Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

All surgical and animal handling procedures were performed as approved under the Guide for the Care and Use of Laboratory Animals (National Research Council) and the Guidelines of the Indiana University School of Medicine Institutional Animal Care and Use Committee. The schematic illustration of the 2-photon imaging setup is shown in Figure 1.

1. Preparing the Animal for Imaging

1. Place sterile gauze, cotton swabs, and autoclaved surgical tools in the surgery area (Table 1). Turn on a Micro Bead Sterilizer for intersurgery sterilization of surgical tools.
2. Anesthetize the mouse (male or female, 1.5–2.5 months, 20–25 g) by intraperitoneal injection (i.p.) of a mixture of ketamine (17.2 mg/mL), xylazine (0.475 mg/mL) and acepromazine (0.238 mg/mL) (6 µL/g body weight). Wait until the proper depth of anesthesia is reached when the animal ceases to respond to a toe pinch stimulus and has no corneal reflex. During the surgery, give a booster dose of 1/3 the original dose of the anesthetic cocktail as needed to maintain the original anesthetic state.
3. Inject buprenorphine subcutaneously (s.c.; 0.05–0.10 mg/kg) as an analgesic agent prior to surgery. Apply protective ointment to the eyes of the animal to prevent corneal drying and cataract formation.
4. Place a GCaMP6s transgenic mouse on a heating pad to prevent hypothermia and cover it with a sterile surgical drape. Stabilize the animal's head with a Head Holding Adaptor for Mice.
5. Shave the head over most of the scalp with a double-edge razor blade. Clean the surgical area with a sterile alcohol preparation pad followed by povidone-iodine solution.

2. Chronic Cranial Window Preparation

1. Make a rectangular cut of the scalp (2 mm x 3 mm) with a pair of scissors on one side (right) of the skull. Push the skin aside with a cotton swab to create an exposure area of >3 mm in diameter (Figure 3A). Remove the connective tissue attached to the skull by gently scraping the skull with a blunt microsurgical blade (Scalpel Blade #10; Figure 3C).
2. Mark the S1 area by gently drawing a circle (3 mm in diameter, center coordination: -1.5 mm from the bregma and 2.5 mm from the midline) on the skull using a dental drill (Figure 2B).
3. Perform the same procedure on the left skull (Figure 2B).
4. Apply a thin layer of Cyanoacrylate Super Glue to the bone to provide a base for dental cement application.
5. Under a dissecting microscope, thin down a circular groove in the skull around the S1 area using a high-speed microdrill (Figure 3D). The purpose is to create a smooth edge for placement of an optical window onto it.
6. Repeatedly apply artificial cerebrospinal fluid (ACSF, room temperature) to the skull periodically during the thinning process to facilitate drilling and dissipate heat. Perform the drilling intermittently to avoid friction-induced overheating. Suck away the bone debris with a house vacuum line or a vacuum pump.
7. After approximately 2/3 depth of the bone is drilled, slowly and carefully thin the remaining 1/3 bone until a circular piece of the skull (bone flap) is completely free from the surrounding skull.
8. Slowly and carefully remove the circular bone flap with a pair of # 5/45 forceps and expose the dura (Figure 2C; Figure 3E).
9. Keep the exposed brain region moist with ACSF (Figure 3E). Avoid any damage to pial vessels, since hemorrhage will alter cerebral blood flow, accelerate brain swelling, and severely degrade imaging quality.
10. Perform the same procedure on the left (contralateral) side of the skull.
11. For assembling the optical window, use optical adhesive to glue and cure between coverglass layers one at a time. If necessary, warm the optical window (50 °C for 12 h) in an incubator to enhance the adhesive bonding. The optical window contains 2 portions: the top portion contains a single round cover glass with a diameter of 5 mm and the bottom portion contains 1–3 round cover glasses with a diameter of 3 mm (Figure 2D).
12. Store the optical window in ethanol (70%, vol/vol) and rinse it with sterile saline before use. Before optical window installation, check the optical window for imperfections, including an incorrect amount of optical adhesive or inaccurate alignment under a stereoscope.
13. Install the optical window over the craniotomy region (Figure 2D). The top portion of the optical window rests on the skull and the bottom portion of the optical window fits within the craniotomized opening and rests on the dura in the presence of CSF (Figure 2D, E).
14. Seal the top portion of the optical window edge to the skull with the Cyanoacrylate Super Glue (Figure 2F; Figure 3F).
15. When the Cyanoacrylate Super Glue has dried, apply black dental cement (Dentsply) to cover the glass edge. Apply dental cement to all the exposed skull and wound margins to block light (Figure 2G-H; Figure 3B).
16. Perform the same procedure on the left side.
17. Allow the animal to recover on the heating pad and return the animal to its home cage for recovery under appropriate monitoring. Provide wet food to facilitate chewing and hydration. To reduce pain, administer buprenorphine s.c. (0.05–2.0 mg/kg) every 8–12 h postinjury for 2 days. Allow the mouse to recover for an additional 7–10 days before imaging.

3. Two-photon imaging

1. On the day of the experiment, anesthetize the animal with ketamine and xylazine (doses and maintenance of anesthesia as described above). Clean the cranial window with 75% (vol/vol) ethanol.
2. Place the mouse under an imaging set-up consisting of a 2P microscope and the Head Holding Adaptor, resting on a heating pad with its head immobilized through ear bars attached to a mouse head holder.
3. Image one side of the optical window at a time. To minimize optical aberrations, make sure that the optical window and right skull are oriented perpendicular to the optical axis of the microscope.
4. Take an initial picture of the cranial window with bright field illumination at 4x magnification as a reference map for registration with higher magnification 2P angiographies (Figure 4A1).
5. Zoom in on the area of interest using 10x magnification (Figure 4A2).
6. Select an area of interest in the S1 area in cortical layers I or II (up to 150 µm below the pial surface). If higher magnification is desired, use a 20X objective.
7. Search for a field of view with multiple neurons. Acquire images using a 2P microscope (Figure 4B). Determine the exposure time and excitation light level (≈910 nm) based on the depth and expression level of GCaMP6s, with weaker signals requiring longer exposure times and concordantly less time resolution. We typically use ~4 µs per pixel exposure times. Acquire images at frame rates of 3–5 Hz at a resolution of 512 × 512 pixels.
8. Start to record the time series. Store coordinates of every region of interest (ROI) relative to the vascular pattern or to fiducial points in low magnification images. Image the same ROI over time. Perform the same procedure to observe calcium dynamics of the S1 area on the left hemisphere. Calculate changes in fluorescence from each ROI.
9. Acquire images either individually or as a time-lapse movie (Figure 4C1-C3, D, E).
10. Study cortical blood flow dynamics in vivo by imaging fluorescent dextran dyes injected into the tail vein of rodents with 2P microscopy. Use vasculatures as landmarks to recognize and image the same ROI in imaging sessions over long periods.

4. Data Processing

1. Use ImageJ and Origin for image analysis. Import image sequences with ImageJ (Figure 5A-B). For individual images (or t-series movies), select an ROI within an imaged neuron and 2 separate nearby regions for background measurement (Figure 5C). Acquire total intensity with an ROI manager (Figure 5C).
   NOTE: Figure 6 shows representative calcium images. Calcium transient (green) and blood vessel (red) show at different time points in seconds (Figure 6A-D) in a mouse at 7 d after the initial window installation. The z-projection of a 3 min recording period for red and green channels (Figure 6E) or green channel (Figure 6F) is also shown. A z-projection displays a compressed image of all images in the movie from which a rough and circle boundary (encompassing the object neuron of interest in all frames) is manually selected as well as the selection of the 2 independent nearby background regions (Figure 6G).
2. From each ROI, calculate the average fluorescence background FB, FB = (FB1+FB2)/2, and the fluorescence changes of the soma (F) expressed as ΔF/F, ΔF/F = (F-FB)/FB.
3. Perform the peak analysis step by step using functions of ImageJ including Baseline Correction, Peak Finding/Determination, Peak Integration, Peak Fitting, and/or Time-Saving Peak Analysis Features.