All procedures described below were approved by the Washington University Animal Studies Committee in compliance with AAALAC guidelines. Imaging studies were performed on transgenic mice expressing GCaMP6f under control of a mouse Thy1 promoter acquired from Jackson Laboratories (JAX Strain: C57BL/6J-Tg(Thy1-GCaMP6f)GP5.5Dkim; stock: 024276). GCaMP6/Thy1 transgenic genotypes were confirmed by PCR using the forward primer 5′-CATCAGTGCAGCAGAGCTTC-3′ and reverse primer 5′-CAGCGTATCCACATAGCGTA-3′. Histology of GCaMP6 fluorescence is shown in Figure S1 . Electrophysiology studies were performed on C57Bl6/J mice (Jackson Laboratories; stock: 664). All mice were male, 12 to 16 weeks of age, weighed 28-36 g, and raised in standard cages in a dedicated mouse facility with a 12hr-12hr light/dark cycle.

Method Details

Animal Preparation for Optical Imaging Fourteen GCaMP6 mice (12-16 weeks of age, 28-36 g) were used for imaging in this study. Genetically modified GCaMP6 mice were studied solely for the purpose of optical calcium imaging, and were otherwise phenotypically normal. Mice were sedated with isoflurane (3% induction, 1% maintenance, 0.5 L/min) and placed in a stereotactic holder. The head was then shaved, and a midline incision made to expose the skull. Body temperature was maintained at 37°C using a temperature controlled heating pad. Chronic cranial windows made of Plexiglas and with pre-tapped holes were fixed to the skull using dental cement (C&B-Metabond, Parkell, Edgewood, NY, USA).

Animal Preparation for Electrophysiology Thirteen C57Bl6/J mice were used to obtain electrophysiological recordings. Mice were given dexamethasone (20 μL 4mg/mL, S.C.) 4 hours prior to surgery, and mannitol (150 μL, 20% manitol, I.P.) immediately prior to surgery. Mice were anesthetized using isoflurane anesthesia (3% induction, 1.5% maintenance). Once anesthetized, lidocaine anesthetic was given locally, scalp hair was removed, a midline incision was made in the scalp, and the scalp was retracted. The periosteal membranes were removed. Two craniectomies (1mm in diameter) were performed over the motor and visual locations determined by stereotactic coordinates derived from the Granger Causality analysis shown in Figure 2 C (Motor cortex = 1.5 mm Left of bregma, 1.6 mm anterior to bregma; visual cortex = 2.6 mm Left of bregma, 3.0 mm posterior to bregma). A third craniotomy was made on the right hemisphere (2.3 mm Right of bregma, 0.9 mm anterior to bregma), and a permanent ground wire was placed and secured with C&B Metabond dental cement (Parkell, Edgewood, NY, USA). A custom-made fixation block with screw threading was attached to the skull with dental cement (to enable head fixation during recording). The left hemisphere craniectomies were covered with a self-healing silicone polymer that allowed silicone electrodes to pass through undamaged. Mice were given S.C. buprenorphine at the end of the procedure for pain control, and mice were given 5 days of recovery time before any recording was performed.

Optical Imaging System Sequential illumination was provided by four LEDs: 470nm (measured peak λ = 454nm (referred to as 454nm LED in this study), LCS-0470-15-22, Mightex Systems, Pleasanton, CA, USA), 530nm (measured peak λ = 523nm, LCS-0530-15-22), 590nm (measured peak λ = 595nm, LCS-0590-10-22), and 625nm (measured peak λ = 640nm, LCS-0625-03-22). The 454nm LED is used for GCaMP excitation, and the 523nm, 595nm, and 640nm LEDs are used for multispectral oximetric imaging. The 523nm LED was also used as an emission reference for GCaMP6 fluorescence in order to remove any confound of hemodynamics in the fluorescence signal (described below). Both the 454nm and 523nm LED light paths were made collinear by using a multi-wavelength beam combiner dichroic mirror (LCS-BC25-0505, Mightex Systems, Pleasanton, CA, USA). For image detection, we used a cooled, frame-transfer EMCCD camera (iXon 897, Andor Technologies, Belfast, Northern Ireland, United Kingdom) in combination with an 85mm f/1.4 camera lens (Rokinon, New York, NY, USA). The acquisition framerate was 16.8Hz per channel, with the overall framerate of the camera as ∼67Hz. This framerate is well above the temporal resolution necessary to adequately characterize hypothesized GCaMP6 activity. To increase frame rate as well as increase SNR, the CCD was binned at 4 × 4 pixels; this reduced the resolution of the output images from full-frame 512 × 512 pixels to 128 × 128 pixels. Both the LEDs and the exposure of the CCD were synchronized and triggered via a DAQ (PCI-6733, National Instruments, Austin, TX, USA) using MATLAB (MathWorks, Natick, MA, USA). The field-of-view was adjusted to be approximately 1 cm2 resulting in an area that covered the majority of the convexity of the cerebral cortex with anterior-posterior coverage from the olfactory bulb to the superior colliculus. The resulting pixels were approximately 78μm x 78μm. To minimize specular reflection from the skull, we used a series of linear polarizers in front of the LED sources and the CCD lens. The secured mouse was placed at the focal plane of the camera. The combined, collimated LED unit was placed approximately 8 cm from the mouse skull, with a working distance of approximately 14cm as determined by the acquisition lens. A 515nm longpass filter (Semrock, Rochester, NY, USA) was placed in front of the CCD to filter out 470nm fluorescence excitation light and a 460/60nm bandpass filter (Semrock, Rochester, NY, USA) was used in front of the excitation source to further minimize leakage of fluorescence excitation light through the 515nm longpass filter. The pulse durations for the LEDs are 20ms, 5ms, 3ms, 1ms for 454nm, 523nm, 595nm, and 640nm, respectively.

Electrophysiology System For electrophysiology recordings, mice were placed on a felt hammock and the skull was fixed to a secure bar via the fixation block. Two 1.5mm 16-channel linear array electrodes (NeuroNexus model number A1x16-5mm-100-703-A16, Ann Arbor, MI, USA) were attached to separate micromanipulators (David Kopf Instruments, Los Angeles, California, USA). Electrodes were painted with DiI (1,1’-Dioctadecyl-3,3,3′,3′-Tetramethylindocarbocyanine Perchlorate; Sigma-Aldrich, St. Louis, MO, USA), and placed into the brain (through the transparent silicone sealant which enabled direct visualization of the cortex) under direct visualization using a surgical stereoscope (Olympus, Tokyo, Japan). Electrode placement was confirmed in three ways: 1) the most superficial contact was visually guided to just under the cortical surface; 2) the electrophysiological signal in the most superficial contact during the transition from noise/air to brain was monitored 3) electrodes were painted with DiI and placement was confirmed with histologic sections of the mouse brain ( Figure 4 A). Local field potentials were recorded using an amplifier with high-pass filter cutoff of 0.02Hz (Intan RDH2132) connected to the recording computer through an acquisition board (OpenEphys), with a reference wire positioned on the right hemisphere contralateral to the electrodes (2.3 mm Right of bregma, 0.9 mm anterior to bregma). All recordings were made in a completely dark room. For each mouse, awake recordings were done first, followed by ketamine/xylazine administration via I.P. injection in the same session (i.e., without removing the electrodes from the brain).

Awake Recordings As described in the electrophysiology system description, awake mouse recordings (imaging and electrophysiology) were performed by mice on a felt hammock with head-fixation, either to the optical window in the case of imaging or to the skull in the case of electrophysiology. The hammock provided a dark, comfortable environment while preventing the awake mouse from applying torque on their restrained head. After recovery from surgery, the mouse was acclimated to the hammock apparatus by a training period consisting of two 20 minute sessions. Acclimation is indexed by a return to normal behavior (e.g., whisking, grooming, and walking with head restrained). Though no accelerometers or other behavioral measures were used to track motion within the pouch during recordings, mice were qualitatively observed to be relaxed with infrequent limb motion after completion of the acclimation protocol. For 7 of the 14 mice, awake imaging was performed for 60 minutes on two separate days, separated by two weeks. For the remaining 7 mice, awake imaging was performed for 60 minutes on a single day. The 60 minute imaging sessions were acquired over 12 5-minute runs. Awake electrophysiology was acquired continuously for 60 minutes in 13 mice.

Anesthetized Recordings For ketamine anesthetized imaging and electrophysiology, mice were anesthetized with I.P. injection of a ketamine/xylazine cocktail (86.9 mg/kg Ketamine, 13.4 mg/kg Xylazine). For dexmedetomidine imaging and electrophysiology, mice were anesthetized with I.P injection of dexmedetomidine (0.5 mg/kg). Anesthetic effect was verified by confirming that the animal was not responsive to a hind paw pinch. The animal was placed and kept on a solid state water circulating heating pad (T/Pump Classic, Stryker Co., Kalamazoo, MI, USA), maintained at 42°C. For 7 of the 14 mice, anesthetized imaging was performed for 45 minutes (the duration of anesthetic effect) on two separate days, separated by two weeks. In the remaining 7 mice, anesthetized imaging was performed for 45 minutes on a single day. The 45 minute imaging sessions were acquired over 9 5-minute runs. Anesthetized electrophysiology was acquired continuously for 45 minutes in 13 mice. We also collected data in which Dexmedetomidine anesthesia was instantaneously reversed with an intraperitoneal injection of atipamezole (0.5 mg/kg). Reversal of dexmedetomidine anesthesia (dex reversal) was confirmed by the resumption of mouse reflexes and typical waking behavior, such as whisking and grooming. 7 mice were imaged in the dex reversal condition for 45 minutes each; electrophysiology was obtained from 5 mice in the dex reversal condition for 45 minutes each.