This protocol describes some methods to quantify lipid droplet formation in human intestinal organoids. It can be used as a high-throughput screening platform to test for drugs that modulate lipid droplet formation. The method we describe is based on a lipid droplet-specific dye and biopsy-derived intestinal organoids.
It thereby provides a stable, accurate, and physiologically-relevant model for lipid droplet formation. This protocol can be used to screen for patient-specific novel drug candidates that modulate lipid droplets formation, for example, in DGAT1-deficient patients. This lipid droplet formation assay can also be applied to other types of organoid cultures to study lipid metabolism in other cell types.
The assay is to a significant extent dependent on the density of the organoid culture. Try to ensure a consistent organoid density across samples and experiments. Visual demonstration of this assay is critical because it is important to show how the organoid should be cultured and how they should be handled to ensure proper analysis of lipid droplet formation.
After preparing the organoids, carefully aspirate the culture medium without disturbing the droplets of basement membrane matrix. Add 500 microliters of cold basal medium to the first well of organoids. Using a P1000 pipette, gently pipette up and down to disrupt the basement membrane matrix droplets with organoids.
Repeat this procedures with the same medium in the next well as required, but do not harvest more than two wells per 500 microliters of basal medium. Collect the organoids in a low-binding 1.5 milliliter microcentrifuge tube, and spin them down in a mini-tabletop centrifuge for 15 to 20 seconds. Add 400 microliters of trypsin to the spun-down organoids.
Incubate in a water bath at 37 degrees Celsius for five minutes. Next, use a P200 pipette to gently pipette the suspension up and down to disrupt the remaining aggregates of cells. Incubate in a water bath at 37 degrees Celsius for five minutes.
When only single cells remain, add one milliliter of basal medium, and spin down the cells in a mini-tabletop centrifuge for 15 to 20 seconds. Aspirate the supernatant completely. Use a P200 pipette to re-suspend the single cells in 200 microliters of fresh hSI-EM, and add an additional 800 microliters of hSI-EM.
Then count the number of cells in suspension and calculate the volume of cells needed for the final cell density. Take out the appropriate volume of suspension and adjust the density to 750 cells per microliter. Add basement membrane matrix to the cell suspension in a ratio of two-to-one.
Gently pipette the suspension to mix being careful to avoid creating bubbles. In a pre-warmed culture plate, seed out three 10-microliter droplets per well if the plate has 24 wells or one five-microliter droplet per well if the plate has 96 wells. Place the plate in an incubator at 37 degrees Celsius with five percent carbon dioxide for 10 to 15 minutes to solidify the droplets.
Meanwhile, pre-warm an appropriate volume of hSI-EM+Y in a water bath at 37 degrees Celsius. After this, carefully add the pre-warmed hSI-EM+Y to each well of the plate. Incubate the cells at 37 degrees Celsius with five percent carbon dioxide.
After two to three days, replace the medium with hSI-EM and make sure to refresh it two to three times a week. First, weigh 0.2 grams of liquid oleic acid at room temperature. Add 1.5 milliliters of culture-grade sterile PBS and heat the mixture to 70 degrees Celsius for one hour making sure to vortex intermittently.
Next, weigh 5.89 grams of fatty acid-free BSA and dissolve it in 33.9 milliliters of PBS. Warm the mixture in a water bath at 37 degrees Celsius until the BSA is fully dissolved. Vortex the oleic acid mixture again to create an emulsion of fine droplets and immediately add it to the BSA solution using a glass pipette.
Keep the mixture at 37 degrees Celsius for 30 minutes until a clear yellowish solution remains. Passage the organoids as previously described and seed out the organoid-derived single cells into a black clear-bottom 96-well plate. On day six of culturing on hSI-EM, replace the culture medium with hSI-EM containing one millimolar of the oleic acid and BSA conjugate.
Incubate the cells for 16 to 17 hours at 37 degrees Celsius in the presence or absence of 0.1 micromolar DGAT1 inhibitor. After this, aspirate the medium without disturbing the basement membrane matrix droplets. Fixate the organoids by adding 100 microliters of four percent neutral buffered formaldehyde to the wells for 30 minutes at room temperature.
Remove the formaldehyde gently and carefully wash the cells with 150 microliters of PBS per well. Stain the cells for lipid droplets with 025 milligrams per milliliter LD540 and DAPI in PBS for 15 minutes at room temperature in the dark. Then wash the wells carefully with PBS.
For overview images of whole organoids use a 40X subjective suited for confocal fluorescent imaging. Set the microscope to image for DAPI and the LD540 dye using the excitation and emission settings shown here. Set the pinhole size to one airy unit for sufficient z-axis resolution.
To image one-half of a spherical organoid set a z-stack to approximately 85 micrometers. Using an appropriate image processing system, transform the z-stack of each organoid to a maximum projection by clicking Image, Stacks, Z Project. Set the threshold of the maximum projection to a level in which there is no LD540 signal visible in the BSA vehicle control sample by clicking Image, Adjust, Threshold.
Use these settings to threshold each image. Then measure the total area of fluorescence for each maximum projection using the function Analyze, Analyze Particles. For proper analysis of lipid droplet formation, the organoids should not be seeded too densely prior to stimulation with oleic acid and subsequent staining.
This is especially of importance for the confocal and plate-reader readout since overlapping organoids might interfere with the fluorescence. After stimulation with one millimolar of oleic acid overnight, lipid droplet formation can be visualized with an inverted bright-field microscope. The accumulation of lipid droplets scatters transmitted light, and therefore the organoids appear darker, whereas non-stimulated organoids have a translucent appearance.
Once the oleic acid-stimulated organoids are fixed and stained, the lipid droplet formation can be visualized using a confocal microscope. As the organoids are 3D structures, a regular epifluorescent microscope is not suitable due to the out-of-focus background signal, so a confocal z-stack is used to characterize LDF in organoids. Quantification of the confocal microscopy samples can also be performed using a fluorescent plate reader normalized to the fluorescent Hoechst signal.
The plate reader assay indicates a significant decrease in the LD540 signal in organoid cells treated with DGAT1 inhibitor compared to untreated organoids. Quantification of LD formation in individual cells can be achieved using a flow cytometer. The gating strategy shown here is for dissociated human intestinal organoids.
Representative histograms for both the SSC-A and the LD540 signal of the final live cell population show that lipid droplet formation will result in an increase in SSC-A due to the formation of intracellular lipid droplets. In addition, LD540 stains for lipids that are stored in the lipid droplets, and the signal will also increase upon the induction of lipid droplet formation. As such, lipid droplet formation is measured as a shift in the mean fluorescent intensity of both SSC-A and LD540.
The most crucial steps of this protocol are the culture of human intestinal organoids and the proper corrugation of oleic acid to BSA. To accommodate more conditions, this assay can be analyzed using a fluorescent plate reader. To correct for the organoid number per well, the LD540 signal was normalized to the DAPI signal.
With this technique, researchers can study lipid droplet formation in organoid systems, and improve various analysis techniques such as electron microscopy to validate their findings.
