This content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. HC is supported by grant P20GM134973 from the National Institutes of Health. KB is supported by a Children&#39;s Hospital Foundation (CHF) and Presbyterian Health Foundation (PHF) grant. We thank the Laboratory for Molecular Biology and Cytometry Research at OUHSC for the use of the Core Facility, which provided confocal imaging.

All animal procedures in this study were approved by the University of Oklahoma Health Sciences Center Institutional Animal Care and Use Committee. Small intestine convenience samples from a preterm non-human primate (NHP, 90% gestation, olive baboon, Papio anubis) were obtained following euthanasia for a separate study (Protocol #101523-16-039-I).
1. Establishment of apical-out enteroid NEC-in-a-dish model
<ol>
	<li>Preparation of media and enteroid treatments 
	NOTE: Once prepared following standard aseptic technique, the media can be stored at 4 &#176;C for up to 1 week.
	<ol>
		<li>Prepare antibiotic-free media (AFM) by mixing 50 mL of human organoid growth media with 50 mL of organoid supplement (see Table of Materials). Prepare 5 mM ethylenediaminetetraacetic acid/1x phosphate-buffered saline (EDTA/PBS) by adding 100 &#181;L of 0.5 M EDTA to 9,900 &#181;L of PBS.</li>
		<li>Chill Dulbecco&#39;s modified eagle&#39;s medium/nutrient Ham&#39;s mixture F12 + 15 mM HEPES Buffer (DMEM/F12) at 2-8 &#176;C.</li>
		<li>Prepare TNF-&#945; stock solution by suspending 100 &#181;g of TNF-&#945; powder in 1 mL of 1x PBS. Further dilute the TNF-&#945; stock to 20 ng/mL and 50 ng/mL concentrations using AFM as the solvent. Prepare LPS stock solution by suspending 2 mg of LPS in 1 mL of 1x PBS. Further dilute stock LPS to 100 &#181;g/mL and 200 &#181;g/mL using AFM as the solvent.</li>
	</ol>
	</li>
	<li>Reversing the polarity of 3D enteroids 
	NOTE: The following procedure is intended for the reversal of a single 24-well flat-bottom tissue culture plate into two 24-well ultra-low attachment tissue culture plates. The tissue culture plates should be warmed to 37 &#176;C for at least 30 min prior to use. The derivation of basolateral-out 3D enteroids was achieved as described previously by many groups35,36, with slight modifications for media (detailed above). The general enteroid derivation procedure does not differ from that of humans. In brief, excised tissue is washed, cut into small fragments, and incubated in cell disruption buffer. The cell solution is then run through a 70 &#181;M cell strainer, resulting in crypts that are plated at a high density.
	<ol>
		<li>Aspirate media from each well of a 24-well plate of established 3D basolateral-out enteroids.</li>
		<li>Add 500 &#181;L of 5 mM ice-cold EDTA/PBS to each well of the 24-well plate and gently mechanically disrupt the basement membrane extract/ECM dome by pipetting up and down with a p200 pipette a minimum of 5x-6x.</li>
		<li>Transfer the contents of four wells to a 15 mL conical tube and add 8 mL of ice-cold EDTA/PBS to the conical tube. Transfer the remaining 20 wells in the same manner.</li>
		<li>Incubate 15 mL conical tubes at 4 &#176;C for 1 h on a rotating platform or shaker at 330 rpm. After the 1 h incubation, centrifuge the conical tubes at 300 x g for 3 min at 4 &#176;C. Remove and discard the supernatant from each tube.</li>
		<li>Combine and wash the cell pellets with 5 mL of DMEM/F12 and distribute the cell suspension into two 15 mL conical tubes.</li>
		<li>Pellet the cells by centrifuging at 300 x g for 3 min at 4 &#176;C. Remove the supernatant and add 12 mL of AFM to each tube, ensuring the cell pellets are resuspended.</li>
		<li>Pipette 500 &#181;L of the enteroid suspension into each well of two 24-well ultra-low attachment plates.</li>
		<li>Incubate the plates at 37 &#176;C and 5% CO2 for 2-5 days or until enteroids have reversed polarity.</li>
		<li>To check enteroid reversal, first establish the confirmatory immunofluorescent staining and the average time to reversal (~3 days), then confirm the polarity reversal using a basic light microscope visualization. Apical-out enteroids are very cellular in appearance, while basolateral-out enteroids are often dominated by the presence of a large central lumen or budding (see Supplementary Figure 1). 
		NOTE: Once the procedure has been standardized, the conversion rate to apical-out typically exceeds 95%. The use of apical-out enteroids is intended as a terminal experiment, though it is theoretically possible to passage apical-out enteroids into a small number of basolateral-out enteroids.</li>
	</ol>
	</li>
	<li>Apical-out NEC-in-a-dish 
	NOTE: Once the enteroids have reversed polarity, use the cultures in subsequent experiments quickly, as their longevity in apical-out conformation has not been confirmed beyond 3-4 days. For the following NEC-in-a-dish model, the enteroids were treated for 24 h, then collected immediately afterward and preserved for downstream experiments.
	<ol>
		<li>Once the enteroids have visually reversed polarity with at least 95% efficiency, remove the plates from the incubator and collect eight wells of a 24-well plate into a 15 mL conical tube. As the apical-out enteroids are in suspension, simply pipette the enteroid/media solution. Centrifuge the 15 mL tube at 300 x g for 5 min at 4 &#176;C.</li>
		<li>Once the cells are pelleted, remove the supernatant and resuspend the cell pellet in 4 mL of AFM mixed with the desired volume (10 &#181;L here to match the highest treatment volume added) of 1x PBS (control).</li>
		<li>Pipette 500 &#181;L of enteroid/PBS/AFM suspension into eight wells of a 24-well ultra-low attachment plate and incubate the plate at 37 &#176;C and 5% CO2 for 24 h.</li>
		<li>Repeat Steps 1.3.1.-1.3.3. for TNF-&#945; (20 ng/mL and 50 ng/mL) and LPS (100 &#181;g/mL and 200 &#181;g/mL) treatments at normoxia. For all hypoxia treatments, repeat Steps 1.3.1.-1.3.3., but place the plate in a separate incubator at 37 &#176;C, 5% CO2, and 1% O2 for 24 h. 
		NOTE: Step 1.3.4. can be done simultaneously with Steps 1.3.1.-1.3.3, but, for clarity, the procedure above&#160;was separated by treatment in order to reduce complexity.</li>
	</ol>
	</li>
</ol>
2. Immunofluorescent staining and confocal microscopy
<ol>
	<li>Preparation of staining solutions
	<ol>
		<li>Prepare 0.1% Triton X-100 by mixing 7 &#181;L of Triton X-100 in 1x PBS to a final volume of 7 mL. Prepare PBST (PBS-Tween) by adding 30 &#181;L of Tween-20 to 1x PBS to a final volume of 30 mL.</li>
		<li>Prepare 10% normal donkey serum (NDS)/PBST by adding 700 &#181;L of NDS to 6.3 mL of PBST. Prepare 1% NDS/PBST by adding 70 &#181;L of NDS to PBST to a final volume of 7 mL. 
		NOTE: Donkey serum was&#160;used here to correlate with the secondary antibody host. However, the blocking serum species should match the secondary antibody species in order to properly block non-specific binding.</li>
	</ol>
	</li>
	<li>Staining (Day 1)
	<ol>
		<li>Transfer the contents of four wells of a 24-well plate to a 1.5 mL microcentrifuge tube. Repeat for all desired wells/treatments, with each treatment in a separate tube. Centrifuge the tubes at 300 x g for 4 min at 4 &#176;C. Once the enteroids are pelleted, remove the supernatant.</li>
		<li>Resuspend the cell pellets in 300 &#181;L of 4% formaldehyde fixative for 30 min at room temperature (RT). After 30 min, centrifuge the tubes at 300 x g for 4 min at 4 &#176;C, remove the supernatant, and wash the pellet with 500 &#181;L of sterile, RT 1x PBS.</li>
		<li>Centrifuge the tubes at 300 x g for 4 min at 4 &#176;C. Remove the supernatant and add 500 &#181;L of 0.1% Triton X-100, and let sit for 1 h at RT.</li>
		<li>After 1 h, centrifuge the tubes at 300 x g for 4 min at 4 &#176;C and remove the supernatant. Wash with 500 &#181;L of 1x PBS and place the tubes on a rotator or shaker at 200 rpm for 15 min at 2-8 &#176;C. Repeat this a total of 4x.</li>
		<li>Centrifuge the tubes at 300 x g for 4 min at 4 &#176;C and remove the supernatant. Add 500 &#181;L of 10% NDS/PBST and incubate at RT for 45 min. During the incubation, prepare primary antibody solutions by combining 20 &#181;L of recombinant anti-villin antibody, 10 &#181;L of E-cadherin antibody, and 1970 &#181;L of 1% NDS/PBST for eight wells of a 24-well plate.</li>
		<li>After the 45 min incubation, centrifuge the tubes at 300 x g for 4 min at 4 &#176;C. Place the tubes on ice. Remove the supernatant and replace with 250 &#181;L of primary antibody solution. Incubate the tubes overnight at 2-8 &#176;C.</li>
	</ol>
	</li>
	<li>Staining (Day 2)
	<ol>
		<li>Centrifuge the tubes at 300 x g for 4 min at 4 &#176;C and remove the supernatant. Add 250-500 &#181;L of PBST to each tube, depending on the size of the pellet, and place on the rotator or shaker at 200 rpm for 1 h at 2-8 &#176;C. Repeat the PBST wash 4x.</li>
		<li>Prepare secondary antibody solution by adding 52 &#181;L of Cy3 donkey anti-rabbit IgG (H + L) to 50% glycerol and 5.2 &#181;L of donkey anti-mouse fluorescent green 488 secondary antibodies to 5142.8 &#181;L of 1% NDS/PBST.</li>
		<li>Following the last PBST wash and centrifugation, remove the supernatant from the tubes. Dispense 200 &#181;L of secondary antibody solution to each tube and incubate in the dark overnight at 2-8 &#176;C.</li>
	</ol>
	</li>
	<li>Mounting stained apical-out enteroids (Day 1)
	<ol>
		<li>Bring glycerol-based mountant containing blue nuclear dye to RT over 1 h.</li>
		<li>Centrifuge the tubes at 300 x g for 4 min at 4 &#176;C. Remove 100 &#181;L of the supernatant and resuspend the cells in the remaining ~100 &#181;L of supernatant. Transfer the cell suspension to 0.5 mL tubes.</li>
		<li>Centrifuge the tubes in a mini centrifuge for 20 s to pellet the cells. Remove the supernatant and resuspend the cell pellets in 100 &#181;L of far-red nucleic acid stain. Incubate the tubes at RT for 10 min.</li>
		<li>Centrifuge the tubes in a mini centrifuge for 20 s to pellet the cells. Remove the supernatant and resuspend the cell pellets in 100 &#181;L of 1x PBS. Repeat for a second wash.</li>
		<li>Centrifuge the tubes in a mini centrifuge for 20 s to pellet the cells and remove 70 &#181;L of the supernatant. Resuspend the cells in the remaining volume of PBS and transfer the contents to a labeled coverslip (24 mm x 60 mm).</li>
		<li>Apply 75 &#181;L of mountant directly onto the specimen and remove any bubbles with a pipette tip. Allow the coverslips to cure overnight (18-24 h) at RT in the dark.</li>
	</ol>
	</li>
	<li>Mounting stained apical-out enteroids (Day 2)
	<ol>
		<li>After curing overnight, apply a thin layer (~15 &#181;L) of glycerol to the coverslips. Mount the coverslip to the glass slide and gently press into place. Tap to remove any bubbles between the coverslip and the slide.</li>
		<li>Allow the coverslip to set at RT in the dark for a minimum of 2 h, before imaging at 20x magnification with a confocal microscope. For microscopic visualization of enteroids using various confocal acquisition methods, refer to Lallemant et al.37.</li>
	</ol>
	</li>
	<li>Immunofluorescent quantification 
	NOTE: This method determines the corrected total fluorescence by removing the background signal using ImageJ software (https://imagej.nih.gov/ij/).
	<ol>
		<li>Open the confocal images in ImageJ and outline the desired area with the region of interest (ROI) tool.</li>
		<li>Set user-defined parameters by navigating to Analyze &#62; Set Measurements. Select Area &#62; Integrated Density &#62; Mean Grey Value under the settings tab.</li>
		<li>Navigate to Analyze &#62; Measure. A pop-up window will appear containing the measurements. Copy and paste these measurements into a spreadsheet.</li>
		<li>To subtract the background signal, select a small non-fluorescent area of the image. Navigate to Analyze &#62; Measure for that region. A pop-up window will appear containing the measurements. Copy and paste the output into a spreadsheet.</li>
		<li>Multiply the area of the selected cell by the average fluorescence of background readings, and subtract the sum from the integrated density to calculate the corrected total cell fluorescence (CTCF). Repeat the steps above for all images of interest, while maintaining records in a spreadsheet or statistical software package (see Table of Materials).</li>
	</ol>
	</li>
</ol>
3. Apical-out NEC-in-a-dish cell viability
<ol>
	<li>Enteroid treatments
	<ol>
		<li>Establish apical-out NEC-in-a-dish model for 24 h, as in Step 1.</li>
		<li>Thaw cell viability assay reagent overnight at 4 &#176;C. On the day of assay, bring the reagent to RT 30 min before use. Invert the reagent to mix.</li>
		<li>Before performing the assay, remove 100 &#181;L of media from each well, leaving the remaining 400 &#181;L. Add 400 &#181;L of cell viability assay reagent to each well.</li>
		<li>Vigorously mix well the contents on a plate shaker at RT for 5 min at 200 rpm to induce cell lysis. Following shaking, incubate the plate at RT for an additional 25 min.</li>
		<li>After 25 min incubation, mix the contents of one well via pipetting and transfer 200 &#181;L of the 800 &#181;L to a single well of a 96-well, opaque, polystyrene, clear-bottomed plate. Repeat for the remaining 600 &#181;L, creating four technical replicates per well. Transfer the remaining contents of each well of the 24-well plate into a 96-well plate in this manner.</li>
		<li>Using a plate reader capable of luminescence, record values at 0.25 ms integration and compare the relative values among treatments. Alternatively, compare treatment luminescence to an adenosine triphosphate (ATP) standard.</li>
	</ol>
	</li>
</ol>

The authors have nothing to disclose and have no conflicts of interest.

The recent development of enteroid models derived from intestinal epithelial crypts allows for a more physiologically relevant in vitro tissue in which to study necrotizing enterocolitis pathogenesis. Despite including all major differentiated cell types of the intestinal epithelium, 3D enteroids are still subject to several significant limitations. The conventional, basolateral-out enteroids are suspended in 3D ECM hydrogel domes, the composition and density of which may limit normal diffusion within the tissue...

Necrotizing enterocolitis (NEC), a severe inflammatory disease of the small intestine occurring in up to 10% of preterm infants, is commonly associated with high morbidity and mortality1,2. Mortality rates approaching 50% in very low birth weight (&#60;1500 g) infants, requiring surgical intervention, are not uncommon3. While the exact etiology of NEC is not currently understood, risk factors, such as formula feeding, are thought to compound with physiological anomalies, such as dysbiosis, an immature intestinal epithelium, and a dysfunctional intestinal barrier, in the development of the disease2,4. Despite significant effort, little progress in the prevention or treatment of NEC has occurred over the last decade5. A novel in vitro method to study NEC and associated intestinal epithelial barrier dysfunction is required to advance understanding of the pathogenesis of the disease, as findings from animal models have, thus far, translated poorly to the bedside6.
A number of in vitro models have been utilized to investigate the mechanisms at play during NEC. The human intestinal epithelial cell line, Caco-2, is among the most commonly utilized in vitro models of NEC7,8. Caco-2 cells emulate brush border morphological features of the small intestine, but, as a cell line, do not differentiate into the wide variety of in vivo cell types, including mucus-producing goblet cells, required for a highly translatable model. HT-29-MTX, human colon adenocarcinoma cells, include a mixed enterocyte and goblet cell phenotype, but still lack crypt-based cell types of the intestinal epithelium9. IEC-6 and IEC-18 are non-transformed cell lines with an immature ileal crypt-like morphology but are not derived from human tissue, limiting their translational capacity. FHs 74-Int and H4 intestinal cell lines are derived from human fetal tissue and do not form tight junctions or polarized monolayers10,11, and thus are immature compared with even the most premature infants susceptible to NEC. Typically, NEC in vitro models utilize lipopolysaccharide (LPS) treatments to induce toll-like receptor 4 (TLR4), a major receptor initiating intestinal inflammation in NEC12. Damage mediated through reactive oxygen species (ROS) treatment, typically via hydrogen peroxide, is often used to induce NEC-like oxidative damage and apoptosis13,14. As the main driver of intestinal inflammation, tumor necrosis factor-alpha (TNF-&#945;), a downstream component of the inflammatory TLR4 signaling, is also commonly utilized in these in vitro models to mimic in vivo pathogenesis15.
Organoids, generated from inducible pluripotent stem cells (iPSCs), have grown in popularity as an in vitro model of the intestine due to their ability to recapitulate the complex in vivo architecture and cell-type composition of the tissue from which they are derived16,17. A related in vitro system, enteroids, are organoids derived from resected intestinal crypts that are more easily established and maintained than iPSC-derived organoids. Enteroids are typically grown in a three-dimensional (3D) extracellular matrix (ECM) with experimental access limited to the basolateral cell surface. Methods, such as microinjection18,19, have been developed to overcome this barrier to the apical surface, but the buildup of sloughed cellular debris and mucus within the lumen renders microinjection both technically difficult and inconsistent. As custom robotic microinjection platforms are not widely accessible20, lab-to-lab variability in technical ability and general technique become significant variables to overcome with microinjection protocols. Two-dimensional (2D) monolayers derived from dissociated 3D enteroids, still comprising all major cell types of the intestinal epithelium, allow access to the apical surface but have traditionally been difficult to maintain without a feeder layer of mesenchymal myofibroblasts21. While cell culture permeable supports can be used to access both the apical and basolateral sides of enteroid monolayers without the use of underlying myofibroblasts, these inserts require excision and mounting of the membrane before use with modalities such as confocal microscopy, resulting in a more technically demanding and difficult process when using traditional microscopy methods22. NEC has been modeled in vitro using traditional 3D enteroids23,24,25 and permeable supports26,27, and intestinal inflammation has recently been replicated with gut-on-a-chip models28,29. While gut-on-a-chip models incorporating microfluidics are, by far, the most advanced and translatable models, this technology is expensive, complex, and inaccessible to most investigators30.
Recent advances in apical-out enteroid techniques have allowed easier access to the apical surface of 3D enteroids without risking damage to the structural integrity of the in vitro epithelium31,32,33. Apical-out enteroids share the cell type composition and barrier function of in vivo intestinal epithelium, but, unlike typical 3D enteroids, the apical surfaces of these cells face the culture medium, allowing for more physiologically relevant studies on nutrient absorption, microbial infection, and luminal secretion31. An additional advantage of apical-out enteroids is the&#160;ability to homogenously distribute experimental agents to enteroids. Varying treatment volumes based on enteroid size is not required, as it is with microinjection, and the ability to maintain these enteroids in suspension culture negates any ECM interference on experimental agent diffusion32.
Necrotizing enterocolitis is a multifactorial disease involving multiple intestinal epithelial cell types and a variety of environmental and pathophysiological factors34. The varied cell composition of intestinal enteroids is a clear improvement over monocultures in modeling a complex disease such as NEC. Interestingly, while a single inflammatory exposure is often sufficient to induce damage in in vitro monocultures, enteroids, as with mouse models23, appear to require a minimum of two inflammatory components to induce NEC-like damage6. Here, we present an apical-out NEC-in-a-dish model, using apical-out enteroids in combination with hypoxia (an important clinical feature of NEC6) and either LPS or TNF-&#945;, as an improved and more physiologically relevant in vitro model to study epithelial responses to NEC-like inflammation and, potentially, identify therapeutic targets. We describe a protocol for reversing the polarity of small intestinal 3D enteroids, as well as an immunofluorescent staining protocol to identify epithelial barrier disruption and junctional protein expression alterations. Finally, we further demonstrate a simple enteroid viability assay to determine the impact of our dual-hit, apical-out NEC-in-a-dish model.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5 M EDTA, pH 8.0</td>
			<td>Fisher Scientific</td>
			<td>15575-020</td>
			<td></td>
		</tr>
		<tr>
			<td>1.5 mL microcentrifuge tubes</td>
			<td>Fisher Scientific</td>
			<td>05-408-129</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL Conical tube</td>
			<td>VWR</td>
			<td>89039-666</td>
			<td></td>
		</tr>
		<tr>
			<td>CellTiter-Glo 3D Cell Viability Assay</td>
			<td>Promega</td>
			<td>G9681</td>
			<td></td>
		</tr>
		<tr>
			<td>Corning Costar Ultra-Low Attachment 24-Well Microplates</td>
			<td>Fisher Scientific</td>
			<td>07-200-602</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover Glass 24 mm x 60 mm</td>
			<td>Thermo Scientific</td>
			<td>102460</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-Mouse IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 488</td>
			<td>Thermo Scientific</td>
			<td>A-21202</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey Anti-Rabbit IgG Antibody, Cy3 conjugate</td>
			<td>Sigma-Aldrich</td>
			<td>AP182C</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle&#39;s Medium/Nutrient Ham&#39;s Mixture F-12 (DMEM-F12) with 15 mM HEPES buffer</td>
			<td>STEMCELL Technologies</td>
			<td>36254</td>
			<td></td>
		</tr>
		<tr>
			<td>E-cadherin antibody (7H12)</td>
			<td>Novus Biologicals</td>
			<td>NBP2-19051</td>
			<td></td>
		</tr>
		<tr>
			<td>Formaldehyde solution 4%, buffered, pH 6.9</td>
			<td>Millipore Sigma</td>
			<td>1004960700</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Sigma-Aldrich</td>
			<td>56-81-5</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>Fiji</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>IntestiCult Organoid Growth Medium (Human)</td>
			<td>STEMCELL Technologies</td>
			<td>06010</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica SP8 Confocal Microscope</td>
			<td>Leica Biosystems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Lipopolysaccharides from Escherichia coli O111:B4, purified by gel filtration chromatography</td>
			<td>Millipore Sigma</td>
			<td>L3012-10MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fisher Scientific</td>
			<td>12-544-7</td>
			<td></td>
		</tr>
		<tr>
			<td>Normal Donkey Serum</td>
			<td>Sigma-Aldrich</td>
			<td>566460</td>
			<td></td>
		</tr>
		<tr>
			<td>Nunc MicroWell 96-Well, Nunclon Delta-Treated, Flat-Bottom Microplate</td>
			<td>Thermo Scientific</td>
			<td>136101</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS (Phosphate Buffered Saline), 1x [-] calcium, magnesium, pH 7.4</td>
			<td>Corning</td>
			<td>21-040-CM</td>
			<td></td>
		</tr>
		<tr>
			<td>Prolong Glass Antifade Mountant with NucBlue</td>
			<td>Fisher Scientific</td>
			<td>P36983</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Anti-Villin antibody [SP145]</td>
			<td>Abcam</td>
			<td>ab130751</td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant Human TNF-&#945; protein 100 &#181;g</td>
			<td>Bio-Techne</td>
			<td>210-TA-100/CF</td>
			<td></td>
		</tr>
		<tr>
			<td>SpectraMax iD3 Multi-Mode Microplate Reader</td>
			<td>Molecular Devices</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Thermo Forma Series II Water-Jacketed Tri-Gas Incubator, 184L</td>
			<td>Fisher Scientific</td>
			<td>3140</td>
			<td></td>
		</tr>
		<tr>
			<td>TO-PRO-3 Iodide (642/661)</td>
			<td>Thermo Scientific</td>
			<td>T3605</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>9002-93-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubes, 0.5 mL, flat cap</td>
			<td>Thermo Scientific</td>
			<td>AB0350</td>
			<td></td>
		</tr>
		<tr>
			<td>Tween-20</td>
			<td>Sigma-Aldrich</td>
			<td>9005-64-5</td>
			<td></td>
		</tr>
	</tbody>
</table>

in vitro apical-out enteroid model of necrotizing enterocolitis

Necrotizing enterocolitis (NEC) is a devastating disease affecting preterm infants, characterized by intestinal inflammation and necrosis. Enteroids have recently emerged as a promising system to model gastrointestinal pathologies. However, currently utilized methods for enteroid manipulation either lack access to the apical surface of the epithelium (three-dimensional [3D]) or are time-consuming and resource-intensive (two-dimensional [2D] monolayers). These methods often require additional steps, such as microinjection, for the model to become physiologically translatable. Here, we describe a physiologically relevant and inexpensive protocol for studying NEC in vitro by reversing enteroid polarity, resulting in the apical surface facing outward (apical-out). An immunofluorescent staining protocol to examine enteroid barrier integrity and junctional protein expression following exposure to tumor necrosis factor-alpha (TNF-&#945;) or lipopolysaccharide (LPS) under normoxic or hypoxic conditions is also provided. The viability of 3D apical-out enteroids exposed to normoxic or hypoxic LPS or TNF-&#945; for 24 h is also evaluated. Enteroids exposed to either LPS or TNF-&#945;, in combination with hypoxia, exhibited disruption of epithelial architecture, a loss of adherens junction protein expression, and a reduction in cell viability. This protocol describes a new apical-out NEC-in-a-dish model which presents a physiologically relevant and cost-effective platform to identify potential epithelial targets for NEC therapies and study the preterm intestinal response to therapeutics.

The use of enteroids to model intestinal inflammation, even within the context of necrotizing enterocolitis, is now common. However, most methods currently utilized either lack access to the apical surface of enteroids, negating the physiological relevance of compounds intended for eventual use as oral therapeutics, or are technically difficult and time-consuming, as with enteroid-derived monolayers. To increase the utility of current in vitro enteroid models of NEC, we reversed the polarity of enteroids, and, i...

Watch this Scientific Journal Video about In Vitro Apical-Out Enteroid Model of Necrotizing Enterocolitis at JoVE.com

In Vitro Apical-Out Enteroid Model of Necrotizing Enterocolitis

This protocol describes an apical-out necrotizing enterocolitis (NEC)-in-a-dish model utilizing small intestinal enteroids with reversed polarity, allowing access to the apical surface. We provide an immunofluorescent staining protocol to detect NEC-related epithelial disruption and a method to determine the viability of apical-out enteroids subjected to the NEC-in-a-dish protocol.

In Vitro Apical-Out Enteroid Model of Necrotizing Enterocolitis

Necrotizing enterocolitis (NEC) is a devastating disease affecting preterm infants, characterized by intestinal inflammation and necrosis. Enteroids have ...

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Research

JoVE Journal

Medicine

2.1K Views.  University of Oklahoma Health Sciences Center and Center for Pregnancy and Newborn Health. This protocol describes an apical-out necrotizing enterocolitis (NEC)-in-a-dish model utilizing small intestinal enteroids with reversed polarity, allowing access to the apical surface. We provide an immunofluorescent staining protocol to detect NEC-related epithelial disruption and a method to determine the viability of apical-out enteroids subjected to the NEC-in-a-dish protocol.

Video: In Vitro Apical-Out Enteroid Model of Necrotizing Enterocolitis

Stem Cell Transplantation in an in vitro Simulated Ischemia/Reperfusion Model

Stem cell transplantation protocols are finding their way into clinical practice1,2,3. Getting better results, making the protocols more robust, and finding new sources for implantable cells are the focus of recent research4,5. Investigating the effectiveness of cell therapies is not an easy task and new tools are needed to investigate the mechanisms involved in the treatment process6. We designed an experimental protocol of ischemia/reperfusion in order to allow the observation of cellular connections and even subcellular mechanisms during ischemia/reperfusion injury and after stem cell transplantation and to evaluate the efficacy of cell therapy. H9c2 cardiomyoblast cells were placed onto cell culture plates7,8. Ischemia was simulated with 150 minutes in a glucose free medium with oxygen level below 0.5%. Then, normal media and oxygen levels were reintroduced to simulate reperfusion. After oxygen glucose deprivation, the damaged cells were treated with transplantation of labeled human bone marrow derived mesenchymal stem cells by adding them to the culture. Mesenchymal stem cells are preferred in clinical trials because they are easily accessible with minimal invasive surgery, easily expandable and autologous. After 24 hours of co-cultivation, cells were stained with calcein and ethidium-homodimer to differentiate between live and dead cells. This setup allowed us to investigate the intercellular connections using confocal fluorescent microscopy and to quantify the survival rate of postischemic cells by flow cytometry. Confocal microscopy showed the interactions of the two cell populations such as cell fusion and formation of intercellular nanotubes. Flow cytometry analysis revealed 3 clusters of damaged cells which can be plotted on a graph and analyzed statistically. These populations can be investigated separately and conclusions can be drawn on these data on the effectiveness of the simulated therapeutical approach.

Stem Cell Transplantation in an in vitro Simulated Ischemia/Reperfusion Model

We demonstrate how to set up an in vitro ischemia/reperfusion model and how to evaluate the effect of stem cell therapy on postischemic cardiac cells.

Stem cell transplantation protocols are finding their way into clinical practice1,2,3. Getting better results, making the protocols more robust, and ...

Assessing Teratogenic Changes in a Zebrafish Model of Fetal Alcohol Exposure

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, including mental retardation due to disordered and damaged brain development. Fetal alcohol spectrum disorder (FASD) is a term used to cover a continuum of birth defects that occur due to maternal alcohol consumption, and occurs in approximately 4% of children born in the United States. With 50% of child-bearing age women reporting consumption of alcohol, and half of all pregnancies being unplanned, unintentional exposure is a continuing issue2. In order to best understand the damage produced by ethanol, plus produce a model with which to test potential interventions, we developed a model of developmental ethanol exposure using the zebrafish embryo. Zebrafish are ideal for this kind of teratogen study3-8. Each pair lays hundreds of eggs, which can then be collected without harming the adult fish. The zebrafish embryo is transparent and can be readily imaged with any number of stains. Analysis of these embryos after exposure to ethanol at different doses and times of duration and application shows that the gross developmental defects produced by ethanol are consistent with the human birth defect. Described here are the basic techniques used to study and manipulate the zebrafish FAS model.

In order to understand the molecular mechanisms of the ethanol-induced developmental damage, we have developed a zebrafish model of ethanol exposure and are exploring the physical, cellular, and genetic alterations that occur after ethanol exposure1. We then seek to find potential interventions and rapidly test them in this animal model.

Fetal alcohol syndrome (FAS) is a severe manifestation of embryonic exposure to ethanol. It presents with characteristic defects to the face and organs, ...

Stretch in Brain Microvascular Endothelial Cells (cEND) as an In Vitro Traumatic Brain Injury Model of the Blood Brain Barrier

Due to the high mortality incident brought about by traumatic brain injury (TBI), methods that would enable one to better understand the underlying mechanisms involved in it are useful for treatment. There are both in vivo and in vitro methods available for this purpose. In vivo models can mimic actual head injury as it occurs during TBI. However, in vivo techniques may not be exploited for studies at the cell physiology level. Hence, in vitro methods are more advantageous for this purpose since they provide easier access to the cells and the extracellular environment for manipulation.
Our protocol presents an in vitro model of TBI using stretch injury in brain microvascular endothelial cells. It utilizes pressure applied to the cells cultured in flexible-bottomed wells. The pressure applied may easily be controlled and can produce injury that ranges from low to severe. The murine brain microvascular endothelial cells (cEND) generated in our laboratory is a well-suited model for the blood brain barrier (BBB) thus providing an advantage to other systems that employ a similar technique. In addition, due to the simplicity of the method, experimental set-ups are easily duplicated. Thus, this model can be used in studying the cellular and molecular mechanisms involved in TBI at the BBB.

Stretch in Brain Microvascular Endothelial Cells cEND as an In Vitro Traumatic Brain Injury Model of the Blood Brain Barrier

In vitro traumatic brain injury models are being developed to reproduce in vivo brain deformation. Stretch-induced injury has been employed for astrocytes, neurons, glial cells, aortic, and brain endothelial cells. However, our system uses a blood brain barrier (BBB) model that possesses properties constituting a legitimate model of the BBB to establish an in vitro&#160;TBI model.

Due to the high mortality incident brought about by traumatic brain injury (TBI), methods that would enable one to better understand the underlying ...

Excitotoxic Stimulation of Brain Microslices as an In vitro Model of Stroke

Examining molecular mechanisms involved in neuropathological conditions, such as ischemic stroke, can be difficult when using whole animal systems. As such, primary or &#39;neuronal-like&#39; cell culture systems are commonly utilized. While&#160;these systems are relatively easy to work with, and are useful model systems in which various functional outcomes (such as cell death) can be readily quantified, the examined outcomes and pathways in cultured immature neurons (such as excitotoxicity-mediated cell death pathways) are not necessarily the same as those observed in mature brain, or in intact tissue. Therefore, there is the need to develop models in which cellular mechanisms in mature neural tissue can be examined. We have developed an in vitro technique that can be used to investigate a variety of molecular pathways in intact nervous tissue. The technique described herein utilizes rat cortical tissue, but this technique can be adapted to use tissue from a variety of species (such as mouse, rabbit, guinea pig, and chicken) or brain regions (for example, hippocampus, striatum, etc.). Additionally, a variety of stimulations/treatments can be used (for example, excitotoxic, administration of inhibitors, etc.). In conclusion, the brain slice model described herein can be used to examine a variety of molecular mechanisms involved in excitotoxicity-mediated brain injury.

Excitotoxic Stimulation of Brain Microslices as an In vitro Model of Stroke

We have developed a brain slice model which can be used to examine molecular mechanisms involved in excitotoxicity-mediated brain injury.&#160;This technique generates viable mature brain tissue and reduces animal numbers required for experimentation, whilst keeping the neuronal circuitry, cellular interactions, and postsynaptic compartments partly intact.

Examining molecular mechanisms involved in neuropathological conditions, such as ischemic stroke, can be difficult when using whole animal systems. As ...

Assessing Specificity of Anticancer Drugs In Vitro

A procedure for assessing specificity of anticancer drugs in vitro using cultures containing both tumor and non-tumor cells is demonstrated. The key element is the quantitative determination of a tumor-specific genetic alteration in relation to a universal sequence using a dual-probe digital PCR assay and the subsequent calculation of the proportion of tumor cells. The assay is carried out on a culture containing tumor cells of an established line and spiked-in non-tumor cells. The mixed culture is treated with a test drug at various concentrations. After the treatment, DNA is prepared directly from the survived adhesive cells in wells of 96-well plates using a simple and inexpensive method, and subjected to a dual-probe digital PCR assay for measuring a tumor-specific genetic alteration and a reference universal sequence. In the present demonstration, a heterozygous deletion of the NF1 gene is used as the tumor-specific genetic alteration and a RPP30 gene as the reference gene. Using the ratio NF1/RPP30, the proportion of tumor cells was calculated. Since the dose-dependent change of the proportion of tumor cells provides an in vitro indication for specificity of the drug, this genetic and cell-based in vitro assay will likely have application potential in drug discovery. Furthermore, for personalized cancer-care, this genetic- and cell-based tool may contribute to optimizing adjuvant chemotherapy by means of testing efficacy and specificity of candidate drugs using primary cultures of individual tumors.

Assessing Specificity of Anticancer Drugs In Vitro

The goal of this protocol is to assess specificity of anticancer drugs in vitro using mixed cultures containing both tumor and non-tumor cells.

A procedure for assessing specificity of anticancer drugs in vitro using cultures containing both tumor and non-tumor cells is demonstrated. The key ...

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a condition that typically involves damage to or loss of the delicate mechanosensory structures of the inner ear. Sophisticated in vitro and ex vivo assays have emerged in recent years, enabling the screening of an increasing number of potentially therapeutic compounds while minimizing resources and accelerating efforts to develop cures for SNHL. Though homogenous cultures of certain cell types continue to play an important role in current research, many scientists now rely on more complex organotypic cultures of murine inner ears, also known as cochlear explants. The preservation of organized cellular structures within the inner ear facilitates the in situ evaluation of various components of the cochlear infrastructure, including inner and outer hair cells, spiral ganglion neurons, neurites, and supporting cells. Here we present the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The careful preparation of these explants facilitates the identification of mechanisms that contribute to SNHL and constitutes a valuable tool for the hearing research community.

Neonatal Murine Cochlear Explant Technique as an In Vitro Screening Tool in Hearing Research

The goal of this protocol is to demonstrate the preparation, culture, treatment, and immunostaining of neonatal murine cochlear explants. The technique can be utilized as an in vitro screening tool in hearing research.

While there have been remarkable advances in hearing research over the past few decades, there is still no cure for Sensorineural Hearing Loss (SNHL), a ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

In Vitro Model of Coronary Angiogenesis

Here, we describe an in vitro culture assay to study coronary angiogenesis. Coronary vessels feed the heart muscle and are of clinical importance. Defects in these vessels represent severe health risks such as in atherosclerosis, which can lead to myocardial infarctions and heart failures in patients. Consequently, coronary artery disease is one of the leading causes of death worldwide. Despite its clinical importance, relatively little progress has been made on how to regenerate damaged coronary arteries. Nevertheless, recent progress has been made in understanding the cellular origin and differentiation pathways of coronary vessel development. The advent of tools and technologies that allow researchers to fluorescently label progenitor cells, follow their fate, and visualize progenies in vivo have been instrumental in understanding coronary vessel development. In vivo studies are valuable, but have limitations in terms of speed, accessibility, and flexibility in experimental design. Alternatively, accurate in vitro models of coronary angiogenesis can circumvent these limitations and allow researchers to interrogate important biological questions with speed and flexibility. The lack of appropriate in vitro model systems may have hindered the progress in understanding the cellular and molecular mechanisms of coronary vessel growth. Here, we describe an in vitro culture system to grow coronary vessels from the sinus venosus (SV) and endocardium (Endo), the two progenitor tissues from which many of the coronary vessels arise. We also confirmed that the cultures accurately recapitulate some of the known in vivo mechanisms. For instance, we show that the angiogenic sprouts in culture from SV downregulate COUP-TFII expression similar to what is observed in vivo. In addition, we show that VEGF-A, a well-known angiogenic factor in vivo, robustly stimulates angiogenesis from both the SV and Endo cultures. Collectively, we have devised an accurate in vitro culture model to study coronary angiogenesis.

In vitro models of coronary angiogenesis can be utilized for the discovery of the cellular and molecular mechanisms of coronary angiogenesis. In vitro explant cultures of sinus venosus and endocardium tissues show robust growth in response to VEGF-A and display a similar pattern of COUP-TFII expression as in vivo.

Here, we describe an in vitro culture assay to study coronary angiogenesis. Coronary vessels feed the heart muscle and are of clinical importance. Defects ...

An In Vitro Hemodynamic Loop Model to Investigate the Hemocytocompatibility and Host Cell Activation of Vascular Medical Devices

In this study, the hemocompatibility of tubes with an inner diameter of 5 mm made of polyvinyl chloride (PVC) and coated with different bioactive conjugates was compared to uncoated PVC tubes, latex tubes, and a stent for intravascular application that was placed inside the PVC tubes. Evaluation of hemocompatibility was done using an in vitro hemodynamic loop model that is recommended by the ISO standard 10993-4. The tubes were cut into segments of identical length and closed to form loops avoiding any gap at the splice, then filled with human blood and rotated in a water bath at 37 &#176;C for 3 hours. Thereafter, the blood inside the tubes was collected for the analysis of whole blood cell count, hemolysis (free plasma hemoglobin), complement system (sC5b-9), coagulation system (fibrinopeptide A), and leukocyte activation (polymorphonuclear elastase, tumor necrosis factor and interleukin-6). Host cell activation was determined for platelet activation, leukocyte integrin status and monocyte platelet aggregates using flow cytometry. The effect of inaccurate loop closure was examined with x-ray microtomography and scanning electron microscopy, that showed thrombus formation at the splice. Latex tubes showed the strongest activation of both plasma and cellular components of the blood, indicating a poor hemocompatibility, followed by the stent group and uncoated PVC tubes. The coated PVC tubes did not show a significant decrease in platelet activation status, but showed an increased in complement and coagulation cascade compared to uncoated PVC tubes. The loop model itself did not lead to the activation of cells or soluble factors, and the hemolysis level was low. Therefore, the presented in vitro hemodynamic loop model avoids excessive activation of blood components by mechanical forces and serves as a method to investigate in vitro interactions between donor blood and vascular medical devices.

Presented here is a protocol for a standardized in vitro hemodynamic loop model. This model allows to test the hemocompatibility of perfusion tubes or vascular stents to be in accordance with ISO (International Organization for Standardization) standard 10993-4.

In this study, the hemocompatibility of tubes with an inner diameter of 5 mm made of polyvinyl chloride (PVC) and coated with different bioactive ...

In vitro Assessment of Myocardial Protection following Hypothermia-Preconditioning in a Human Cardiac Myocytes Model

Ischemia/reperfusion-derived myocardial dysfunction is a common clinical scenario in patients after cardiac surgery. In particular, the sensitivity of cardiomyocytes to ischemic injury is higher than that of other cell populations. At present, hypothermia affords considerable protection against an expected ischemic insult. However, investigations into complex hypothermia-induced molecular changes remain limited. Therefore, it is essential to identify a culture condition similar to in vivo conditions that can induce damage similar to that observed in the clinical condition in a reproducible manner. To mimic ischemia-like conditions in vitro, the cells in these models were treated by oxygen/glucose deprivation (OGD). In addition, we applied a standard time-temperature protocol used during cardiac surgery. Furthermore, we propose an approach to use a simple but comprehensive method for the quantitative analysis of myocardial injury. Apoptosis and expression levels of apoptosis-associated proteins were assessed by flow cytometry and using an ELISA kit. In this model, we tested a hypothesis regarding the effects of different temperature conditions on cardiomyocyte apoptosis in vitro. The reliability of this model depends on strict temperature control, controllable experimental procedures, and stable experimental results. Additionally, this model can be used to study the molecular mechanism of hypothermic cardioprotection, which may have important implications for the development of complementary therapies for use with hypothermia.

The distinct effects of different degrees of hypothermia on myocardial protection have not been thoroughly evaluated. The goal of the present study was to quantify the levels of cell death following different hypothermia treatments in a human cardiomyocyte-based model, laying the foundation for future in-depth molecular research.

Ischemia/reperfusion-derived myocardial dysfunction is a common clinical scenario in patients after cardiac surgery. In particular, the sensitivity of ...