Culture of Adult Transgenic Zebrafish Retinal Explants for Live-cell Imaging by Multiphoton Microscopy

Note: Zebrafish were raised and maintained in the Notre Dame Zebrafish facility in the Freimann Life Sciences Center. The methods described in this manuscript are approved by the University of Notre Dame Animal Care and Use Committee and are in compliance with the statement for the use of animals in vision research by the Association for Research in Vision and Ophthalmology.

1. Solutions

1. Prepare 70% ethanol to sterilize the tissue culture hood and any equipment/reagents that are transferred into the tissue culture hood.
2. Add 2 mL of 2-phenoxyethanol to 1 L of system water (1:500 2-phenoxyethanol).
3. Prepare 0.1 mM NaHCO₃, pH 8.0. Use a 10 mL syringe and a 0.2 µm pore-size syringe filter to sterilize the solution in a sterilized tissue culture hood.
   NOTE: NaHCO₃ is used to efficiently spread the cell and tissue adhesive across the coverslip surface of fluorodishes (see step 3.2).
4. Prepare 1.0 M CaCl₂ and 1.0 M MgCl₂. Sterilize with a 10 mL syringe and 0.2 µm pore-size syringe filter in a sterilized tissue culture hood.
5. To prepare Hank's-balanced Salt Solution (HBSS), add sterile CaCl₂ and sterile MgCl₂ to 1x HBSS without Ca²⁺/Mg²⁺, without phenol red at a final concentration of 1 mM each. Work in a sterile environment.
6. To prepare culture medium, mix 50% 1x Minimum Essential Medium (MEM) without phenol red, 25% HBSS containing CaCl₂ and MgCl₂ (see section 1.4), 25% Horse Serum (HS), 10 units/mL penicillin, and 10 µg/mL streptomycin. Work in a sterile environment.
   NOTE: Avoid medium containing phenol red as it autofluoresces, thereby affecting signal to noise ratios.²³
7. Prepare 10 mL of 1% low melting point agarose in 1x MEM without phenol red. Melt agarose/1x MEM using a microwave. Prepare small volumes of agarose (e.g., 10 mL) as repetitive reheating will change ion concentrations due to fluid evaporation.
8. Prior to culturing, sterilize 1.5 mL microcentrifuge tubes by autoclaving.

2. Light-damage Paradigm

1. Dark-adaptation
   1. Place 2 – 3 Tg[gfap:nGFP]^mi2004 zebrafish (or other transgenic zebrafish of interest) at 6 – 14 months of age into an environment devoid of light for 14 d. For further detail see reference^{2, 6, 24, 25}.
2. Position the tank containing 2 – 3 dark-adapted transgenic zebrafish in system water between two fluorescent lamps that emit light of 2,800 lux^{2, 6, 24, 25}.
3. Expose zebrafish to constant intense light for 35 h. During the light exposure, assure that the water temperature is maintained between 31 – 33 °C.

3. Preparation for Culturing (On the Day of Retinal Isolation)

1. Using 70% ethanol, sterilize the tissue culture hood and the components/tools that are transferred into the tissue culture hood (e.g., flasks containing MEM and HBSS, pipettes, sterile pipette tips, etc.).
2. Preparation of fluorodishes
   1. Coat one fluorodish for each retina with cell and tissue adhesive (see Table of Materials).
   2. Dilute the cell and tissue adhesive in 0.1 mM NaHCO₃ to a final concentration of 70 µg/mL, add 50 µL to each fluorodish and spread the solution across the center of the fluorodish with a 200 µL pipette tip (approximately 1 – 1.2 cm diameter).
   3. Incubate the coated fluorodishes for 1 – 3 h at RT (alternatively, incubate O/N at 4 °C).
   4. Remove the solution and rinse 3x with 500 µL of 1x MEM for each wash. To avoid the coated fluorodishes from drying out, maintain them in MEM until retinal explants are mounted.
3. Prepare culture medium as described in step 1.6. The culture medium can be stored for up to 1 week at 4 °C.

4. Isolation and Culturing of Retinal Explants

NOTE: The protocol outlined below is for the isolation of the dorsal retina, which is the retinal region that is predominantly lesioned by the described light-damage paradigm. Therefore, damage-induced proliferation and the associated event of interkinetic nuclear migration occur in the dorsal retina. However, the isolation procedure can be adjusted to yield retinal regions according to the specific requirements of the researcher/research question.

1. Euthanize one light-damaged transgenic zebrafish at a time in 1:500 2-phenoxyethanol.
2. Remove MEM from one fluorodish with a 1,000 µL pipette so that only a thin film of fluid remains.
3. Transfer the zebrafish onto a dry paper towel, remove the eye with a curved pair of Dumont forceps (forceps #5, 45° angle) and transfer it onto the fluorodish.
4. Using a stereomicroscope, orient the eye with the pupil onto the cover slip of the fluorodish so that the back of the eye with the optic nerve is visible (Figure 1D).
5. With a pair of McPherson-Vannas scissors remove the optic nerve, cutting close to the back of the eye. In addition, remove connective tissue lining the outside of the eye.
6. Hold the eye between its nasal and temporal side with a pair of #5 forceps while making an incision at the optic stalk by piercing with one scissor blade through the lamina cribrosa and cutting along both the nasal and temporal sides of the eye (see Figure 1E, segmented line).
7. Using two pairs of #5 forceps, one for holding the dorsal side of the retina and the other pair to separate the ventral from the dorsal retina, pull the tissue.
   NOTE: It is advisable to orient the dorsal retina so that the lens and the ganglion cell layer face the coverslip while the sclera is facing upwards.
8. Remove the sclera from the dorsal retina with one pair of #5 forceps, while holding the lens that is connected to the retina with a second pair of #5 forceps (Figure 1H, I).
9. To remove the lens, use McPherson-Vannas scissors and cut behind the lens without damaging the retina (Figure 1I, J). Sometimes, the lens separates in step 4.7. In this case, remove the sclera carefully with forceps by holding the retina at a cut edge with a second pair of #5 forceps.
10. Remove the vitreous while removing the lens without damaging the retina. Flatten the retina with the ganglion cell layer facing the cover slip of the fluorodish (Figure 1L, M).
11. Surround the retina with 10 µL of 1% low melting point agarose and let the agarose solidify.
12. Watch that the liquid agarose does not lift the retina as even a slight elevation might affect the ability to focus deeply into the tissue. If lifting is observed, use a pipette to remove agarose. Let residual agarose set before attempting to add more.
13. Repeat step 4.12 several times before adding 1% low melting point agarose to cover the entire fluorodish. Once the agarose has solidified, add 1.5 mL culture medium.
14. Maintain the retinal explant culture in a 5% CO₂/air environment set at 32 °C for approximately 12 h to allow the retina to recover from the stress incurred by the isolation procedure. Set the temperature to 32 °C to maintain retinas at the same temperature as light-treated zebrafish (see step 2.3).

5. Multiphoton Microscopy

NOTE: The experiments performed in this manuscript were optimized for a multiphoton microscope equipped with an infrared laser (see Table of Materials), a 40X Apo long distance water immersion objective (N.A. 1.15), a galvanometer scanner and an environmental chamber that contains an insert for four 35 mm Petri dishes. The images were acquired with a non-descanned detector (R-NDD).

1. Prior to imaging, equilibrate the environmental chamber to achieve a 5% CO₂/air atmosphere. Make sure that empty Petri dishes are inserted into the holder to avoid leakage of gas into the room.
2. Turn on the microscopy system.
3. Once the environmental chamber is equilibrated, add refractive index liquid onto the 40X Apo long distance water immersion objective (N.A. 1.15).
   NOTE: The refractive index liquid with optical properties similar to water is used to avoid evaporation of water during long-term imaging.
4. Place fluorodishes with retinal explants into the chamber. Using brightfield light, position the specimen into the light path and bring the midregion of the dorsal retina into the plane of focus.
5. Use GFP epifluorescent light to focus on gfap:nGFP-positive Müller glia nuclei (Figure 2A, C).
   NOTE: If explants are not mounted flat or agarose accumulated under the explant, it will be difficult to focus onto the gfap:nGFP-positive nuclei or they will fluoresce dimly.
   1. Check whether moving to a different region within the same retinal explant will overcome the focusing issue. Otherwise move to a different retinal explant.
6. In the image acquisition software, open the 'A1 MP GUI', the 'TiPad', the 'A1 Compact GUI' and the 'ND acquisition' windows. For multiphoton imaging, ensure that the 'IR NDD' option is chosen in the 'A1 Compact GUI'.
7. In the 'setting' field, select IR-DM for the 1^st dichroic mirror and choose the band pass filter 525/50 to acquire GFP fluorescence.
8. Switch on the IR laser in the window labeled 'A1MP GUI'. It will take a few minutes for the laser to be ready. Set the wavelength to 910 nm to excite GFP fluorescence and align the laser by clicking the 'Auto alignment' button in the 'A1 MP GUI' window.
9. Ensure that room and equipment lights are switched off or covered before opening the shutter in the 'A1 MP GUI' to avoid overexposure of the photomultiplier tube. To reduce noise levels, house the microscope in a darkened environment.
10. Acquire images of a field of view of 300 x 300 pixels, at a zoom of two, and a pixel dwelling time of 4.8 µs/pixel. Roughly set up the laser power by changing the 'acquisition area' in the 'A1MP GUI' and the gain in the 'A1 Compact GUI' window.
11. Setting up the z-stack
    1. Focus on the ganglion cell layer to set the top focal plane of the z-stack in the 'z'-subwindow within the 'ND acquisition' window. Some gfap:nGFP-positive cells are typically located in the ganglion cell layer, which helps to identify the basal limit of the retina (Figure 2A, D).
    2. Move the focal plane through the level of the ONL (Figure 2A, B), which is characterized by the presence of dimly labeled gfap:nGFP-positive cells that are round and enlarged relative to their counterparts in the INL (Figure 2A, C).
    3. Set this plane as the bottom of the z-stack. Ensure that the entire ONL will be imaged (Figure 2A, B).
    4. When experiencing focal plane shifts that require re-adjusting during the imaging period, double-click on the middle position in the 'z' subwindow to assign it as the 'home' position. Change to 'symmetric mode defined' and click 'relative'.
    5. Set the z-step size between 0.7 to 1 µm.
12. Z-intensity correction:
    1. Apply z-intensity corrections to compensate for loss of pixel intensity due to light scattering when imaging in deep layers of the tissue.
    2. To set up the correction, open the 'z-intensity correction' window. To set the 'z-stack range', choose 'From ND'.
    3. Click on the bottom focal plane in the 'z-intensity correction' window (in this case, corresponds to the ganglion cell layer) and set the laser intensity ('acquisition area' in the 'A1MP GUI') and gain (A1 Compact GUI).
    4. Click the arrow next to the 'z-values' in the 'z-intensity correction' window to confirm the settings that are subsequently shown under 'device settings' in the 'z-intensity correction' window for the chosen focal plane.
    5. Repeat the process for the middle and top planes, increasing the laser power and gain. Additional focal planes can be added if necessary. See Table 1 for specific laser and gain settings for experiments in Figures 2 – 4.
    6. Set the 'acquisition area' in the 'A1 MP GUI' window. Avoid selecting an acquisition area larger than 15 and a gain higher than 126 at the start of imaging to circumvent photobleaching and increased noise levels.
       NOTE: As lasers and photomultiplier tubes differ between microscopy systems, test laser and gain settings to obtain optimal imaging conditions for the microscope set up while avoiding photobleaching.
    7. Choose 'relative intensity correction' in the 'z-intensity correction' window.
13. In the 'timeseries' subwindow in the 'ND acquisition' window, set the duration to 8 h and the interval to 'no delay'. Then, make sure to click the 'Run z-correction' in the 'z-stack' sub window in the 'ND acquisition' window button to acquire the 3-D timeseries.
14. Maintain retinal explants at a temperature of 27 – 29 °C throughout the duration of image acquisition.
15. Throughout the image acquisition period, if necessary, readjust the power level and gain in order to maintain image quality for post-imaging analysis. For adjustments perform steps 5.11.2 – 5.11.4.
16. If the focal plane shifts, pause or stop the run and perform step 5.11 again.
    NOTE: If 'relative z-correction' was chosen in step 5.12.7 and step 5.11.4 was performed it should not be necessary to readjust power and gain levels, unless extensive photobleaching occurred.

6. Analysis of Velocity

1. Extract the time, setting a 'region of interest' on the image. Use the 'time measurement' tool and export the time values to a spreadsheet.
2. Crop a region that contains a dividing gfap:nGFP-positive nucleus.
3. Choose the cropped region so that it contains at least one nucleus that does not undergo INM in order to set a reference point to subsequently measure the distances that the dividing nucleus migrated in relation to the basal INL. NOTE: To choose a nucleus that remains in the basal INL, it helps to prepare a 3-D reconstruction of the timeseries.
4. Alternatively, if a gfap:nGFP-positive cell is observed in the ONL throughout the acquisition period, use a vertical line of fixed length spanning from the ONL nucleus to the basal INL in order to identify a reference point.
5. Using the 'show slices view' function, generate orthogonal projections. Subsequently, change the mode from 'slice' to 'maximum intensity projection'.
6. Turn off the 'xy' view of the orthogonal maximum projection. Depending on the orientation at which the migrating/dividing nucleus is best visible, also turn off either the 'xz'- or the 'yz'-view.
7. Click on the remaining image with the right mouse button and extract the 'xz' or 'yz'-image series with the 'Create new document from this view' function. If necessary, rotate the image.
8. Using the 'manual measurement' function, draw a horizontal line across the image at the bottom level of the nucleus that remains in the basal INL and does not undergo INM (= reference point; Figure 4A – E, red horizontal line).
9. Measure the distance between the reference line and the basal point of the migrating nucleus for the timeseries using the 'line measurement' tool in the analysis software (see Figure 4A – E).
10. Once the nuclear envelope breaks down, measure the basal position of the soma if it is identifiable following the diffusion of GFP into the entire cell.
11. Using a spreadsheet software, plot the distance a nucleus migrated against the time passed (Figure 4F).
    1. To determine the apical migration velocity (v_a), graph the distances traveled for the period before nuclear envelope breakdown occurs (Figure 4G).
    2. Select the data series within the chart, right-click the mouse and insert a linear regression curve including the corresponding function 'y=mx+c'. The slope 'm' in the function represents the velocity, v_a (Figure 4G).
    3. Repeat steps 6.11.1 and 6.11.2 for the first phase of rapid basal migration to determine the basal migration velocity, v_b (Figure 4H).