Infecting Primary Mouse Midbrain Neurons with Engineered Adeno-Associated Viruses

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

NOTE: Cell culture solutions should be prepared using sterile procedures 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 it 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 it 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 an ascorbic acid stock solution by adding 352 mg of ascorbic acid to sterile distilled H2O and bringing it to a final volume of 20 mL. If necessary, heat in a 37 °C bath to dissolve. 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 a 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) solutions by adding 1 mL of NGS to 9 mL of 1x PBS for a 10% solution and 100 µL of NGS to 9.9 mL of 1x PBS to make a 1% solution.
11. Prepare a glutamate stock solution (100 mM) by adding 735 mg of L(+)-glutamic acid to sterile distilled H2O and bringing it 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 bringing it 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. Cover the dishes with lids and place them in a 37 °C incubator for 1 h.
3. Aspirate the 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 a 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 the 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.

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 it into a 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 it on the ventral side. Stabilize the head with forceps by placing and penetrating near the eyes/snout. The forceps should be angled downward at ~45° to avoid penetrating the mesencephalon.
9. Using the forceps on 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 them by grabbing them with the forceps and lifting them 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 with 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-tituration 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 the total yield of cells per timed pregnant mouse is approximately 1 million cells.

1. Using a hemocytometer, perform a cell count and then dilute the suspension to 2,000 cells/µL using a 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 prevent 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 it back into the 37 °C incubator. Perform half-medium changes twice weekly 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 it with 1 mL of serum-free DMEM containing hSyn-GCaMP6f. Then, place the dishes back into the 37 °C incubator for 1 h.
3. Aspirate the serum-free medium containing AAVs and replace it with 3 mL of cell culture medium. Place the dishes back into the 37 °C incubator. We have found that 5-7 days of AAV infection allows for ideal levels of GCaMP expression. Throughout this period of viral infection, continue to change the medium every 2-3 days.

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

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 the medium back into the 37 °C incubator. Transport the dish with the recording buffer to the confocal microscope.
3. Start the imaging software. Proceed to the 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 the 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 the 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 on 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 it back into the 35 mm Petri dish containing recording buffer until the day of imaging is completed.