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Ketoprofen

Manufactured by Performance Health

Ketoprofen is a non-steroidal anti-inflammatory drug (NSAID) used in laboratory settings. It has analgesic, anti-inflammatory, and antipyretic properties. Ketoprofen is commonly used in research applications to study its effects on pain, inflammation, and fever.

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7 protocols using ketoprofen

1

Anesthesia and Electrophysiology in Rats

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Animals were anesthetized with 5% isoflurane, and then placed in a
stereotaxic apparatus where anesthesia was maintained with 1.5% isoflurane. Two
stainless steel screws were implanted for EEG recording; one placed above neocortex
(Bregma +1 mm A/P, +3.0 mm M/L) and the other above the hippocampus (Bregma −2.5
mm A/P, +3.2 mm M/L). Two stainless steel electrodes (Cooner Wire, Chatsworth, CA) were
implanted into the neck muscle for recording EMG activity. Rats received post-surgical
antibiotic (Neo-Predef, Pharmacia & Upjohn Company, New York, NY) and analgesic
(5 mg/kg; Ketoprofen, Patterson Veterinary, Devens, MA), and were allowed to recover for
3 days before testing.
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2

Oxycodone Self-Administration in Rats

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Rats were anesthetized using 2.5% isoflurane and implanted with a silastic catheter (ID, 0.012 in OD, 0.025 in. Access Technologies, Skokie, IL) into the right jugular vein for intravenous delivery of oxycodone. The catheter was connected to a cannula which exited through the skin on the dorsal surface in the region of the scapulae. Ketoprofen (Patterson Veterinary, Devens, MA; 5mg/kg s.c. of 5 mg/ml) and Enrofloxacin (Norbrook, Northern Ireland; 5 mg/kg s.c. of 5 mg/ml) were provided at the time of surgery and a second dose was given 12 hrs later. In addition, antibiotic/analgesic powder (Neopredef, Kalamazoo, MI) was applied around the chest and back incisions. Rats were subsequently singly-housed and allowed to recover for 5 days prior to self-administration training. Intravenous catheters were manually flushed with Gentamicin (5 mg/kg i.v. of 5 mg/ml) in heparinized saline every day during recovery to maintain catheter patency.
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3

Intravenous Cocaine Self-Administration in Rats

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Rats were anesthetized using 2.5% isoflurane and implanted with a silastic catheter with an inner diameter (ID) of 0.012 in., and an outer diameter (OD) of 0.025 in. (Access Technologies, Skokie, IL) in the right jugular vein for intravenous delivery of cocaine. The catheter was connected to a cannula which exited through the skin on the dorsal surface in the region of the scapulae. Ketoprofen (Patterson Veterinary, Devens, MA; 5mg/kg s.c. of 5 mg/ml) and Enrofloxacin (Norbrook, Northern Ireland; 5 mg/kg s.c. of 5 mg/ml) were provided at the time of surgery and a second dose was given 12 h later. In addition, antibiotic/analgesic powder (Neopredef, Kalamazoo, MI) was applied around the chest and back incisions. Rats were subsequently singly housed and allowed to recover for 7 days prior to self-administration training. Intravenous catheters were manually flushed with saline every 2–3 days during recovery to maintain catheter patency. After recovery, rats were randomly assigned to three groups: ShA, LgA, or IntA.
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4

Surgical Implantation of Intravenous Catheter in Rats

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Rats used for self-administration experiments were anesthetized using ketamine
(100 mg/kg) and xylazine (10 mg/kg), and implanted with an intravenous (i.v.) silastic
catheter placed into the right jugular vein. Rats received post-surgical antibiotic
(Neo-Predef, Pharmacia & Upjohn Company, New York, NY) and analgesic (5 mg/kg;
Ketoprofen, Patterson Veterinary, Devens, MA) and recovered for 3 days prior to
training.
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5

Intravenous Cocaine Self-Administration in Rats

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Rats were anesthetized using 2.5% isoflurane and implanted with a silastic catheter with an inner diameter (ID) of 0.012 in., and an outer diameter (OD) of 0.025 in. (Access Technologies, Skokie, IL) in the right jugular vein for intravenous delivery of cocaine. The catheter was connected to a cannula which exited through the skin on the dorsal surface in the region of the scapulae. Ketoprofen (Patterson Veterinary, Devens, MA; 5mg/kg s.c. of 5 mg/ml) and Enrofloxacin (Norbrook, Northern Ireland; 5 mg/kg s.c. of 5 mg/ml) were provided at the time of surgery and a second dose was given 12 h later. In addition, antibiotic/analgesic powder (Neopredef, Kalamazoo, MI) was applied around the chest and back incisions. Rats were subsequently singly housed and allowed to recover for 7 days prior to self-administration training. Intravenous catheters were manually flushed with saline every 2–3 days during recovery to maintain catheter patency. After recovery, rats were randomly assigned to three groups: ShA, LgA, or IntA.
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6

Surgical Procedures for Rodent Neurophysiology

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After a minimum 7 day acclimation period, rats
used for self-administration experiments were anesthetized with ketamine
(100 mg/kg) and xylazine (10 mg/kg) and implanted with an i.v. silicone
catheter (ID, 0.012 in OD, 0.025 in Access Technologies, Skokie, IL)
inserted into the jugular vein that exited through the skin of the
dorsal scapulae region. Rats received postsurgical antibiotic (Neo-Predef,
Pharmacia & Upjohn Company, New York, NY) and analgesic (5 mg/kg;
Ketoprofen, Patterson Veterinary, Devens, MA) and recovered for 3
days prior to training.
For voltammetry experiments, rats were
anesthetized with intraperitoneal (i.p.) urethane (1.5 g/kg) and implanted
with a jugular vein catheter before being placed in a stereotaxic
apparatus. Once in the apparatus, rats were implanted with a bipolar
stimulating electrode (Plastics One, Roanoke, VA) aimed at the VTA
(+5.2 P, +1.1 L, −7.5 to −8.0 V). A carbon fiber microelectrode
was implanted within the core of the NAc (−1.3 A, +1.3 L, −6.5
to −7.0 V), and a reference electrode was implanted in the
contralateral cortex (−2.5 A, −2.5 L, −2.0 V).
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7

Optogenetic Manipulation of Insular Cortex

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Rats were kept under continuous isoflurane anesthesia (3–5% induction, 1–3% maintenance, 1L/min oxygen) throughout the surgery as previously described (McGinnis et al. 2019 ). An adeno-associated viral vector containing Channelrhodopsin (AAV5-CamKIIα-hChR2(H134R)-EYFP; UNC Vector Core, Chapel Hill, NC) was bilaterally microinjected (1μL/side, 0.1μL/min over 10min) into the agranular insular cortex (AIC) using a Neurostar StereoDrive (Germany) with the following coordinates (in mm, relative to bregma): 2.76AP, ± 3.50ML, 5.10DV. Injectors were left in place for an additional 5min after injection. Rats were given 2mL of warmed sterile saline and 3mg/kg ketoprofen (I.P.; Patterson Veterinary, Devens, MA) at the end of the surgery and were individually housed until 1 week after surgery when sutures were removed and housing pairs were re-established. A total of 4 weeks was allowed for virus expression and transport of hChR2 to BLA terminal fields prior to experimentation. Injection sites were confirmed by visualizing EYFP in coronal slices of the AIC using fluorescence microscopy postmortem. Rats were excluded if there was unintended delivery or spread outside of the AIC.
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