Digital stereotaxic instrument

Further, 6–8 weeks female NCRNU athymic mice were purchased from Taconic Biosciences. All animals were housed in the American Association for Laboratory Animal Care–accredited Animal Resource Center at Moffitt Cancer Center. All animal procedures and experiments were carried out under protocols approved by the Institutional Animal Care and Use Committee of the University of South Florida and Moffitt Cancer Center. All animal studies were performed in accordance with relevant guidelines and regulations of University of South Florida and Moffitt Cancer Center. For orthotopic model, xenograft tumors were established by intracranially injecting 1 × 105 indicated GSCs in a 2–3 μL volume of PBS in the right striatum of mice (n = 5/group) on a Stoelting Digital Stereotaxic Instrument (Stoelting, Wood Dale, IL, USA). Seven days after implantation, the mice were randomized into distinct groups for treatment (as indicated in figures). TMZ, 50 mg/kg/day; 5-Aza, 5 mg/kg/day; Lomeguatrib, 20 mg/kg/day. For survival studies, animals were followed every day until they lost 20% of body weight or had trouble ambulating, feeding or grooming.

After the experimental mice were anesthetized by isoflurane inhalation, they were placed on a digital stereotaxic instrument (Stoelting Co., United States), the heads of the mice were fixed, and the balance was adjusted using anterior tooth adapters and ear bars on both sides. Their skin was cut, and their skulls were exposed. The mouse hippocampal CA1 area coordinates were AP-1.9 mm, ML ± 1.4 mm, DV-1.3 mm, and the hole punch was lightly drilled. After redetermining the position, a microinjection system (Stoelting Co., United States) was used to control a glass electrode microinjector to slowly inject rAAV-EF1a-IFITM3-P2A-mCherry or rAAV-EF1a-mCherry into the bilateral hippocampal CA1 area. The total volume was 300 nL, and the speed was 23 nL/min for about 15 min. After the microinjection stopped, there was a 10 min wait to allow the virus to spread evenly to the tissue. After the needle was slowly pulled out, the head skin was sutured, the surgical area was smeared with aneriodine after surgery, 0.1 mL of 4% ropivacaine was injected locally into the incision, and lidocaine ointment was applied. After the mice were awake for 1 h, they were put back into their cage and were free to move around and eat and drink water.

Rats were exposed to 10% chloral hydrate (300 mg/kg, ip) to induce anesthesia followed by immobilization in a digital stereotaxic instrument (Stoelting, USA). A hole was drilled in the skull after a midline incision. Drugs were injected slowly with a Hamilton microsyringe into one of the LV under stereotactic guidance (Brega: 1.2±0.4 mm, deep: 3.2±0.4 mm, right to median sagittal plane: 1.4±0.2 mm) over 15 min. An additional 15 min was allowed to prevent drug diffusion. Animals were kept warm and allowed to recover from anesthesia. When the rats were completely awake, they were returned to the cage to rest.