Experiment Summary

Genetically encoded fluorescent biosensors are versatile tools for studying brain metabolism and function in live tissue. The genetic information for these biosensors can be delivered into the brain by stereotaxic injection of engineered adeno-associated viruses (AAVs), which can selectively target different cell types depending on the capsid serotype and/or the viral promoter. Here, we describe a protocol for intracranial injections of two viral vectors encoding the metabolic biosensor Peredox and the calcium biosensor RCaMP1h. When combined with 2-photon microscopy and fluorescence lifetime imaging, this protocol allows the simultaneous quantitative assessment of changes in the cytosolic NADH/NAD+ ratio and the intracellular Ca²⁺ levels in individual dentate granule cells from acute hippocampal slices.

Fig. 1 Workflow diagram for biosensor expression in the mouse hippocampus using intracranial injections of adeno-associated viruses.

Materials and Reagents

1. Wiretrol^® II disposable micropipets and plunger
2. MicroFil 28 Gauge, length: 67 mm, non-metallic syringe needle
3. Electrophysiology pipette holder
4. Surgical tape
5. 70% isopropyl alcohol wipes
6. 30 Gauge needle
7. Parafilm
8. Q-tips
9. 3 ml syringe
10. Laboratory marker
11. Mineral oil
12. Experimental animals
13. Viral vectors
14. Sterile 0.9% NaCl saline solution
15. Ice block

Equipment

1. Dumont # 5 forceps
2. P10 pipette
3. Flaming/Brown micropipette puller
4. Digital stereotaxic instrument
5. Microsyringe pump (World Precision Instruments, model UMP3) and controller (World Precision Instruments, model: SYS-MICRO4)
6. DC temperature controller
7. Heating pad (FHC, catalog number: 40-90-2-02) and thermistor probe (FHC, catalog number: 40-90-5D-02)
8. Surgical lamp, dual arm gooseneck illuminator
9. Precision stereo zoom binocular microscope

Procedure

A. Make micropipettes for injections

1. Pull a micropipette from a 10 µl glass capillary tube using a micropipette puller. The micropipettes (from both halves of the capillary) should have a total length of 4.0-4.5 cm, and a taper length (i.e., neck-to-tip) of ~0.5 cm.
2. Break the tip of the micropipette using Dumont #5 forceps.

B. Prepare the viral mix

1. Thaw aliquots of the two desired AAVs on ice.
2. Mix the viral samples at a 1:1 ratio and let the viral mix equilibrate for 10 min. The viral mix is prepared and stored on ice (4°C) until use.

C. Load viral mix into micropipette

1. Fully load one micropipette with mineral oil using a 3 ml syringe with the MicroFil 28 Gauge needle.
2. Place the plunger and the pipette holder onto the microinjector system.
3. Place the plunger first and then place the pipette holder with the side steel tube inlet resting on the plastic mount attached to the stereotaxic instrument.
4. Mount the micropipette in the micropipette holder on the stereotaxic instrument, which must be connected to the microsyringe pump controller.
5. Ensure that the plunger enters the open end of the loaded micropipette. Ideally, this should result in some of the mineral oil coming out of the sharp end of the micropipette.
6. Using the microsyringe pump controller, carefully eject 500 nl of the mineral oil to further confirm that the tapered end of the micropipette is sufficiently open for injections, and that the micropipette does not move with the injector. Eject more mineral oil until 300-500 nl remain in the pipette.
7. Pipette viral mix (typically 4-6 µl to inject 6 pups) onto the parafilm away from the oil drop.
8. Under visual control with the stereo zoom microscope, use the stereotaxic instrument to place the sharp end of the micropipette into the viral suspension.
9. Use the microsyringe pump controller to withdraw the viral mix into the micropipette; avoid taking up air bubbles.
10. Move the injection holder away.

D.Induce cryoanesthesia

1. Place the mouse pups at postnatal day 1 or 2 (P1-P2) in a clean cage and keep them wrapped in a paper towel.
2. Place one mouse at a time between two paper towels on top of an ice block (3-4°C) for 2-5 min.
3. Lean a plastic bag filled with crushed ice over the paper towel covering the mouse.
4. Confirm anesthesia by gently squeezing a paw and monitor the lack of withdrawal reflex after 5 min, then every two minutes.

E. Perform intracranial injections

1. Place another ice block touching the metallic end of the inhalator attached to the stereotaxic instrument, and secure it laterally with the blunt ends of small support bars (Figure 1).

Fig. 1 Typical setup for cryoanesthesia and intracranial injections.

2. Transfer the pup from the ice block used for cryoanesthesia to the ice block secured in the stereotaxic instrument, and position the mouse flat in prone position.
3. Gently secure the pup to the ice block using surgical tape (Figure 2).
4. Using the stereo microscope, visually locate the zero coordinate point (lambda) and mark it with a laboratory marker.

Fig. 2 Schematic representation of a mouse pup under a stereo microscope.

5. Under the stereo microscope, lower the micropipette on this zero coordinate point—tip touching the skull.
6. Set coordinates to zero on the digital manipulator attached to the stereotaxic instrument.
7. Move the micropipette up to avoid breaking the pipette tip, and then move it to the desired anterior-posterior (AP, Y-axis) and medial-lateral (ML, X-axis) coordinates, using the digital manipulator.
8. Gently touch the skull with the tip of the micropipette, and set the dorso-ventral coordinate (DV, Z-axis) on the manipulator to zero.
9. Move up the pipette and mark the location in the skin with a laboratory marker.
10. Make a hole in this spot for micropipette injection by pricking the skin and piercing through the skull with a 30 G needle.

For hippocampal injections, two injections per hemisphere are done in the following sites:

Fig. 2 Coordinates for intracranial injections (mm)

11. Lower the micropipette to the desired depth (Z-axis).
12. Use the microinjector system to inject 150 nl of virus at a rate of 50 nl/min.
13. Wait 1-2 min before withdrawing the micropipette; then withdraw slowly to avoid backflow of the virus to the surface.
14. Before the next injection and monitoring under the stereoscope, confirm that the pipette is not clogged by ejecting 10 nl of viral mix from the pipette.
15. Perform the second injection and repeat step 10.
16. Repeat the same process to inject the other hemisphere (Steps E10-E15).
17. After finishing all injections, dispose the micropipette into a BSL-2 waste container for sharps, and remove the pipette holder and Wiretrol II plunger.

F. Post-injection recovery

1. After injections, place the pup on a heating pad until the mouse begins to move again and the skin color returns to normal.
2. Place the pups in a clean cage until they recover their normal movement.
3. Return the injected pups to their original cage.
4. Fill a surgery card and a surgery log to notify the veterinarians about the procedure.
5. Perform a subcutaneous injection of Ketoprofen (10 mg/kg), once a day for 3 days.
6. Fluorescence can be verified in acute hippocampal slices after two weeks (Figure 3).

Figure 3. Intracranial injection of AAVs induces biosensor expression in the mouse hippocampus.