All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.
1. Preparation of Instruments, Solutions, and Media
Note: All information about materials, reagents, tools, and instruments reported in the protocol are specified in the Table of Materials.
- Autoclave all dissection instruments and store them in a sterile area. Use the following instruments: 4 standard forceps (2 long and 2 short) and 2 scissors, as well as 2 forceps (1 long and 1 short) and 1 scissor for the dissection; keep an extra set as a backup.
- Prepare 100 mL of sterile phosphate-buffered saline (PBS)/1% fetal bovine serum (FBS) solution to use during washes, immunolabeling procedures, and cell sorting steps. Keep the solution chilled at 4 °C.
Note: Do not add sodium azide (NaN3) to the solution, as it can be toxic to live cells.
- Prepare 100 mL of PBS/1% FBS with 99 mL of PBS and 1 mL of FBS.
- Prepare 100 mL of sterile neural cell medium supplemented with 3% FBS (see Table of Materials) for use as collection and culture medium. Keep cell culture medium sterile at 4 °C. Only warm it to room temperature (RT) prior to use.
- Prepare 100 mL of neural cell medium supplemented with 3% FBS using 97 mL of neural cell medium and 3 mL of FBS.
- Pre-chill collection tubes (15-mL tubes) pre-coated with 5 mL of collection medium by placing them in an ice bucket. Only use polypropylene tubes to prevent the cells from adhering to the tube surface.
- Place 40-mm dishes, 70-µm nylon strainers, syringes, pestles for the cell strainer, and sterile polypropylene tubes in the biosafety cabinet. Sterilize all items prior to the procedures and maintain them in a sterile manner throughout the procedures.
Note: All steps after the collection of the retinae will be performed in the biosafety cabinet.
2. Preparation of the Retinal Cell Suspension
- Place the collected eye on the base plate of a dissection microscope to begin the corneal dissection. Dissect each eye individually.
- Carefully hold the globe at the optic nerve base using forceps. For this step, use one long and one short standard forceps.
- Use a 30-gauge (30G) sharp needle to puncture the cornea. This will allow the aqueous humor to evacuate from the eye, making it easier to hold the eye with the forceps.
- Hold the cornea with the forceps and use scissors to make a small incision in the cornea. Gently peel the cornea and sclera using the forceps. When the globe is peeled halfway, roll the retina and lens out using the forceps. Discard the cornea, sclera, and lens.
Note: This method ensures that the retina completely detaches from the remainder of the eye.
- Place the retina in a small 40-mm Petri dish containing PBS/1% FBS.
Note: As an alternative to the Petri dish, a cell culture dish can be used. Make sure to always keep the retinae moist, as in physiological conditions.
- Wash each retina three times with fresh sterile PBS/1% FBS within the biosafety cabinet.
- Place up to 12 retinae on a sterile 70-µm nylon strainer moistened with PBS/1% FBS. Using the back end of a 10-mL syringe, gently macerate the retinae using a circular movement to detach the cells.
- Alternatively, use a pestle for the cell strainer maceration or use enzymatic digestion with a combination of 15 IU/mL papain, 5 mM L-cysteine, and 200 U/mL DNase I for 15 min at 37 °C, followed by inactivation with PBS/10% FBS.
Note: No changes in the percentage of cells recovered were observed when enzymatic digestion was compared to maceration with either the back end of the syringe or the pestle.
- To transfer the isolated cells, place the sterile 70-µm nylon strainer over the polypropylene collection tube. Pass the collected cells through the strainer using a P1000 pipette. Rinse the strainer (step 3.6) with PBS/1% FBS to release any remaining cells and transfer them to the collection tube.
- Add PBS/1% FBS to achieve a final volume of 1 mL per retina. Centrifuge the cell suspension for 7 min at 200 x g and RT.
Note: Depending on the number of macerated retinae (< 6 retinae), the cell pellet may be too small to be visible.
- Discard the cell supernatant and resuspend the cell pellet in PBS/1% FBS using a ratio of 1 mL per 5 retinae.
- Count the cells using a hemocytometer.
- Clean a glass hemocytometer and coverslip with 70% alcohol. Gently swirl the tube containing the cells to ensure that the retina cell suspension is evenly distributed. Place 20 µL of 0.4% trypan blue in a microcentrifuge tube and mix with 20 µL of the cell suspension.
- Mix gently and apply 10 µL of the 0.4% trypan blue/cell suspension mix to the hemocytometer, filling both chambers; capillary action will draw the 0.4% trypan blue/cell suspension mix under the coverslip. Using a microscope, count all trypan blue-negative cells; this number represents the live cells.
- Determine cell viability using the following formula: live cell count / total cell count = % viability.
Note: If the viability of the cells is less than 95%, a fluorescent dye is required to discriminate cell viability during fluorescence-activated cell sorting (FACS).
- Use a fluorescent viability dye that is non-permeant to live cells. Wash the cells with PBS to remove any trace of serum. Add 1 µL of fluorescent dye for cell viability discrimination per 5.0 x 106 cells in a 100-µL final volume. Incubate at RT for 15 min, protected from light. Add 10x the volume of PBS/1% FBS. Centrifuge for 5 min at 200 x g and RT.
- If necessary, incubate the cell suspension overnight at 4 °C. If this is done, fill the cell suspension tube with neural cell medium rather than PBS/1% FBS. Place the tube horizontally and keep it at 4 °C.
3. Immunolabeling the Retinal Cells
- Use the following conversion to determine the volume of antibody per cell number: 2 µL of antibody per 5.0 x 106 cells in a 100-µL volume.
- Wash the cells and maintain them in PBS/1% FBS. Take a small aliquot of cells (5.0 x 106 cells) to use as a negative control (unlabeled) at the time of the sort set up.
Note: This negative control is critical for the proper calibration of the cell sorter.
- To minimize the non-specific binding of antibodies to cells that express the Fcγ receptors II and III, add 1 µL of an anti-mouse CD16/32 antibody per 1.0 x 106 cells in a 50-µL final volume using PBS/1% FBS. Incubate for 10 min at RT.
- Add the antibody cocktail to the sample and mix gently by pipetting. Incubate for 30 min in an ice bucket. Ensure that the ice bucket is covered, as light could compromise the experiment due to the photobleaching of the tagged fluorophores.
- Prepare the antibody cocktail using the following fluorescently tagged anti-mouse antibodies: CD90.2 AF (Alexa fluor)-700, CD48 PE (Phycoerythrin)-Cyanine7, CD15 PE, and un-tagged CD57.
Note: For each antibody, use the following concentrations per 5.0 x 106 cells: CD90.2 AF700, 1 µg; CD48 PE-Cyanine7, 0.4 µg; CD15 PE, 0.02 µg; and un-tagged CD57, 0.4 µg.
- Use volumes as per the following calculations (based on the commercially available antibodies listed in the Table of Materials): total of 5.0 x 107 cells to be labeled; 2 µL per 5.0 x 106 cells = 20 µL of each antibody; 4 different antibodies = 80 µL of antibody cocktail; final volume = [(5.0 x 107 total cells)/5.0 x 106 cells] x 100 = 1,000 µL; 80 µL Abs + 50 µL of anti-mouse CD16/32 + 870 µL PBS/1% FBS = 1,000 µL.
Note: This combination provided the optimal combination of fluorochromes for the instrument configuration. The following parameters summarize the emission and excitation: AF-700, emission of 719 nm when excited with a 638-nm red diode laser; PE-Cyanine7, emission of 767 nm when excited with a 488-nm blue diode laser; and PE, emission of 575 nm when excited with a 488-nm blue diode laser.
- After the 30-min incubation, bring volume up to 5 mL using PBS/1% FBS. Wash the samples by centrifuging them for 7 min at 200 x g and RT. Repeat the procedure once to remove all unbound antibodies.
- Add 2 µL of secondary antibody (0.1 µg), which will bind the CD57 of 5.0 x 106 cells. Incubate for 30 minutes in an ice bucket, as in step 3.4. Wash the cells twice, as in step 3.5.
- Label the cells with a secondary antibody if an un-tagged primary antibody was used in step 3.4.
Note: The secondary antibody of choice for this configuration is listed in the Table of Materials; it has an emission wavelength of 421 nm. Use it with a bandpass filter of 450/50 nm when excited with the violet diode laser at 405 nm.
- Use volumes as per the following calculations (based on the commercially available antibody listed in the Table of Materials): 2 µL per 5.0 x 106 cells = 20 µL of secondary antibody; final volume = [(5.0 x 107 total cells)/5.0 x 106 cells] x 50 = 500 µL; 20 µL of Abs + 480 µL PBS/1% FBS = 500 µL.
- Keep the labeled cells in PBS/1% FBS. Count the cells using a hemocytometer; use a final retinal cell concentration of 3.0 - 4.0 x 107 cells/mL.
Note: Do not use cell culture medium to dilute the cells because the phenol red can increase the autofluorescence, thereby reducing the resolution between negative and positive cells. The final cell concentration per volume highly depends upon the volume flow rate of the instrument configuration.
- Use single-color controls during setup to minimize fluorescence spillover.
Note: This step, also known as compensation, is applied to correct the noise created by the combination of fluorophores. Polystyrene microspheres in PBS/0.1% BSA/2 mM NaN3 with the capacity to bind fluorescently tagged immunoglobulin (Ig) isotypes from multiple species provide positive controls to set the compensation.
- Place 3 drops of the polystyrene microspheres into sterile FACS tubes, allotting one per fluorophore. Add 1 µg of the respective fluorophore to each tube. Incubate for 15 min at RT, protected from light. Add 3 mL of PBS/1% FBS to the sample tubes. Centrifuge for 5 min at 200 x g and RT. Carefully remove the supernatant and resuspend in 250 µL of PBS/1% FBS.
- To set up the negative controls, use polystyrene microspheres that do not bind to Ig. If necessary, carry out this step the day before.