Single-cell RNA-Seq of Defined Subsets of Retinal Ganglion Cells

All procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at Northwestern University.

1. Preparation of Solutions for Electrophysiology (4 h)

1. Make 0.1% DEPC-treated H₂O by adding 1 mL of diethyl pyrocarbonate (DEPC) to 999 mL of reverse osmosis-purified H₂O. Mix thoroughly and let the mixture incubate for 1 h at room temperature (RT). Then, autoclave the DEPC-mixed H₂O for 15 min on a liquid cycle. Let the DEPC-treated H₂O cool at RT.
2. Make the extracellular solution by mixing one bottle of Ames' medium and 1.9 g (23 mM) of sodium bicarbonate into 1 L of H₂O. Bubble the extracellular solution with
   95% O₂/5% CO₂ and maintain it at a pH of 7.3-7.4.
3. Generate the intracellular solution by combining 125 mM K-gluconate, 2 mM MgCl₂,10 mM EGTA, 10 mM HEPES, and 0.1% DEPC-treated H₂O. Store it in 1 mL aliquots at -20 °C. Add 10 µM of fluorescent tracer at the beginning of each experiment.
4. Make enzyme solution by adding 10,000 units of collagenase and 83 mg of hyaluronidase to 4.15 mL of extracellular solution. The enzyme solution should be stored in 50 µL aliquots at -20 °C.

2. Preparation of Retinal Tissue (2 h)

NOTE: All procedures in this section should be performed under dim red illumination

1. Dark-adapt animals for at least 1 h prior to dissection. Perform all procedures under dim red illumination.
2. Euthanize the animals by CO₂ asphyxiation and enucleate the eyeballs into a Petri dish with previously oxygenated extracellular solution.
3. Poke the cornea with a needle and cut it away by cutting with ophthalmic scissors at the border of the cornea and sclera¹⁹.
4. Remove the lens using #5 forceps. Gently make a tear in the sclera with the forceps and sever the optic nerve where the retina and sclera meet. Carefully finish removing the sclera from the retina.
5. Remove the transparent vitreous using #5 forceps; once removed, the vitreous appears as a gelatinous substance stuck to the forceps. Slice the retinas in half (so that there are 4 pieces/animal) and store them in oxygenated extracellular solution at RT until use.
6. When ready to mount the tissue in the recording chamber, place a piece of retina to incubate in enzyme solution diluted in 500 µL of oxygenated extracellular solution. Incubate in a Petri dish for 2 min at RT on a shaker.
   1. Wash the piece of retina in oxygenated extracellular solution and place the tissue in a glass-bottom recording chamber; use a plastic transfer pipette with the tip cut off to allow for the retina to be transferred without causing damage to the tissue.
7. Use forceps to carefully flatten the tissue with the photoreceptor layer facing down. Remove excess fluid using a pipette. Anchor the tissue using a platinum ring with nylon mesh.
   NOTE: This method could also be used to prepare the tissue for the isolation of RNA from labeled amacrine and bipolar cells.
8. Fill the chamber with oxygenated extracellular solution and mount it onto a microscope stage. Perfuse tissue with oxygenated extracellular solution at 2-4 mL/min.

3. Visualization and Targeting of GFP+ Retinal Ganglion Cells (10 min)

NOTE: All procedures in this section should be performed under dim red illumination

1. Before beginning, pull glass micropipettes (OD: 1.2 mm, ID: 0.69 mm) for electrophysiological recordings using a micropipette puller. Use the following protocol for the electrodes (please note that the parameters should be adjusted accordingly to achieve the desired resistance and will vary across pullers and with different glass): Heat: Ramp +10; Pull: 0; Vel: 23; Delay: 1; Pressure: 500; Program loop: 5 times. Ensure that the tips are ~1 µm in diameter, with resistances of 2-4 MΩ for targeting large cells and of 5-7 MΩ for targeting smaller cells.
2. Observe the ganglion cell layer using Infrared Differential Interference Contrast (IR-DIC) optics (Figure 1A). Identify GFP+ Retinal Ganglion Cells (RGCs) using epifluorescence (~480 nm) (Figure 1B).
3. Locate the pipette filled with intracellular solution in DIC. Apply slight positive pressure and zero any voltage offsets on the amplifier.
4. Press the glass micropipette against a GFP+ cell and apply negative pressure to form a GΩ seal between the pipette and the cell membrane. Apply test voltage command steps (e.g., 5 mV) to monitor the seal resistance. After forming a stable seal, rupture the membrane by applying brief pulses of negative pressure to gain whole-cell access.
5. Wait 1-2 min for the dendrites of the cell to fill with fluorescent tracer.
   NOTE: The cell can be morphologically typed by examining the morphology in epifluorescence (Figure 1C). In the case of melanopsin-expressing RGCs, dendritic stratification in the inner plexiform layer is visualized by examining the dendrites filled with fluorescent tracer under epifluorescent illumination and determining whether they stratify far from the soma in the OFF sublamina (M1 ipRGCs), near the ganglion cell layer in the ON sublamina (M2 & M4 ipRGCs), or both (M3 ipRGCs). This observation, combined with soma size (M4s have distinctly large somas compared to all other ipRGC subtypes), allow for the identification of cell type^20,21,22. Thus, this technique allows for the identification of cell type in vitro prior to RNA isolation. This method could be modified for other cell type identification protocols involving either dendritic morphology or cellular physiology.

4. Cell Isolation (2 min)

1. Before beginning, set the tabletop microcentrifuge to 2,000 x g. Prepare a sample-expelling apparatus by connecting tubing (OD: 3/32 in, ID: 1/32 in) with a 1 cc syringe.
2. Place 0.2 mL PCR tubes containing 10 µL of lysis buffer and 1% β-mercaptoethanol on ice. Prepare a 1 cc syringe containing DEPC-treated H₂O to rinse the pipette tips. Prepare a container of dry ice to freeze the lysis buffer after sample collection.
3. Carefully extract the cytoplasmic content of the cell pipette by applying negative pressure using a 10 mL syringe; all cytoplasmic content, including organelles, should be extracted if possible.
   1. Monitor the extraction in DIC by visualizing the cell body decreasing in size. After extracting the cytoplasmic contents, lift the pipette carefully off of the tissue and quickly remove the pipette from the solution.
4. Quickly remove the pipette from the head-stage holder and rinse the pipette tip briefly with DEPC-treated H₂O using a 1 mL syringe. Connect the pipette to a 1 mL syringe via tight-fitting tubing to expel the sample.
5. Immediately expel the cells into 10 µL of lysis buffer 1 containing 1% β-mercaptoethanol in 0.2 mL PCR tubes.
   NOTE: The entire aspirate with the cells should be expelled gently so as to not introduce bubbles.
   1. Briefly centrifuge the tube in a tabletop mini centrifuge at 2,000 x g for 10 s. Immediately flash-freeze the samples for 5 min on dry ice. After freezing, store them at -80 °C for up to two weeks for best results; the samples may last longer, but it is recommended that they are processed as quickly as possible.

5. RNA Purification (30 min)

1. Before beginning, set up a magnetic separator device by taping the top part of an inverted P20 or P200 tip holder to the 96-well magnetic stand²³.
2. Prepare fresh 70% ethanol (EtOH) – approximately 1 mL per sample will suffice. Remove the RNA magnetic beads from 4 °C storage and thaw them at RT for at least 30 min.
   NOTE: No more than 8 samples should be processed at one time, as many steps in this protocol rely on efficiency and quick handling.
3. Once the magnetic beads are at RT, vortex for 30 s to ensure that the solution is well mixed.
   NOTE: The beads use an RNA-specific buffer to selectively bind RNA, and they allow for the removal of other cellular waste when employed with a magnetic plate stand.
4. Thaw cells at RT for 1 min, and then add 5 µL of RNase-free H₂O to each sample; pipette up and down. Add 22 µL of RNA beads to each tube and pipette thoroughly to mix. Incubate the samples at RT for 5 min to allow the RNA to interact and bind with the magnetic beads.
5. Place the tubes on a magnetic separator device and let sit for 8 min; before proceeding, ensure that the supernatant is clear. Observe the beads from one pellet and be sure to not detach it during pipetting.
6. Remove the supernatant from the samples and add 150 µL of 70% EtOH. Remove the EtOH and repeat the wash twice more.
7. Allow the samples to air dry for 6 min. Check intermittently to see if more EtOH has collected at the bottom of the tube. Remove it accordingly.
8. While the samples are drying, prepare 10X reaction buffer by combining 19 µL of lysis buffer 2 and 1 µL of RNase Inhibitor (40 U/µL). Briefly spin it down and keep it on ice.
9. Once the samples are dry and the bead pellets no longer appear glossy, remove the tubes from the magnetic separator and add 9.5 µL of RNase-free H₂O to rehydrate the samples. Place the samples on ice and add 1 µL of 10x reaction buffer to each sample.

6. Reverse Transcription (10 min)

NOTE: Before beginning, thaw the necessary reagents for reverse transcription (RT; except for the enzyme) on ice. These include: primer II, buffer 1, oligonucleotide, and RNase inhibitor.

1. To each tube, add 2 µL of primer II (AAGCAGTGGTATCAACGCAGAGTACT₍₃₀₎N_-1N, for which N_-1 can be A, C, or G and N can be A, C, G, or T; 12 µM). Place the tubes in a thermocycler that has been preheated at 72 °C for 3 min.
2. During incubation, prepare the RT master mix. For each reaction, add 4 µL of buffer 1 (250 mM Tris-HCl, pH 8.3; 375 mM KCl; and 30 mM MgCl₂), 1 µL of oligonucleotide (48 µM), and 0.5 µL of RNase inhibitor (40 U/µL).
3. Immediately following the incubation, place the tubes on ice for 2 min.
4. Add 2 µL per reaction of the reverse transcriptase (100 U/µL) to the master mix and pipette thoroughly. Add 7.5 µL of master mix to each tube and mix by gently pipetting. Briefly spin the tubes to collect the contents at the bottom and place them into a preheated thermocycler with the following program: 42 °C for 90 min, 70 °C for 10 min, and a hold at 4 °C.
5. Store the tubes at -20 °C O/N before proceeding, although it is recommended that the samples be carried through the amplification step before being stored for long periods of time; other sources suggest O/N storage at 4 °C would also be acceptable at this step²⁴.

7. cDNA Amplification (2.5 h)

NOTE: Before beginning, thaw PCR buffer and PCR primer on ice and spin the tubes down in a tabletop mini centrifuge before making the PCR master mix.

1. For each reaction, prepare a PCR master mix containing 25 µL of PCR buffer, 1 µL of PCR primer (12 µM), 1 µL of DNA polymerase, and 3 µL of nuclease-free H₂O. Add the DNA polymerase last, just prior to the addition of master mix to the samples.
2. Add 30 µL of master mix to each tube and spin at 2,000 x g for 10 s to collect the contents at the bottom of the tubes.
3. Place the tubes in a preheated thermocycler with the following program: 95 °C for 1 min; 34 cycles of 98 °C for 10 s, 65 °C for 30 s, and 68 °C for 3 min; 72 °C for 10 min; and a hold at 4 °C.
   NOTE: The number of cycles for this PCR has been increased to 34, differing from the suggested manufacturer's instructions. After repeat testing, this cycle number was found to produce consistently reliable results.
4. Store the tubes at -20 °C for up to one year before proceeding.

8. Purification of Amplified cDNA (30 min)

1. Before beginning purification, bring the DNA beads and elution buffer to RT for at least 30 min. Prepare fresh 80% EtOH; 1 mL per sample should suffice. Add 1 µL of 10X lysis buffer to each sample.
2. Vortex the DNA beads for 30 s and add 50 µL of DNA beads to each sample. Mix thoroughly by pipetting, and then briefly spin them down at 2,000 x g for 10 s. Incubate the tubes at RT for 8 min.
3. Place the tubes on the magnetic separation device for 5 min. Gently pipette the supernatant up and down twice and allow the samples to sit for 2 min. While the samples are on the magnetic device, remove the supernatant and discard.
4. Add 150 µL of freshly made 80% EtOH to each sample and allow them to sit at RT for 30 s. Remove the EtOH and repeat the EtOH wash once.
5. Briefly spin the samples and place them back on the magnetic separator for 1 min. Remove any remaining EtOH and allow the samples to air dry for 5 min.
6. Check the samples intermittently to see if any EtOH has collected and remove it with a pipette.
7. Once the bead pellet no longer appears glossy, but before cracks begin to appear, remove the tube from the magnetic separator and add 17 µL of elution buffer. Gently pipette up and down to resuspend the beads completely.
8. Incubate the resuspended samples at RT for 2 min.
9. Briefly spin the samples to collect all liquid at the bottom and place them on the magnetic separator for 2 min. Transfer 15 µL of clear supernatant to a 1.5 mL RNase-free tube and store it at -20 °C.
10. Examine quality and size of the cDNA library with a high-sensitivity bioanalyzer chip using 1 µL of cDNA. The ideal size of a cDNA library is 0.3-2 Kb, with a concentration of at least 10 ng/µL (Figure 2).
    NOTE: You may choose to employ PCR as a way to screen samples to ensure the expression of known markers before proceeding with sample preparation.

9. Determine Concentrations and Tagment cDNA (20 min)

1. Before beginning, bring the assay reagent, dilution buffer, and standards to RT for 30 min. Prepare a master mix containing 1 µL of assay reagent and 199 µL of dilution buffer per reaction. Aliquot 199 µL of this mixture into 500 µL assay tubes. Add 1 µL of sample and mix thoroughly.
2. Aliquot 190 µL of master mix to tube 1 & 2. Add 10 µL of standard 1 to tube 1 and 10 µL of standard 2 to tube 2. Vortex all sample tubes and standard tubes for 5 s; incubate at RT for 3 min.
3. Determine concentration of samples with a high-sensitivity fluorometer. Ensure the correct sample amount has been input by selecting the "calculate stock solution" option and highlighting "1 µL". Dilute each sample in a separate 1.5 mL tube to a final concentration of 0.2 ng/µL in RNase-free H₂O.
   NOTE: Whereas the manufacturer's instructions suggest that one should begin with 1 ng/µL total of DNA, we have used a smaller starting amount and have also adjusted the protocol to account for this amendment.
4. In new 0.2 mL PCR tubes, pipette 2.5 µL of buffer 2. Add 1.25 µL of cDNA to the appropriate tube for a total of 250 pg. Finally, add 1.25 µL of tagmentation mix to each tube and pipette thoroughly to mix; the transposase within the tagmentation mix will fragment the DNA into short strands and anneal the adapters to either end of each strand, for later use by the sequencing instrument.
   NOTE: The volumes used for tagmentation and index coupling are a fraction of the amount suggested by the manufacturer's instructions. This not only allows for the conservation of reagents, but it also has consistently produced tagmented samples in our experience.
5. Centrifuge the tubes on a countertop microcentrifuge for 1 min.
6. Place the tubes in a preheated thermocycler with the following program: 55 °C for 10 min and a hold at 10 °C.
   NOTE: The length of this incubation is 10 min, as opposed to the proposed 5 min from the manufacturer's instructions.
7. Immediately after this program is complete, remove the tubes and add 1.25 µL of tagmentation neutralizing buffer to each. Pipette well to mix and incubate at RT for 5 min. Complete this step promptly, as the transposase remains active until the buffer neutralizes the enzyme and stops the reaction.

10. Index Coupling and Purification (1 h)

NOTE: Before beginning, bring the DNA beads and resuspension buffer to RT for at least 30 min. Decide which indices to use for each of the samples.

NOTE: These indices will be attached to the respective 5' and 3' ends of the fragmented DNA for the identification of samples following sequencing. Ensure that no two pairings are the same for samples that may be sequenced together. For example, if sample 1 will use indices white 1 and orange 1, sample two should use white 1 and orange 2 or white 2 and orange 2, but never the same combination of indices. This kit contains 4 distinct white and 6 distinct orange indices. All of the different possible combinations allow up to 24 samples to be pooled in one sequencing lane. Although we typically only pool 10 samples in a lane, one could also use the kit containing 24 indices, which would allow for the pooling of 96 samples in a single lane of sequencing, if desired.

1. To each tube, add 1.25 µL of the left index and 1.25 µL of the right index for that specific sample. Add 3.75 µL of PCR master mix and pipette well to mix.
2. Centrifuge in a countertop microcentrifuge for 1 min.
3. Place the tubes in a preheated thermocycler with the following program: 72 °C for 3 min; 95 °C for 30 s; 12 cycles of 95 °C for 10 s, 50 °C for 30 s, and 72 °C for 1 min; 72 °C for 5 min; and a hold at 4 °C.
   NOTE: The first 95 °C step was changed from 98 °C, which was suggested by manufacturer's instructions. Also, the cycle details were adjusted so that the temperatures and time lengths allowed for the optimal tagmentation of our samples. The final 72 °C incubation was also added to our protocol and was not included in the manufacturer's instructions.
4. Briefly spin the tubes to collect the contents at the bottom. Vortex the DNA beads for 30 s, and then add 30 µL to each tube and mix thoroughly by pipetting.
5. Incubate the samples at RT for 5 min.
6. Place the samples on a magnetic separator for 2 min. Pipette the supernatant up and down twice and incubate samples for 1 min.
7. Remove and discard the clear supernatant. Add 150 µL of 80% EtOH to each sample and remove. Repeat the EtOH wash once.
8. Allow the samples to air dry for 10 min. Check intermittently to see if any EtOH has collected at the bottom of the tubes and remove as necessary.
9. Once a crack begins to appear in the DNA bead pellet, remove the sample from the magnetic separator and add 27.5 µL of resuspension buffer to rehydrate the pellet. Ensure that the pellet is completely resuspended, and then incubate at RT for 2 min.
   NOTE: The volume used for the resuspension buffer was modified to account for the smaller amount of starting material in our protocol and differs from the suggested volume in the manufacturer's instructions.
10. Place the tubes on the magnetic separator for 2 min. Transfer 25 µL of clear supernatant into an RNase-free tube and store at -20 °C.
    NOTE: Do not proceed with the manufacturer's instructions to complete library normalization. The samples are successfully prepared following this step and are prepared for sequencing as is.
11. Analyze the tagmentation of the samples using a bioanalyzer chip and 1 µL of each sample.
    NOTE: The analysis of smears will be conducted as earlier; however, cDNA should now be detected at a size range of 0.2-1 Kb (Figure 3). Smears below this point likely represent the degradation of samples, while larger smears suggest incomplete tagmentation.

11. Pooling of Samples (10 min)

1. Obtain concentrations of tagmentation samples with the high-sensitivity fluorometer from earlier and determine the concentration in nM.
   NOTE: This calculation can be computed with the use of an internet conversion tool and relies on the average fragment length from the bioanalyzer trace. To determine the average fragment length, observe the trace for each sample individually and determine the size of the average DNA fragment. This may also be calculated when examining the bioanalyzer trace by highlighting the range of the smear and examining the molarity calculated by the program.
2. Combine samples such that the pool contains the same concentration of each sample. Do not dilute samples; the pool should be only as dilute as the lowest-concentrated sample and would ideally have a total concentration of at least 15 nM.
3. Store the pooled samples at -20 °C prior to sequencing; it is suggested that samples undergo sequencing within 1 week of pooling. Perform paired-end, 100 bp sequencing with a HiSeq platform.