Dental cement

For cranial window surgery, transgenic mice were anesthetized with an intraperitoneal injection of zoletil (0.04 mg/g body weight) and xylazine (0.5 µg/g body weight). Dexamethasone (4 mg/kg) was administered subcutaneously 5 min before surgery to prevent tissue stress and cerebral edema. Viscotears moisturizing gel (Novartis Healthcare) was applied to prevent eye drying. Mice were fixed in a stereotaxic frame (Stoelting) and 37 °C body temperature was maintained by a heating plate (Physitemp). 3 mm craniotomy over the PtA (centered 1.0 mm lateral and 1.7 mm posterior to the Bregma) [3 (link)] was performed as described previously [39 (link)]. A 5-mm round glass coverslip (Menzel, Thermo Fisher) was attached to the skull using cyanoacrylate glass glue (Henkel). A Neurotar head post (Neurotar Ltd., Helsinki, Finland) was cemented to the skull with dental cement (Stoelting) and was later used for head fixation in the Mobile Home Cage system (MHC, Neurotar Ltd., Helsinki, Finland).
Two weeks after surgery Fos-EGFP and Fos-Cre-GCaMP mice were head-fixed in the MHC each day (5 to 40 min) for two weeks for habituation to imaging conditions. In this system, a head-fixed mouse can move around a lifted MHC and freely explore its environment [40 (link)].

Rats (n = 18; n = 8 males, 10 females) were anesthetized with a cocktail of ketamine, xylazine, and acepromazine (80 mg/kg, 1.6 mg/kg, and 5 mg/kg, respectively, i.m.) until a deep plane of anesthesia was attained (abolition of the foot pinch withdrawal reflex). Rats were placed on a stereotaxic frame and the skull was exposed via blunt dissection. Guide cannulas (7.9 mm length, 22 gauge, Plastics One) were placed above the fourth ventricle, 2.5 mm anterior to the occipital suture, on the midline, 6.0 mm below the skull surface. The cannulae were affixed to the skull with 3 screws (0–80 × 3–32, Plastics One) and dental cement (Stoelting Co.) and closed with obturators (0.014–0.36 mm, Plastics One) after suturing. Rats were returned to their home cage (singly housed) and administered daily fourth ventricular (i.c.v.) PBS (0.1 M; in mM: 115 NaCl, 75 Na₂HPO₄, 7.5 KH₂PO₄) to maintain cannula patency and to allow for acclimation to handling. Rats were allowed a minimum of 4 days of recovery prior to experimentation. After 5 days of HFD exposure, all rats were euthanized and the cannula placement was confirmed with i.c.v. administration of the dye, cresyl violet, and gross visual inspection.

A 250 μl polypropylene reservoir was placed over the mucosal graft such that the reservoir had good contact with the surrounding skull. The reservoir was attached to the skull using cyanoacrylate and tested for leaks using sterile saline. After implantation of a screw (Morris Precision screws and parts– 000x 3/32 Flat self tap screws), dental cement from Stoelting was applied to the skull to fix the reservoir (Fig 1D).

Electrode placement was performed 60 days after CCI injury, as previously described [44 (link)]. Using a stereotaxic micromotor drill (Stoelting) equipped with a 0.7 mm carbon steel burr drillbit (FST), two holes were drilled through the skull for reference electrodes at coordinates (1.00 ML, 1.00 AP) and (−1.00 ML, −1.00 AP), along with a ground electrode (−1.00 ML, −5.00 AP). Two more holes were drilled partially through the skull, and screws were inserted as anchors. A 0.125 mm diameter platinum–iridium electrode coated in Teflon (Plastics One, Roanoke, VA, USA) was implanted intracranially within 0.5 mm from the surface of the dura. Dental cement (Stoelting, Wooddale, IL, USA) was applied to secure the electrode. Animals were excluded from the experiment if they had profuse hemorrhage during EEG implementation or if they had lost more than 20% of their body weight throughout the study. Sham animals were subjected to craniectomy surgery and electrode implantation. Electrodes were connected to a commutator (Plastics One) using EEG cables (Plastics One), and then to an amplifier (EEG100C, BioPac) with a gain of 5000, a 100 Hz low-pass filter, a 0.5 Hz high-pass filter, and a 500 Hz sampling rate, and they were recorded continuously for two months using BioPac’s AcqKnowledge software, version 4.0.