Isoflurane inhalation

For morphological studies, 3 mice/group were anesthetized with isoflurane inhalation (Abbott Scandinavia AB, Sweden) and perfused transcardially with 0.1 M phosphate buffer + 0.3% heparin (+37°C), followed by perfusion with a fixative solution (2% paraformaldehyde and 0.25% glutaraldehyde (Sigma-Aldrich, USA) in 0.1 M phosphate buffer, +4°C). After perfusion, the brains were dissected and split into two hemispheres. The left hemispheres were used for immunohistochemistry, whereas the isolated hippocampi from the right hemispheres were examined under light and transmission electron microscopy. Coronal sections of the dorsal hippocampus were obtained from the anatomical areas between 1.65 and 2.48 mm posterior to the bregma, as defined by the Allen Mouse Brain Atlas (Figures 3A, 4A) (22 (link)). For biochemical assays, the remaining 5–6 animals/group were anesthetized with an overdose of pentobarbital (Nordvace, Sweden), after which their brains were quickly removed; both the left and right hippocampi were isolated on pre-chilled plates and immediately frozen until analysis.

Rat surgery was performed by Dr. Jwo at the Chang Gung Memorial Hospital, Keelung [1 (link)]. In brief, following anesthetization using isoflurane inhalation (Abbott, USA), laparotomy was performed via upper median incision. The peritoneal cavity was exposed aseptically. After dividing the small bowel between the proximal jejunum 3 cm from the Treitz ligament and midgut 50% length of whole intestine, end-to-end jejunojejunostomy anastomosis was performed using 7-0 polypropylene sutures between both severed ends of proximal and distal jejunum to process adaptation segment and continue the intestinal integrity. To establish the neointestinal regeneration model, 0.5 cm donor of autologous intestine, accompanied by mesenteric vessels, was retrieved from the mid-portion of the severed intestinal tissue. Unnecessary redundant tissue was removed, with only 0.5 cm donor intestine being preserved and intubated by a 6 cm soft silastic Penrose tube for neointestinal regeneration. Donor intestinal segments were fixed upon the tube with 7-0 polypropylene sutures. An intraluminal tube served as a tissue scaffold and drained out through the abdominal wall via bilateral tube-enterostomies created in both flanks of the rats. Animals were sacrificed at 1, 4, and 12 weeks after surgery using isoflurane to anesthetization followed by CO₂ inhalation for 5 min.

The rat was anesthetized with 1.5% isoflurane inhalation (Abbott Laboratories, North Chicago, IL, USA). Vital signs, including temperature and respiration rate, were monitored throughout the entire surgical process. A heating pad was used to maintain a constant body temperature at about 37 ± 0.5 °C. The anesthesia depth was assessed by hind paw pinch and respiration rate.
After the rat was anesthetized, an abdominal incision of about 2 cm was made. One pair of cardiac pacing wires (A&E Medical, Farmingdale, NJ) used as electrodes for EMG recording, were fixed in the external oblique muscles of the abdomen^{37 (link)}. The electrodes were spaced 0.5 cm apart, and the connecting wires were tunneled underneath the skin and externalized at the back of the neck. After the surgery, the rat was placed in an individual cage in order to prevent the externalized electrode sires being chewed off by the other rat. Rats were given 5 days for a complete recovery before the experiment was initiated.

Following the acclimatization period rats receive a high-fat diet (HFD) containing 40% digestible energy from lipids (Specialty Feeds SF00-219, Glen Forrest, WA, Australia) for 9 weeks, as described in our previously published study [13 (link)]. Throughout the study, animals could access food and water ad libitum. Animals were then maintained on the HFD and treated for a further six weeks with a daily i.p. injection, with either vehicle (0.9% isotonic saline solution containing 0.75% Tween 80: n = 9–10), or 3 mg/kg body weight of AM251 (Cayman Chemicals, Ann Arbour, MI, USA) dissolved in the vehicle solution (n = 9–10). Following treatment, rats were anesthetized with 3% isoflurane inhalation (Abbott Laboratories, Illinois, United States) with each animal undergoing surgical removal of skeletal muscle, cardiac blood was then collected confirming death, with all other major organs including fat pads were removed post-mortem, weighed, and stored at −80 °C for further analyses.

C57BL/6J or CD1 mice were used to obtain wild-type mouse embryos. Timed mating was used to obtain embryos at the appropriate stages. Pregnant mice were anaesthetised to unconsciousness through isoflurane inhalation (Abbot Laboratories, Sittingbourne, UK) before cervical dislocation. Embryos were transferred into Leibovitzʼs 15 (L-15 Gibco, Thermo Fisher Scientific, Grand Island, NY, and Paisley, Scotland) before decapitation, then further dissection.

All animal study protocols and procedures were approved by the Animal Care Ethics Committee of the Tokyo Medical and Dental University. All experiments were carried out in accordance with the approved guidelines by Science Council of Japan for proper conduct of animal experiments. We used 16 male nude mice (BALB/c Slc-nu/nu, Sankyo Labo Service Corporation, Tokyo, Japan) that were 8 weeks old for this experiment. Isoflurane inhalation (Abbott Laboratories, Queenborough, UK) was used to anesthetize mice. After making an incision in the skull skin, the periosteum was carefully removed and two critical sized bone defects were created on the calvaria (3.75 mm diameter) using a trephine bur (ACE Dental Implant System, Brockton, MA, USA). The cell-transferred amniotic membrane was then trimmed and transplanted to cover the bone defect and the wound was closed using a 7–0 silk suture (Mani, Tochigi, Japan).