Primary Cell Cultures to Study the Regeneration Potential of Murine Müller Glia after MicroRNA Treatment

Procedures involving animal subjects have been approved by the Institutional Animal Care and Use Committee (IACUC) at SUNY College of Optometry.

NOTE: This culture protocol consists of three phases: growth, transfection, and conversion phase. A summary of the overall protocol with the timeline is given in Figure 1.

1. Preparation of media and all required reagents

NOTE: All steps need to be carried out in an A2 or B2 biosafety cabinet (BSC). During the growth phase, a high-serum growth medium is used which consists of a basal neuronal medium supplemented with epidermal growth factor (EGF). For the conversion phase, a low-serum neurophysiological basal medium supplemented with neuronal supplements is used to ensure neuronal differentiation and survival.

1. Prepare growth medium (used during growth phase) by supplementing 200 mL of basal neuronal medium with 20 mL of fetal bovine serum (FBS, 10%), 1 mL of 200 mM L-glutamine (0.5%), 2 mL of penicillin/streptomycin (1%), and 2 mL of N2 supplement (1%). Perform sterile filtration (filter units with 0.22 µm pore size). Pre-warm the medium in 37 °C metal bead bath before use.
2. Prepare neuronal medium (used during conversion phase) following manufacturer's instructions (see Table of Materials) by supplementing 100 mL of serum-free neurophysiological basal medium with 2 mL of B27 neuronal supplement (2%), 1 mL of N2 supplement (1%), 20 µL of 100 ng/mL brain-derived neurotrophic factor (BDNF, reconstitute in 0.1% bovine serum albumin (BSA) in phosphate buffered saline (PBS, final concentration 20 ng/mL), 20 µL of 100 ng/mL glia cell-derived neurotrophic factor (GDNF, reconstitute in sterile Hanks' Balanced Salt Solution [HBSS], final concentration 20 ng/mL), 500 µL of 100 mg/mL Dibutyryl-cAMP (reconstitute in DMSO), 70 µL of 50 ng/mL ascorbic acid (reconstitute in sterile PBS), and 1.5 mL of penicillin/streptomycin. Perform sterile filtration (filter units with 0.22 µm pore size). Pre-warm medium in 37 °C metal bead bath before use.
   NOTE: These media can be stored at 4 °C, for 1 month.
3. Reconstitute Papain, DNase I, and Ovomucoid reagents required for retinal dissociation following manufacturer's protocol. Aliquot 750 µL of Papain in sterile 1.5 mL tubes, 75 µL of DNase I in sterile 0.6 mL tubes, and 750 µL of Ovomucoid protease inhibitor in 2 mL tubes. Freeze Papain and DNase I aliquots at -20 °C and keep Ovomucoid aliquots at 4 °C to avoid degradation of the reagents. Thaw at room temperature right before use.
   NOTE: Papain, DNase I, and Ovomucoid are found in a kit called Papain Dissociation System (see Table of Materials).
4. Reconstitute poly-L-ornithine (Poly-O) and Laminin for coverslip coating if immunofluorescent labeling is performed following the datasheet instructions (Poly-O: 0.1 mg/mL in sterile water; Laminin: dilute 1.2 mg/mL at 1:50 in DMEM). Aliquot Poly-O and Laminin (2.5 mL) and freeze aliquots at -20 °C. Thaw at room temperature right before use.
   ​NOTE: Step 1.4. is required only if immunofluorescent labeling and laser-scanning microscopy is performed.

2. Mice and tissue extraction

NOTE: For these reprogramming studies, the Ascl1^CreERT:tdTomato^STOPfl/fl mouse was created by crossing an Ascl1^CreERT mouse (Ascl1-CreERT: Jax # 012882) with a tdTomato^STOPfl/fl mouse (B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J: Jax # 007914). This mouse has a mixed background (C57BL/6, S129, and ICR strain). The genotype of this mouse is shown in Figure 1A. All strains can be used for this protocol.

1. Wear gloves and sanitize workspace, including dissection microscope and all fine tools (Dumont #5 fine and Dumont #7 curved forceps, fine scissors, and Vannas scissors) with 70% ethanol. Sanitize a 10 cm silicone-coated black dissection dish by exposing it to UV light for 20 min.
2. Preparation of dishes and plates for tissue extraction
   1. Prepare a 24-well plate for eye collection and sample separation (two eyes per mouse in one well, labeled with mouse IDs) with ~1 mL of cold HBSS (4 °C) per well. Place the 24-well plate on ice.
   2. Fill a sterile 10 cm Petri dish (for washing) and the sanitized silicone-coated dissection dish with several mL of cold HBSS (4 °C) to ensure that the tissue is fully covered with HBSS.
   3. Prepare a sterile 1.5 mL tube with 1 mL of 70% ethanol.
   4. Prepare a 12 well culture plate by labeling the plate with culture number, date, strain, and all the required information.
3. Euthanize P12 mice using any approved method.
4. Eye removal
   1. Gently hold the mouse head with the thumb and index finger around the eye.
   2. Using Dumont #7 curved forceps, gently go behind the eyeball and clip the optic nerve. Carefully take the eye out.
      NOTE: If adult mice are used, do not use curved forceps, and do not pull the eye. Instead cut carefully around the eye globe using fine scissors; cut the optic nerve but do not cut the eye itself. Use forceps to carefully remove the eyeball.
5. Eyeball cleaning
   1. Dip the eyeball briefly into an ethanol-containing tube to avoid carryover of bacteria from the animal.
   2. Wash the eyeball briefly in the 10 cm Petri dish before placing it into the 24 well plate on ice.
6. Repeat steps 2.4 and 2.5 for the other eyes. Keep the well plate on the ice during the process. Place two eyes from one animal in the dissection dish placed under a dissection microscope with a light source.
7. Retina extraction
   1. Fix one eyeball by grabbing the optic nerve and the surrounding connective tissue around the sclera with Dumont #5 fine forceps and press it carefully against the dissection dish (cornea upward).
   2. Make a hole in the center of the cornea using a 30 G needle to allow easier access for the Vannas scissors.
   3. Dissect the cornea around the ciliary body using Vannas scissors and remove cornea, lens, iris, and vitreous body carefully with Dumont #5 fine forceps. Figure 2A illustrates an eye cup with the retina inside.
   4. Dissect the sclera with Vannas scissors until the optic nerve is reached. Clip the optic nerve and carefully extract the retina using Dumont #5 fine forceps.
   5. Use a second pair of Dumont #5 fine forceps to push against the retina and allow complete removal of the vitreous body. Figure 2B illustrates two extracted retinas.
8. Transfer and wash
   1. Cut about 2.5 cm of the tip of a sterile transfer pipette to enlarge the diameter. Using this tip, pick up (suck in) whole retinas without damaging the tissue.
   2. Transfer the retinas into a new sterile Petri dish with cold HBSS (4 °C) and rock the dish (back and forth, left, and right).
   3. Using the transfer pipette tip, carefully push the retinas around to wash off retinal pigment epithelial (RPE) cells.
      NOTE: Alternatively, the entire retina with lens and vitreous body can be removed from the eye cup. Then, the lens and vitreous body can be removed from the extracted retina. If the retina is ripped, make sure to collect all pieces for dissociation; otherwise, there will be insufficient amounts of tissue to grow confluent cell layers.
9. Immediately place the isolated retina in a new, clean well of the 24-well plate filled with 1 mL of HBSS. Keep the 24-well plate on the ice during the dissection process.
10. Repeat steps 2.7-2.9 to isolate the second retina.

3. Retina dissociation

NOTE: All following steps (until cell harvest) need to be carried out in an A2 or B2 biosafety cabinet (BSC).

1. Prepare the Papain/DNase I dissociation mixture as follows.
   1. For six retinas, add 75 µL of DNase I into the tube containing 750 µL of Papain (from step 1.3) and mix carefully (dissociation mixture).
   2. For individual sample preparation that is required for this protocol, split the total volume of 825 µL into three aliquots: 275 µL of the mixture in one 1.5 mL tube per mouse (two retinas). Calculate the required amounts of DNase I and Papain accordingly.
      NOTE: Up to six retinas can be dissociated in one tube of Papain/DNase I mixture. However, individual sample preparation results in fewer clumps and better cell growth than in combined samples.
2. Transfer two retinas to Papain/DNase I dissociation mixture. Use a transfer pipette with an enlarged tip diameter (step 2.8.1), pick up the retinas, wait until the retinas settle at the bottom of the tip, and then release the retinas without excessive HBSS into the tube containing Papain/DNase I mixture.
3. Place on a nutator and incubate it for 10 min in a cell culture incubator (37 °C, 5% CO₂).
4. Dissociate the cells by carefully pipetting up and down (about 20-30 times) with a 1 mL pipette. After cells are dissociated (i.e., resulting in a homogenous solution with no chunks), add 275 µL of Ovomucoid protease inhibitor from the Papain Dissociation Kit to neutralize the Papain. Mix gently by pipetting up and down.
   NOTE: If six retinas were dissociated in 825 µL of Papain/DNase I mixture, 825 µL of Ovomucoid is required.
5. Centrifuge the mixture at 300 x g for 10 min at 4 °C.
6. Add epidermal growth factor (EGF, 1 µL per 1 mL of the growth medium, reconstituted at 200 µg/mL in PBS) to the calculated volume of growth medium (1 mL per mouse) pre-warmed at 37 °C.
   NOTE: Depending on the experimental design, the proliferation marker 5-ethynyl-2'-deoxyuridine (EdU), as well as 4-Hydroxytamoxifen (4-OHT) or other required factors can be added at the beginning of the culture period.
7. Remove the tubes carefully from the centrifuge. Do not touch the pellet at the bottom of the tube.
8. Remove the supernatant carefully and entirely. Resuspend the cell pellet with 500 µL of EGF-supplemented growth medium.
9. Transfer the cell suspension into one well of the labeled 12-well plate (Figure 2C). Rinse the tube with another 500 µL of the EGF-supplemented growth medium and add it to the well (total volume of 1 mL per well).
10. Repeat steps 3.8 and 3.9 with all other samples.
11. Rock the well plate three times carefully (back and forth; left, and right). Place the plate into the incubator (37 °C, CO₂).
    ​NOTE: If transgenic mice are used, perform genotyping for every animal (Figure 2D). Identify Cre recombinase positive and negative mice and label the plate accordingly. For this protocol, only cells of Cre recombinase positive reporter mice were used for the next steps. Cells of Cre negative specimens are frozen and used for other applications.

4. Growth phase

NOTE: The growth phase has a duration of about 4-5 days (Figure 1B). For adding liquids to wells containing cells, the pipette needs to point to the wall of the well and the liquid needs to be released slowly to avoid cell detachment. Do not pipette directly on top of the cells.

1. One day after dissociation, remove the medium and add 1 mL of fresh EGF-supplemented growth medium.
2. On day 3, remove the medium and add 1 mL of HBSS (room temperature) to remove cell debris. Rock gently back and forth left to right. Remove HBSS, repeat the wash step, and add 1 mL of pre-warmed EGF-supplemented growth medium.
3. Monitor the cells every day and evaluate their growth status until the cells reach 90%-100% confluency. Figure 3 shows an example of good MG growth over time. Check for possible contamination or cell death (Supplementary Figure 1). Discard contaminated cultures.
   NOTE: To monitor and record cell state, take images at various magnifications using a light microscope with an attached camera and 4x, 10x, or 20x objectives. In this study, a fluorescence microscope is used.

5. Preparation of coverslips with poly-L-ornithine (Poly-O) and Laminin coat

NOTE: This step is only necessary if immunofluorescent labeling and confocal laser-scanning microscopy are performed. Round glass coverslips (12 mm diameter) are required for proper imaging. The coating protocol can also be found in the neuronal medium datasheet (see Table of Materials).

1. Place sterile coverslips carefully in the center of every well of a 24-well plate using sterile Dumont #2AP forceps.
   NOTE: Place coverslips in the center of the well. Placing coverslips close to the wall of a well will cause surface tension issues for the following steps.
2. Thaw a 2.5 mL Poly-O aliquot at room temperature and place 100 µL of it in the center of each coverslip.
3. Incubate the well plate for 30 min in a 37 °C incubator.
4. Remove Poly-O and wash the well with the coverslip three times with ~1 mL of sterile water.
5. Let the well plate dry overnight in the BSC. Thaw 2.5 mL of Laminin at 4 °C overnight.
6. The next morning, add 100 µL of Laminin in the center of each coverslip and incubate for 4 h in a 37 °C incubator.
7. Remove the Laminin carefully and entirely.
8. Keep the plate at 4 °C if passaging cannot be performed immediately.
   ​NOTE: The coated coverslips in prepared plates can be kept for a few days at 4 °C.

6. Cell passage to remove neuronal survivors

NOTE: Cell passage is required to remove neuronal cells, not to increase the cell population. Glia divide only a few times and will not grow further after passage. Do not dilute cell suspensions. The cells of one confluent well of a 12-well plate can be distributed onto one well of a 12-well plate or two wells of a 24-well plate. When coated coverslips are used, only about one-third of the coverslip is coated. Therefore, six coverslips sitting in a 24-well plate, with confluent cells (~80%-90%) can be obtained from one well of confluent cells of a 12-well plate. Other ratios can be chosen as well to increase or decrease cell density. For this protocol, one Cre+ reporter mouse is used [one experiment, two treatments: miR-25 or control-miR; technical replicates n = 3 (three coverslips per treatment), biological replicate n = 1]. The number of technical and biological replicates can be defined differently depending on the experimental design.

1. Check whether the cells are 90%-100% confluent (also at the margin of the well; Figure 3E,F).
2. Remove the medium and add 1 mL of HBSS (room temperature) to wash. Gently rock the plate (back and forth, left and right). Remove HBSS completely.
3. Add 500 µL of a pre-warmed trypsin-containing solution (pre-warmed at 37 °C in a metal bead bath) to detach the cells from the well. Rock gently (back and forth, left to right) and incubate for 2 min in a 37 °C incubator.
4. Move the plate from the incubator to BSC. While tilting, aspirate the trypsin-containing solution and disperse it carefully and slowly over the well several times until the cells detach completely. Hold the plate against the light and make sure no cells are left at the bottom.
5. Transfer this cell suspension to a sterile 1.5 mL tube. Centrifuge at 300 x g for 8 min at 4 °C.
6. Remove the supernatant and carefully resuspend the cell pellet by adding 600 µL (100 µL per well for 6 wells/coverslips) of pre-warmed growth medium (supplemented with EGF; 1:1000). and pipetting up and down ~30-40 times.
   NOTE: The cells can be frozen at this time at -80 °C or in liquid nitrogen and defrosted following standard protocols for thawing cell lines. If cells are frozen, they will not be resuspended in EGF-supplemented medium (growth medium). They will be resuspended in basic medium (without EGF) and freezing solution (1/1 ratio). Steps 6.1.1 to 6.6.2 describe the steps for freezing the cells. If no cells are frozen, continue with step 6.7.
   1. Prepare freezing solution by mixing 100 µL of DMSO and 400 µL of FBS (total volume of 500 µL per well/sample).
   2. Resuspend the cell pellet in 500 µL of pre-warmed growth medium. Add the cell suspension to 500 µL of DMSO/FBS freezing solution (total volume of 1 mL). Keep tubes on ice until all samples are processed. Freeze cells at -20 °C for 1 h, and then store at -80 °C.
7. Seed cells by placing 100 µL of the 600 µL cell/media suspension (step 6.6) in the center of six coated coverslips (see step 5) of the 24-well plate. Place the plate carefully in the incubator and let the cells settle.
   NOTE: 100 µL of the 600 µL cell suspension (harvested from one well of a 12-well plate) will result in 90%-100% cell confluency, which is required for transfection.
8. Check the cells after 3 h to see whether they have settled on the coverslip. Add 400 µL of growth medium supplemented with EGF.
   ​NOTE: Cells are usually ready for transfection the following day (Figure 3G). If not confluent (90%-100%), leave them for another day. If still not confluent, do not use them for transfection. If other downstream applications are conducted, such as miRNA profiling, RNA-Seq, RT-qPCR, or western blot, cells need to be passaged into 12-well plates (1:1 ratio; no plate treatment required) and harvested for RNA/protein extraction.

7. Transfection

NOTE: The transfection phase consists of a 3 h phase in transfection medium only (transfections procedures are described in the transfection manual that comes with the transfection reagent) and an elongated phase in which transfection reagent and miRNAs are still present, but neuronal medium with required supplements is added (total duration is 2 days; Figure 1B). In this protocol, six wells will be transfected: three wells will receive the reprogramming miRNA miR-25 and three wells will receive the control miRNA.

1. Check whether the cells reached 90% confluency and record/image the cells before transfection.
2. Remove the growth medium and add 500 µL of HBSS (room temperature) to wash the cells.
3. Remove HBSS and add 400 µL of reduced serum medium used for transfections. Place the plate back into a 37 °C incubator.
4. Prepare the transfection mixture by following the instructions of the manufacturer's transfection reagent protocol. Make two mixtures: transfection reagent mix and miRNA mix (see Figure 1B for illustration).
   1. Prepare the transfection reagent mixture: For a 24-well plate, 49 µL of reduced serum medium and 1 µL of transfection reagent are required for one well (50 µL in total). For six wells, 294 µL of reduced serum medium and 6 µL of transfection reagent are combined and mixed well by gently pipetting up and down.
   2. Prepare miRNA mimic mixture: For a 24-well plate, a total volume of 50 µL of the mimic mixture is required for one well. Three wells will receive the control miRNA and the other three wells will be treated with miR-25. For this protocol, a 200 nM final concentration is used.
      1. For the miR-25 treatment (three wells), 150 µL of reduced serum medium and 3 µL of miR-25 mimics (100 µM stock solution) are combined and mixed well by gently pipetting up and down. Incubate for 5 min.
         For the control mimics treatment (three wells), 150 µL of reduced serum medium and 3 µL of control-miRNA mimics (100 µM stock solution) are combined and mixed well by gently pipetting up and down. Incubate for 5 min.
         NOTE: The volume of miRNA mimics depends on the dilution factor and concentration needed (20-500 nM). miRNA inhibitors (antagomiRs), or other molecules including plasmids, can be used as well. Also, combinations of molecules are possible to be transfected.
5. Combine miRNA mimic mixture and transfection reagent mixture. Mix carefully by pipetting slowly and thoroughly by gently pipetting up and down. Incubate for 20 min at room temperature (per manufacturer's instructions).
6. Add the above transfection mixture dropwise and slowly on top of the cells, close to the media surface in the well using a 20 µL pipette. Rock the plate gently (back and forth, left to right).
7. Incubate in a 37 °C incubator for 3 h.
8. After 3 h, add neuronal medium to the wells (500 µL per well) supplemented with 4-Hydroxytamoxifen (5 mM stock reconstituted with 2.58 mL of ethanol, 250 nM final concentration) to activate the Cre recombinase and 5-ethynyl-2'-deoxyuridine (EdU, 10 mM stock solution reconstituted with 2 mL of DMSO, 1 µM final concentration) to track cell proliferation.
   CAUTION: 4-Hydroxytamoxifen and EdU are known to be human carcinogens, teratogens, and mutagens. Read Material Safety Data Sheet before use and wear gloves, goggles, and lab coat. Reconstitute both reagents according to the manufacturer's recommendations. Do not inhale the substance/mixture. Tightly close after use. Waste material must be disposed of following national and local regulations. Wash hands and face after working with the substance.
9. Incubate in a 37 °C incubator for 2 days (Figure 1B).

8. Cell conversion

NOTE: The cell conversion phase has a duration of about 5-6 days (Figure 1B), but longer periods are possible.

1. Check cells daily for successful induction of tdTomato expression, potential cell death, and/or contamination. Cell density does not change (Figure 4). First faint red fluorescent-labeled cells can be observed 1 day after 4-Hydroxytamoxifen treatment. A fluorescent microscope is required to monitor and image the cells.
   NOTE: In this study, a fluorescence microscope is used for live imaging. Images are taken with 4x, 10x, or 20x objectives.
2. Two days after transfection, remove the medium and add 500 µL of pre-warmed neuronal medium supplemented with 4-Hydroxytamoxifen and EdU in each well.
3. Replace the medium with a fresh medium every other day until the cells are harvested.
4. Take live images for evaluation and assessment of the number of red fluorescent (Ascl1 expressing) cells and their morphological changes (Figure 4 and Figure 5A-C).

9. Cell harvest: fixation for immunofluorescent labeling

NOTE: Cells can be harvested for other downstream applications, including bulk or scRNA-Seq, RT-qPCR, or western blot.

1. Remove the medium and add 500 µL of cold HBSS (4 °C) per well and rock the plate gently (back and forth, left to right). Remove HBSS.
2. Fix the cells by adding 500 µL of 2% Paraformaldehyde (PFA) and incubate for 20 min at room temperature.
3. Perform immunofluorescent labeling according to established protocols.