Quantifying Spontaneous Ca²⁺ Fluxes and their Downstream Effects in Primary Mouse Midbrain Neurons

All procedures involving the use of animal subjects have been approved by the Texas A&M University Institutional Animal Care and Use Committee (25^th Nov 2019; AUP# 2019-0346).

NOTE: Preparation of cell culture solutions should be done using sterile procedure in a biological safety cabinet and filtered at 0.2 µm to prevent contamination.

1. Preparation of solutions and culture medium

1. Prepare laminin coating solution by diluting 20 µL of 1 mg/mL laminin stock into 2 mL of sterile distilled H₂O. Prepare on the day of dissection.
2. Prepare 10% equine (horse) serum (ES) stop solution by adding 5 mL of ES to 45 mL of 1x Hank’s Balanced Salt Solution (HBSS). Sterile filter using a 0.2 µm filter system or syringe filter tip. Store at 4 °C.
3. Prepare 4% bovine serum albumin (BSA) stock solution by adding 2 g of BSA powder to 1x phosphate-buffered saline (PBS) and bringing to a final volume of 45 mL. Sterile filter using a 0.2 µm filter system or syringe filter tip. Store at 4 °C.
4. Prepare papain stock solution by diluting papain to 3 mg/mL in 1x HBSS. Sterile filter using a 0.2 µm filter system or syringe filter tip. Store at -20 °C.
5. Prepare deoxyribonuclease (DNase) solution by adding 20 mg of DNase powder to sterile H₂O and bringing to a final volume of 20 mL. Sterile filter using a 0.2 µm filter system or syringe filter tip. Store at -20 °C.
6. Prepare ascorbic acid stock solution by adding 352 mg of ascorbic acid to sterile distilled H₂O and bringing to a final volume of 20 mL. Heat in 37 °C bath to dissolve if necessary. Sterile filter using a 0.2 µm filter system or syringe filter tip. Store at -20 °C.
7. Prepare Cell Culture Medium by adding the following to 50 mL of Neurobasal medium: 500 µL of Glutamax (100x), 500 µL equine serum, 1 mL of B-27, 100 µL of ascorbic acid, 500 µL of penicillin-streptomycin, 50 µL of kanamycin and 50 µL of ampicillin. Sterile filter using a 0.2 µm filter system. Store at 4 °C.
8. Prepare 0.01% Triton X-100 Solution by adding 1 mL of Triton X-100 into 9 mL of 1x PBS to make a 10% solution. As Triton X-100 is viscous, pipette slowly to allow tip to fill completely. Heat in 37 °C bath to dissolve if necessary. Store at 4 °C.
9. To dilute 10% Triton X-100 stock to 0.01%, perform 3 serial 1:10 dilutions. Dilute 1 mL of 10% stock into 9 mL of 1x PBS to make 1% solution. Dilute 1 mL of 1% solution into 9 mL of 1x PBS to make 0.1% solution. Dilute 1 mL of 0.1% solution into 9 mL of 1x PBS to make a 0.01% solution.
10. Prepare 10% and 1% normal goat serum (NGS) solution by adding 1 mL of NGS to 9 mL of 1x PBS for a 10% solution. Add 100 µL of NGS to 9.9 mL of 1x PBS to make 1% solution.
11. Prepare glutamate stock solution (100 mM) by adding 735 mg of L(+)-Glutamic acid to sterile distilled H₂O and bring to a final volume of 50 mL. Solubility at this concentration will be an issue. Adding small volumes (100 µL) of 1 M hydrochloric acid is sufficient to increase solubility.
12. Prepare NBQX stock solution (10 mM) by adding 50 mg of NBQX to sterile distilled H₂O and bring to a final volume of 13 mL.

2. Preparation of culture dishes and coverslips (Done the day before dissection)

NOTE: We have found that combining three coating agents, poly-L-lysine, poly-L-ornithine, and laminin allows for ideal cell adhesion and viability.

1. Place 10 35 mm Petri dishes in a biological safety cabinet. Place two circular 12 mm coverslips in each dish and fill with 70% EtOH for 10 min. Use a vacuum line to aspirate the remaining EtOH from each dish, allowing the EtOH to evaporate completely.
2. Pipette ~90-100 µL of 0.1% poly-L-lysine solution onto each coverslip, making sure that the entire coverslip is covered by the poly-L-lysine solution. Cover the dishes with lids and place in a 37 °C incubator for 1 h.
3. Aspirate remaining poly-L-lysine solution from each coverslip and rinse with sterile H₂O.
4. Repeat steps 2.2 – 2.3 with 0.1% poly-L-ornithine solution.
5. Again, repeat steps 2.2 – 2.3 with 0.01% laminin solution. Place in a 37 °C/5% CO₂ incubator until ready for cell plating on the following day.

3. Mouse embryonic dissections

NOTE: We use between 4 to 6 timed pregnant mice per culture. While much of the dissection process occurs outside of a biological safety cabinet it is still important to maintain sterile procedure. Plentiful use of 70% EtOH on surfaces near the dissection microscope and on surgical tools is ideal. A mask may also be worn during the dissection to further prevent contamination. Additionally, we use 4 separate antibiotics in the culture medium, so contamination is unlikely. However, if use of antibiotics is problematic, this dissection setup could be moved inside a sterile hood. To preserve cell viability all dissection solutions should be pre-chilled at 4 ˚C, and dissections should be completed as quickly as possible. We do not perform the dissections on ice. The method for dissection of mouse embryonic midbrain neurons is identical to previously described methods^9,10.

1. Prepare a space on a bench near a dissection microscope with an absorbent pad and spray liberally with 70% EtOH.
2. Spray two 100 x 15 mm glass Petri dishes and one 50 x 10 mm glass Petri dish with 70% EtOH and allow EtOH to evaporate. Once evaporated, place 50 mL of sterile 1x HBSS into each 100 x 15 mm Petri dish.
3. Submerge surgical scissors, forceps, and microtome blade in 70% EtOH for 10 min minimum to sterilize. Place instruments on the absorbent pad to dry.
4. Using CO₂ followed by cervical dislocation, euthanize 2-3 month old timed pregnancy mice on embryonic day 14.
5. Spray the abdomen of the euthanized mice with 70% EtOH. Using forceps grab the lower abdomen and open the abdominal cavity using surgical scissors. Start cutting near where the forceps are holding the abdomen, making lateral cuts on each side until the abdominal wall can be folded back and the uterus is clearly visible.
6. Using surgical scissors, cut both ends of the uterine horn. Then remove the uterus and place into Petri dish with 1x HBSS.
7. Using straight-tip forceps carefully remove embryos from the uterus. Leave embryos in HBSS throughout this process. Using either the forceps or a microtome blade, quickly decapitate embryos by cutting near the neck. Making as level a cut as possible.
8. Under a dissecting microscope, move an embryo head to a dry 50 mm Petri dish and place on the ventral side. Stabilize the head with forceps by placing and penetrating near the eyes/snout. Forceps should be angled downward at ~45° to avoid penetrating the mesencephalon.
9. Using the forceps in the other hand, carefully remove the translucent layer of skin and skull just before the prominent ridge of the mesencephalon. Start near the midline and remove skin and skull caudally until the mesencephalon is fully exposed.
10. Hold the forceps perpendicular to the exposed mesencephalon with one tip between the cortex and mesencephalon and the other near the cerebellum. Press down and pinch the forceps together to remove the entire midbrain. The midbrain segment should be approximately 0.5 mm thick. Place the midbrain segment into the second Petri dish filled with fresh 1x HBSS. Repeat this process for each embryo.
11. Using the dissection microscope, position the brain segment with the ventral side facing up. If the meninges are still attached, carefully remove it by grabbing with the forceps and lifting up and away from the brain segment.
12. The brain segment should have 4 visible quadrants. Place the segment in such a manner that the two smaller quadrants are positioned superior to the two larger quadrants. There is a prominent ridge separating the superior two (small) quadrants from the inferior two (large) quadrants.
13. Using the forceps pinch and separate the superior quadrants from the inferior quadrants, and then discard the superior quadrants. The remaining inferior quadrants will have excess tissue laterally on the dorsal side, this tissue will look less opaque than the remaining ventral tissue. Remove the less dense dorsal tissue and discard. The remaining segment should contain both the Substantia nigra pars compacta (SNc) and the ventral tegmental area (VTA).
14. Using the forceps cut the remaining ventral tissue segment into 4 smaller pieces and using a 1mL wide bore pipette transfer these segments in a 15 mL conical tube with 1x HBSS. Keep the conical tube with brain segments on ice throughout the procedure.
15. Repeat this process for all remaining brain segments.

4. Dissociation of cells

1. Enzymatic digestion of cells
   1. Carefully aspirate the HBSS from the 15 mL conical tube containing midbrain segments, leaving the segments at the bottom of the tube.
   2. Add ~800 µL of papain solution to the tube and place in a 37 °C incubator for 7 min. Resuspend cells by flicking the tube and replace to the 37 °C incubator for an additional 7 min.
   3. With a wide-bore 1 mL pipette tip remove only the midbrain segments into a 1 mL aliquot of DNase. Allow the segments to reach the bottom of the aliquot or about 1 min of exposure.
   4. With a wide-bore 1 mL pipette tip remove only the midbrain segments into a 15 mL conical tube containing 2 mL of stop solution. Allow segments to settle at the bottom of the tube and repeat the rinse in an additional conical tube filled with stop solution.
2. Mechanical trituration of cell suspension
   1. In the second stop solution rinse tube, using a wide-bore 1 mL pipette tip, pipette the cells up and down 10 times until there are no large tissue segments visible. It is important to avoid over trituration for minimal cell lysis.
   2. Slowly pipette 300 µL of 4% BSA solution to the bottom of the 15 mL conical tube containing brain segments. Carefully remove the pipette tip to maintain a suspension layer. Centrifuge at 0.4 x g for 3 min. Then carefully aspirate the supernatant and resuspend cells in 400 µL of cell culture medium.

5. Plating the cells

NOTE: Based on experience, about 100,000 viable cells per embryo are collected. 2-3 month old timed pregnant mice typically have litter sizes of 8-10 embryos; therefore, a rough estimate for total yield of cells per timed pregnant mouse is approximately 1 million cells.

1. Using a hemocytometer preform a cell count and then dilute the suspension to 2,000 cells/µL using cell culture medium. Triturate briefly to mix.
2. Remove coverslips with laminin solution from step 2 from the incubator and aspirate the remaining laminin solution from the coated coverslips using a vacuum. Plate quickly to avoid the coverslips from drying completely. Pipette 100 µL (2.0 x 10⁵ cells/coverslip) onto each coverslip and place Petri dishes into a 37 °C incubator for 1 h.
3. Carefully add 3 mL of cell culture medium to each dish and place back into the 37 °C incubator. Preform half medium changes 2 times per week for 2 weeks.

6. Infection of cell culture at 14 DIV with adeno-associated viral (AAV) vectors

1. For each dish prepare 1 mL of serum free DMEM medium with 1 µL of hSyn-GCaMP6f AAV (1.0 x 10¹³ titer)
2. Aspirate the cell culture medium from each dish and replace with 1 mL of serum free DMEM containing hSyn-GCaMP6f. Place dishes back into the 37 °C incubator for 1 h.
3. Aspirate the serum free medium containing AAVs and replace with 3 mL of cell culture medium. Place dishes back into the 37 °C incubator. We have found that 5-7 days of AAV infection allows for ideal levels of GCaMP expression. Continue to change medium every 2-3 days throughout this period of viral infection.

7. Live confocal Ca²⁺ imaging between 19-21 DIV

NOTE: As mentioned in step 6.3, imaging can be done between 5-7 days following viral infection. This is the ideal window to achieve visible expression of the fluorophore at levels which allow for detection of spontaneous Ca²⁺ activity.

1. Preparation of recording buffers
   1. To make 1 L of HEPES recording buffer, add: 9.009 g of NaCl, 0.3728 g of KCl, 0.901 g of D-glucose, 2.381 g of HEPES, 2 mL of 1 M CaCl₂ stock solution, and 500 µL of 1 M MgCl₂ stock solution to 800 mL of sterile distilled H₂O. Bring the pH to 7.4 with NaOH. Bring to a final volume of 1 L.
   2. To make 200 mL of 20 µM glutamate recording buffer, dilute 40 µL of 100 mM glutamate stock solution into 200 mL of HEPES recording buffer described above.
   3. To make 200 mL of 10 µM NBQX recording buffer, dilute 200 µL of 10 mM NBQX stock solution into 200 mL of HEPES recording buffer.
2. Confocal imaging
   1. Fill a sterile 35 mm Petri dish with 3 mL of recording buffer.
   2. Remove a 35 mm Petri dish with infected cultures from the 37 °C incubator. Using fine tip forceps, carefully grab the edge of one coverslip and transfer it quickly into the Petri dish filled with recording buffer. Place the remaining coverslip in medium back into the 37 °C incubator. Transport the dish with recording buffer to the confocal microscope.
   3. Start the imaging software. Proceed to next step while it initializes.
   4. Start the peristaltic pump and place the line into the recording buffer. Calibrate the speed of flow to be 2 mL/min.
   5. Transfer the infected coverslip from the 35 mm Petri dish into the recording bath.
   6. Using the 10x water immersion objective and BF light, find the plane of focus and look for a region with a high density of neuron cell bodies. Switch to the 40x water immersion objective and using BF light refocus the sample.
   7. In the “Dyes list” window within FluoView select AlexaFluor 488 and apply it.
   8. AAV expression can be variable; therefore, in order to prevent overexposure and photobleaching of the fluorophores, start with low HV and laser power settings. For the AlexaFluor 488 channel, set the high voltage (HV) to 500, the gain to 1x, and offset to 0. For the 488 laser line set the power to 5%. In order to increase the effective volume imaged in the z-plane, increase the pinhole size to 300 µm. Use the “focus x2” scanning option to optimally adjust emission signals to sub-saturation levels. From here, settings can be adjusted until ideal visibility of each channel is achieved.
      NOTE: To accurately capture the full range of Ca²⁺ fluxes with GCaMP, adjust the baseline HV and laser power settings in order to allow for an increase in fluorescent intensity without oversaturating the detector.
   9. Once microscope settings are optimized, move the stage in order to locate a region with multiple cells displaying spontaneous changes in GCaMP6f fluorescence and focus to the desired plane for imaging.
   10. Use the “Clip rect” tool to clip the imaging frame to a size that can achieve a frame interval of just under 1 second. This is necessary to set the imaging interval at 1 frame per second.
   11. Set the “Interval” window to a value of 1.0 and the “Num” window to 600.
       NOTE: In order to deliver different recording buffers at the desired time point (300 s), it is important to calibrate the latency of the pump to deliver the new solution to the bath. This will be dependent on the solution perfusion rate (2 mL/min) and the length of the line used to pump solution.
   12. To capture a t-series movie select the “Time” option and then use the “XYt” scanning option to begin imaging.
   13. Watch the imaging progress bar and move the line from the HEPES recording buffer into the 20 µM glutamate recording buffer at the appropriate time point (e.g., if the latency of the pump is calibrated to deliver solution at 60 s, move the line into the glutamate buffer at 240 frames in order to deliver glutamate at 300 s).
   14. When imaging is complete, select the Series Done button and save the finished t-series movie. Continue to perfuse 20 µM Glutamate for an additional 5 min, so that the cultured neurons have been exposed to glutamate for a total of 10 min. Repeat this process for each coverslip to be imaged.
   15. Following the additional 5 min exposure to 20 µM Glutamate, remove the coverslip from the bath and place back into the 35mm Petri dish containing recording buffer until the day of imaging is completed. When finished, proceed to step 8.
3. Ca²⁺ trace analysis
   1. Perform image analysis in ImageJ. Install the BIO-FORMATS plugin for ImageJ, which will allow .OIB image files to open.
   2. In the ImageJ toolbar, click Analyze | Set Measurements, and select the box for Mean gray value (MGV).
   3. In ImageJ, open a t-series movie as a hyperstack.
   4. Drag the slider for the movie and identify the frame with maximal glutamate response to visualize all neurons that respond to glutamate. Use the polygon tool to trace all visible neuron cell bodies, adding their ROIs to the “ROI manager” list.
   5. When finished tracing and adding ROIs, select all ROIs within the ROI Manager window and use the Multi measure selection in the more list of options. Copy and paste these data into a spreadsheet. Complete this process for all movies to be analyzed.
   6. For each ROI, convert the raw MGV data from each frame to ΔF/F₀ values using the equation: ΔF/F₀ = [F(t) – F₀] / F₀. Where F(t) = MGV of any given frame, and F₀ = average baseline MGV of ~10 frames where no Ca²⁺ fluxes are present.
   7. Using a statistical software such as OriginPro 2020, converted ΔF/F₀ traces can be made into line graphs. The “Peak analyzer” function can be used (or similar function if using a different software) to measure the peak amplitude of glutamate response, latency to respond to glutamate, and area under the curve.

8. Immunostaining of cultures

NOTE: Following fixation with formalin, coverslips can be stored in 1x PBS at 4 °C until ready to be processed for immunostaining. Primary and secondary antibody incubation was done in a serial manner, as such incubation with anti-Caspase-3 primary antibody and its complementary secondary antibody preceded incubation with the anti-TH primary antibody and its complementary secondary antibody.

1. Immediately following glutamate exposure, place the coverslip back into its 35 mm Petri dish, aspirate the recording buffer, and add 3 mL of 10% formalin. Let sit for 40 min at room temperature (RT).
2. Rinse the dish 3 times with 1x PBS.
3. Aspirate the PBS and permeabilize cells in 1 mL of 0.01% Triton X-100 in PBS for 2 min.
4. Rinse the dish 3 times with 1x PBS.
5. Aspirate the PBS and block cells in 1 mL of 10% NGS in PBS for 40 min.
6. Rinse the dish 3 times with 1x PBS.
7. Add 1 µL of rabbit anti-Caspase-3 primary antibody to 1 mL of 1% NGS in PBS (1:1000 dilution). Aspirate PBS from the dish and replace with the primary antibody solution. Place on a shaker and incubate for 1.5 h at RT.
8. Rinse the dish 3 times with 1x PBS.
9. Add 1 µL of goat anti-rabbit AlexaFluor 488 secondary antibody to 1 mL of 1% NGS in PBS (1:1000 dilution). Aspirate PBS from the dish and replace with the secondary antibody solution. Place on a shaker and incubate for 1 h at RT. Moving forward protect the samples from light.
10. Rinse the dish 3 times with 1x PBS.
11. Repeat steps 8.7 – 8.10, but using the chicken anti-TH primary antibody (1:1000) in step 8.7 and the goat anti-chicken AlexaFluor 594 secondary antibody (1:1000) in step 8.9.
12. Following the final PBS rinse, place 30 µL of mounting medium onto a microscope slide. Using forceps grab a coverslip from the 35 mm Petri dish and place the coverslip with the cells facing down into the mounting medium. Both coverslips will fit on a single microscope slide if placed correctly. Place in a dry, dark area and allow the mounting medium to dry overnight.

9. Confocal imaging of immunostained cultures

1. Confocal imaging
   1. Start the imaging software. Place the sample onto the microscope stage.
   2. With the microscope eyepieces, using a 20x magnification objective and epifluorescent light with a TRITC filter, focus the sample and search for a TH+ cell body.
   3. Once locating a TH+ cell body, center it in the field of view and then move to the 60x magnification objective.
   4. Select the AlexaFluor 488 and AlexaFluor 594 dyes in the “dye list” window.
   5. As with live imaging, start with low HV, gain, offset, and laser power settings to prevent photobleaching. Use the “focus x2” scan option to assess the fluorescent intensity of each channel and adjust accordingly. As these images will later be quantified for fluorescent intensity it is necessary to keep the imaging settings consistent across all fields of view. Therefore, it is best to look at a few examples of each condition to get an idea of the range of fluorescent intensity across samples.
   6. Once ideal imaging settings are determined, select the “focus x2” scan option and move the cell of interest into the center of the field of view. Increase the digital zoom to 3x using the “zoom” slider.
   7. Using the focus knob, find the plane of focus with the brightest fluorescence and capture a single plane XY image. Save the image to finish.
   8. Switch back to the 20x magnification objective to search for another TH+ cell. Repeat this process until the desired number of cells have been sampled from each condition.
2. Image analysis
   1. In the ImageJ toolbar, click Analyze | Set Measurements, and select the boxes for Area, Integrated density, and Mean gray value.
   2. Open an image as a hyperstack with each channel separated by dragging and dropping into the ImageJ toolbar or selecting the image via the file menu.
   3. Use the TH channel (594 nm) to draw ROIs around the cell body. Using the polygon tracing tool in ImageJ, closely trace the outer edge of the cell body. Where the distance between the cell membrane and cell nucleus is the smallest, trace a straight line through the cytosol to the edge of the nucleus and then closely follow the outline of the nucleus in order to exclude it. Then trace a straight line back to the external membrane, bordering the initial line as closely as possible, and continue to follow the outline of the cell body until the ROI is complete.
   4. Using the keyboard shortcut “T” or using the toolbar menu path Analyze | Tools | ROI Manager, open the ROI manager and add the ROI that was just drawn to the list.
   5. Select the window of the caspase-3 channel (488 nm), and then select the added ROI in the “ROI Manager” list.
   6. In the ROI Manager window, select the Measure button. The results window will appear with the measurements set previously. Copy these to a spreadsheet and repeat this process for each cell.