Carprofen

Rats were anesthetized with ketamine hydrochloride (70 mg/kg) and xylazine (5 mg/kg) injected intraperitoneally. To reduce pain and inflammation, rats were injected with carprofen (Zoetis) subcutaneously prior to all surgical procedures. Rats were placed in a stereotaxic frame, and an incision was made to expose bregma and lambda. Six anchor screws (Stoelting) were inserted into the skull anterior to the targeted injection site. A small craniotomy was made to target the LC (AP, −12 mm; ML, −1.25 mm from bregma), and a 32-gauge infusion needle attached to a 10 μl Hamilton syringe (Hamilton) was stereotaxically lowered to the target depth (5.5 mm below the pial surface) at an angle of 20° posterior to vertical (Quinlan et al., 2019 (link)). A total volume of 1 μl of virus was infused at a rate of 0.1 μl/min. The needle was held in place for 5 min after the completion of the infusion to allow the virus to diffuse, then slowly raised and removed from the brain. A Ø400 µm core, 0.39 NA, fiber-optic cannula (Thorlabs) was then stereotaxically placed just above the injection site (5.4–5.45 mm below the pial surface) and cemented in place with acrylic. The incision was closed with sutures, and a topical antibiotic cream was applied to the incision site. Rats received 3–7 d of recovery before training was resumed.

Animal experiments were performed with approval of the local State Review Board of Saarland, Germany, and conducted following the national and European guidelines for the ethical and human treatment of animals. Preparation of the bacterial inoculum and infection of the animals were carried out as described [34 (link)], with minor modifications. Briefly, 100 µL bacterial suspensions containing ~10⁷ colony forming units (CFU) were administered intravenously by retro-orbital injection into female, 8- to 10-week-old C57BL/6N mice (Charles River, Sulzfeld, Germany) that were anesthetized by isoflurane inhalation (3.5%; Baxter, Unterschleißheim, Germany). Immediately after infection, mice were treated with a dose of carprofen (5 mg/kg; Zoetis, Berlin, Germany), and at four days post infection, mice were sacrificed, and livers and kidneys were removed. The organs were weight adjusted and homogenized in PBS (Thermo Fisher, Dreieich, Germany), and serial dilutions of the homogenates were plated on blood agar plates to enumerate the CFU rates in the organs.

Pan02 cells were orthotopically implanted in the pancreas of five-week-old C57B1/6 mice (National Laboratory Animal Center, Taipei City, Taiwan). All experimental procedures and animal care protocols were performed in accordance with the guidelines approved by the IACUC of the National Health Research Institute (approved protocol number: NHRI-IACUC-107113-A).
For surgery, the mice were anesthetized by the inhalation of 1–2% isoflurane in oxygen. Carprofen (5 mg/kg, Zoetis, Inc., Taipei City, Taiwan) was injected subcutaneously for analgesia. Hair around the surgical site was shaved, and the pancreas was exteriorized through a small incision in the left flank. Pan02 cells (0.5 × 10⁶) suspended in 50 μL of phosphate-buffered solution (PBS) were injected into the tail of the pancreas using a 28-gauge needle and a syringe [38 ]. The pancreas was then tucked back into the abdominal cavity, and the incision was closed using wound clips and size 3-0 nylon suture (UNIK Surgical Sutures Mfg. Co., Ltd., New Taipei City, Taiwan).

Cranial window surgery was performed as previously described (Mostany and Portera-Cailliau, 2008 (link); Mostany et al., 2013 (link); Voglewede et al., 2019 (link)). To prevent brain swelling and inflammation mice were injected with carprofen (5.0 mg/kg; s.c.; Zoetis Inc., Parsippany-Troy Hills, NJ, USA) and dexamethasone (0.2 mg/kg; s.c.; MWI, Boise, ID, USA) after anesthesia induction and before any incision was made. Mice were anesthetized with isoflurane (5.0% for induction, 1.5%–1.7% for maintenance), placed in a stereotaxic frame, and a 4-mm craniotomy was performed with a pneumatic dental drill over S1BF, centered at 3 mm lateral to the midline and 1.95 mm caudal to bregma. A 5-mm glass coverslip (#1; Electron Microscopy Sciences, Hatfield, PA, USA) was placed over the intact dura and secured using cyanoacrylate glue and dental acrylic (Lang Dental Mfg. Co, Inc., Wheeling, IL, USA) to the skull. A custom-made titanium bar (9.5 mm × 3.2 mm × 1.1 mm) was cemented within the dental acrylic for securing the mouse to the microscope’s stage. A 3-week recovery time was allowed before in vivo imaging.

Kiss1Cre/Cas9-GFP female animals (>2 mo) were checked for estrous cycles for >10 days before surgery; only mice with regular 4–5 day cycles were used. Mice were anesthetized with 1.5–2% isoflurane. AVPV injections were targeted to 0.55 mm posterior to Bregma, ±0.2 mm lateral to midline, and 4.7 and 4.8 mm ventral to dura. Arcuate injections were targeted to 1.5–1.7 mm posterior to Bregma, ±0.2 mm lateral to midline, and 5.9 mm ventral to dura. 100 nl virus injected bilaterally at the target coordinates at ~5 nl/min. The pipette was left in place for 5 min after injection to allow viral diffusion into the brain. Carprofen (Zoetis, Inc., 5 mg/kg, sc) was given before and 24 hr after surgery to alleviate postsurgical pain. Estrous cycle monitoring continued after surgery for up to 8 weeks. Stereotaxic hits were defined as ≥70% infection rate in both hemispheres; only bilateral hits were included for in vivo evaluation of reproductive parameters.