Our protocol can be used to visualize the structural complexity of organoids, allowing mapping of identity, sulfate, and distribution in these 3D structures at the single-cell resolution. Our quick three-day protocol is fully optimized for organoids of various origin and uses a straightforward sample preparation, that includes a non-toxic optical clearing step and a silicon-based mounting method. Although our protocol is straightforward, some of them are critical steps such as the organized handling, or the slide preparation, can be better explained through visual demonstration than through text.
To recover 100 to 500 micrometer diameter organoids grown in basement membrane extract in 24 well plates, wash each weld to be harvested with PBS without disrupting the 3D matrices, and place the plate on ice. Add one milliliter of ice-cold recovery solution to each well and place the plate on a horizontal shaker at four degrees Celsius for 30 to 60 minutes. Dip one milliliter pipette tips into a 1%BSA in PBS solution and pipette up and down two times to coat each tip with BSA.
Next, add five milliliters of 1%PBS BSA to one 15 milliliter tube per condition, and invert the tube two to three times before discarding the solution, to coat the inside of the tube. To collect the organoids, use a coated tip to gently re-suspend the well contents five to 10 times, and pull all of the organoids from each condition in a single coated 15 milliliter tube. Rinse each well with one milliliter of ice-cold 1%PBS BSA to ensure that all of the organoids have been collected, and transfer the washes to the appropriate tubes.
Bring the final volume in each tube up to 10 milliliters with cold PBS, and sediment the organoids by centrifugation to obtain a tight palette without a visible layer of 3D matrix. To fix the organoids, use a coated one milliliter pipette tip to carefully re-suspend each pellet in one milliliter of ice cold para formaldehyde, and fix at four degrees Celsius for 45 minutes. Gently re-suspend the organoids halfway through the fixation time.
After 45 minutes, add 10 milliliters of ice-cold PBS plus tween 20 to each tube and gently mix by inversion, before placing the tubes at four degrees Celsius for 10 minutes. To block the organoids, at the end of the incubation, spin down the samples and re-suspend the pellets in at least 200 microliters of ice-cold organoid washing buffer per well to be plated. Then transfer the organoids to individual wells of a 24 wells suspension plate for a 15 minute incubation at four degrees Celsius.
For immunolabeling add 200 microliters of organoid washing buffer into an empty reference well, and allow the organoids to settle to the bottom of the plate. When the organoids have settled, tilt the plate at a 45 degree angle to allow removal of all but the last 200 microliters of wash buffer. Next, add 200 microliters of organoid washing buffer containing the primary antibodies of interest, and place the plate overnight at four degrees Celsius with mild rocking and shaking at 40 revolutions per minute.
The next morning, add one milliliter of organoid washing buffer to each well, and allow the organoids to settle to the bottom of the plate for three minutes. When the organoids have settled, remove all but the the last 200 microliters from each well, and wash the organoids with three two-hour washes with one milliliter of fresh organoid washing buffer, and mild rocking and shaking per wash. After the third wash, allow the organoids to settle at the bottom of the plate for three minutes before removing all but the last 200 microliters of organoid washing buffer from each well.
Add 200 microliters of organoid washing buffer containing the appropriate secondary antibodies to each well for an overnight incubation at four degrees celsius with mild rocking and shaking. The next morning, wash the organoids with three two-hour washes in one milliliter of fresh organoid washing buffer per wash, as demonstrated. After the last wash, transfer the organoids into one 1.5 milliliter tube per well, and collect the organoids by centrifugation.
For optical clearing of the organoids, remove as much wash buffer from each tube as possible, without disrupting the organoids, and use a modified 200 microliter pipette tip to add at least 50 microliters of FUnGI to each pellet. After a 20 minute incubation at room temperature the organoids can be stored for up to one week at four degrees Celsius, or for up to six months at minus 20 degrees Celsius. To prepare slides for organoid imaging by confocal microscopy, fill a 10 milliliter syringe with a silicone sealant and attach a modified 200 microliter pipette tip to the syringe.
Use the syringe to draw a rectangle of one by two centimeters in the middle of a slide, and use a second modified 200 microliter pipette tip to place cleared organoids into the middle of the rectangle. To place a cover slip over the organoids, place the left side of the cover slip down first, before slowly lowering the cover slip from left to right until there is no trapped air. Then gently apply pressure to all of the edges of the cover slip to firmly attach it to the silicone sealant.
To image the organoids, place the slide onto the stage of a confocal laser scanning microscope, and select a multi immersion 25 X objective for confocal imaging. Set the microscope to the appropriate acquisition settings, selecting a low laser power to reduce photo bleaching. Use the Z stack mode to define the lower end upper bounds, and set the Z step size to optimal.
When imaging large organoid structures, or multiple organoids together, use the tiling mode with a 10%overlap and indicate the area of interest. When all of the parameters have been set, obtain a 3D rendered representation of the imaging in the imaging software, and optimize the brightness, contrast, and 3D rendering properties. Then export RGB snapshots of the results as TIFF files.
The strength of 3D imaging compared to 2D imaging is illustrated by these images of mouse mammary gland organoids that were generated as demonstrated. The central layer of these representative organoids consists of columnar shaped K8/K18 positive luminal cells, and the outer layer contains elongated K5 positive basil cells, recapitulating the morphology of the mammary gland in vivo. This polarized organization is challenging to appreciate from a 2D optical section of the same organoid.
Another example of a complex structure that is impossible to interpret without 3D information is the network of MRP2 positive canaliculi that facilitate the collection of the bile fluid of human liver organoids. Moreover, the obtained quality and resolution allows for semi-automated segmentation and image analysis. Thus, total cell numbers and the presence of markers can be quantified in specific cellular subtypes in whole organoids.
By segmenting the nuclei of an entire organoid containing 140 cells, three cells that display a high positivity for the Ki67 cell cycle marker can be identified. The optical clearing agent, FUnGI, outperforms uncleared and fructose-glycerol clearing in fluorescent signal quality deep within an organoid. With FUnGI cleared organoids demonstrating an overall enhanced fluorescence intensity compared to uncleared organoids.
Note that any leftover BME in the sample may result in reduced antibody penetration and can lead to high background signal. In addition, cystic organoids should not be optically cleared if they collapse. With slight adaptations to the protocol, the samples can be used with super resolution confocal, multi photon, and light sheet microscopy.
