This method is a valuable tool in the field of cellular biology for assessing cellular ROS levels in living primary cells cultivated in vitro as organoids using a fluorogenic probe. The main advantage of this technique is that ROS can be qualitatively visualized in intact intestinal organoids by microscopy and quantitatively analyzed in dissociated cells by flow cytometry using 96-well plates. This method can be applied to other organoid models and alternative fluorescent sensors might be used to analyze different cellular pathways.
Before applying this protocol for the first time, the experimenter should become familiar with the organoid culture and passaging. Although our protocol is well-adapted to screening approaches, beginners should start with a few samples. Demonstrating the procedure will be Sophie Dulauroy, an engineer from the laboratory.
After sacrificing an 8-to-10-week-old Lgr5-GFP mouse, collect five to eight centimeters of the jejunum encompassing region between the duodenum and ileum, and place it in cold DPBS supplemented with antibiotics on ice. Next, clean the intestinal content by flushing with five to 10 milliliters of cold DPBS containing antibiotics. Then using ball tip scissors, open the intestine longitudinally, and using forceps, transfer the tissue into a Petri dish containing cold DPBS with antibiotics.
Shake the tissue inside the dish to rinse it. Next, grab the intestine by aspiration using a plastic Pasteur pipette and transfer it into a 15 milliliter tube containing 10 milliliters of cold 10 millimolar EDTA. Invert the tube three times and incubate it on ice for 10 minutes.
After the incubation, transfer the tissue in a tube containing 10 milliliters of DPBS and vortex for two minutes. Immediately add 10 microliters of the fraction in a Petri dish and use a microscope to assess the quality of the fraction. After two transfers in DPBS containing tubes with two-minute vortexing between each transfer, incubate the intestine in EDTA on ice for five minutes as demonstrated previously.
Following three successive transfers in tubes containing DPBS with three-minute vortexing between each transfer, combine the best fractions into a 50 milliliter tube by inverting the selected tubes three times and filtering through a 70 micron cell strainer. Then spin the crypts, discard the supernatant and disrupt the pellet mechanically before adding five milliliters of cold DMEM F12. Count the number of crypts present in a 10 microliter aliquot manually under a microscope.
After counting, spin the crypts again and carefully remove the supernatant. Then mechanically disrupt the pellet and add growth culture medium to the tube to obtain a concentration of 90 crypts per microliter. Next, add two volumes of undiluted basement membrane matrix or BMM to get a final concentration of 30 crypts per microliter and carefully pipette the suspension up and down without introducing air bubbles into the mix.
For flow cytometry analysis, plate 10 microliters of the crypts BMM mix as a dome at the center of each well of a pre-warmed round bottom 96-well plate. And for imaging, deposit 10 microliters of the mix as a thin layer in a pre-warmed microslide eight-well chamber. After allowing the BMM to solidify at room temperature for five minutes, place the plates in an incubator at 37 degrees Celsius and 5%carbon dioxide.
After 15 minutes, add 250 microliters of growth medium into each well, taking care not to detach the BMM, and place the plate in the incubator. To visualize oxidative stress by confocal microscopy, add one microliter of n-acetylcysteine stock solution in the corresponding wells of the microslide eight-well chamber plated with organoids. After a one-hour incubation, add one microliter of tert-butyl hydroperoxide stock solution in the corresponding wells and incubate for another 30 minutes.
Next, add one microliter of the 1.25 millimolar dilution of the fluorogenic probe per well, followed by one microliter of 1.25 milligrams per milliliter Hoechst solution. After another 30-minute incubation, remove the medium without disturbing the BMM and gently add 250 microliters of warm DMEM without phenol red. Image the organoids using a confocal microscope equipped with a thermic chamber and gas supply that detects the fluorogenic probe.
Using the positive control, set up the laser intensity and time exposure for the ROS signal, and check that this signal is lower in the negative control. Next, using an eyepiece, screen the slide to identify the organoids expressing GFP and adjust the laser intensity. Set up a z-stack of 25 micrometers and define positions to obtain a stitched image of the whole organoid to get a section of the organoids showing one layer of cells.
Add one microliter of the n-acetylcysteine stock solution in the negative control wells of the 96-well round bottom plate containing the organoids. After a one-hour incubation, add one microliter tert-butyl hydroperoxide stock solution in the corresponding wells and incubate for 30 minutes. Then using a multi-channel pipette, remove the medium without disturbing the attached BMM and transfer it to another 96-well round bottom plate.
Keep this plate aside. Next, add 100 microliters of trypsin and using a multi-channel pipette, pipette up and down five times to destroy the BMM. After a short five-minute incubation, dissociate the organoids by pipetting up and down a second time.
Spin the plate and discard the supernatant by inverting the plate. Add the medium previously collected in another 96-well plate back into the corresponding wells and resuspend the cells by pipetting up and down five times. Next, add one microliter of the fluorogenic probe and incubate for 30 minutes.
Then spin the plate again and resuspend the cells with 250 microliters of DAPI solution. Transfer the samples in flow cytometry tubes and keep the tubes on ice. Optimize the forward and side scatter voltage settings on unstained control and laser voltages for each fluorophore using monostained samples.
Then using an appropriate gating strategy, collect a minimum of 20, 000 events. Representative confocal images of ROS staining and organoids showed that in the presence of the NAC inhibitor, only the signal from the dead cells contained in the lumen of the organoid is visible. In the non-treated organoid, the basal ROS levels can be seen, particularly in GFP positive cells, proving that stem cells produce higher ROS than differentiated cells.
GFP positive cells present a more significant cytoplasmic signal with the inducer in the presence of the fluorogenic probe, demonstrating that ROS levels increase particularly in stem cells after treatment. Flow cytometry analysis of ROS production in the intestinal organoids shows that basal ROS levels decrease after stimulation with inhibitor and increase after challenge with inducer. Cells pretreated with inhibitor and then stimulated with inducer present lower levels than those stimulated with inducer alone.
A similar analysis on stem cells gated as GFP positive shows a 3.5 fold decrease in ROS level upon inhibitor treatment and a fourfold increase upon inducer treatment over non-stimulated cells. When attempting this procedure, users should remember to limit cell stress during organoid manipulation and dissociation. Additionally, positive and negative controls should always be included when analyzing ROS.
Following this procedure, the analysis can be complemented by immunofluorescence staining on fixed intact organoids, cell sorting, and gene expression analysis to get additional insights on ROS regulation. After watching this video, you should know how to grow murine intestinal organoids, perform live imaging analysis of intact organoids, and process intestinal organs for flow cytometry analysis.
