The authors would like to acknowledge Dr. Ellen Lauchnor, Dr. Phil Stewart, and Bengisu Kilic for their previous work and assistance with the O2 microsensors; Andy Sebrell for training in organoid culture and micromanipulation; Lexi Burcham for assistance in organoid culture, media preparation, data recording, and organization; and Dr. Susy Kohout for general advice in electrophysiology. We would like to thank Dr. Heidi Smith for her assistance with imaging and acknowledge the Center for Biofilm Engineering Bioimaging Facility at Montana State University, which is supported by funding from the National Science Foundation MRI Program (2018562), the M.J. Murdock Charitable Trust (202016116), the US Department of Defense (77369LSRIP &#38; W911NF1910288), and by the Montana Nanotechnology Facility (an NNCI member supported by NSF Grant ECCS-2025391).
Special thanks to the entire Unisense team who made this work possible, especially Dr. Andrew Cerskus, Dr. Laura Woods, Dr. Lars Larsen, Dr. Tage Dalsgaard, Dr. Line Daugaard, Dr. Karen Maegaard, and Mette Gammelgaard. Funding for our study was provided by the National Institutes of Health grants R01 GM13140801 (D.B., R.B.) and UL1 TR002319 (K.N.L), and a Research Expansion Award from the Montana State University Office for Research and Economic Development (D.B.). Figure 1A was created with BioRender.

This protocol requires 3D organoids of at least 200 &#181;m in diameter that have a distinct lumen and that are embedded in an artificial extracellular matrix (ECM, e.g., Matrigel). Human gastric tissues for organoid derivation were obtained with approval from the Institutional Review Board of Montana State University and informed consent from patients undergoing upper endoscopy at Bozeman Health (protocol # 2023-48-FCR, to D.B.) or as exempt whole stomach or sleeve gastrectomy specimens from the National Disease Research Interchange (protocol #DB062615-EX). Information about the organoid lines and passage numbers used for this study is provided in Table 1, and the media composition is listed in Table 2. Refer to previously published protocols for the generation and maintenance of gastrointestinal organoid lines6,26,27.
1. Preparation of human gastric organoids for pH profiling
<ol>
	<li>Initiate and maintain gastric organoid cultures using standard protocols. Maintain organoids on 24-well plates in 500 &#181;L of expansion media per well (Table 2). Passage established organoid lines every 5-7 days, transferring to a 35 mm glass-bottom dish in preparation for pH profiling experiments. 
	NOTE: Organoid cultures for our experiments are derived from gastric gland preparations, which are obtained from adult tissue as described above. We use a collagenase tissue digestion method to isolate these glands before they are suspended in ECM, as previously described25,28,29.</li>
	<li>From actively growing organoid cultures at passages 1-15, select wells that contain at least 100 organoids (approximately 2 million cells) with diameters between 200 and 700 &#181;m for transfer and expansion on 3.5 mm Petri dishes.</li>
	<li>While preparing organoids for plating (steps 1.4-1.8), allow extracellular matrix (ECM) aliquot(s) to thaw on ice for at least 45 min. Maintain ECM on ice throughout this protocol to prevent gelation. Prewarm cell culture plates by placing them in a 37 &#176;C, 5% CO2 incubator.</li>
	<li>Remove the media from the wells and harvest the gastric organoid cultures by pipetting ice-cold PBS onto each ECM droplet and scratching the gel with the tip of a P1000 pipette. Pipette the PBS with the ECM fragments containing organoids into a 15 mL conical tube.</li>
	<li>Centrifuge the tube at 200 &#215; g at 4 &#176;C for 5 min. ECM fragments containing the organoids will be visible as a layer at the bottom of the tube. Carefully aspirate the supernatant and pipette 350 &#181;L of 0.25% trypsin-EDTA into each tube, mixing gently by pipetting up and down. Incubate the tubes in a 37 &#176;C water bath for 2-5 min.</li>
	<li>Following incubation with trypsin-EDTA, add 600 &#181;L of ice-cold DMEM with penicillin/streptomycin to each tube and vigorously pipette up and down at least 40x. Centrifuge at 200 &#215; g at 4 &#176;C for 5 min. Aspirate the supernatant. To set up cultures for pH measurements, resuspend the cell pellet in ice-cold, liquid ECM at a 1:4 v/v ratio of organoid pellet to ECM for plating.</li>
	<li>For each sample, plate 40 &#181;L of liquid ECM containing organoids/organoid fragments in a thin horizontal line along the diameter of a 35 mm glass-bottom dish. 
	NOTE: Dispensing the gel onto the plate in a line instead of a round droplet enables easier access to individual organoids by thinning out the culture.</li>
	<li>Allow the gel to polymerize for 15-30 min; then, carefully add 2 mL of organoid expansion media to the plate by pipetting along the edge of the well to avoid disturbing the gel.</li>
	<li>Replace media every other day. Allow 4-8 days for a minimum of 10 organoids to grow to a minimum diameter of 400 &#181;m.</li>
	<li>To proceed with a pH profiling experiment, ensure that each culture contains at least 10 organoids that meet the following criteria: &#62;200 &#181;m diameter, healthy appearance (little to no dark material visible within organoids observed with a brightfield microscope), and not overcrowded by other organoids.</li>
</ol>
2. Unpacking and calibration of microelectrodes
NOTE: To enable microscale measurements, a separate reference electrode is used in addition to the pH sensor microelectrode rather than using an integrated (and hence bulkier) design. Both pH microelectrode and reference electrode must be stored wet. Do not allow exposure to air for more than 10 min at a time. Determine the appropriate tip size for the application. Here, we used a potentiometric pH microelectrode with a beveled tip diameter of 25 &#181;m.
<ol>
	<li>With the microelectrode still in its protection tube, visually inspect the tip for any damage.</li>
	<li>Connect the reference electrode to the pH electrode cable via the connector. Then, connect the microelectrode to the amplifier and the amplifier to a grounded PC with the software via a USB cable (Figure 1A).</li>
	<li>Fill two 50 mL conical tubes 2/3 with 70% ethanol and deionized H2O (diH2O). Place the microelectrode and the reference electrode in the diH2O tube, ensuring that both tips appear submerged at least 1 cm in the liquid. 
	NOTE: Tall beakers can also be used for this and similar steps.</li>
	<li>Allow the electrodes to equilibrate for ~10 min. 
	NOTE: The procedure can also be paused at this step and resumed later as the electrodes may be stored in diH2O.</li>
	<li>While the electrodes are equilibrating, open the software (red icon) on the computer workstation. Under the Settings tab, ensure that the box&#160;next to the microelectrode is checked,&#160;indicating that it is properly connected and&#160;recognized by the software. Click Start&#160;Experiment on the top left of the window and enter your desired file name and destination. In the Sensor calibration &#38;&#160;experiment&#160;settings window, click Clear all points to&#160;prepare the software for a new calibration.</li>
	<li>Fill two 50 mL conical tubes 2/3 with pH 4.01 and pH 9.21 calibration buffers. Gently blot the protective tubing and the reference electrode dry with a delicate laboratory wipe.</li>
	<li>Starting with the electrodes in the pH 4.01 buffer, enter 4.01 as the known pH value. Once the mV reading has stabilized, select add point. Confirm that the signal is stable at ~380 mV. After the point has been added, place both electrodes back in diH2O to rinse, then blot dry again.</li>
	<li>Repeat the previous step with the 9.21 buffer and click add point when the signal is stable (~83 mV). Check that the microelectrodes respond linearly between pH 4.01 and 9.21; therefore, only a two-point calibration curve is necessary. Ensure that the resulting line between these points has a negative slope of 50-70 mV/pH-unit. Click Save &#38;&#160;use calibration. 
	NOTE: You are now ready to begin recording measurements30.</li>
</ol>
3. pH profiling of human gastric organoids
NOTE: The following protocol is described for a right-handed user. CAUTION: Disable all power-saving options on your PC as ongoing measurements will be disrupted if the PC enters sleep mode.
<ol>
	<li>Assemble a ring stand with a clamp on the left-hand side of a stereomicroscope to hold the reference electrode. Position a micromanipulator attached to a heavy lab stand on the right side of the microscope (Figure 1A).</li>
	<li>Carefully remove the microelectrode from its protection case by laying it flat on the bench and pulling the case off in a swift, quick motion. 
	CAUTION: The microelectrodes are incredibly fragile and care should be taken to ensure that the tip does not come into contact with stiff, solid material. For best results, perform measurements on a sturdy benchtop free of equipment that could cause vibration of the bench or unwanted movement of the electrodes.</li>
	<li>Mount the microelectrode on a micromanipulator and arrange the stereoscope and micromanipulator in such a way that the microelectrode may advance toward the culture dish without hitting the objective or other parts of the microscope.</li>
	<li>Position the culture dish containing the organoids to have adequate visualization of the first organoid that is to be profiled (Figure 1B).</li>
	<li>Lightly secure the reference electrode to the clamp on the ring stand to the left of the stereoscope. Position the reference electrode so that it rests in the medium surrounding the ECM. Take care to ensure that it will not disrupt the organoids as the stage is moved around between measurements.</li>
	<li>Looking directly at the culture plate (not into the microscope), advance the tip of the microelectrode until it is sufficiently submerged in the media. Under the Data logger tab in the software,&#160;click the Start button (single triangle pointing to&#160;the right) to begin recording. Ensure that the&#160;Calibrated checkbox is selected on the right&#160;side of the screen.&#160; 
	NOTE: Allow ~10 s for each reading to stabilize before advancing to the next region.</li>
	<li>Before entering the ECM, ensure that the electrode tip is visible under the microscope and that it is positioned to advance linearly toward the first organoid of interest. Slowly advance the electrode into the gel, without making contact with any organoids. Record at least three pH readings and calculate the average.</li>
	<li>Position the microelectrode so that it is prepared to advance toward the first organoid of interest along the axis perpendicular to the surface. Allow the electrode to make a minor indentation on the epithelial surface without penetrating (Figure 1C). 
	NOTE: This step will also provide insight into whether the organoid will remain in place or whether it may move about more freely in the ECM.</li>
	<li>With one swift motion, advance the microelectrode into the organoid. If the organoid moves around the microelectrode or moves away, attempt a slightly different angle, or back it up against another organoid or the plastic rim surrounding the coverglass bottom of the dish. Measure the luminal pH (three individual times) of 10 different organoids for each experimental condition.&#160;When finished recording measurements, click the square Stop button. 
	CAUTION: Estimate the diameter of the organoid to decide how far to advance the electrode into the organoid lumen, taking care not to pierce through it. For increased accuracy, measure the diameter with your microscope&#39;s camera software, if available. NOTE: If the tip of the microelectrode becomes noticeably coated with debris following one or multiple measurements, wash the microelectrode in cell dissociation solution, PBS, EtOH, and then diH2O before proceeding.</li>
</ol>
4. Motorized profiling (optional)
NOTE: This option requires a micromanipulator that is mounted on a mechanical motor stage, which is ultimately controlled by computer software via a motor controller31.
<ol>
	<li>Open the Profiling software (brown icon) and start a new experiment under the Profiling tab.</li>
	<li>The settings for the z-axis will automatically recognize Motor when a motor apparatus is properly connected. 
	NOTE: Movement in the x and y directions is still controlled manually with this setup.</li>
	<li>Locate the Profile interaction tab30. Before hitting start, ensure that it is possible to quickly transition to looking at the microscope eyepiece to ensure organoid entry at the desired distance. Go to Profile settings and do the following:
	<ol>
		<li>Indicate that the Start distance is 0 &#181;m. 
		CAUTION: If the epithelial shell is particularly tough on a given organoid, the organoid may be pushed away or deformed instead of penetrated. If pushing is the case, continue advancing the electrode but note the point at which entry occurs, as the software cannot record this automatically.</li>
		<li>Judging the size of the organoid being profiled, indicate an End distance that is no more than &#190; way through an organoid&#39;s estimated diameter. 
		NOTE: This particular step is designed to prevent damage to the electrode tip.</li>
		<li>Indicate the desired step size&#8212;100 &#181;m as shown in Figure 2E. Ensure that the minimum step size matches the size of the electrode tip (e.g., 25 &#181;m for a 25 &#181;m pH-25 electrode tip).</li>
		<li>Indicate a safe position to which the motor will return the electrode tip between profiles. 
		NOTE: This should be a comfortable height above the sample (outside of the organoid) where the sensor can be safely moved sideways in the x- and y-directions with the manual micromanipulator. Leave the Sensor angle at its default setting.</li>
		<li>To allow the electrode to equilibrate, indicate at least 3 s under Wait before measure(s).</li>
		<li>Set the Measure period(s) to at least 1 s. The measurements over this period will be averaged. 
		NOTE: If performing measurements in an environment high in electrical noise, it may be helpful to indicate a longer period.</li>
		<li>Set the Delay between(s) to at least 1 s.</li>
		<li>Set the preferred number of Replicates to be measured at each depth.</li>
		<li>Begin recording measurements by hitting the Start button.</li>
	</ol>
	</li>
</ol>
5. Cleaning of electrodes
<ol>
	<li>Place the electrode to be cleaned back in its protection tube.</li>
	<li>Flush the electrodes with diH2O after measurements.</li>
	<li>Flush the electrodes with 70% ethanol for a couple of minutes.</li>
	<li>Rinse the electrodes with pH 4.01 buffer.</li>
	<li>Rinse&#160;the electrodes once more with diH2O before&#160;proceeding with measurements.</li>
</ol>
6. Storage of electrodes
NOTE: Both electrodes are to be stored at room temperature in a low-activity location, safe from accidental damage.
<ol>
	<li>After using the pH microelectrode, carefully slide it horizontally back into its protection tube (sliding along the lab bench for horizontal support).</li>
	<li>Holding the microelectrode upright (tip pointed upward), gently fill the protection tube with sterile water. Plug the protection tube with the stopper the electrode came with. Ensure that any holes in the protection tube are sealed with electrical tape. Store in a shatter-proof box until the next use. 
	NOTE: It is recommended to use the original box that the electrodes came in as it will contain protective inserts designed to keep electrodes in place when stored.</li>
	<li>Store the reference electrode in a beaker or graduated cylinder filled with a 3M KCl solution. Cover the beaker/graduated cylinder with parafilm to prevent evaporation of the solution. 
	NOTE: If using a graduated cylinder, it is recommended to secure it to the lab bench with tape to prevent accidental movement.</li>
</ol>
7. Methyl red injection (optional)
NOTE: Methyl red is a colorimetric indicator dye that can be used to validate the microelectrode measurements.
<ol>
	<li>Backfill a 2 &#181;L glass capillary with sterile mineral oil, load it onto a micromanipulator-controlled nL autoinjector, and subsequently fill it with a solution containing 0.02% methyl red and 150 mM HCl. 
	NOTE: The solution should appear red/pink while in the capillary due to the acidity of the HCl.</li>
	<li>Inject organoids of at least 300 &#181;m in diameter with 9.2 &#181;L of the solution25.</li>
	<li>Capture images or video using a camera adapted to the stereomicroscope (Figure 2G).</li>
</ol>

Precise Luminal pH Measurement in 3D Gastric Organoids with Microelectrodes

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D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+synthetic+hydrogel,+vitrogel((r))+organoid-3,+improves+immune+cell-epithelial+interactions+in+a+tissue+chip+co-culture+model+of+human+gastric+organoids+and+dendritic+cells.">A synthetic hydrogel, vitrogel((r)) organoid-3, improves immune cell-epithelial interactions in a tissue chip co-culture model of human gastric organoids and dendritic cells.</a> Front Pharmacol. 12, 707891 (2021).</li><li>Sebrell, T. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+analysis+of+human+gastric+epithelial+spheroids+reveals+spontaneous+rupture,+rotation+and+fusion+events.">Live imaging analysis of human gastric epithelial spheroids reveals spontaneous rupture, rotation and fusion events.</a> Cell Tissue Res. 371 (2), 293-307 (2018).</li><li>Sensortrace suite user manual. 3.3. Unisense Available from: <a target="_blank" href="https://unisense.com/wp-content/uploads/2021/10/SensorTrace-Suite-Manual.pdf">https://unisense.com/wp-content/uploads/2021/10/SensorTrace-Suite-Manual.pdf</a> (2023)</li><li>Microprofiling system user manual. Unisense Available from: <a target="_blank" href="https://unisense.com/wp-content/uploads/2021/09/2023.11-MicroProfiling-System-2.pdf">https://unisense.com/wp-content/uploads/2021/09/2023.11-MicroProfiling-System-2.pdf</a> (2023)</li><li>Wolffling, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Egf+and+bmps+govern+differentiation+and+patterning+in+human+gastric+glands.">Egf and bmps govern differentiation and patterning in human gastric glands.</a> Gastroenterology. 161 (2), 623-636 (2021).</li><li>Boccellato, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polarised+epithelial+monolayers+of+the+gastric+mucosa+reveal+insights+into+mucosal+homeostasis+and+defence+against+infection.">Polarised epithelial monolayers of the gastric mucosa reveal insights into mucosal homeostasis and defence against infection.</a> Gut. 68 (3), 400-413 (2019).</li><li>Mccracken, K. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modelling+human+development+and+disease+in+pluripotent+stem-cell-derived+gastric+organoids.">Modelling human development and disease in pluripotent stem-cell-derived gastric organoids.</a> Nature. 516 (7531), 400-404 (2014).</li><li>Schumacher, M. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+murine-derived+fundic+organoids+in+studies+of+gastric+physiology.">The use of murine-derived fundic organoids in studies of gastric physiology.</a> J Physiol. 593 (8), 1809-1827 (2015).</li><li>. Unisense Available from: <a target="_blank" href="https://unisense.com/products/ph-microelectrode/">https://unisense.com/products/ph-microelectrode/</a> (2024)</li><li>Mccracken, K. W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Wnt/beta-catenin+promotes+gastric+fundus+specification+in+mice+and+humans.">Wnt/beta-catenin promotes gastric fundus specification in mice and humans.</a> Nature. 541 (7636), 182-187 (2017).</li><li>Schreiber, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+situ+measurement+of+ph+in+the+secreting+canaliculus+of+the+gastric+parietal+cell+and+adjacent+structures.">In situ measurement of ph in the secreting canaliculus of the gastric parietal cell and adjacent structures.</a> Cell Tissue Res. 329 (2), 313-320 (2007).</li><li>Xu, H., Li, J., Chen, H., Wang, C., Ghishan, F. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nhe8+plays+important+roles+in+gastric+mucosal+protection.">Nhe8 plays important roles in gastric mucosal protection.</a> Am J Physiol Gastrointest Liver Physiol. 304 (3), G257-G261 (2013).</li><li>Gawenis, L. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impaired+gastric+acid+secretion+in+mice+with+a+targeted+disruption+of+the+nhe4+na+/h++exchanger.">Impaired gastric acid secretion in mice with a targeted disruption of the nhe4 na+/h+ exchanger.</a> J Biol Chem. 280 (13), 12781-12789 (2005).</li><li>Lewis, O. L., Keener, J. P., Fogelson, A. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+physics-based+model+for+maintenance+of+the+ph+gradient+in+the+gastric+mucus+layer.">A physics-based model for maintenance of the ph gradient in the gastric mucus layer.</a> Am J Physiol-Gastrointest Liver Physiol. 313 (6), G599-G612 (2017).</li><li>Okkelman, I. A., Neto, N., Papkovsky, D. B., Monaghan, M. G., Dmitriev, R. I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+deeper+understanding+of+intestinal+organoid+metabolism+revealed+by+combining+fluorescence+lifetime+imaging+microscopy+(flim)+and+extracellular+flux+analyses.">A deeper understanding of intestinal organoid metabolism revealed by combining fluorescence lifetime imaging microscopy (flim) and extracellular flux analyses.</a> Redox Biol. 30, 101420 (2020).</li><li>Pleguezuelos-Manzano, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Establishment+and+culture+of+human+intestinal+organoids+derived+from+adult+stem+cells.">Establishment and culture of human intestinal organoids derived from adult stem cells.</a> Curr Protoc Immunol. 130 (1), e106 (2020).</li><li>Guimera, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+minimally+invasive+microsensor+specially+designed+for+simultaneous+dissolved+oxygen+and+ph+biofilm+profiling.">A minimally invasive microsensor specially designed for simultaneous dissolved oxygen and ph biofilm profiling.</a> Sensors (Basel). 19 (21), 4747 (2019).</li></ol>

The authors have no conflicts of interest to disclose.

Limited access to the luminal space of organoids has severely restricted our understanding of the physiological dynamics of this microenvironment. A reliable tool for functional analyses of luminal physiology will expand our ability to leverage organoids as in vitro models for physiology, pharmacology, and disease research. Organoids are highly tunable, physiologically relevant models with the added potential to replicate genetic variability within the human population. Existing methods for pH measurement inside...

Organoids-miniature multicellular structures derived from stem cells-have revolutionized our ability to study human physiology and are starting to replace animal models, even in regulatory settings1. Since the initial description of intestinal organoids by Sato et al. in 2009, organoid technology has become immensely popular2. A large number of studies have characterized the cellular composition and function of organoid models in great detail3,4,5,6. However, the luminal space of these 3D multicellular structures remains largely undefined7,8. The lumen is the central cavity of organoids derived from mucosal tissues that is surrounded by the apical portions of polarized epithelial cells. Since cellular secretion and absorption predominantly occur at the apical epithelial surface, the luminal microenvironment of organoids is controlled by these important physiological processes. Currently used organoid models demonstrate variations in cell signaling patterns, overall stemness, metabolite concentration gradients, and environmental conditions9. Understanding organoid luminal physiology is therefore necessary for the accurate modeling of organ function and pathology. Unfortunately, the relative inaccessibility of the lumen significantly hinders functional analyses of luminal physiology in 3D organoids10.
The ability to examine pH profiles is especially important in the stomach, which is notorious for having the steepest proton gradient in the body, ranging from approximately 1-3 in the lumen, to near neutral at the epithelium11,12,13.There remains a significant gap in our understanding of the microscale maintenance of the gastric pH gradient, and the relevance of organoid models in recapitulating this dynamic milieu across the gastric mucus layer. Traditional approaches for the analysis of organoid pH have involved the use of pH-sensitive dyes, which can be fluorescent or colorimetric indicators. McCracken et al. used a luminal injection of SNARF-5F-a ratiometric pH indicator-into organoids to analyze a drop in luminal pH in response to histamine treatment. Such dyes can be incorporated into the culture media, allowing for real-time, non-invasive monitoring of pH. Not only do pH-sensitive dyes require complex calibration steps that contribute to poor reliability and accuracy with measurements, but such dyes also tend to operate within specific detection ranges that may not be representative of the full pH range within the microenvironment of interest14,15. It could be considered reasonable, however, to use indicator dyes for confirmatory experiments. Optical nanosensors that use fluorescent optode-based, pH-sensing approaches have also been developed; however, such sensing techniques require microscopic imaging and are also susceptible to photobleaching, phototoxicity, as well as imaging bias16,17. Additionally, Brooks et al. have 3D-printed multiwell plates containing microelectrodes on top of which organoids may be plated18. This approach, however, does not allow for measurements directly inside the organoid lumen.
Electrode-based pH measurements can achieve improved accuracy compared to other methods, as well as provide real-time pH monitoring. In addition, pH electrodes mounted on micromanipulators allow for superior spatial resolution of pH measurements as the precise location of the electrode tip can be finely controlled. This enables the highest possible flexibility and reproducibility in the analyses of organoid models. The electrodes used here are miniaturized pH microelectrodes that operate based on the diffusion of protons through selective pH glass that surrounds a thin platinum wire. The microelectrode is connected to an external Ag-AgCl reference electrode and then connected to a high-impedance millivolt meter. The electrical potential between the two electrode tips when submerged in the same solution will reflect the pH of the solution19. Such microprofiling systems have been used in the metabolic analysis of biofilms20,21, planktonic algae22, human sputum samples23, and even in mesenchymal stem cell spheroids24. Both our lab and Murphy et al. have previously used micromanipulator-controlled O2 microelectrodes to evaluate the oxygen concentrations in the luminal spaces of organoids. Murphy et al. paired this method with mathematical modeling to reveal an oxygen gradient within their spheroids. Our group was able to find reduced luminal oxygen levels in tissue-derived gastric organoids compared to the surrounding extracellular matrix25.
Here, we provide a detailed method for the manual microelectrode profiling of the luminal pH in spherical gastrointestinal tract organoids that will enable enhanced physiological understanding of their complex luminal microenvironment. We anticipate that this technique will add a new dimension to the exploration of organoid physiology through real-time, high-resolution measurements of pH levels at a microscale. Furthermore, the following protocol could be easily adapted for the analysis of O2, N2O, H2, NO, H2S, redox, and temperature in various types of organoid models. Physiological profiling serves as a valuable tool for optimizing organoid culture conditions to better mimic in vivo environments, thereby enhancing the relevance and utility of organoid models in biomedical research.

<table><tbody><tr><td>3 M KCl</td><td>Unisense</td><td></td><td></td></tr><tr><td>5 mL Wobble-not Serological Pipet, Individually Wrapped, Paper/Plastic, Bag, Sterile</td><td>CellTreat</td><td>229091B</td><td></td></tr><tr><td>10 mL Wobble-not Serological Pipet, Individually Wrapped, Paper/Plastic, Bag, Sterile</td><td>CellTreat</td><td>229092B</td><td></td></tr><tr><td>15 mL Centrifuge Tube - Foam Rack, Sterile</td><td>CellTreat</td><td>229412</td><td></td></tr><tr><td>24 Well Tissue Culture Plate, Sterile</td><td>CellTreat</td><td>229124</td><td></td></tr><tr><td>25 mL Wobble-not Serological Pipet, Individually Wrapped, Paper/Plastic, Bag, Sterile</td><td>CellTreat</td><td>229093B</td><td></td></tr><tr><td>35 mm Dish | No. 1.5 Coverslip | 20 mm Glass Diameter | Uncoated</td><td>MatTek</td><td>P35G-1.5-20-C</td><td></td></tr><tr><td>50 mL Centrifuge Tube - Foam Rack, Sterile</td><td>CellTreat</td><td>229422</td><td></td></tr><tr><td>70% Ethanol</td><td>BP82031GAL</td><td>BP82031GAL</td><td></td></tr><tr><td>70 &amp;mu;m Cell Strainer, Individually Wrapped, Sterile</td><td>CellTreat</td><td>229483&amp;nbsp;</td><td></td></tr><tr><td>1,000 &amp;micro;L Extended Length Low Retention Pipette Tips, Racked, Sterile</td><td>CellTreat</td><td>229037</td><td></td></tr><tr><td>Amphotericin B (Fungizone) Solution</td><td>HyClone Laboratories, Inc</td><td>SV30078.01</td><td></td></tr><tr><td>Biosafety Cabinet</td><td>Nuaire&amp;nbsp;</td><td>NU-425-600</td><td>Class II Type A/B3</td></tr><tr><td>Bovine Serum Albumin</td><td>Fisher Bioreagents</td><td>BP1605-100</td><td></td></tr><tr><td>Cell recovery solution</td><td>Corning</td><td>354253</td><td>Cell dissociation solution</td></tr><tr><td>DMEM/F-12 (Advanced DMEM)</td><td>Gibco</td><td>12-491-015</td><td></td></tr><tr><td>Dulbecco&#039;s Modification of Eagles Medium (DMEM)</td><td>Fisher Scientific</td><td>15017CV</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>HyClone Laboratories, Inc</td><td>SH30088</td><td></td></tr><tr><td>G418 Sulfate</td><td>Corning</td><td>30-234-CR</td><td></td></tr><tr><td>Gentamycin sulfate</td><td>IBI Scientific</td><td>IB02030</td><td></td></tr><tr><td>HEPES, Free Acid</td><td>Cytiva</td><td>SH30237.01</td><td></td></tr><tr><td>HP Pavillion 2-in-1 14&quot; Laptop Intel Core i3</td><td>HP</td><td>M03840-001</td><td></td></tr><tr><td>Hydrochloric acid</td><td>Fisher Scientific</td><td>A144C-212</td><td></td></tr><tr><td>Incubator</td><td>Fisher Scientific</td><td>11676604</td><td></td></tr><tr><td>iPhone 12 camera</td><td>Apple</td><td></td><td></td></tr><tr><td>L-glutamine</td><td>Cytiva</td><td>SH3003401</td><td></td></tr><tr><td>Large Kimberly-Clark Professional Kimtech Science Kimwipes Delicate Task Wipers, 1-Ply</td><td>Fisher Scientific</td><td>34133</td><td></td></tr><tr><td>M 205 FA Stereomicroscope</td><td>Leica</td><td></td><td></td></tr><tr><td>Matrigel Membrane Matrix 354234</td><td>Corning</td><td>CB-40234</td><td></td></tr><tr><td>MC-1 UniMotor Controller</td><td>Unisense</td><td></td><td></td></tr><tr><td>Methyl red</td><td></td><td></td><td></td></tr><tr><td>MM33 Micromanipulator</td><td>Marzhauser Wetzlar</td><td>61-42-113-0000</td><td>Right handed</td></tr><tr><td>MS-15 Motorized Stage</td><td>Unisense</td><td></td><td></td></tr><tr><td>Nanoject-II</td><td>Drummond</td><td>3-000-204</td><td>nanoliter autoinjector</td></tr><tr><td>Penicillin/Streptomycin (10,000 U/mL)</td><td>Gibco</td><td>15-140-148</td><td></td></tr><tr><td>pH Microelectrodes</td><td>Unisense</td><td>50-109158, 25-203452, 25-205272, 25-111626, 25-109160</td><td>SensorTrace software is not compatible with Apple computers</td></tr><tr><td>Reference Electrode</td><td>Unisense</td><td>REF-RM-001652</td><td>SensorTrace software is not compatible with Apple computers</td></tr><tr><td>SB 431542</td><td>Tocris Bioscience</td><td>16-141-0</td><td></td></tr><tr><td>Smartphone Camera Adapter</td><td>Gosky</td><td></td><td></td></tr><tr><td>Specifications Laboratory Stand LS</td><td>Unisense</td><td>LS-009238</td><td></td></tr><tr><td>Trypsin-EDTA 0.025%, phenol red</td><td>Gibco</td><td>25-200-056</td><td></td></tr><tr><td>UniAmp</td><td>Unisense</td><td>11632</td><td></td></tr><tr><td>United Biosystems Inc&amp;nbsp;MINI CELL SCRAPERS 200/PK</td><td>Fisher</td><td>MCS-200</td><td></td></tr><tr><td>Y-27632 dihydrochloride</td><td>Tocris Bioscience</td><td>12-541-0</td><td></td></tr><tr><td>&amp;micro;Sensor Calibration Kit</td><td>Unisense/ Mettler Toledo</td><td>51-305-070, 51-302-069</td><td>pH 4.01 and 9.21, 20 mL packets</td></tr></tbody></table>

profiling luminal ph in three-dimensional gastrointestinal organoids using microelectrodes

The optimization and detailed characterization of gastrointestinal organoid models require advanced methods for analyzing their luminal environments. This paper presents a highly reproducible method for the precise measurement of pH within the lumina of 3D human gastric organoids via micromanipulator-controlled microelectrodes. The pH microelectrodes are commercially available and consist of beveled glass tips of 25 &#181;m in diameter. For measurements, the pH microelectrode is advanced into the lumen of an organoid (&gt;200 &#181;m) that is suspended in Matrigel, while a reference electrode rests submerged in the surrounding medium in the culture plate. 
Using such microelectrodes to profile organoids derived from the human gastric body, we demonstrate that luminal pH is relatively consistent within each culture well at ~7.7 &plusmn; 0.037 and that continuous measurements can be obtained for a minimum of 15 min. In some larger organoids, the measurements revealed a pH gradient between the epithelial surface and the lumen, suggesting that pH measurements in organoids can be achieved with high spatial resolution. In a previous study, microelectrodes were successfully used to measure luminal oxygen concentrations in organoids, demonstrating the versatility of this method for organoid analyses. In summary, this protocol describes an important tool for the functional characterization of the complex luminal space within 3D organoids.

Secretion of acid is a crucial function of the human stomach. However, to what extent acid secretion can be modeled in organoids is still a matter of debate6,32,33,34. We therefore developed the protocol detailed above to accurately measure acid production in gastric organoids. Notably, we used unstimulated adult stem cell-derived organoids cultured under standard expansion conditions that had ...

Author Spotlight: Gastric Epithelial Cell Responses in Helicobacter pylori infection

The present protocol describes pH measurements in human tissue-derived gastric organoids using microelectrodes for spatiotemporal characterization of intraluminal physiology.

Profiling Luminal pH in Three-Dimensional Gastrointestinal Organoids Using Microelectrodes

The optimization and detailed characterization of gastrointestinal organoid models require advanced methods for analyzing their luminal environments. This ...

profiling-luminal-ph-three-dimensional-gastrointestinal-organoids

Research

JoVE Journal

Medicine

548 Views.  Montana State University. The present protocol describes pH measurements in human tissue-derived gastric organoids using microelectrodes for spatiotemporal characterization of intraluminal physiology. 

Video: Author Spotlight: Gastric Epithelial Cell Responses in Helicobacter pylori infection

Real-Time, Semi-Automated Fluorescent Measurement of the Airway Surface Liquid pH of Primary Human Airway Epithelial Cells

In recent years, the importance of mucosal surface pH in the airways has been highlighted by its ability to regulate airway surface liquid (ASL) hydration, mucus viscosity and activity of antimicrobial peptides, key parameters involved in innate defense of the lungs. This is of primary relevance in the field of chronic respiratory diseases such as cystic fibrosis (CF) where these parameters are dysregulated. While different groups have studied ASL pH both in vivo and in vitro, their methods report a relatively wide range of ASL pH values and even contradictory findings regarding any pH differences between non-CF and CF cells. Furthermore, their protocols do not always provide enough details in order to ensure reproducibility, most are low throughput and require expensive equipment or specialized knowledge to implement, making them difficult to establish in most labs. Here we describe a semi-automated fluorescent plate reader assay that enables the real-time measurement of ASL pH under thin film conditions that more closely resemble the in vivo situation. This technique allows for stable measurements for many hours from multiple airway cultures simultaneously and, importantly, dynamic changes in ASL pH in response to agonists and inhibitors can be monitored. To achieve this, the ASL of fully differentiated primary human airway epithelial cells (hAECs) are stained overnight with a pH-sensitive dye in order to allow for the reabsorption of the excess fluid to ensure thin film conditions. After fluorescence is monitored in the presence or absence of agonists, pH calibration is performed in situ to correct for volume and dye concentration. The method described provides the required controls to make stable and reproducible ASL pH measurements, which ultimately could be used as a drug discovery platform for personalized medicine, as well as adapted to other epithelial tissues and experimental conditions, such as inflammatory and/or host-pathogen models.

We present a protocol to make dynamic measurements of the airway surface liquid pH under thin film conditions using a plate-reader.

In recent years, the importance of mucosal surface pH in the airways has been highlighted by its ability to regulate airway surface liquid (ASL) ...

Biochemistry

Real-time Measurement of Epithelial Barrier Permeability in Human Intestinal Organoids

Advances in 3D culture of intestinal tissues obtained through biopsy or generated from pluripotent stem cells via directed differentiation, have resulted in sophisticated in vitro models of the intestinal mucosa. Leveraging these emerging model systems will require adaptation of tools and techniques developed for 2D culture systems and animals. Here, we describe a technique for measuring epithelial barrier permeability in human intestinal organoids in real-time. This is accomplished by microinjection of fluorescently-labeled dextran and imaging on an inverted microscope fitted with epifluorescent filters. Real-time measurement of the barrier permeability in intestinal organoids facilitates the generation of high-resolution temporal data in human intestinal epithelial tissue, although this technique can also be applied to fixed timepoint imaging approaches. This protocol is readily adaptable for the measurement of epithelial barrier permeability following exposure to pharmacologic agents, bacterial products or toxins, or live microorganisms. With minor modifications, this protocol can also serve as a general primer on microinjection of intestinal organoids and users may choose to supplement this protocol with additional or alternative downstream applications following microinjection.

This protocol describes the measurement of epithelial barrier permeability in real-time following pharmacologic treatment in human intestinal organoids using fluorescent microscopy and live cell microscopy.

Advances in 3D culture of intestinal tissues obtained through biopsy or generated from pluripotent stem cells via directed differentiation, have resulted ...

Developmental Biology

In vitro Monitoring of Extracellular pH in Real-Time

Early accumulation of neutrophils (PMN) is a hallmark of acute intestinal inflammation. This acute inflammation is either resolved or progresses to chronic inflammation. Without efficient PMN clearance at sites of infiltration, PMN can accumulate and contribute to chronic inflammatory conditions, including the intestinal diseases ulcerative colitis (UC) and Crohn's Disease (CD). The pH in the distal colon in individuals with active UC can range between a pH of 5 and 6, whereas healthy individuals maintain colonic pH in the range of 6.8-7.4. Extracellular pH has been shown to influence both intestinal epithelial cells and the infiltrating immune cells. More specifically, extracellular acidosis significantly impacts PMN. At pH below 6.5, there are increases in the production of H2O2, inhibition of apoptosis, and increases in the functional lifespan of PMN. Given the significant presence of PMN and extracellular acidification at sites of inflammation, we developed a novel model that allows for the monitoring of extracellular pH during PMN transepithelial migration in real time. Here, we describe this model and how it can be utilized to measure both the apical and basal pH during PMN trafficking. This model can be utilized to monitor extracellular pH under a wide range of conditions; including, hypoxia, PMN transepithelial migration, and for extended periods of time.

In vitro Monitoring of Extracellular pH in Real-Time

This article represents a useful in vitro assay to measure changes in extracellular pH during neutrophil (PMN) transepithelial migration (TEM)

Early accumulation of neutrophils (PMN) is a hallmark of acute intestinal inflammation. This acute inflammation is either resolved or progresses to ...

Investigating Intestinal Barrier Breakdown in Living Organoids

Organoids and three-dimensional (3D) cell cultures allow the investigation of complex biological mechanisms and regulations in vitro, which previously was not possible in classical cell culture monolayers. Moreover, monolayer cell cultures are good in vitro model systems but do not represent the complex cellular differentiation processes and functions that rely on 3D structure. This has so far only been possible in animal experiments, which are laborious, time consuming, and hard to assess by optical techniques. Here we describe an assay to quantitatively determine the barrier integrity over time in living small intestinal mouse organoids. To validate our model, we applied interferon gamma (IFN-&#947;) as a positive control for barrier destruction and organoids derived from IFN-&#947; receptor 2 knock out mice as a negative control. The assay allowed us to determine the impact of IFN-&#947; on the intestinal barrier integrity and the IFN-&#947; induced degradation of the tight junction proteins claudin-2, -7, and -15. This assay could also be used to investigate the impact of chemical compounds, proteins, toxins, bacteria, or patient-derived probes on the intestinal barrier integrity.

Here we describe a technique to quantify the barrier integrity of small intestinal organoids. The fact that the method is based on living organoids enables the sequential investigation of different barrier integrity modulating substances or combinations thereof in a time-resolved manner.

Organoids and three-dimensional (3D) cell cultures allow the investigation of complex biological mechanisms and regulations in vitro, which previously was ...

Optical Quantification of Intracellular pH in Drosophila melanogaster Malpighian Tubule Epithelia with a Fluorescent Genetically-encoded pH Indicator

Epithelial ion transport is vital to systemic ion homeostasis as well as maintenance of essential cellular electrochemical gradients. Intracellular pH (pHi) is influenced by many ion transporters and thus monitoring pHi is a useful tool for assessing transporter activity. Modern Genetically Encoded pH-Indicators (GEpHIs) provide optical quantification of pHi in intact cells on a cellular and subcellular scale. This protocol describes real-time quantification of cellular pHi regulation in Malpighian Tubules (MTs) of Drosophila melanogaster through ex vivo live-imaging of pHerry, a pseudo-ratiometric GEpHI with a pKa well-suited to track pH changes in the cytosol. Extracted adult fly MTs are composed of morphologically and functionally distinct sections of single-cell layer epithelia, and can serve as an accessible and genetically tractable model for investigation of epithelial transport. GEpHIs offer several advantages over conventional pH-sensitive fluorescent dyes and ion-selective electrodes. GEpHIs can label distinct cell populations provided appropriate promoter elements are available. This labeling is particularly useful in ex vivo, in vivo, and in situ preparations, which are inherently heterogeneous. GEpHIs also permit quantification of pHi in intact tissues over time without need for repeated dye treatment or tissue externalization. The primary drawback of current GEpHIs is the tendency to aggregate in cytosolic inclusions in response to tissue damage and construct over-expression. These shortcomings, their solutions, and the inherent advantages of GEpHIs are demonstrated in this protocol through assessment of basolateral proton (H+) transport in functionally distinct principal and stellate cells of extracted fly MTs. The techniques and analysis described are readily adaptable to a wide variety of vertebrate and invertebrate preparations, and the sophistication of the assay can be scaled from teaching labs to intricate determination of ion flux via specific transporters.

Optical Quantification of Intracellular pH in Drosophila melanogaster Malpighian Tubule Epithelia with a Fluorescent Genetically-encoded pH Indicator

Cellular ion transport can often be assessed by monitoring intracellular pH (pHi). Genetically Encoded pH-Indicators (GEpHIs)&#160;provide optical quantification of intracellular pH in intact cells. This protocol details the quantification of intracellular pH through cellular ex vivo live-imaging of Malpighian tubules of Drosophila melanogaster with pHerry, a pseudo-ratiometric genetically encoded pH-indicator.

Epithelial ion transport is vital to systemic ion homeostasis as well as maintenance of essential cellular electrochemical gradients. Intracellular pH ...

Biology

Establishment of 3D Morphogenesis in a Canine IBD Gut-on-a-Chip System

Three-Dimensional Morphogenesis in Canine Gut-on-a-Chip Using Intestinal Organoids Derived from Inflammatory Bowel Disease Patients

Canine intestines possess similarities in anatomy, microbiology, and physiology to those of humans, and dogs naturally develop spontaneous intestinal disorders similar to humans. Overcoming the inherent limitation of three-dimensional (3D) organoids in accessing the apical surface of the intestinal epithelium has led to the generation of two-dimensional (2D) monolayer cultures, which expose the accessible luminal surface using cells derived from the organoids. The integration of these organoids and organoid-derived monolayer cultures into a microfluidic Gut-on-a-Chip system has further evolved the technology, allowing for the development of more physiologically relevant dynamic in vitro intestinal models.
In this study, we present a protocol for generating 3D morphogenesis of canine intestinal epithelium using primary intestinal tissue samples obtained from dogs affected by inflammatory bowel disease (IBD). We also outline a protocol for generating and maintaining 2D monolayer cultures and intestine-on-a-chip systems using cells derived from the 3D intestinal organoids. The protocols presented in this study serve as a foundational framework for establishing a microfluidic Gut-on-a-Chip system specifically designed for canines. By laying the groundwork for this innovative approach, we aim to expand the application of these techniques in biomedical and translational research, aligning with the principles of the One Health Initiative. By utilizing this approach, we can develop more physiologically relevant dynamic in vitro models for studying intestinal physiology in both dogs and humans. This has significant implications for biomedical and pharmaceutical applications, as it can aid in the development of more effective treatments for intestinal diseases in both species.

Author Spotlight: Development and Application of a Canine IBD Gut-on-a-Chip Model for 3D Intestinal Morphogenesis Studies 

Integration of canine intestinal organoids and a Gut-on-a-Chip microfluidic system offers relevant translational models for human intestinal diseases. Protocols presented allow for 3D morphogenesis and dynamic in vitro modeling of the gut, aiding in the development of effective treatments for intestinal diseases in dogs and humans with One Health.

Canine intestines possess similarities in anatomy, microbiology, and physiology to those of humans, and dogs naturally develop spontaneous intestinal ...

3D Culturing of Organoids from the Intestinal Villi Epithelium Undergoing Dedifferentiation

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is compartmentalized into crypts and villi: the stem and proliferating cells are confined to the crypts, whereas the villi epithelium contains only differentiated cells. Hence, the normal intestinal crypts, but not the villi, can give rise to organoids in 3D cultures. The procedure described here is applicable only to villus epithelium undergoing dedifferentiation leading to stemness. The method described uses the Smad4-loss-of-function:&#946;-catenin gain-of-function (Smad4KO:&#946;-cateninGOF) conditional mutant mouse. The mutation causes the intestinal villi to dedifferentiate and generate stem cells in the villi. Intestinal villi undergoing dedifferentiation are scraped off the intestine using glass slides, placed in a 70 &#181;m strainer and washed several times to filter out any loose cells or crypts prior to plating in BME-R1 matrix to determine their organoid-forming potential. Two main criteria were used to ensure that the resulting organoids were developed from the dedifferentiating villus compartment and not from the crypts: 1) microscopically evaluating&#160;the isolated villi to ensure absence of any tethered crypts, both before and after plating in the 3D matrix, and 2) monitoring the time course of organoid development from the villi. Organoid initiation from the villi occurs only two to five days after plating and appears irregularly shaped, whereas the crypt-derived organoids from the same intestinal epithelium are apparent within sixteen hours of plating and appear spherical. The limitation of the method, however, is that the number of organoids formed, and the time required for organoid initiation from the villi vary depending on the degree of dedifferentiation. Hence, depending upon the specificity of the mutation or the insult causing the dedifferentiation, the optimal stage at which villi can be harvested to assay their organoid forming potential, must be determined empirically.

The procedure describes isolation of the villi from the mouse intestinal epithelium undergoing dedifferentiation to determine their organoid forming potential.

Clonogenicity of organoids from the intestinal epithelium is attributed to the presence of stem cells therein. The mouse small intestinal epithelium is ...

Culture Methods to Study Apical-Specific Interactions using Intestinal Organoid Models

The lining of the gut epithelium is made up of a simple layer of specialized epithelial cells that expose their apical side to the lumen and respond to external cues. Recent optimization of in vitro culture conditions allows for the re-creation of the intestinal stem cell niche and the development of advanced 3-dimensional (3D) culture systems that recapitulate the cell composition and the organization of the epithelium. Intestinal organoids embedded in an extracellular matrix (ECM) can be maintained for long-term and self-organize to generate a well-defined, polarized epithelium that encompasses an internal lumen and an external exposed basal side. This restrictive nature of the intestinal organoids presents challenges in accessing the apical surface of the epithelium in vitro and limits the investigation of biological mechanisms such as nutrient uptake and host-microbiota/host-pathogen interactions. Here, we describe two methods that facilitate access to the apical side of the organoid epithelium and support the differentiation of specific intestinal cell types. First, we show how ECM removal induces an inversion of the epithelial cell polarity and allows for the generation of apical-out 3D organoids. Second, we describe how to generate 2-dimensional (2D) monolayers from single cell suspensions derived from intestinal organoids, comprised of&#160;mature and differentiated cell types. These techniques provide novel tools to study apical-specific interactions of the epithelium with external cues in vitro and promote the use of organoids as a platform to facilitate precision medicine.

Here we present two protocols that allow the modeling of intestinal apical-specific interactions. Organoid-derived intestinal monolayers and Air-Liquid Interface (ALI) cultures facilitate the generation of well-differentiated epithelia accessible from both luminal and basolateral sides, whereas polarity-inverted intestinal organoids expose their apical side and are amenable to high throughput assays.

The lining of the gut epithelium is made up of a simple layer of specialized epithelial cells that expose their apical side to the lumen and respond to ...

Culture of Piglet Intestinal 3D Organoids from Cryopreserved Epithelial Crypts and Establishment of Cell Monolayers

Intestinal organoids are increasingly being used to study the gut epithelium for digestive disease modeling, or to investigate interactions with drugs, nutrients, metabolites, pathogens, and the microbiota. Methods to culture intestinal organoids are now available for multiple species, including pigs, which is a species of major interest both as a farm animal and as a translational model for humans, for example, to study zoonotic diseases. Here, we give an in-depth description of a procedure used to culture pig intestinal 3D organoids from frozen epithelial crypts. The protocol describes how to cryopreserve epithelial crypts from the pig intestine and the subsequent procedures to culture 3D intestinal organoids. The main advantages of this method are (i) the temporal dissociation of the isolation of crypts from the culture of 3D organoids, (ii) the preparation of large stocks of cryopreserved crypts derived from multiple intestinal segments and from several animals at once, and thus (iii) the reduction in the need to sample fresh tissues from living animals. We also detail a protocol to establish cell monolayers derived from 3D organoids to allow access to the apical side of epithelial cells, which is the site of interactions with nutrients, microbes, or drugs. Overall, the protocols described here is a useful resource for studying the pig intestinal epithelium in veterinary and biomedical research.

Here, we describe a protocol to culture pig intestinal 3D organoids from cryopreserved epithelial crypts. We also describe a method to establish cell monolayers derived from 3D organoids, allowing access to the apical side of epithelial cells.

Intestinal organoids are increasingly being used to study the gut epithelium for digestive disease modeling, or to investigate interactions with drugs, ...

3D Culture of Organoids from Murine Intestinal Crypts and Single Stem Cell

The 3D Culturing of Organoids from Murine Intestinal Crypts and a Single Stem Cell for Organoid Research

At present, organoid culture represents an important tool for in vitro studies of different biological aspects and diseases in different organs. Murine small intestinal crypts can form organoids that mimic the intestinal epithelium when cultured in a 3D extracellular matrix. The organoids are composed of all cell types that fulfill various intestinal homeostatic functions. These include Paneth cells, enteroendocrine cells, enterocytes, goblet cells, and tuft cells. Well-characterized molecules are added into the culture medium to enrich the intestinal stem cells (ISCs) labeled with leucine-rich repeats containing G protein-coupled receptor 5 and are used to drive differentiation down specific lineages; these molecules include epidermal growth factor, Noggin (a bone morphogenetic protein), and R-spondin 1. Additionally, a protocol to generate organoids from a single erythropoietin-producing hepatocellular receptor B2 (EphB2)-positive ISC is also detailed. In this methods article, techniques to isolate small intestinal crypts and a single ISC from tissues and ensure the efficient establishment of organoids are described.

Author Spotlight: The 3D Culturing of Organoids from Murine Intestinal Crypts and a Single Stem Cell for Organoid Research  

We describe a protocol to isolate murine small intestinal crypts and culture intestinal 3D organoids from the crypts. Additionally, we describe a method to generate organoids from a single intestinal stem cell in the absence of a sub-epithelial cellular niche.

At present, organoid culture represents an important tool for in vitro studies of different biological aspects and diseases in different organs. Murine ...

Profiling Luminal pH in Three-Dimensional Gastrointestinal Organoids Using Microelectrodes

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Three-Dimensional Morphogenesis in Canine Gut-on-a-Chip Using Intestinal Organoids Derived from Inflammatory Bowel Disease Patients

3D Culturing of Organoids from the Intestinal Villi Epithelium Undergoing Dedifferentiation

Culture Methods to Study Apical-Specific Interactions using Intestinal Organoid Models

Culture of Piglet Intestinal 3D Organoids from Cryopreserved Epithelial Crypts and Establishment of Cell Monolayers

The 3D Culturing of Organoids from Murine Intestinal Crypts and a Single Stem Cell for Organoid Research