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Method Article

Isolation of Primary Murine Retinal Ganglion Cells (RGCs) by Flow Cytometry

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DOI:

10.3791/55785

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July 5th, 2017

 ,  ,  ,  ,  ,  ,  , 
1Department of Ophthalmology, Hamilton Eye Institute, University of Tennessee Health Science Center, 2Department of Microbiology, Immunology and Biochemistry, University of Tennessee Health Science Center, 3Department of Anatomy and Neurobiology, University of Tennessee Health Science Center, 4Department of Pharmaceutical Sciences, University of Tennessee Health Science Center

In This Article

Summary

Millions of people suffer from retinal degenerative diseases that result in irreversible blindness. A common element of many of these diseases is the loss of retinal ganglion cells (RGCs). This detailed protocol describes the isolation of primary murine RGCs by positive and negative selection with flow cytometry.

Abstract

Neurodegenerative diseases often have a devastating impact on those affected. Retinal ganglion cell (RGC) loss is implicated in an array of diseases, including diabetic retinopathy and glaucoma, in addition to normal aging. Despite their importance, RGCs have been extremely difficult to study until now due in part to the fact that they comprise only a small percentage of the wide variety of cells in the retina. In addition, current isolation methods use intracellular markers to identify RGCs, which produce non-viable cells. These techniques also involve lengthy isolation protocols, so there is a lack of practical, standardized, and dependable methods to obtain and isolate RGCs. This work describes an efficient, comprehensive, and reliable method to isolate primary RGCs from mice retinae using a protocol based on both positive and negative selection criteria. The presented methods allow for the future study of RGCs, with the goal of better understanding the major decline in visual acuity that results from the loss of functional RGCs in neurodegenerative diseases.

Introduction

RGCs are terminally differentiated neurons, and therefore, primary cells are required for experimentation. The development of a protocol for the isolation and enrichment of primary murine retinal ganglion cells (RGCs) is fundamental to revealing the mechanisms of RGC health and degeneration in vitro. This is especially important for studies that seek to generate potential therapies to promote RGC function and to minimize their death. The degeneration of RGCs is associated with retinal degenerative diseases, such as glaucoma, diabetic retinopathy, and normal aging. Although the specific cellular mechanisms underlying RGC loss are unclear, a series of risk factors have been identified. Lack of oxygenation at the optic nerve head1,2,3 causes RGC death4 and acts as the disturbance of the homeostasis between the activation of excitatory and inhibitory receptors within individual RGCs5,6. A series of challenges impede progress towards the use of these cells for in-depth studies. First, the number of RGCs present in a murine retina is small. RGCs account for less than 1% of total retinal cells7,8,9. Second, most RGC-specific markers are intracellular proteins10,11,12. Selection based upon these markers leaves the cells non-viable, which precludes downstream functional analyses. Finally, currently available protocols are lengthy and lack standardization13,14. Early RGC isolation protocols were based on immunopanning methods. Barres et al.15 adapted the classic immunopanning technique and added a second step, which excluded monocytes and endothelial cells from the bulk of retinal cells prior to positive selection based upon immunopositivity to anti-thymocyte antigen (aka Thy1), a cell-surface marker. Years later, Hong et al. combined magnetic bead isolation techniques with cell sorting strategies to isolate RGCs with higher purity16. The use of magnetic beads is still used in many scientific applications. Together, magnetic beads and flow cytometry protocols improved the purity of isolated cells. However, these purification systems have not yet been standardized for the isolation of murine RGCs from dissociated retinae.

Flow cytometry is a powerful analytical method that measures the optical and fluorescence characteristics of cell suspensions. Cells are analyzed both quantitatively and qualitatively with a high level of sensitivity, providing a multi-dimensional analysis of the cell population. Cellular discrimination is based upon two main physical properties: cell size or surface area and granularity or internal complexity17. A multi-dimensional analysis can be performed by combining antibodies tagged with fluorochromes that have similar excitation wavelengths and different emissions. Flow cytometry is fast, reproducible, and sensitive. Multitpe lasers permit even greater multi-dimensional analyses of single cells by flow cytometry. Thus, it is an attractive methodology for the study of cytological specimens. Fluorescence activated cell sorting (FACS) uses the multi-dimensional phenotypic differences identified by flow cytometry to sort individual cells into distinct subpopulations.

In the last decade, multiple surface and intracellular proteins have been identified as potential biomarkers for the selection of cells, including neurons. Initial studies that sought to isolate RGCs from rats used Thy1 as a ganglion cell marker. Unfortunately, Thy1, aka CD90, has multiple isoforms in other rodent species18,19,20 and is expressed by multiple retinal cell types19,20, making it a non-specific marker for RGCs. Another surface marker, CD48, is found on monocytic populations in the retina, including macrophages and microglia. Using these two surface markers, a modified RGC signature-Thy1+ and CD48neg cells-was developed15,16,21,22. Unfortunately, these two selection criteria are not sufficient to select for a highly enriched RGC population. To address this unmet need, a flow cytometry protocol was developed23 based on multi-layered positive and negative selection criteria using known cell surface markers to enrich and purify primary murine RGCs.

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Protocol

All procedures detailed in the following protocol were approved by the Institutional Animal Care and Use Committee (IACUC) review board at the University of Tennessee Health Science Center (UTHSC) and followed the Association for Research in Vision and Ophthalmology (ARVO) Statements for the Use of Animals in Ophthalmic and Vision Research, in addition to the guidelines for laboratory animal experiments (Institute of Laboratory Animal Resources, Public Health Service Policy on Humane Care and Use of Laboratory Animals).

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.

  1. 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.
  2. Prepare 100 mL of sterile PBS/1% 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.
    1. Prepare 100 mL of PBS/1% FBS with 99 mL of PBS and 1 mL of FBS.
  3. Prepare 100 mL of sterile neural cell medium supplemented with 3% FBS (see the 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.
    1. Prepare 100 mL of neural cell medium supplemented with 3% FBS using 97 mL of neural cell medium and 3 mL of FBS.
  4. 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.
  5. 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. Enucleation

Note: A total of 10 young (5-7 weeks old) C57BL/6J mice were used in this experiment to isolate 1.0 x 106 RGCs with the phenotype CD90.2+CD48negCD15negCD57neg.

  1. Euthanize the mice using CO2 inhalation followed by cervical dislocation. Always use a secondary euthanasia method to ensure that the euthanized animal is dead.
    Note: Isofluorane can be used as an alternative to CO2. Ketamine is not recommended, as it is associated with an increase in the intraocular pressure (IOP) when used as an inducer of anesthesia24,25.
  2. Insert forceps under the globe of the eye, grip the optic nerve, and pull up; the globe will be enucleated, with the optic nerve intact. Place collected eyes in a vial containing PBS and keep the vial on ice until the next step.
    Note: Any personnel can carry out this step after receiving the appropriate training from the Laboratory Animal Care Unit (LACU).

3. Preparation of the Retinal Cell Suspension

  1. Place the collected eye on the base plate of a dissection microscope to begin the corneal dissection. Dissect each eye individually.
  2. Carefully hold the globe at the optic nerve base using forceps. For this step, use one long, and one short standard forceps.
  3. Use a 30-gauge (30G) sharp needle to puncture the cornea to allow the aqueous humor to evacuate from the eye, making it easier to hold the eye with the forceps.
  4. 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.
  5. 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.
    1. Wash each retina three times with fresh sterile PBS/1% FBS within the biosafety cabinet.
  6. 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.
    1. 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.
  7. 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.
  8. 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.
    1. Discard the cell supernatant and resuspend the cell pellet in PBS/1% FBS using a ratio of 1 mL per 5 retinae.
  9. Count the cells using a hemocytometer.
    1. 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.
    2. 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.
    3. 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%, the use of fluorescent dye for cell viability discrimination is required during FACS.
  10. 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.
  11. Incubate the cell suspension overnight at 4 °C, if necessary. If this is done, fill up the cell suspension tube with neural cell medium rather than PBS/1% FBS. Place the tube horizontally and keep it at 4 °C.

4. Immunolabeling the Retinal Cells

  1. 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.
  2. 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 setup.
    Note: This negative control is critical for the proper calibration of the cell sorter.
  3. 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.
  4. 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.
    1. Prepare the antibody cocktail using the following fluorescently tagged anti-mouse antibodies: CD90.2 AF-700, CD48 PE-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.
      1. 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.
  5. 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 antibody.
  6. Add 2 µL of secondary antibody (0.1 µg), which will bind the CD57 of 5.0 x 106 cells. Incubate for 30 min in an ice bucket, as in step 4.4. Wash the cells twice, as in step 4.5.
    1. Label the cells with secondary antibody if an un-tagged primary antibody was used in step 4.4.
      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.
    2. 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.
  7. 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.
  8. 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.
    1. 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.
    2. To set up the negative controls, use polystyrene microspheres that do not bind to Ig. Carry out this step the day before, if necessary.

5. Cell Sorting Strategy

Note: Specific instructions for instrument setup for FACS with 355 nm, UV; 405 nm, violet; 488 nm, blue; and 640 nm, red lasers with 2, 2, 5, and 3 fluorescence channel distribution, respectively. The operating software was DIVA version 8.0.1. Cell sorting was performed with a 70 μm nozzle, 70 psi sheath pressure, 87.5 frequency, 48.6 amplitude with first drop breakoff at 333, gap asetting of 6, sort precision set to four-way purity with default (32) purity mask, and drop delay adjusted to 42.98 using beads.

  1. Use 15-mL polypropylene conical tubes as collection vessels. Add 5 mL of the collection medium (cell culture medium) to each tube and rotate the tube to coat the walls.
    Note: This step prevents the cells from sticking to the sides of the collection tube. As an alternative, tube can be coated with undiluted FBS.
  2. Take into consideration the efficiency of the cell sorter to estimate the final yield of cells.
    Note: The efficiency of the sort varies based on the flow cytometer used. RGCs with the CD90.2+CD48negCD15negCD57neg phenotype comprise about 1% of all retinal cells.
  3. Have the cell sorting performed by a trained operator (usually in a core facility) . Ensure that the instrument is cleaned and sterilized with 70% ethanol prior to sample acquisition. Carefully wipe the outside of collection tubes with 70% ethanol as well.
  4. If the cells have settled or aggregated, pipette the cells up and down and filter the suspension through a sterile 40-µm nylon strainer prior to acquisition. Make sure to control the temperature and keep it at 4 °C while the cell sorting is being performed; failure to do so will reduce the cell yield.
  5. Discuss the details of the sort with the cell sorter operator prior to performing the experiments.
    Note: At the time of experiment, the cell sorter operator will have clicked a series of buttons from the Workspace toolbar of the acquisition software for the cytometer. These are the Browser, Cytometer, Inspector, Worksheet, and Acquisition Dashboard. The strategy of cell sorting used here is known as a 2-way sort. Two populations are collected: the enriched RGCs and all other cell types. If additional populations are to be collected, up to two additional populations can be isolated as a 4-way sort. If additional populations will be collected, the cell sorter requires different collection tubes and setup.
  6. Perform compensation to correct for spectral overlap from the fluorophores.
    Note: This process can be done manually by adjusting each setting, or it can be done automatically as part of the software in use. Most software for cell sorting equipment have the option of an automated compensation. It is considered the gold standard and the most accurate type of compensation. The negative (no fluorophore) sample and the single controls prepared with polysterene microspheres (Compensation beads) specifically manufactured to bind all relevant monoclonal antibodies used in the experiment will be used in this step.
    1. Select Experiment > Compensation Setup > Create Compensation Controls. Add the fluorophore-specific controls from the list displayed in the screen. Click OK.
      Note: A compensation tube for each of the controls will be displayed.
      1. Use an unlabeled control (no fluorophore, step 4.2) to verify the forward-scattered light (FSC), the side-scattered light (SSC), and to gate the initial population (P1).
    2. Install the negative control tube onto the cytometer and click Load. Verify that the population of interest is displayed and select it. This is population 1 (P1). Right-click the P1 gate and select Apply to All Compensation Controls. Click Record Data. Once the recording is done, click Unload and remove the tube.
    3. Install the next tube onto the cytometer, as displayed on the screen. Repeat step 5.6.2 until all data from the controls have been recorded.
    4. Select Experiment > Compensation Setup > Calculate Compensation. Save the setup and name the experiment. Click Link & Save.
      Note: Compensation is a necessary step, as it removes the spectrum overlap between the detectors.
    5. If manual compensation is needed, do so by adjusting the means of the positive signals so that they equal the negatives for each of the fluorochromes used.
      Note: This step is done by the cell sorter operation and may take 30 min.
  7. Gating strategy ( Figure 3A).
    1. Set up P1 by plotting FSC versus SSC; the FSC is indicative of the size and the SSC indicates the internal complexity of the cells. See Figure 3A.
    2. Draw a pseudocolor plot of SSC-H (height) versus SSC-W (width) using the selected population in step 5.7.1 (P1); the pseudocolor plot allows for the visual representation of the density of cells relative to one another.
      Note: A lower cell density is represented by blue and green, whereas red and orange areas represent high cell density.
      1. Perform this step to collect single cells only; the single cell gate is called P2. See Figure 3A.
    3. Repeat step 5.7.2 to plot FSC-H versus FSC-W using P2. Ensure that the selection of single cells as "doublets" or cell clumps contains both a positive and a negative marker, providing false positivity.
    4. Draw a pseudocolor plot of CD90.2 versus CD48. Select all CD90.2+CD48neg cells to eliminate monocytes from the RGC enrichment; call this gate CD90.2+CD48neg, or P3. See Figure 3A.
    5. Using the selected CD90.2+CD48neg population or P3 gate, draw a pseudocolor plot of CD57 versus CD15 to eliminate amacrine cell contaminants from the RGC enrichment. Gate the CD15negCD57neg population. See Figure 3A.
  8. Define the populations to be collected for the sample in the "Sort Layout" window and proceed with the cell sorting; the sample to collect is CD90.2+CD48negCD15negCD57neg.
    Note: The population to be sorted is selected from the drop-down Add menu in the sort window. After adding the population, it will ask for target events or how many events are to be collected. At any time, the sort layout can be edited by clicking the Sort Location field containing the population. 0.9% ± 0.3 of RGCs with the phenotype CD90.2+CD48negCD15negCD57neg are obtained from young (5-7 weeks old) and 0.5% ± 0.3 and old (>12 months old) C57BL/6J mice23.
  9. Using 25,000 cells, perform a purity check23. See Figure 3B-E.
    Note: The purity check is the process to re-analyze the sorted cells to verify the accuracy of the cell sort. A small aliquot of sorted cells will be loaded onto the cytometer to verify the effectiveness of the sort.

6. Confirmation on RGC Intracellular Markers

  1. Intracellular labeling.
    1. Fix sorted cells for 1 h and permeabilize at 4 °C to make the cells metabolically inactive and to allow the penetration of intracellular antibodies.
    2. Dilute the following antibodies in the permeabilization solution: anti-RNA binding protein with multiple splicing (RBPMS) at a 1:100 dilution in a 100 µg/mL stock; anti-synuclein gamma (SNCG), 1:100 in a 1 mg/mL stock; brain-specific homeobox/POU domain protein 3A (BRN3A), 1:100 in a 200 µg/mL stock; and anti-neuron-specific class III beta tubulin (TUJ1), 1:100 in a 1 mg/mL stock. Incubate the cells and antibody solutions for 1 h in a covered ice bucket.
    3. Wash the samples twice with PBS/1% FBS.
    4. Resuspend the cells in the appropriate AF488-tagged secondary antibody at a 1:200 dilution for 30 min in an ice bucket.
    5. Wash the samples as in step 6.1.3. Keep the cells in PBS/1% FBS (250 µL) until ready to analyze.

7. Validation of the Cell Sort by qPCR Analysis

Note: See Figure 4.

  1. RNA extraction23.
    1. Extract the RNA from 5.0 x 105 sorted cells by cell lysis and homogenization followed by the addition of chloroform.
    2. Transfer the upper, colorless phase to a clean microtube for alcohol precipitation followed by extract concentration in a spin column. Perform DNAse digestion on the column.
    3. Wash the column with RNase-free water for RNA elution.
    4. Assess the RNA concentration by spectrophotometry.
  2. cDNA synthesis and pre-amplification.
    1. Use 100 ng of RNA material and mix with a solution containing reverse transcriptase enzyme, recombinant ribonuclease inhibitor (to avoid RNA degradation), magnesium chloride (MgCl2), and deoxynucleotides.
    2. Mix the tube contents and incubate for 10 min at 25 °C followed by 60 min at 42 °C. Finish the reaction with a 5-min incubation at 85 °C. Store the resulting cDNA at -20 °C until ready to use.
      Note: cDNA material can be stored at -20 °C for up to a month if the pre-amplification step cannot be carried out immediately.
  3. Pre-amplification of cDNA.
    1. Pre-amplify cDNA material using a series of primers specific for multiple retinal cell types including: retinal ganglion cells, amacrine cells, astrocytes, Müller, bipolar, horizontal, photoreceptors, and retinal pigment epithelial cells, as detailed in Reference 23 and in the Table of Materials.
      Note: The pre-amplification reaction is prepared to increase the sensitivity of detection for quantification. The pre-amplification reaction contains a mixture of the primer mixes from the Table of Materials; 2.5 µL of cDNA, which started at 100 ng of RNA; reverse transcriptase enzyme; and nuclease-free water in a final volume of 10 µL.
    2. Carry out the enzyme activation step at 95 °C for 10 min, followed by 14 cycles at 95 °C for 15 s, followed by 60 °C for 4 min. Dilute the pre-amplified material 1:10 in Tris EDTA buffer and keep it at -20 °C until ready for use.
  4. 7.4 qPCR reaction.
    1. Prepare all qPCR reactions in a 10 µL final volume using the diluted, pre-amplified cDNA (2.5 µL), primers (listed in the Table of Materials), nuclease-free water, and a concentrate with DNA polymerase and deoxynucleotides.
    2. Use the following instrument conditions to run the qPCR: a hold step of 50 °C for 2 min, followed by 95 °C for 10 min. Run a total of 40 cycles at 95 °C for 15 s followed by 60 °C for 1 min. Perform all measurements in replicates of three.
    3. Perform relative quantification using the comparative threshold (CT) after determining the values of CT for the housekeeping gene and the target genes in each sample. Calculate the relative fold change (Rq) using the following equation: Rq = 2T –∆C, where ∆CT = CT target gene – CT reference gene23.
      Note: The CT is defined as the PCR cycle at which the fluorescent signal of the reporter dye crosses an arbitrarily placed threshold26. The CT is inversely related to the amount of amplicon (PCR product) in the reaction.

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Results

The in-depth study of RGCs is impeded by many factors, including their low frequency and the lack of a robust and standardized methodology for their isolation. Figure 1 shows the methodology used for retinae isolation. Variations in the enucleation procedure exist based on the type of analysis, such as if the enucleation is part of in vivo experimentation27. Enucleation in this protocol is performed on euthanized mice. As show...

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Discussion

FACS is the technique of choice to purify cell populations. Other isolation methods include immunopanning, magnetic beads, and complement fixation depletion. The advantage of FACS over these other methodologies is based on the simultaneous identification of cell-surface markers with varying degrees of intensity. The fluorescent intensity of the molecule is proportional to the amount of protein expression. Until now, the isolation of RGCs was based solely on Thy1 (CD90) positivity and CD48 negativity15

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Disclosures

The authors declare that they have no competing financial interests.

Acknowledgements

The authors would like to thank Mr. Tim Higgins, Senior Illustrator from the Department of Microbiology, Immunology and Biochemistry, for technical video assistance; Dr. Matthew W. Wilson for discussions and the members of the Jablonski and Morales-Tirado laboratories for their helpful comments. This work was supported by the Alcon Research Institute Young Investigator Award (VMM-T), the University of Tennessee Research Foundation (VMM-T), the National Eye Institute EY021200 (MMJ), the Gerwin Fellowship (VMM-T); the Gerwin Pre-doctoral Fellowship (ZKG), the Department of Defense Army Medical Research and Materiel Command (VMM-T), and the Unrestricted Grant from Research to Prevent Blindness.

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Anti-mouse CD15 PEBioLegend125606Clone MC-480
Anti-mouse CD48 PE-Cy7BioLegend103424Clone HM48-1
Anti-mouse CD57Sigma AldrichC6680-100TSTClone VC1.1
Anti-mouse CD90.2 AF700BioLegend105320Clone 30-H12
Brilliant Violet 421 Goat Anti-mouse IgGBioLegend405317Clone Poly4053
Purified Anti-mouse CD16/32BioLegend101302FcgRII/III block, Clone 93
Zombie Aqua BioLegend423102Live cell/ Dead cell discrimination
Fetal Bovine SerumHycloneSH30071.03U.S. origin
AbC Total Antibody Compensation Bead KitThermo Fisher ScientificA10497Multi-species Ig
Neurobasal MediumThermo Fisher Scientific21103049Add serum to media prior to culture.
Phosphate-Buffered Saline (PBS)Thermo Fisher Scientific10010049Saline solution
Dissection MicroscopeOlympusSZ-PT ModelStereo Microscope
Sorvall CentrifugeThermo ScientificST 16RAll centrifugation performed  at RT
Base Plate – Dissection PanFisher ScientificSB15233FIMA wax plate can also be used
ForcepsAesculap5002-74 ½ inches
Iris Scissors, StraightAesculap13605 ½ inches
Falcon 15 mL conical tubesFisher Scientific352097Polypropylene tubes
Falcon 50 mL conical tubesFisher Scientific352098Polypropylene tubes
BD FACS TubesFisher Scientific352003Polypropylene tubes
40 mm dishesMidSciTP93040Tissue culture treated
70 μm nylon strainerMidSci70ICSsterile
40 μm nylon strainerMidSci40ICSsterile
 BD 10 mL syringeFisher Scientific301604Disposable Syringe without needle
PestlesMidSciPESTsterile
Wheaton VialsFisher Scientific986734No Liner
BD 30 G needleFisher Scientific3051281 inch
Hausser Scientific Bright-Line Glass Counting ChamberFisher Scientific0267151BHemocytometer
Gibco Trypan blue 0.4% SolutionFisher Scientific15250061Viability Dye
Eppendorf tubesFisher Scientific05-402-251.5mL
EVOS Floid Cell ImagingThermo Fisher Scientific447113Fluorescence Imaging with a 20X objective
100% EthanolFisher Scientific04-355-452Used to make 70% Ethanol
Pipet-Lite LTS Pipette L-1000XLS+Rainin17014282LTS Pipette
Pipet-Lite LTS Pipette L-200XLS+Rainin17014391LTS Pipette
Pipet-Lite LTS Pipette L-20XLS+Rainin17014392LTS Pipette
Rack LTS 1000 mL – GPS-L1000SRainin17005088Blue Rack Sterile Tips
Rack LTS 250 mL – GPS-L250SRainin17005092Green Rack Sterile Tips
Rack LTS 20 mL – GPS-L10SRainin17005090Red Rack Sterile Tips
FACSAria II Cell SorterBD BiosciencesN/ACustom order
LSR II CytometerBD BiosciencesN/ACustom order
Abca8aThermo Fisher ScientificMm00462440_m1Müller cells
Aldh1alThermo Fisher ScientificMm00657317_m1Müller cells
Aqp4Thermo Fisher ScientificMm00802131_m1Astrocytes
Calb2Thermo Fisher ScientificMm00801461_m1Amacrine, Horizontal
Cd68Thermo Fisher ScientificMm03047340_m1Retinal Pigment Epithelial Cells
Gad2Thermo Fisher ScientificMm00484623_m1Amacrine
HprtThermo Fisher ScientificMm01545399_m1House keeping gene
Lhx1Thermo Fisher ScientificMm01297482_m1Horizontal
Lim2Thermo Fisher ScientificMm00624623_m1Horizontal
NrlThermo Fisher ScientificMm00476550_m1Photoreceptors
Ntrk1Thermo Fisher ScientificMm01219406_m1Horizontal
Pcp4Thermo Fisher ScientificMm00500973_m1Bipolar, Amacrine
Pou4f1Thermo Fisher ScientificMm02343791_m1Retinal Ganglion Cells
Prdx6Thermo Fisher ScientificMm00725435_s1Astrocytes
PrkcaThermo Fisher ScientificMm00440858_m1Bipolar
Prox1Thermo Fisher ScientificMm00435969_m1Horizontal
PvalbThermo Fisher ScientificMm00443100_m1Amacrine
RbpmsThermo Fisher ScientificMm02343791_m1Retinal Ganglion Cells
Rom1Thermo Fisher ScientificMm00436364_g1Photoreceptors
Rpe65Thermo Fisher ScientificMm00504133_m1Retinal Pigment Epithelial cells
Slc1a3Thermo Fisher ScientificMm00600697_m1Astrocytes
Slc6a9Thermo Fisher ScientificMm00433662_m1Amacrine
SncgThermo Fisher ScientificMm00488345_m1Retinal Ganglion Cells
Tubb3Thermo Fisher ScientificMm00727586_s1Retinal Ganglion Cells
VimThermo Fisher ScientificMm01333430_m1Müller cells
Taqman Universal Master MixThermo Fisher Scientific4440047qPCR Reagent
miRNeasy Mini KitQiagen217004RNA Isolation
SuperScript VILO cDNA Synthesis KitThermo Fisher Scientific11754250cDNA synthesis
Taqman PreAmp Master MixThermo Fisher Scientific4391128Pre-Amplification step
BD Cytofix/ CytopermBD Biosciences554714Fixation/ Permeabilization Buffer
BD Perm/ WashBD Biosciences554723Permeabilization Solution
RBPMSSanta Cruz Biotechnologysc-86815intracellular antibody
SNCGGene TexGTX110483intracellular antibody
BRN3ASanta Cruz Biotechnologysc-8429intracellular antibody
TUJ1BioLegend801202intracellular antibody

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Cell IsolationPrimary Murine CellsCell SortingImmunolabelingCD90.2 PositiveCD48 NegativeCompensation ControlsViable Cell Population
Isolation of Primary Murine Retinal Ganglion Cells (RGCs) by Flow Cytometry
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