Characterizing 3D in vitro model at cellular and subcellular resolution is crucial for the full exploitation, but can be challenging. Therefore, we have provided complimentary protocol for the staining and subcellular resolution imaging of fixed 3D in vitro models ranging from 100 micrometers to several millimeters. 3D structures are radial, and should be carefully manipulated.
Before pipetting, check the location of your 3D structures within tube to avoid any aspiration. As we have adapted classical techniques for the embedding of small structures such as organoids or spheroids, visualized instructions can help first time users perform the task more easily. Demonstrating the procedure with me will be Laura Francols, a lab tech from the Pathology Research Platform.
To prepare samples for 3D whole mount staining, use a one milliliter pipette tip pre-coded with BSA to add a one milliliter 3D structure suspension to a 500 microliter tube of PBS. When the 3D structures have settled to the bottom of the tube, carefully replace the PBS with 500 microliters of permeabilization blocking solution for a one hour incubation at room temperature with gentle horizontal agitation at 30 to 50 revolutions per minute. At the end of the incubation, allow the 3D structures to sediment again before carefully washing the 3D structures two times in one milliliter of 0.1%PBS supplemented with BSA for three minutes per wash.
After the last wash, label the 3D structures with 250 microliters of primary antibody in PBS for two to three days with gentle horizontal agitation at four degrees Celsius. At the end of the incubation, wash the sample with five three minute and two 15 minute washes in one milliliter of 0.1%PBS BSA per wash. After the last wash, label the 3D structures with 250 microliters of the appropriate secondary antibody in PBS for 24 hours at four degrees Celsius with gentle horizontal agitation protected from light.
The next day, label the 3D structures with 250 microliters of the appropriate concentration of Hoechst 3342 for a two hour incubation at four degrees Celsius with gentle horizontal agitation. At the end of the incubation, wash the 3D structures five times for three minutes and two times for 15 minutes in one milliliter of PBS per wash as demonstrated. After the last wash, transfer the 3D structures to a 96 well black polystyrene microplate, and gently add 200 microliters of clearing solution onto the structures, avoiding bubbles.
Then place the plate covered with aluminum foil at four degrees Celsius for at least 48 hours. To embed large 3D cell culture models, use a BSA coated one milliliter pipette tip to carefully transfer the 3D structures to a flat bottom glass tube with a PTFE lined bottle cap. After the structures have settled, carefully replace the PBS with 70%ethanol for a 30 minute incubation at room temperature.
At the end of the incubation, replace the 70%ethanol with one milliliter of ready to use eosin Y solution. Flick the tube and stain the structures for at least 30 minutes. At the end of incubation, dehydrate the structures with three successive 30 minute washes in one milliliter of 100%ethanol per wash.
After the last dehydration, clear the 3D structures with three successive 30 minute washes in one milliliter of xylene per wash under a chemical hood. After the last xylene immersion, place a piece of xylene soaked biopsy pad into one compartment of a white microtwin tissue cassette, and use a BSA coated two milliliter plastic Pasteur pipette to transfer the 3D structures onto the pad. Cover the 3D structures with another xylene soaked biopsy pad to hold the 3D structures in place and close the cassette.
When all samples have been transferred, place the cassettes in a 65 degrees Celsius paraffin bath for 30 minutes before placing the cassettes in a fresh 65 degrees Celsius paraffin bath overnight. The next morning, add liquid paraffin to one 65 degrees Celsius warmed embedding mold per sample, and place one paraffin embedded sample into each mold. Gently agitate the molds until all of the 3D structures drop to the bottoms, and use 65 degrees Celsius warmed fine forceps to position the 3D structures in the center of each mold.
Chill the mold until the paraffin forms a thin solid layer, securing the 3D structure in place. Place a tissue cassette and add hot paraffin over the mold. Remove the mold when the paraffin has completely solidified.
To embed small 3D cell culture models, gently wash the 3D structures three times in one milliliter of TBS per wash. After the last wash, remove as much TBS as possible without touching the structures, and add two drops of reagent two from a commercial paraffin embedding kit. Mix gently by tapping, and add two drops of reagent one with tapping until the gel solidifies.
Use fine forceps to transfer the gel from the well of the preassembled cassette from the kit, and dehydrate the gel embedded sample with 30 minute ascending ethanol and xylene incubations in a fume hood as indicated. After the last xylene immersion, place the cassette in a 65 degrees Celsius paraffin bath overnight before using fine forceps to transfer the paraffin embedded sample into the center of a liquid paraffin loaded 65 degrees Celsius warmed embedding mold. Then secure the 3D structure with one more layer of paraffin as demonstrated for the large 3D cell culture models.
Three-dimensional whole mount staining and confocal microscopy provide visual information about the cellular composition and spatial position of the 3D culture models to a field of depth of up to 200 microns. The high resolution that can be obtained for 3D structures using this protocol allows the application of cellular and subcellular segmentation algorithms for quantification of the cell number and the detection of various cell markers within different cell subtypes. 3D whole mounts analysis can be applied to visualized cells up to 200 micron in depth, regardless of the whole structures.
On the contrary, 2D section analysis can provide insights into cells of any size.
