Atc2000

We measured hemodynamic changes in the heart by pressure-volume (P-V) loop study using ADV500 system, Transonic Scisense, USA (20). We followed the best practice guidelines for invasive hemodynamic measurement in mice described by Lindsey et al. (47 (link)). We anesthetized mice with 1–2% isoflurane, intubated it and connected it to a ventilator (MidiVent, Harvard Apparatus, USA). Then, we opened the chest and created a fine hole in the apex area of left ventricle (LV) using a 27-gaug needle and immediately inserted a 1.2F 4.0 mm catheter probe (Transonic Scisence, USA). The probe was advanced slowly into the LV in order to avoid contact with the LV and septal walls, which was ensured by observing the P-V loop formation. Since body temperature may contribute to the hemodynamic measurements, we maintained the body temperature at 37°C by a temperature controller (ATC2000, World Precision Instruments, USA). Admittance technique and Wei's equation were used to calculate volume (48 (link)). Because heart rate could affect hemodynamic changes, we kept the heart rate between 450 and 500 beats per minute by adjusting anesthesia. Hemodynamic parameters of diastolic function were determined by averaging at least 20 cardiac cycles following a baseline scan, which was performed after proper probe placement was confirmed using the LabChart Pro PV Loop Analysis Add-On.

Surgeries and preparation of animals for experiments were performed as previously described (Rasmussen et al., 2020 (link)). Mice were anesthetized with an intraperitoneal injection of fentanyl (0.05 mg/kg body weight; Hameln), midazolam (5.0 mg/kg body weight; Hameln), and medetomidine (0.5 mg/kg body weight; Domitor, Orion) mixture dissolved in saline. The depth of anesthesia was monitored by the pinch withdrawal reflex throughout the surgery. Core body temperature was monitored using a rectal probe and temperature maintained at 37-38°C by a feedback-controlled heating pad (ATC2000, World Precision Instruments). Eyes were protected from dehydration during the surgery with eye ointment (Oculotect Augengel). The scalp overlaying the longitudinal fissure was removed, and a custom head-fixing head-plate was mounted on the skull with cyanoacrylate-based glue (Super Glue Precision, Loctite) and dental cement (Jet Denture Repair Powder) to allow for subsequent head fixation during video-oculographic tracking. Mice were returned to their home cage after anesthesia was reversed with an intraperitoneal injection of flumazenil (0.5 mg/kg body weight; Hameln) and atipamezole (2.5 mg/kg body weight; Antisedan, Orion Pharma) mixture dissolved in saline, and after recovering on a heating pad for 1 h.

Mice of either sex were implanted with a custom-built head plate (stainless steel, 1.1 g; Fig 1). The animals were anesthetized with isoflurane, and kept on a feedback-controlled heating pad (ATC2000, World Precision Instruments, Inc.). Hair overlying the skull was shaved. Next, the skin and the muscles over the central part of the skull (approximately between the bregma and lambda) were removed. The skull was thoroughly washed with saline, followed by cleaning with a 3% solution of hydrogen peroxide. The head plate (Fig 1) was held in its intended position by an alligator clip, while dental cement (Super-Bond C&B; Sun Medical, Japan) was applied to attach it to the skull. The exposed part of the skull within the hole in the head plate was covered with Kwik-Cast (World Precision Instruments, Inc.). The entire procedure was typically completed in < 40 minutes. Post-operative pain was prevented by administering a non-steroidal anti-inflammatory agent (Rimadyl) before the procedure and on the three following days.

All mice were first implanted with a custom metal head plate. To do so, the animals were anesthetized with isoflurane, and were kept on a feedback-controlled heating pad (ATC2000, World Precision Instruments, Inc.). Hair overlying the skull was shaved and the skin and the muscles over the central part of the skull were removed. The skull was thoroughly washed with saline, followed by cleaning with sterile cortex buffer. The head plate was attached to the bone posterior to bregma using dental cement (Super-Bond C&B; Sun Medical). For electrophysiological experiments, we covered the exposed bone with Kwik-Cast (World Precision Instruments, Inc.), trained the animals in the behavioral task in the following weeks, and subsequently performed a craniotomy over the frontal cortex for lowering the silicon probes. For fiber photometry and optogenetic experiments, after the head plate fixation, we made a craniotomy over the target area (mPFC or VTA) and injected viral constructs followed by implantation of the optical fiber, which was secured to the head plate and skull using dental cement. Post-operative pain was prevented with Rimadyl on the three following days.