barrier characteristics, some systems incorporate
parallel microchannels separated by a microporous membrane, enabling
modeling of both the vascular and intestinal luminal environment within
the same device. However, to simulate the hypoxic environment of the
gut that enables growth of anaerobic bacteria, gases must be strictly
separated to create the anaerobic environment on the intestinal luminal
side, while providing oxygen to the intestinal cells from below. The
latter feature has proven technically challenging and the few published
models that have succeeded^[58]19 are highly complex setups that so far
have not translated from laboratory pilots to more widely applicable
systems.
To our knowledge, only three suggested solutions have been reported
that support a 3D intestinal epithelium co-cultured with obligate
anaerobic bacteria for several days. One model was published by
Jalili-Firoozinezhad et al. in 2019 and consists of a PDMS chip lined
with Caco-2 and endothelial cells, which allows for the culture of
human microbiota for up to three days^[59]13. Hypoxic conditions
(O[2] < 2%, gas-phase equivalent) were maintained in the upper luminal
channel by placing the chip in a box filled with nitrogen gas, while
perfusing the lower vascular channel with oxygenated medium, to supply
the intestinal cells with oxygen through the porous membrane. The
authors demonstrated proof of replication of Bacteroides fragilis (B.
fragilis) in the anaerobic chip and measured oxygen levels below 1%
during the three-day culture period. Jalili- Firoozinezhad et al.
further demonstrate the performance of the model by culturing primary
ileal epithelium and gut microbiota. A second model, published by Shin
et al. in 2019, likewise consisted of a modified PDMS chip but with a
thicker upper part and a glass coverslip underneath the chip to control
the oxygen flux through the gas-permeable PDMS. The authors
computationally simulated the oxygen levels in the intestinal chip
using measurements from experiments with a chemical oxygen scavenger in
the medium. The authors estimated that the two obligate anaerobes
Bifidobacterium adolescentis and Eubacterium halili remain viable for
up to seven days, although no evidence of bacterial replication was
provided^[60]14. In 2023, Liu et al. reported a soft lithography PDMS
model where oxygen levels were also computer simulated and
reoxygenation of the upper channel was prevented by attaching a glass
coverslip to the outside of the chip^[61]15. Similarly, no evidence of
bacterial replication and real-time oxygen measurements during
co-culture were reported.
Common to the above models is the generation of anaerobic conditions in
the chip by liquid-air passive diffusion of oxygen. This is however an
equipment-intensive, slow process that requires encapsulation of the
chip in sealed nitrogen containers that additionally limits the
possibility to access the chip for material and data extraction.
In this study, we propose a novel approach to designing an in vitro
flow model that enables the co-culture of intestinal cells and
anaerobic bacteria. Instead of relying on oxygen permeable PDMS chips
and the necessary encapsulation in anaerobic incubators, we use hard,
oxygen impermeable flow chambers combined with a novel method for fast
online anaerobization of media to the upper (intestinal lumen) channel.
The flow model supports co-culture with obligate anaerobic bacteria for
at least five days, without compromising the viability of the cell
layer. The model can be used to study host–microbe interactions and
novel treatments against obligate anaerobic pathogens, for which in
vitro models are urgently needed^[62]20. Using this model, we
demonstrate, to our knowledge for the first time, sustained
colonization by C. difficile on a live 3D intestinal epithelium over
several days. Additionally, we monitored the bacterium’s response to
vancomycin treatment in vitro, which reproduced the clinically
challenging ability of C. difficile to survive standard antibiotic
treatment in vivo. Contrary to earlier reported and commercial models,
the reported concept is readily applicable in standard cell culture
laboratories and generates highly stable co-cultures with even highly
oxygen-sensitive bacteria.
Results
Establishment of the hypoxic intestinal lumen environment
To establish a model closely simulating the physiology and
microenvironment of the human colon, we aimed to create a system that
incorporates the shear stress conditions and the low-oxygen environment
present in the colon, while supporting the growth and maintenance of an
intestinal epithelial cell culture. We chose the dual flow channel
principle for the model. Contrary to PDMS dual-channel chips, however,
in which media gas content is controlled by the gas outside of the chip
due to the gas-permeable PDMS, we opted for a hard-plastic system that
would eliminate the need for encapsulation in gas containers (Fig.
[63]1A, B). We intended to establish such a stand-alone system that
would be compatible with physiological wall shear stress of at least
0.1 dyn/cm^2 and media oxygen levels below 1%, both of which approach
the conditions in the intestine^[64]21–[65]23. Realizing such a system
required a novel method for fast and effective online deoxygenation of
flow media, before reaching the chamber. Our solution exploits the fast
diffusion of oxygen through silicone rubber and the highly
oxygen-attractant properties of antioxidant liquids^[66]24. An
ultrathin silicone tube is coiled within a container filled with a
strong aqueous antioxidant solution (Fig. [67]1C). Passing liquid media
through this system rapidly depletes dissolved oxygen, even at high
flow rates (Fig. [68]2). The dual flow chamber (DFC) was made by
mounting two Ibidi® sticky slides® back-to-back with a thin, porous,
transparent (track-etched) polyester membrane in between (Fig. [69]1A,
B, Supplementary Fig. S[70]1). This design created apical and
basolateral flow channels within the DFC, with the outer walls acting
as effective oxygen barriers, while oxygen readily diffuses across the
culture membrane from the lower channel to the intestinal cell culture.
Oxygen concentrations were measured in the media exiting the
anaerobization unit (AU) to test the deoxygenation efficiency, and in
the media exiting single- and DFCs to monitor reabsorption of oxygen
during flow chamber passage (Fig. [71]2A–C, E). As demonstrated in Fig.
[72]2, a rate of oxygen depletion was achieved that allowed flow rates
necessary for simulating the physiological liquid shear of the
intestine. Several factors influence the deoxygenation efficiency,
including media flow rate and tube wall thickness, length, and diameter
(Fig. [73]2B, C, E).
Fig. 1. Design of the dual flow chamber and system setup.
[74]Fig. 1
[75]Open in a new tab
A Three-dimensional images of the dual flow chamber (DFC) showing the
entire DFC with luer connectors (Ai) and a cross-section showing the
microporous membrane in brown (Aii). B Real life images of the DFC
during an aerobic experiment. The DFC dimensions without luer
connectors are 75 × 25 × 4 mm. The DFC is placed in an upright position
to prevent the accumulation of air bubbles. C Schematic presentation of
the system setup placed in a standard CO[2] incubator. Cell culture
media (Ci) is pumped via the flow pump (Cii) through the anaerobization
unit (AU) containing the silicone coil submerged in an antioxidant
solution (Ciii) which is directly connected to the apical channel of
the DFC (Civ) via a stainless-steel tube. Aerobic cell media is
supplied through the basolateral channel (Cv) using the same flow pump
(Cii). Cell media exiting the DFC is collected in the liquid waste
container (Cvi). Figure was made with Biorender.com and Microsoft
Powerpoint.
Fig. 2. Oxygen levels and dependency on flow rate, tube length and chamber
conditions.
[76]Fig. 2
[77]Open in a new tab
A Schematic presentation of the single and the dual flow chamber (DFC)
showing the connection sites of the oxygen sensor (indicated "Inlet"
and “Exit"). Inlet measurements were performed by connecting the oxygen
sensor to the exit site of the anaerobization unit (AU) and exit
measurements by connecting the oxygen sensor to the exit of the flow
chambers. The AU and the flow chambers are connected with
stainless-steel tubing to prevent reabsorption of oxygen into the cell
media. B Inlet oxygen percentage measured using different lengths of
the AU silicone tube coil and at different flow rates. C Oxygen
percentage at inlet and exit sites in a single flow chamber (µ-slide I
luer 0.4mm, Ibidi) at different flow speeds, showing the relationship
between oxygen percentage and flow rate. D Oxygen percentage in
effluent from DFCs with Caco-2 cells cultured under anaerobic
conditions for 6 days with a flow of 320 µl/min. The data is based on
an hourly average measured in three individual flow chambers (Mean
±range). Exit measurements represent the highest oxygen levels in the
apical channel, as oxygen is slowly reabsorbed into the medium during
passage through the channel. E Reabsorption of oxygen into the apical
flow channel of a dual flow chamber, when the flow is stopped for
different periods of time.
To provide deoxygenized media at flow rates of 120–640 µl/min,
necessary to generate the desired 0.1–0.6 dyn/cm² in DFCs, a tube with
luminal diameter of 0.99 mm, wall thickness of 0.31 mm, and length of
≥150 cm was used in the AU (Fig. [78]2B).
To avoid variations due to the consumption of oxygen by the living
epithelium, oxygen reabsorption kinetics in media during channel
passage was measured in a single-channel flow chamber without cells,
with channel dimensions matching the DFC’s apical channel (Fig.
[79]2A). The results indicated a flow rate of 320 µl/min (shear stress
of 0.3 dyn/cm²) as a sweet spot with both low oxygen levels and little
difference between exit and inlet measurements (Fig. [80]2C).
Measurements of oxygen levels passing through DFCs with 7-days matured
Caco-2 epithelium confirmed consistently low anaerobic conditions over
the following six-days, reaching levels between 0.1-1% (Fig. [81]2D).
During flow interruptions, oxygen concentration measurements revealed
that flow could be paused for up to 60 s before exit levels exceeded
1%, and up to 5 min before surpassing 3% (Fig. [82]2E). Oxygen levels
decreased to <1% within a few minutes after resuming the flow.
Caco-2 cells matured in the DFC demonstrate in vivo characteristics of the
intestine
For the first iteration of the model, we selected the human epithelial
cell line Caco-2, a gold standard in many gut-on-a-chip models. After
confirming that the model met the requirements for shear stress (SS)
and oxygen levels, the characteristics of the Caco-2 cell layer
cultured in the DFC under both aerobic and anaerobic conditions were
investigated and compared to static cultures in cell culture inserts.
Caco-2 cells cultured for 13 days in the DFC, formed a complex 3D
epithelium with crypt- and villus-like structures, reaching up to
135 µm in height (Fig. [83]3A, [84]B). Under anaerobic conditions in
the apical channel, a greater variation in the height of the cell layer
ranging between 50-115 µm was observed (Fig. [85]3B, Supplementary Fig.
S[86]2). By comparison, the average height of the static cultures after
13 days was approximately 20 µm (Fig. [87]3B). Under both aerobic and
anaerobic flow, the mature Caco-2 layer stained positive for the tight
junction protein occludin and the cytoskeleton component F-actin,
indicating establishment of an intestinal barrier (Fig. [88]3A). To
assess mucus production, the cultured tissue was stained for neutral
and acidic mucins, and immunostained for MUC2, the primary gel-forming
mucin in the colon. Both aerobic and anaerobic flow conditions revealed
a higher signal of neutral mucins compared to the static control (Fig.
[89]3C). Due to the height of the cell layer, it was difficult to make
firm conclusions on the presence of acidic mucins, but some blue stains
did appear under all three culture conditions (Fig. [90]3C,
Supplementary Fig. S[91]3). The presence of MUC2 by immunostaining was
detected in the Caco-2 cells cultured under aerobic and anaerobic
conditions in the DFC. MUC2 appeared mostly in vesicles within some of
the cells at the top of the cultured epithelium. The low signal
indicated minimal MUC2 production by the cells. Subsequent ELISA
analysis confirmed this, showing MUC2 levels below the lower detection
limit (0.78 ng/mL, data not shown). The Caco-2 epithelium was found to
maintain viability for at least 21 days under anaerobic conditions
(Supplementary Fig. S[92]4).
Fig. 3. Characterization of the cultured Caco-2 epithelium 13 days
post-seeding.
[93]Fig. 3
[94]Open in a new tab
A Representative confocal laser scanning microscopy z-stack images of
Caco-2 cells grown under static (cell culture inserts) or flow
conditions (dual flow chamber), showing the three-dimensional structure
of the cultured epithelium. The anaerobization unit was connected to
the apical inlet on day 7 post-seeding to create an oxygen gradient
throughout the cell layer (−O[2]) and compared with experiments where
both channels were kept aerobic (+O[2]). Cells were stained for Mucin 2
(MUC2), the tight junction protein Occludin (OCLN) and F-actin.
Z-slices indicate cross sections based on z-stacks. Images are
representative of two individual experiments with three images of each
condition. B Heigh in micrometers (µm) of the cultured Caco-2
epithelium grown under the three different conditions. The height was
determined based on confocal microscopy z-stacks. Error bars indicate
standard deviation. Statistics were made using a one-way ANOVA with
multiple comparisons. C Staining of neutral (Shiff’s reagent) and
acidic (Alcian blue) mucins. For mucin expression at earlier time
points, see Supplementary Fig. S[95]3. Three representative pictures
were taken for each condition.
Transcriptomic profiling reveals enterocyte maturation in the Caco-2 DFC
model
To comprehensively investigate the transcriptional profile of the
intestinal epithelium cultured in the system, we conducted
transcriptomic analyses on Caco-2 cells cultured for 13 days under both
aerobic and anaerobic conditions in the apical channel of the DFC
model, and statically under aerobic conditions in cell culture inserts.
Differentially expressed genes (DEGs) were identified and analyzed by
pathway enrichment analysis using Metascape^[96]25.
Among the 817 upregulated genes in the aerobic DFC model relative to
the static model, several were associated with the ‘HDACs deacetylate
histones’ Reactome pathway, including multiple histone genes (Fig.
[97]4A, B). Upregulation of histone genes, a marker of entry into the S
phase of the cell cycle^[98]26, may indicate an increase in the number
of actively dividing cells in the aerobic DFC, aligning with the
observed increase in 3D growth compared to the static model.
Additionally, pathways related to small molecule transport
(R-HSA-382551 and R-HSA-425407) and lipid metabolism (R-HSA-556833)
were enriched, pointing to enhanced absorptive function of the
enterocytes. Conversely, among the 1182 downregulated genes, pathways
related to cilia development and function (R-HSA-5617833 and
R-HSA-5620912) were significantly enriched. Cilia, membrane-bound
sensory organelles that detect signals such as shear stress, are
typically present in quiescent cells and are generally absent in
rapidly renewing tissues, including the intestinal epithelium^[99]27.
Fig. 4. Transcriptomic characterization of the Caco-2-based DFC model.
[100]Fig. 4
[101]Open in a new tab
Caco-2 cells were cultured for 13 days in the DFC model, both
aerobically (DFC + O[2]) and anaerobically (DFC-O[2]), as well as in
cell culture inserts under aerobic conditions (Static), all in
triplicates. RNA sequencing was then performed to analyze gene
expression profiles. A Differentially expressed genes (DEGs) between
Caco-2 cells cultured in the aerobic DFC and the static cell culture
inserts, as well as between cells cultured in the anaerobic and aerobic
DFC models. B Pathway enrichment analysis of Reactome gene sets for up-
and downregulated genes in the aerobic DFC model compared to the cell
culture inserts, and in the anaerobic DFC model compared to the aerobic
DFC model. Here, top 5 enriched pathways are shown, unless fewer
pathways were significantly enriched. C Clustered heatmap showing
scaled marker gene expression profiles of six major intestinal cell
types: goblet cells, paneth cells, enteroendocrine cells (EEC),
intestinal stem cells (ISC), absorptive enterocytes (AE), and
absorptive coloncytes (AC). This comparison includes data from Caco-2
models of this study, a Caco-2-based 3D gut model from Cheng et
al.^[102]22 and single-cell and bulk RNA sequencing data of intestinal
cell types (goblet, paneth, EEC, distal enterocytes, and proximal
enterocytes) and tissues (colon and small intestine (SI)) derived from
the Human Protein Atlas^[103]57.
By contrast, only 174 genes were differentially expressed between the
anaerobic and aerobic DFC models. The subtle difference in
transcriptional activity suggests that the Caco-2 cells are provided
adequate amounts of oxygen for sustaining a vital state of growth,
despite the only oxygen source is the media flowing beneath the culture
(Fig. [104]4A). Still, the upregulation of several
metallothionein-encoding genes (R-HSA-5661231), which are involved in
metal homeostasis and protection against oxidative stress, alongside
the downregulation of genes related to the electron transport chain
(R-HSA-611105), indicates a hypoxic response. Interestingly,
significant transcriptional induction of hypoxia-inducible factor-1
(HIF1A) mRNA was not observed, although previous studies have reported
increased HIF-1α protein levels under hypoxic conditions^[105]13
(Supplementary Fig. S[106]5). Such a response is widely recognized as
being vital for maintaining intestinal barrier integrity and regulating
nutrient absorption within the intestinal epithelium^[107]28.
Additionally, several pathways related to mitosis (R-HSA-3214815,
R-HSA-2467813, R-HSA-156711, and R-HSA-174178) were also enriched among
the downregulated genes, indicating a decrease in cell proliferation
compared to the aerobic DFC model (Fig. [108]4B).
Although derived from the colon, Caco-2 monolayers exert many of the
same properties as absorptive enterocytes of the small
intestine^[109]29. To determine whether DFC conditions promote
maturation toward a
more differentiated intestinal epithelium, their transcriptomic
profiles were compared to various intestinal cell types and tissues. To
determine the cell type resemblance, we analyzed marker gene expression
for six major intestinal cell types: goblet cells, Paneth cells,
enteroendocrine cells (EEC), intestinal stem cells (ISC), absorptive
enterocytes (AE), and absorptive colonocytes (AC). Our data was
compared with single-cell RNA-sequencing and bulk RNA-sequencing
datasets, as well as a recently described Caco-2-based anaerobic 3D gut
model^[110]30. The clustered heatmap (Fig. [111]4C) reveals that all
the Caco-2 models most closely resemble (proximal) enterocytes of the
small intestine, as reported in the literature^[112]29,[113]31.
Notably, the anaerobic DFC model exhibited higher expression of
AE-specific markers (APOA4, ALPI, MTTP, RBP2, MAF, and LCT) compared to
the static model, indicating a stronger AE profile. Expression levels
of commonly reported genes in intestinal models, including VIL1 and
genes associated with epithelial barrier integrity (TJP1, OCLN), were
also within the range of in vivo human intestinal transcriptional
profiles (Supplementary Fig. S[114]5). Low expression of MUC2,
previously demonstrated by immunostaining and ELISA, was confirmed at
the transcriptional level, consistent with a small intestinal
enterocyte phenotype and in line with data from the Caco-2-based
anaerobic chip model described by Cheng et al. Overall, the
transcriptional profile of our anaerobic DFC model closely mirrors that
of the model by Cheng et al., demonstrating that a similar Caco-2
culture is achieved in our DFC model as in the more complex, anaerobic
PDMS microfluidic-based systems.
Co-culture of obligate anaerobes and Caco-2 cells in the DFC
To test if the model supports growth of obligate anaerobes, it was used
to simulate colonization with the obligate anaerobic bacterial species
C. difficile. The obligate anaerobe B. fragilis, a prominent member of
the normal microbiota, was included to represent the commensal
population^[115]32–[116]34. Bacteria were inoculated seven days
post-seeding of the Caco-2 cells in the DFC. After inoculation,
bacterial colonization was monitored both macro- and microscopically
and by plating the effluent from the apical channel daily. Three days
post inoculation, areas resembling biofilm were visible to the naked
eye, and the bacterial counts in the effluent exceeded 10^7 CFU/ml for
B. fragilis and 10^5 CFU/ml for C. difficile, demonstrating that the
system provides favorable growth conditions for these obligate
anaerobic microorganisms (Fig. [117]5A, Supplementary Fig. S[118]6).
Following bacterial colonization, the viability of the intestinal
epithelium was examined by viability staining and confocal laser
scanning microscopy (CLSM). This analysis showed that a confluent layer
of viable Caco-2 cells was present after the 5-day colonization with C.
difficile and B. fragilis (Fig. [119]5D).
Fig. 5. Colonization with C. difficile and B. fragilis.
[120]Fig. 5
[121]Open in a new tab
The cultured Caco-2 cell epithelium was infected with C. difficile and
B. fragilis 7 days post-seeding in the dual flow chamber (DFC). On the
third day post-infection (DPI), the bacteria-colonized epithelia were
treated with 6 µg/mL vancomycin (VAN) or left untreated (CTRL). A
Bacteria shed from the simulated infection is monitored in CFU/mL
measured in the effluent from the apical channel of the DFC before and
after vancomycin treatment. Data represent the mean ± standard
deviation (SD) from three biological replicates. B Bacteria associated
with the colonized epithelium harvested 5 DPI. The numbers of viable
bacteria are shown in log[10] CFU per cm^2 of the epithelium. Data
represent the mean ± standard deviation (SD) from three biological
replicates. Statistical comparisons were made using an unpaired
Wilcoxon test. C Oxygen levels (% O[2], gas-phase equivalent) was
measured at the DFC apical channel exit site before and at 5 DPI. Black
lines indicate mean values. Statistical comparisons were made using a
paired t test. D Confocal laser scanning microscopy images of LIVE/DEAD
stained epithelium 5 DPI. Live cells stain green, dead cells stain red.
The images are combined from z-stacks. Images are based on one
experiment. E Gram stain of effluent containing shed bacterial clumps
from the apical channel of the DFC 5 DPI. The gram-positive C.
difficile stains blue while the gram-negative B. fragilis stains red.
Vancomycin treatment of the C. difficile colonized epithelium
To assess the DFC model’s applicability for testing antimicrobial
treatment regimens, vancomycin treatment experiments were conducted.
Vancomycin, a common antibiotic used to treat C. difficile infections,
is effective against C. difficile, while B. fragilis tolerates the
drug^[122]34,[123]35. Treatment with vancomycin (6 ug/ml) was initiated
three days post-infection (DPI). After two days of treatment, the CFU
count of C. difficile in the effluent had reduced approximately
300-fold (Fig. [124]5A), but some bacteria were still present in the
aggregates harvested from the chamber (Fig. [125]5E). Interestingly, a
10-fold reduction in the amount of B. fragilis was observed in the
effluent, compared to the untreated control (Fig. [126]5A). However,
when harvesting the cell layer to enumerate the sessile population, the
same amount of B. fragilis was present in the vancomycin treated DFC,
compared to the untreated control (Fig. [127]5B). The amount of C.
difficile associated with the cell layer was reduced nearly 100-fold
following vancomycin treatment, compared to the control (Fig. [128]5B).
Despite this, high numbers of C. difficile were still present in the
vancomycin treated DFC ( > 10,000 CFU/cm2). After treatment, the
majority of C. difficile cells were observed in close association with
B. fragilis aggregates, as indicated in Gram stains of supernatant
samples collected from the apical channel of DFC (Fig. [129]5E).
Measurements of the oxygen concentration at the apical channel exit
before and after the five-day experiment showed a mean decrease in
oxygen levels from 0.6 to 0.2% (Fig. [130]5C). This result shows that
the barrier properties to oxygen remains intact and indicates an
uncompromised epithelium despite bacterial colonization.
Invasion of the intestinal epithelium
To further explore the C. difficile and B. fragilis intestinal
colonization strategy and assess the effect of vancomycin treatment,
transmission electron microscopy (TEM) was performed on treated and
non-treated, colonized epithelia. In the non-treated control, TEM
revealed numerous bacteria located both extra- and intracellularly
(Fig. [131]6, i and ii). In the vancomycin-treated bacterial
co-culture, only a few bacteria were observed extracellularly, but
numerous intracellular bacteria were observed by TEM (Fig. [132]6, iii
and iv). The extracellular colonies contained what appeared to be lysed
bacteria (Fig. [133]6, iii). Additionally, TEM analysis revealed a
consistent layer of microvilli on the lumen of the cultured epithelium
at 5 days post-infection (Fig. [134]6, iii and iv).
Fig. 6. Transmission electron microscopy of bacterial colonization of
cultured Caco-2 epithelium.
[135]Fig. 6
[136]Open in a new tab
Caco-2 epithelia cultured for 7 days in dual flow chambers (DFCs) were
inoculated with C. difficile and B. fragilis and at 3 days
post-infection (DPI) treated with 6 µg/ml vancomycin (VAN) or left
untreated (CTRL). Bacteria and cultured Caco-2 epithelium were
harvested 5 DPI. Bacteria were found both extra- (i, iii) and
intracellular (ii, iv) in both the vancomycin-treated (iii, iv) and
untreated (i, ii) DFCs. Red arrows indicate areas with colonizing
bacteria. White arrows indicate microvilli on the surface of the Caco-2
cells. The black arrow indicates lysed bacteria.
Discussion
In this study, we present a novel in vitro model that allows for the
co-culture of intestinal epithelium with obligate anaerobic bacteria
for several days. The model is suitable for a variety of applications,
such as studying host–bacteria interactions and preclinical testing of
antimicrobial agents and can be used for simulating anaerobic
environmental biofilms as well. Anaerobic conditions (<1% O[2]) can be
maintained in the apical channel in principle indefinitely as it
depends on an effective online oxygen depletion method specifically
invented for the model^[137]24. The approach eliminates the need for
time-consuming pre-anaerobization of media, encapsulation in
N[2]-chambers, and breaking of seals when inspecting the chip or
replacing media. The dual flow chamber (DFC) is oxygen-tight meaning
that the DFC and attached anaerobization unit (AU) can be placed and
handled in the open environment without risking oxygen leakage into the
system, and the DFC culture can be maintained in a standard CO[2]
incubator. Furthermore, the AU can be individually adapted to fit the
desired oxygen levels in other parts of the gastrointestinal tract or
for simulating limited oxygen environmental biofilms, e.g. by
shortening the silicone tube running through the AU, as demonstrated in
this study.
The Caco-2 cell line is able to differentiate when exposed to liquid
shear stress (SS), creating both villus-like structures and in some
cases detectable mucus^[138]36–[139]38, making it an excellent choice
for establishing intestinal models of the small intestine. In the DFC,
Caco-2 cells can be cultured for at least 21 days during which the
cells differentiate to an epithelium with gut-like structures including
villi, microvilli, crypts and mucus. The morphology and differentiation
of Caco-2 cells cultured under low SS is well studied^[140]39 as
gut-on-chip models often use a shear rate of
0.02 dyn/cm^2 ^[141]13–[142]15. However, it can be argued that this is
significantly lower than the physiological SS present in the gut. One
study has reported SS values in the small intestine of guinea pigs of
up to 1.2 dyn/cm^2 ^[143]21, while others suggest that the shear stress
of the small intestine reaches levels between 0.1-10 dyn/cm^2 during
peristaltic movements^[144]22. However, the true SS of the human colon
is yet to be explored. Only a few studies have characterized intestinal
cells cultured under these higher, more physiological SS levels. In one
intestinal chip, the highest reported SS was 0.033 dyn/cm^2^[145]13,
while another study investigated the effect of SS between
0–0.03 dyn/cm^2 in the Hele–Shaw cell culture device^[146]38. The
latter study demonstrates that the highest applied SS led to the
highest levels of F-actin and microvilli, while the barrier integrity
was optimal at SS values between 0.019-0.26 dyn/cm^2. It is important
to note that the study used a single-channel chip, and cells might
behave differently in a dual-channel system. In the DFC model presented
in this study, a robust and differentiated layer of Caco-2 cells
developed and was maintained, capable of withstanding a liquid SS of at
least 0.3 dyn/cm², achieved by gradually increasing the flow rate
during the first seven days of culture. The continuous SS of up to
0.3 dyn/cm^2 in the DFC likely contributed to inducing the morphology
of the cultured epithelium, which often reached 50-140 µm thickness,
exhibiting villus-like structures and tight junctions at 13 days of
maturation. This structure was achieved without application of active
mechanical stretching of the membrane as applied in some PDMS
gut-on-a-chip models^[147]13, though the thin polyester membrane used
in the DFC has some flexibility that could, during the applied
sequential flow, accommodate some level of sequential stretching. It
remains unclear whether membrane stretching is critical for the
induction of villus-like structures in Caco-2 cells. In a study by
Dogan et al., a liquid SS of up to 1.3 dyn/cm^2 was applied to Caco-2
layers in a microtiter-well-based model without mechanical
stretching^[148]37. This approach resulted in similar morphological
features as observed in PDMS models with mechanical stretching^[149]13.
The authors concluded that seeding density and flow rate are the two
main features affecting the morphology of the cell layer^[150]37.
Another critical aspect of SS is its influence on the induction of
virulence factors in adherent bacteria. In a study by Alsharif et al.,
the expression of virulence factors in E. coli increased with
increasing SS^[151]40. Another study by Park et al., showed that SS in
the range between 0.17-0.34 dyn/cm^2 promotes the biofilm formation of
bacteria e.g. P. aeruginosa^[152]41. This, along with its role in
intestinal epithelial development and maturation, highlights the
importance of using physiologically correct fluid shear when studying
host–microbe interactions in vitro^[153]38.
Because mucus is an essential part of the human intestine, the presence
of both neutral and acidic mucins in the cell layer was investigated by
staining with Shiff’s reagent and Alcian blue, respectively. An
increasing amount of neutral mucin staining was found in the cultures
during the 13 days culture period, with a higher signal in the DFC than
in the static culture. The gel-forming glycoprotein MUC2 is a major
component of the intestinal mucus layer—particularly in the colon—where
it plays a key role in bacterial colonization^[154]42. Previous studies
have suggested that hypoxia directly promotes MUC2 expression through
HIF-1α^[155]43. In the DFC, anaerobic conditions led to increased MUC2
protein levels in the Caco-2 cells according to immunostaining;
however, increased MUC2 could not be detected by ELISA. RNA-sequencing
also indicated low basal expression levels compared to in vivo data,
suggesting the levels do not reflect physiological conditions. The
absence of a thick, multi-layered mucus lining above the intestinal
epithelium is a known issue in Caco-2-based models. Recently, it has
been shown that co-culturing the goblet cell HT29-MTX and Caco-2
produces a more sufficient mucus layer than Caco-2 cells in
monoculture^[156]44,[157]45. Furthermore, recent studies have
demonstrated that culturing Caco-2 under air–liquid interface
conditions with vasointestinal peptide leads to the formation of a
functional mucus layer^[158]46. Similar approaches are currently being
explored with our DFC model to further approach a colon-like
epithelium.
Studies have shown that Caco-2 cells can differentiate into various
major intestinal cell types when xenografted into mice, with
indications that this differentiation also occurs when cultured in
gut-mimicking microfluidic models^[159]36,[160]47. To assess whether
this maturation towards a more advanced intestinal epithelium occurs in
our model, we used transcriptomic analysis to compare our DFC model
with intestinal single-cell and bulk transcriptomic profiles. Overall,
based on marker gene expression, our results align with previous
studies, indicating that Caco-2 models predominantly resemble
absorptive enterocytes^[161]29. However, our transcriptomic analysis
does not resolve distinct intestinal cell subpopulations, so the
presence of other cell types cannot be excluded. For example, the
stronger MUC2 signal observed under anaerobic DFC conditions may
suggest the presence of goblet-like phenotypes in a small subset of
cells, possibly obscured in the bulk RNA-seq data. The downregulation
of cilia-related pathways, combined with the upregulation of pathways
associated with nutrient absorption, suggests that DFC conditions drive
cells toward a more absorptive and less quiescent phenotype compared to
the static model, thus more closely resembling a mature, rapidly
renewing intestinal epithelium^[162]27. Relatively few genes were
differentially expressed under anaerobic versus aerobic DFC conditions,
with anaerobic conditions specifically showing upregulation of
metallothionein-encoding genes and downregulation of genes involved in
cell proliferation and the electron transport chain. These findings
indicate that while anaerobic DFC conditions lead to some cellular
adjustments compared to maintaining the DFC fully aerobic, such as
reduced proliferation and hypoxic adaptation, the overall absorptive
phenotype of the cells remains consistent. Interestingly, although a
clear hypoxic response was evident, we observed no significant
upregulation of HIF1A mRNA, a central mediator of hypoxia signaling.
This likely reflects our sampling—taken 6 days into anaerobic culture
(13 days post-seeding)—missing an earlier transient HIF1A mRNA peak
that had already returned to baseline, as reported in prior
studies^[163]48,[164]49. Nonetheless, it remains possible that HIF1α
protein is still stabilized and active under these conditions, given
that its regulation is primarily post-translational during
hypoxia^[165]50. Overall, the observation confirms that the conditions
generated in the anaerobic DFC support the growth and maintenance of a
viable and physiological epithelium, and provide a suitable environment
for anaerobic co-culture. Comparison of the transcriptomic profile with
a recently described anaerobic 3D gut model reveals strong similarities
in the expression of marker genes for intestinal cell types^[166]30,
demonstrating that a similar Caco-2 culture is achieved in our system
as in more complex, PDMS-based microfluidic models. These parallels
suggest a common trend toward small intestine-like characteristics in
microfluidic Caco-2-based gut models, highlighting the need to also
implement these models using cell lines that more closely resemble the
colon, where the microbiota is concentrated and intestinal pathogens
commonly manifest^[167]3.
C. difficile is a major health concern, for which in vitro models to
study its interaction with the human host are critically
needed^[168]51. C. difficile is an obligate anaerobic bacterium^[169]52
that has not previously been cultured long-term (>48 h) in an in vitro
model on a viable layer of intestinal cells. Controlling the oxygen
levels at the epithelium surface to permit growth of this organism is a
challenging task that was made possible here with the alternative DFC
design presented, combined with the online AU. To test if our model
supported co-cultures of epithelial cells and C. difficile in the
presence of a normal gut microbiota bacterium, we co-inoculated DFCs
with C. difficile and B. fragilis. Subsequent treatment of simulated
infection with vancomycin was performed to demonstrate the models’
applicability as a treatment test platform. Vancomycin treatment in
vivo typically clears C. difficile symptoms, but approximately 25% of
patients experience recurrent infection within three months^[170]53,
suggesting that C. difficile has yet unrecognized mechanisms to survive
this treatment. In the untreated control, both B. fragilis and C.
difficile multiplied rapidly during the first three days of the
infection (based on CFU/mL in the effluent), demonstrating that the low
oxygen levels at the apical cell surface are sufficiently low to
support culture of these obligate anaerobes. After this time point,
bacterial shedding reached a steady state until the termination of the
experiment on day five. As expected, the amount of C. difficile
decreased following vancomycin treatment; however, vegetative bacteria
were still present in the DFC after two days of treatment. Dense
bacterial aggregates were observed in the vancomycin treated DFC, and
while a direct protective role of these aggregates was not demonstrated
in this study, the close proximity of C. difficile to B. fragilis
within dense aggregates may contribute to persistence, as bacteria
embedded in biofilm clumps are able to survive vancomycin far exceeding
the bactericidal levels of their planktonic state^[171]54. Few to no
planktonic C. difficile were found in the DFC effluent after two days
of treatment with vancomycin but filamentous gram-positive rods were
identified embedded in shed bacterial aggregates. Additionally,
numerous bacteria were observed by TEM to be located in intracellular
niches. Other pathogens, such as uropathogenic E. coli, are known to
invade epithelial cells, creating intracellular bacterial communities
that are less susceptible to antibiotics^[172]55,[173]56. However,
limited information is available about the intracellular lifestyle of
B. fragilis and C. difficile. Recent studies have shown that C.
difficile spores can localize to the intracellular space of intestinal
epithelial cells, suggesting a potential mechanism for escaping
antibiotic treatment^[174]57. The model presented here provides an
excellent opportunity to further explore potentially
invasive/intracellular lifestyles of C. difficile and other obligate
anaerobes to enhance our understanding of the mechanisms underlying
persistent and recurring intestinal infection. Although both C.
difficile and B. fragilis are generally classified as obligate
anaerobic bacteria, they have been shown to replicate under microoxic
conditions^[175]58,[176]59; however, their growth rate might be
limited. While the oxygen levels achieved in the DFC fall within a
physiologically relevant range^[177]23, tolerable for obligate
anaerobic species such as C. difficile and B. fragilis, it remains
unclear whether more oxygen-sensitive microbes requiring strictly
anoxic conditions (<0.1%) can colonize and grow in this environment.
These conditions, however, could be achieved by optimizing the length
of the silicone tubing used in the AU, in conjunction with adjusting
the flow rate, as illustrated in Fig. [178]2B.
In conclusion, we present an accessible and robust in vitro model for
co-culturing of intestinal epithelial cells and obligate anaerobic
bacteria. This model not only enables detailed studies of direct
host–microbe interactions within a more complex, gut-mimicking, low
oxygen environment but also provides a valuable platform for evaluating
novel treatments for intestinal infections that may advance therapeutic
strategies in the field.
Methods
Fabrication of the dual flow chamber
The dual flow chamber (DFC) was made by attaching two sticky-slide I
Luer (Ibidi®) on both sides of a 12 µm thick tissue culture treated
transparent polyester (PET) membrane with a pore size of 0.45 µm and a
porosity of 0.6% (it4ip, 2000M12/640N453). To ensure a leakage tight
construction, an even amount of pressure was applied to the
sticky-slides using screw clamps and plastic blocks with predrilled
holes for the inlet and exit for 10–15 s (see supplementary Fig.
S[179]1). Both sides of the DFC were filled with 70% ethanol and left
for 20 min to sterilize followed by curing of the glue at 37 °C
overnight.
Anaerobization unit (AU)
To ensure a continuous supply of anoxic media, an ascorbate solution
was made by adding 2 g of sodium L-ascorbate (Sigma-Aldrich) to 100 mL
of a 0.1 M NaOH solution (Sigma-Aldrich). Inside the solution, a 150 cm
long silicone tube (Helixmark®; mat.no: 456350287, inner diameter (ID):
0.99 mm, wall thickness (WT): 0.31 mm) was placed. The tube dimensions
were chosen to ensure optimal exchange of oxygen. To prevent
reabsorption of oxygen into the media, which happens extremely fast,
the anoxic media leaving the anaerobization unit (AU) was connected to
the DFC and/or oxygen sensor using stainless steel tubing. Before
connecting to the DFC, culture media were flushed through the AU at a
flow rate of 320 µL/min for at least one hour to stabilize the system.
Cell culture
Caco-2 HTB-37™ cells (ATCC) were cultured in Dulbecco’s modified
eagles’ media (DMEM; Gibco) supplemented with 20% heat-inactivated
fetal bovine serum (HI FBS; Biowest) and 1% penicillin-streptomycin
(PS; Gibco) (Stock: 10,000 units/mL Penicillin and 10,000 µg/mL
Streptomycin). Cells were split when 60-90% confluent (twice–thrice
weekly) and used for experiments in passage 32-54. All cultures were
maintained in a standard CO[2] incubator at 5% CO[2] and 37 °C.
Prior to seeding of cells in DFCs and cell culture inserts (Falcon;
353090), the apical (upper) side of membranes was coated with Collagen
Coating Solution (SAFC®; 125-50) by applying the solution, incubating
at 37 °C for 30 min, and washing thrice with phosphate-buffered saline
(PBS). In DFCs, Caco-2 cells were seeded on the membranes at a density
of 1-2x10^5 cells/cm^2 and left to adhere for four hours before
starting the flow at 15 µL/min (shear stress (SS): 0.014 dyn/cm^2). One
day post seeding (DPS) the flow was set to 60 µL/min (SS:
0.057 dyn/cm^2) and left at this flow rate until 6 DPS. On the 6^th
day, the flow was changed to pulsating flow (1 min flow followed by a
9-min break) with peak flow rates of 320 µL/min (SS: 0.3 dyn/cm^2). At
7 DPS, medium was shifted to DMEM with reduced HI FBS at 2%, and flow
set to a contiuous 320 µL/min (SS: 0.3 dyn/cm2) to maintain stable
anaerobic conditions in the apical channel. The same flow conditions
were applied at the apical and basolateral channels for all time
points. Shear stress was calculated using Eq. [180]1 as advised by
Ibidi^[181]60:
[MATH: Shear stressSS=0,0072×131.6×flow rate(ml/min)
:MATH]
1
For the static cultures, the cell medium was changed every third day
and shifted to serum-reduced medium on day 7 post-seeding.
Oxygen measurements
Oxygen measurements were performed using an O[2] MicroOptode installed
in a PEEK flow cell connected to a single-channel O[2] UniAmp
(UniSense, Denmark). Measurements were made every 10-60 s by flowing
media through the flow cell at a flow rate between 30-1440 µL/min.
Measurements were logged using SensorTrace Suite Logger software
(UniSense, Denmark). The sensor was calibrated using the manufacturers’
guidelines. For oxygen measurement with the DFC, the sensor was
connected to the exit site using stainless steel tubing to prevent
diffusion of oxygen into the media.
The oxygen percentage at the inlet of the DFC depends on the length of
the silicone coil and flow rate, thus these can be adapted to fit the
desired oxygen concentration. For <0.5% oxygen at the inlet, the media
must be in the silicone coil for at least 2 min. The relationship
between tube length, flow rate, and achieved oxygen concentration is
outlined in Fig. [182]2B, [183]C. The length of the coil can then be
adapted to fit the desired flow rate, by using the following formula
(Eq. [184]2):
[MATH: Lenght(cm)=minutes×flow(ul/min)7.38 :MATH]
2
The above key is for the silicone tubing applied here and will diverge
depending on silicone material and tubing dimensions.
Immunofluorescence staining
Before staining, cells were washed with Hank’s balanced salt solution
(HBSS; Gibco) thrice and fixated with a 10% neutral buffered formalin
solution (Sigma-Aldrich) for 15 min. Formalin was removed and cells
were washed thrice with PBS. Cells were permeabilized with 0,1% Triton
X-100 in PBS for 15 min, washed thrice with PBS and blocked with 5%
bovine serum albumin (BSA; Sigma-Aldrich) in PBS for 60 min. Cells were
stained at room temperature with a mixture of 2 µg/mL of Occludin
Monoclonal Antibody, Alexa Fluor 488 (Invitrogen; OC-3F10) and 2 µg/mL
MUC2 Antibody, Alexa Fluor 405 (Novus Biologicals; 944/152) in 1% BSA
in PBS for 3 h. Cells were washed with PBS and F-actin stained with
100 nM Acti-stain™ 555 Phalloidin (Cytoskeleton, Inc.) for 30 min
before washing thrice with PBS. The membranes were removed from the
sticky slides using scalpels and assembled in silicone frames on
microscopy slides, mounted with Fluorescence Mounting Medium (Dako) and
sealed with a coverslip.
Mucus staining
The Alcian Blue/PAS staining kit (Artisan; AR16992-2) was used to
visualize mucins in formalin-fixated Caco-2 cells. For staining of
neutral mucins, cells were first incubated with periodic acid for
2 min, followed by washing thrice with PBS. Cells were then incubated
with Shiff’s reagent for 10 min and washed with PBS thrice to remove
excess staining solution. For staining of acidic mucins, cells were
incubated with Alcian Blue for 15 min, followed by washing once with
distilled water and trice with 3% acetic acid. Membranes were stored in
PBS until imaging to prevent drying.
Viability staining
Viability of cells was visualized using LIVE/DEAD™
Viability/Cytotoxicity Kit (Invitrogen; L3224) by adding 2 µL
Calcein-AM and 2 µL Ethidium homodimer-1 to 2 mL of HBSS before
transferring it to the apical channel of the DFC. Cells were stained
for 30 min followed by a 15-min fixation step with formalin. The cells
were washed thrice with HBSS before each step. The membranes were
removed from the sticky slides using scalpels and assembled in silicone
frames on microscopy slides, mounted with SlowFade^TM Diamond Antifade
Mountant with DAPI (Invitrogen; [185]S36968) and sealed with a
coverslip.
RNA isolation
To isolate RNA from Caco-2 cells grown in DFCs, culture medium was
aspirated, and ice-cold lysis buffer (4 M guanidinium thiocyanate
(GITC), 0.02 mM Tris-HCl (pH 7.5), 10 mM NaAcetate pH 4.5, 25 mM EDTA,
0.1% Triton X-100, 2 mM DTT) was added. The cells were sheared from the
DFC by rapidly moving a syringe, fitted into the Luer port of the
apical channel outlet, up and down. The lysate was then transferred to
RNase-free 1.5 mL tubes and snap-frozen in liquid nitrogen. RNA
extraction was subsequently performed using a phenol-chloroform method.
Briefly, 300 µL of lysate was mixed with 150 μL of solution 2 (10 mM
Na-acetate pH 4.5, and 2% SDS), 700 μL of acidic phenol (pH 4.5) and
300 μL of chloroform. Tubes were inverted and heated at 80 °C for
3-4 min, then cooled on ice. After centrifugation at 10,000g for 5 min,
the aqueous phase was transferred to 96% ethanol with Na-acetate
(37.5 mM) and precipitated overnight (ON). RNA was pelleted by
centrifugation (20,000g for 45 min), washed in ice-cold ethanol,
resuspended in RNase-free H[2]O, and stored at −20 °C or −80 °C.
RNA-sequencing
RNA samples from Caco-2 cultures grown in DFCs were depleted of
ribosomal RNA using the NEBNext rRNA Depletion Kit (Human/Mouse/Rat).
RNA sequencing library preparation was performed with the NEBNext Ultra
II Directional RNA Library Prep Kit for Illumina (New England Biolabs)
and paired-end sequencing was conducted on a NovaSeq 6000 System
(Illumina). Raw paired-end reads were aligned to the human genome
assembly GRCh38/hg38 using STAR (version 2.7.11a). Primary alignments
were filtered, sorted, and indexed using samtools (version 1.19). For
feature counting, only protein-coding genes were considered. Read
counts within exons were determined using featureCounts from the
subread package (version 2.0.6), with multimapping reads included and
counted fractionally. Differential gene expression analysis was
conducted with edgeR (version 4.2.1), identifying genes with |log2(FC)|
> 2 and FDR < 0.05 as significantly differentially expressed. Pathway
enrichment on significant DEGs was conducted using the webtool
MetaScape with Reactome gene sets^[186]25. For marker gene expression
analysis, lineage marker genes from Burclaff et al.^[187]61 were used.
Normalized gene expression data (normalized transcript per million,
nTPM) from the “RNA single cell type data” and “RNA consensus tissue
gene data” datasets from The Human Protein Atlas were used for
comparison^[188]62,[189]63. Clustered heatmaps were generated with the
R package pheatmap (version 1.0.12), using Ward’s method (“ward.D2”)
for hierarchical clustering.
Colonization with Clostridioides difficile and Bacteroides fragilis
Caco-2 cells were matured aerobically for seven days in the DFC, as
previously described. On the day of inoculation, the media were
replaced with DMEM containing 2% HI FBS, and the AU was connected to
the inlet of the apical channel. Antibiotic-free media was flushed
through both channels of the DFC for one hour before infection.
Bacterial suspensions of C. difficile (ATCC 700057) and B. fragilis
(ATCC 25285), grown anaerobically overnight on 5% blood agar plates,
were adjusted to an optical density at 600 nm (OD600) of 0.2 in
deoxygenated Hanks’ Balanced Salt Solution (HBSS, Gibco). A mixture of
B. fragilis and C. difficile was made from the solutions (OD ratio 1:9)
and introduced through the apical channel at a flow rate of 320 µl/min
until the suspension reached the top of the outlet reservoir. The
colony forming unit (CFU) per mL in the inoculum was 5*10^6 for C.
difficile and 1*10^8 for B. fragilis. This ratio was selected to
promote co-colonization by both bacterial species while reflecting a
predominance of the commensal microbiota, as represented by B.
fragilis. The discrepancy between CFU counts can be explained by the
difference in optical absorption, as C. difficile CFU are >100 times
lower than B. fragilis, when adjusting to the same OD600. The flow was
then stopped for 10 min to allow for initial bacterial attachment,
after which the flow rate was resumed at 320 µl/min. Daily effluent
samples from the apical channel were plated on selective agar plates
(CHROMID® C. difficile agar plates and brain heart infusion agar plates
with 6 µg/mL vancomycin (Bactocin®, MIP Pharma)) to measure CFU. On day
3, the medium in DFCs allocated for antibiotic treatment was changed to
DMEM with 2% HI FBS containing 6 µg/mL vancomycin. On day 5, loose
aggregates were aspirated, and the cell layers of the DFCs were
harvested using Trypsin-EDTA (0.25%; Biowest) with 0.1% Triton X-100,
then plated for CFU enumeration. Gram staining was performed on loose
aggregates harvested from the apical channels. For microscopy (confocal
laser scanning microscopy (CLSM) or transmission electron microscopy
(TEM)), the infected cell layers were not harvested but instead
prepared according to the appropriate protocol.
Microscopy
Live cultures of Caco-2 cells were imaged using an Olympus CKX53
inverted microscope connected to an Olympus SC50 camera and using
Olympus cellSens software. PAS-stained cultures were imaged using a
Leica DM4 B microscope using LAS X software. Cells were imaged in the
middle of the flow chamber. Gram-stained aggregates harvested from the
apical channel of the flow chambers were imaged using a Leica DM3000
LED microscope connected to a Flexacam C1 and using LAS X software.
Immunofluorescence stained and LIVE/DEAD stained cultures were imaged
using either a Nikon AX or an Olympus FV1000 MPE CLSM. Cultures were
imaged between the inlet and the center of the flow chamber.
Transmission electron microscopy
The Caco-2 cells were matured in flow chambers for seven days followed
by inoculation with C. difficile and B. fragilis and treatment with
vancomycin as described previously. The cell cultures were fixed using
a 2% glutaraldehyde solution in 0.04 M phosphate buffer. The membranes
with adhering cell cultures were cut out from the flow chambers, washed
with 0.1 M phosphate buffer, and stained with 1% osmium for 90 min.
Following staining, the membranes were washed in phosphate buffer and
water, serially dehydrated in ethanol and acetone, and infiltrated with
TAAB 812 Embedding Resin (T030, TAAB).
Ultrathin (60 nm) sections were cut on a Leica Ultracut UCT microtome.
The sections were collected on Formvar support film copper grids
(FF2010-CU-50, Electron Microscopy Sciences). The grids were stained
with 3% uranyl acetate for 15 min. at 60 °C and 3% lead citrate (Leica
Ultrostain 2) for 6 min at room temperature. The cells were
photographed using a JEM-1400 Plus transmission electron microscope,
equipped with Quemsa TEM CCD camera and Radius imaging software.
Statistics
Statistical analyses were performed using R (version 4.3.2). Normality
of the differences in paired oxygen concentration measurements was
assessed using the Shapiro-Wilk test. Based on this, a paired t-test
was conducted using the compare_means function from the ggpubr package
(method = “t.test”). For the comparison of CFUs between treated and
untreated DFCs, a non-parametric test (Wilcoxon rank-sum test) (method
= “wilcox.test”) was applied.
Resource availability
Lead contact
For further information, professor Thomas Emil Andersen
(thandersen@health.sdu.dk) can be contacted.
Supplementary information
[190]Supplementary Materials^ (6.3MB, docx)
Acknowledgements