Imaging Neurotransmitter Release Using Cell-Based Neurotransmitter Fluorescent Engineered Reporters

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

1. CNiFER Implantation into Mouse Cortex

1. Sterilize all surgical tools in an autoclave before surgery. Prepare a semi-sterile field for surgery by wiping with 70% ethanol and laying down a clean lab diaper.
2. Prepare the CNiFER injection pipet by pulling a glass capillary (inner diameter of 0.53 mm) on a vertical electrode puller. Use a pair of no. 5 fine-tip forceps to break the tip of the electrode to a diameter of ~40 μm.
   Note: This is best accomplished under a stereo zoom microscope with a graticule.
3. Anesthetize an adult (postnatal day 60-90) C57BL/6 mouse with isoflurane: 4% (v/v) for induction and 1.5 to 2.0% (v/v) for maintenance. Use tail or toe pinch to make sure that the mouse is fully anesthetized.
   Note: Re-pinch periodically and assess whisker twitching throughout surgery to reassess depth of anesthesia.
4. Cover the eyes with ophthalmic ointment to prevent drying. Mount the mouse in a stereotaxic frame with ear bars. Maintain the mouse body temperature at 37 °C using a heat pad regulated by a rectal probe.
5. Shave an area approximately 5 mm by 12 mm with an animal electric shaver. Apply Betadine followed by 70% (v/v) isopropanol. Use a scalpel blade to cut and remove the skin over the skull surface. Use a scalpel blade to remove the periosteum from the surface of the skull. Expose and clean the surface of the skull, as described for stereotaxic surgery.
6. Lower an empty glass pipet to bregma and record the antero-posterior (A/P) and medio-lateral (M/L) coordinates. Referring to the mouse brain atlas, calculate the position of the injection site. Shift the pipet to the target site and mark the skull for subsequent window formation.
   Note: The site of injection and window depend on the region to be studied and the distribution of neurotransmitter or peptide releasing projections in the cortex. For instance, in a recent publication, the stereotaxic coordinates +1.0 to +2.0 mm A/P and +1.0 to +2.0 mm M/L were used to inject CNiFER cells into frontal cortex for in vivo imaging of dopamine release during classical conditioning.
7. Form a 2 mm x 3 mm thinned-skull window.
   Note: The bone in the window should be 15-20 μm thick. The small white spots in the bone should not be visible when the skull surface is moistened, if the bone is sufficiently thinned.
8. Place an artificial cerebrospinal fluid (ACSF) soaked sponge on the window to keep it moist while preparing cells to inject.
9. Harvest the CNiFER clone that was grown in a T75 flask to ~80% confluency. Aspirate the media and wash the cells with sterile phosphate-buffered saline (PBS).Note: Trypsin is omitted for these steps.
10. Remove PBS and use 10 ml of ACSF to dislodge cells from the bottom of the flask. Triturate cells to dissociate cell clumps. Centrifuge and resuspend the pellet in 100 μl of ACSF. Centrifuge for 30 sec at 1,400 x g and remove the supernatant, leaving a pellet covered in ACSF. This step leaves a clump of cells in suspension.
11. Backfill the injection pipet prepared in step 1.2 with mineral oil, load the pipet onto a nanoinjector, and advance the plunger to eject a small bead of oil. Put 5 μl of CNiFER cell suspension onto a strip of plastic paraffin film near the mouse preparation. Draw up either the CNiFERs or control CNiFER cells into the pulled pipette.
12. Move the pipette to the target X and Y coordinates, i.e., A/P and M/L noted in step 1.5. Lower the pipette, piercing the thinned skull, and continue to ~200-400 μm below the skull surface, to deposit CNiFER cells in layers 2/3 of cortex.
13. Inject ~4.6 nl of CNiFER cells at the deepest site with the nanoinjector, note movement in the oil and cell interface and then wait for 5 min for the cells to dispense. Withdraw the pipette ~100 μm and inject another ~4.6 nl of CNiFER cells, wait 5 min. Then withdraw the pipette slowly and gently to prevent backflow of the CNiFERs. Repeat injections at one or more adjacent sites.
14. Repeat injection steps 1.8-1.12 with control HEK293 cells (i.e., HEK293/TN-XXL/Gqi5 clone lacking G protein-coupled receptor [GPCR]). Separate the CNiFER and control cell injection sites by ~200 μm.
15. After completing cell implantations, rinse the thinned-skull window with ACSF and wait for the skull to dry. Apply a drop of cyanoacrylate glue (see Materials) over the window and quickly place a pre-cut sterile cover glass on top of the glue. Gently push the cover glass against the skull for a few seconds. Let the glue dry for 2 min.
16. Seal the edges of the cover glass with dental cement and form a well around the window to hold water for the dipping objective.
17. For immobilizing the mouse's head during imaging, attach a custom-built head-bar with a small drop of cyanoacrylate glue behind the window. Let the glue dry thoroughly and then add additional dental cement to further reinforce the custom-built head-bar.
18. Cover the rest of the skull surface, except for the window, with a layer of dental cement. Make sure the edges of the skin are covered by cement and let it dry for 20 min.
19. Following the surgery, stop isoflurane administration and leave the mouse on a heating pad in a cage until it fully recovers from anesthesia. Inject 5% (w/v) glucose in saline (subcutaneous [s.c.]) for rehydration and 0.05 to 0.1 mg/kg buprenorphine (intraperitoneal [i.p.], instant release) for post-operative analgesia.
    Note: To minimize potential immunological reaction to the human CNiFERs, inject the mouse daily with 20 μl/100 g cyclosporine (i.p.) starting the day before injection of CNiFERs.
20. Return the mouse to its home cage for food and water.

2. In Vivo Imaging of CNiFER Clones

Note: Live imaging is conducted with mice using a two-photon microscope and a head-fixed apparatus. No anesthesia is needed during imaging sessions. When imaging animals in the awake state, limit head restraint to only a few hours at a time to reduce stress levels. Return the animal to it home cage between imaging sessions for food and water. Potential stress is minimized by darkening the room lights and surrounding part of the mouse in an enclosure.

1. On the day after surgery, mount the mouse on an imaging platform by screwing the metal head-bar implanted on the skull to the head-fixation frame.
   Note: When imaging awake mice, the imaging session should not exceed a few hours due to the potential stress induced by the head-restraint device.
2. Place the imaging platform with the head-restrained mouse in a two-photon imaging microscope equipped with a 10X (0.30 NA) and 40X (0.80 NA) water immersion objectives.
3. Insert filter cube for fluorescence resonance energy transfer (FRET) imaging (Enhanced Cyan Fluorescent Protein [eCFP] and Citrine) that has a dichroic mirror at 505 nm and band-pass filters that span 460 nm to 500 nm for measuring eCFP and 520 nm to 560 nm for measuring Citrine.
4. Add ACSF to the well containing thinned-skull window and lower the water immersion objective into the ACSF. Use the eyepiece in conjunction with a mercury lamp and green fluorescent protein (GFP) filter cube to locate the surface of the cortex and vasculature below the window.
   Note: The pattern of the vasculature helps locate and image the same region over repeated days of imaging. Switch to the 40X water immersion objective to locate CNiFERs by manually focusing on surface of the cortex over the cells using the GFP filter cube and a mercury lamp.
5. Set up for two-photon imaging. Select the appropriate light path for two-photon imaging. For a typical commercial system, use the software to switch to two-photon imaging mode and redirect light to the photomultiplier tubes (PMTs) in the non-descanned detectors. Turn on the near infrared femtosecond pulsed laser, select a wavelength of 820 nm and a power setting of 5-15%. 
   Note: 5% power typically provides ~25 mW at the specimen.
6. Set the PMT1 & PMT2 voltage close to the maximum value, typically 700-1,000 V depending on the PMT. Set the gain to 1 for each channel and zero the z position for the objective.
7. Lower the objective ~100 to 200 μm from the cortical surface and start the x-y scan. Adjust the laser power, gain, and PMT voltage for each channel, i.e., eCFP and Citrine, to optimize the signal-to-noise ratio of the CNiFER fluorescence.
8. Use the zoom feature in the software to restrict the image to a region that contains the CNiFER cells as well as a background region. Use a scan rate no slower than one frame every 2 sec (0.5 Hz) at 4 μs per pixel. Adjust the line averaging for suitable signal-to-noise ratio, e.g., Kalman 2-line averaging.
9. Draw a region-of-interest (ROI) around CNiFER cells, surrounding about 3 to 4 cells per plane. Set up real-time analysis of ROI average intensities. Start acquisition to monitor CNiFER fluorescence over time.
10. Collect fluorescence from CNiFERs before and during experimental manipulations, e.g., electrical stimulation, Channelrhodopsin-2 (ChR2) stimulation, behavior, as determined by the user.
11. When the imaging experiment is completed, return the mouse to its home cage. Repeat the imaging across days, as desired. When re-imaging the cells refer to the previously acquired low-magnification vasculature image to orient back to the same imaging field (step 2.4).
    Note: Implanted CNiFERs can be imaged for at least 7 days.