Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

The overall goal of this procedure is to isolate 3D in vitro basement matrix embedded intestinal organoids and to subsequently fix and immunolabel the microtubules and centrosomal proteins. 3D in vitro organoids are ideal models for studying key biological questions in cell and developmental biology. But localizing endogenous proteins and structures in 3D models is more complex than in 2D cultures.

Importantly, the fixative needs to preserve the delicate 3D architecture while also preserving the antibody antigenicity. The presented methods include isolation, fixation, and immunolabeling a 3D in vitro intestinal organoids. But the fixation and immunolabeling protocols can also be used in ex vivo isolated tissue.

The main advantage of the presented formaldehyde methanol fixation protocol is that it preserves the 3D structures while also preserving antigenicity. It enables good penetration and clearance of post-fixed tevel antibodies for effective labeling of microtubules, actin filaments and binding proteins and several centrosomal proteins including ninein. Demonstrating the procedure will be doctor Deborah Goldspink, a formal postdoc in my lab, and doctor Zoe Matthews, a co-author.

Following the isolation of intestinal crypts and villi, isolated epithelial structures can be fixed for immunostaining. The isolated crypts can alternately be seated into matrix domes which will then form organoids. After approximately five to seven days of incubation, organoids will have formed.

Use 500 microliters of RT-PBS to wash the wells containing the basement matrix domes with organoids. Then, add 250 microliters of cold cell recovery solution to each well. Next, using a P1000 micropipette, scrape the basement matrix domes and carefully pipette up and down throughout the well to break up and remove the basement matrix from the plastic.

Make sure the recovery solution is cold to help break down the matrix. When scraping, only use a P1000 micropipette so you don't break up the organoids and that they remain intact. Collect the supernatant in 1.5 milliliter low-binding microcentrifuge tubes.

Then, invert the tube several times and check under a microscope at times 50 magnification that the organoids have been isolated, are free moving and are not in clumps. Pellet the organoids by centrifugation at 1, 000 g and room temperature for five minutes. Then, remove the recovery reagent and proceed immediately to fixation.

To fix previously isolated organoids with formaldehyde and methanol, resuspend the organoids in negative 20 degrees Celsius formaldehyde methanol fixative solution. Incubate the organoids at minus 20 degrees Celsius in a freezer for 15 minutes, inverting the tube every five minutes. Then, pellet the organoids by centrifugation at 1, 000 g for five minutes.

Remove the fixative and then add the washing solution consisting of either PBS with 1%secondary antibody species serum or PBS with 0.1%detergent and 1%serum. After pelleting the sample as before, remove the washing solution and resuspend the sample in fresh washing solution. Use a tube rotator to wash the cells for a total of one hour, pelleting the organoids by centrifugation every 15 minutes and replacing the wash solution.

To block the intestinal organoid samples, add 10%secondary antibody species serum to PBS. Pellet the organoids at 1, 000 g for five minutes and remove the supernatant. Add one milliliter blocking solution to each sample to be incubated in the different antibodies.

Then, incubate the samples and blocking solution on a tube rotator at room temperature for one hour. Next, dilute the primary antibodies in PBS containing 10%serum and 0.1%detergent. Then, after spinning the samples and removing the blocking solution, use the primary antibody solution to resuspend the organoid pellet.

Rotate the tubes at 20 RPM and four degrees Celsius overnight to keep the organoids in suspension. The following day, bring the samples back to room temperature on the tube rotator for one hour. After spinning the sample, remove the primary antibody solution, add one milliliter of washing solution to each tube and resuspend the organoid pellets.

Then, immediately remove the solution by centrifugation. Add one milliliter of fresh wash solution and mix the cells on a tube rotator at 20 RPM for two hours. Next, dilute the secondary antibodies in PBS with 1%serum and 0.1%detergent.

Then, after pelleting the tissue, resuspend the pellets in 200 microliters of secondary antibody solution. Incubate the tubes on a tube rotator at room temperature for one hour. After spinning and removing the supernatant, resuspend the pellet in wash solution and immediately centrifuge the samples to pellet the organoids.

Then, remove the supernatant and resuspend the pellet in one milliliter of fresh wash solution. Mix the samples on a tube rotator at room temperature for 1.5 to two hours, changing the wash solution every 20 to 30 minutes as previously described. To mount the organoids, after spinning and removing all the wash solution, add two drops of hard-setting mounting medium with antifade reagent to the pellet.

Cut the end of a P200 micropipette and carefully resuspend the pellet in the mounting medium. When resuspending fixed and stained organoids in mounting medium, be careful not to introduce any bubbles. If bubbles are present, allow them to float to the surface and then avoid transferring them to the slide.

Use the micropipette to dispense the organoids with mounting medium in a line along the center of a microscope slide. Then, carefully place a cover glass over the top and avoid generating bubbles. Place the glass slide in a slide book and store in a fridge overnight for the mounting medium to set before analyzing on a confocal microscope.

Shown here are images from fraction two of isolated small intestine tissue containing both villi and crypts and a sample from fraction three containing mainly crypts. As seen in these images, good preservation and labeling of microtubules and actin in both villi and crypts was achieved with the formaldehyde methanol fixation and immunolabeling protocol described in this video. Small intestinal organoids were generated and grown in basement matrix for three weeks or longer, then isolated as demonstrated in this video.

The differentiated cells within the organoid villus domains contain stable apicobasal microtubules that were well labeled in most cases. EB1 could also be seen along the microtubule lattice. Shown here is an organoid at the cyst stage and at an early stage of crypts development.

Both samples were fixed in formaldehyde methanol and immunolabeled for microtubules and ninein, which demonstrates good microtubule preservation and labeling at apical non-centrosomal microtubule organizing centers. Once mastered, this technique can be performed with multiple samples and over two days. While attempting this procedure, it is important to carefully scrape wells when collecting organoids to avoid disrupting their 3D structure.

Also, be careful when adding or removing solutions to prevent the loss of organoids throughout the staining procedure. After watching this video, you should have a good understanding of how to fix, immunolabel and mount isolated organoids and other epithelial structures such as intestinal crypts. Don't forget that working fixative reagents in this procedure can be extremely hazardous.

Risk assessments of this procedure should be performed and appropriate containment measures and PPE should always be taken while performing this procedure.