Prior to the surgical procedures, the animals were sedated with an intramuscular (im) injection of ketamine (10 mg/kg; Vedco, Saint Joseph, MO, USA) and medetomidine (0.015 mg/kg; Zoetis, Kalamazoo, MI, USA). Tracheal intubation was performed and the animals were placed on inhaled isoflurane (1–4.5%; Piramal Healthcare, Nashville, TN, USA) in oxygen (delivered at 1.0 L/min). Homeostatic monitoring (respirations, vital signs) was performed according to locally established procedures. After completion of the surgical intervention, atipamezole hydrochloride (im, 0.15 mg/kg; Zoetis) was administered and the animals were extubated and returned to their home cages. The animals were visually monitored cage side at 15-min intervals until full recovery from anesthesia. For analgesia and post-operative care, animals received, at minimum, buprenorphine (im, 0.03 mg/kg twice daily for 1.5 days; Patterson Veterinary, Mendota Heights, MN, USA), carprofen (subcutaneously or orally, 2.2 mg/kg twice daily for 2 days; Zoetis), and ceftriaxone (im, 50 mg/kg once daily for 5 days; Patterson Veterinary). Following completion of the carprofen, all animals were given ketofen (im, 2 mg/kg once daily for 3 days; Zoetis).
Ketofen
Manufactured by Zoetis
Sourced in United States
Ketofen is a product developed by Zoetis for use in veterinary settings. It is a laboratory equipment designed to perform specific functions. The core function of Ketofen is to provide a tool for conducting certain tests or analyses, but no further details on its intended use can be provided in an unbiased and factual manner.
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Lab products found in correlation
7 protocols using ketofen
1
Anesthetic and Analgesic Protocol for Surgical Procedures in Animals
2
Stereotactic Surgery for PPTg Implantation
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The drive was lowered into the craniotomy such that the tetrodes extending from its base targeted 100–200 µm above the dorsal surface of the PPTg (2.5 mm from brain surface). The drive was then affixed to the skull using two small screws, a luting, and dental acrylic. One screw was soldered to a ground wire connected to the drive.
In all surgeries, after the acrylic hardened, a topical antibiotic was applied to the scalp around the drive implant, the isoflurane was turned off and oxygen alone was delivered to the animal to gradually alleviate anesthesia. Immediately following surgery, animals were administered sterile 0.9% saline for rehydration and an analgesic (5 mg/kg Ketofen; Zoetis). Post-operative care, including analgesic and antibiotic administration, continued for up to 5 days after surgery and animals were closely monitored for signs of distress.
In all surgeries, after the acrylic hardened, a topical antibiotic was applied to the scalp around the drive implant, the isoflurane was turned off and oxygen alone was delivered to the animal to gradually alleviate anesthesia. Immediately following surgery, animals were administered sterile 0.9% saline for rehydration and an analgesic (5 mg/kg Ketofen; Zoetis). Post-operative care, including analgesic and antibiotic administration, continued for up to 5 days after surgery and animals were closely monitored for signs of distress.
3
Analgesic Efficacy in Piglet Castration
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At 6 days of age (±1 day) piglets were injected intramuscularly with one of 3 treatments; 0.4 mg/kg meloxicam (Meloxicam solution for injection 5 mg/mL, Putney, Inc., Portland, ME, USA), 2.2 mg/kg flunixin meglumine (Banamine-S®, Merck Animal Health, Summit, NJ, USA) or 3 mg/kg ketoprofen (Ketofen®, Zoetis, Inc., Kalamazoo, MI, USA). Treatment groups were assigned using a random number generator (Microsoft Excel 2016, Microsoft Corporation). The doses were chosen based on existing EU labels for piglets at castration (meloxicam and ketoprofen) or existing USA label dose for other indications in pigs (flunixin). Two hours after drug administration, the piglets were processed (defined in this study as only castrated and tail-docked). Piglets were restrained to expose the anogenital region of the piglet, while a second person performed the procedure. An incision was made on each side of the scrotum using a scalpel, the testicles were pulled from the surrounding tissue and the scalpel was used to cut the testicles free. The tail was then docked using standard tail clippers. Both the castration site and tail were sprayed with betadine to disinfect the wounds.
4
Bilateral CAP Injection in Hamsters
5
Tracing IWAT Afferent Innervation in Hamsters
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Siberian hamsters (n = 10) were anesthetized with ketamine/xylazine (100 mg/kg, 10 mg/kg; i.p.) and ventral lateral 2 cm incisions were made to expose both IWAT pads. The neuronal tracer FB (1.0%; EMS-CHEMIE GmbH, Gross-Umstadt, Germany) was injected with a microsyringe into eight separate loci (2 μl/locus) of the left and right IWAT. After the last injection, the incision was closed with sterile wound clips and ketofen (5 mg/kg, s.c.; Fort Dodge Animal Health, Fort Dodge, IA) was administered for 3 d postinjection to minimize postoperative discomfort.
One week later, CL (0.2 ng/kg in 0.9% sterile saline) was injected into one IWAT pad across eight separate loci with nearly simultaneous intra-contralateral IWAT injections of the saline vehicle (2 μl/locus for a total of 16 μl) served as a within animal control. Injections of drug or vehicle were counterbalanced by side to control for possible side bias of IWAT afferent innervations. The skin was gently pinched closed, and sterile saline-soaked gauze was laid over top of the incision site for 60 min until hamsters were sacrificed and DRGs processed for c-Fos immunohistochemistry.
One week later, CL (0.2 ng/kg in 0.9% sterile saline) was injected into one IWAT pad across eight separate loci with nearly simultaneous intra-contralateral IWAT injections of the saline vehicle (2 μl/locus for a total of 16 μl) served as a within animal control. Injections of drug or vehicle were counterbalanced by side to control for possible side bias of IWAT afferent innervations. The skin was gently pinched closed, and sterile saline-soaked gauze was laid over top of the incision site for 60 min until hamsters were sacrificed and DRGs processed for c-Fos immunohistochemistry.
6
Polyethylene Particle Application to Mouse Calvarium
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A polyethylene particle stock solution was prepared to deliver 2.5 mg of particles in 15 µl of solution. Polyethylene particles (S-395 N1, Shamrock Technologies Inc., Newark N. J.), mean diameter 5 µm, were washed 6 times in 70% ethanol. Two ml of wet particles were then suspended in 95% ethanol. Particles were placed over the calvarium using a method described by von Knoch10 (link). Specifically, mice were anesthetized with 2% isoflurane (Vetone, Meridian, ID) delivered in oxygen using a precision vaporizer (Summit Anesthesia Solutions, Bend, OR). Ophthalmic ointment (Puralube Vet Ointment, Dechra Veterinary Products, Overland Park, KS) was applied to the eyes and Ketofen (5 mg/kg, Fort Dodge Animal Health, Fort Dodge, IA) was injected subcutaneously to alleviate postsurgical discomfort. The surgical area (top of head) was shaved, cleaned with a series of washes (Betadine Scrub, Betadine Solution, and 70% ethanol), and a one cm skin incision was made over the calvarium in the sagittal plane. The skin was retracted and 15 µl of particle solution delivering 2.5 mg of particles in ethanol were applied by pipette on top of the exposed calvarial surface between bregma and lambda. The incision was closed with 7 mm surgical staples (Reflex 7 Wound Closure System). Ketofen (5 mg/kg) was administered once daily for two days following surgery.
7
Stereotaxic Implantation of 3V Cannulae
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Cannulae were stereotaxically implanted into the 3V, as described previously (Day & Bartness, 2004 (link)). Briefly, the animals were anesthetized with isoflurane 2%, and the fur at the top of the head was removed to expose the area to be incised. After exposure of the skull, a hole was trephined at the intersection of bregma and the midsagittal sinus and the guide cannula (26 gauge stainless steel; Plastics One) was positioned using the following stereotaxic coordinates: level skull, anterior‐lateral from bregma, 0 mm; medial‐lateral from midsagittal sinus, 0 mm; and dorsal–ventral, −5.5 mm from the top of the skull, which targeted placement just above the 3V. The guide cannula was secured to the skull with 3/16 mm jeweler's screws, cyanoacrylate glue, and dental acrylic. A removable obturator (Plastics One) sealed the opening in the guide cannula throughout the experiment, except when it was removed for the injections. After the surgeries, the hamsters were transferred to clean biohazard cages and received subcutaneous injections of ketofen (5 mg/kg; Fort Dodge Animal Health), an analgesic, for 3 days. They also received apple slices to supply readily consumed calories and water.
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