Animals. All procedures were approved by the Institutional Animal Care and Use Committee of the Scripps Research Institute and were carried out on male EP3R–/– mice backcrossed to C57BL/6 background over more than eight generations and on WT littermates.
Food-Intake Measurements. Mice were fed ad libitum with mouse breeder diet (S-2335 Mouse Breeder, gross energy kcal (1 kcal = 4.18 kJ)/g 4.39, protein % 17.50, fat % 11.72, fiber % 3.36; Harlan Teklad, Madison, WI) and separated in two groups on each age (n = 10 each group). Food intake was monitored every hour during 24 h. For daily consumption, food and body weight were monitored twice per day at the onset of the dark and light period (6:00 a.m. and 6:00 p.m.) for 9 days. Mice were deprived of food for 24 h on day 4. Observation of food consumption was evaluated for 5 additional days after the food deprivation. Body weight was normalized for metabolic demands of body mass according to Kleiber's function (g weight loss/g baseline weight0.75).
Body Weight and Fat Distribution. At the end of this period, mice were anesthetized, and intraabdominal fat pads (gonadal, retroperitoneal, and mesenteric), liver, subcutaneous fat pad (inguinal and the groin), and brown adipose tissue were dissected and weighed.
Glucose-Tolerance Tests. A glucose-tolerance test was performed at the onset of the light cycle (6:00 a.m.). Mice were weighed and fasted for 24 h before the glucose-tolerance test. Access to drinking water was allowed during this period. On the day of the test, baseline glucose levels and body weight were determined before challenge with a glucose load of 1.5 mg of glucose per gram of body weight (D-glucose, anhydrous; Sigma–Aldrich, St. Louis, MO) dissolved in sterile distilled water (0.75 g of D-glucose, anhydrous in 10 ml of sterile water). The mouse was restrained by holding the excess skin at the base of the neck between the technician's thumb and forefinger. The mouse's tail was left hanging out and placed on a glass slide, and a segment of 
1 mm in length was cut off the tip of the tail by using a sharp razor blade. A small drop (5 µl) of blood was placed on the glucometer test strip (Home Diagnostics, Fort Lauderdale, FL). After a 5-second developing time, the baseline blood glucose value was recorded (in mg per deciliter), and the mouse was returned to his home cage. After the baseline glucose measurement, the mouse was injected i.p. by using a 1-ml syringe and a 27-gauge needle. The time of the injection was noted, and 15-, 30-, 60-, and 120-min postinjection blood glucose measurements were performed again. It was sometimes necessary to remove a scab that formed at the initial tail-cut site to collect the second blood sample (n = 6 for each group). After the end of the study, mice were returned, and food and water was provided at libitum.
Insulin-Resistance Test. The effects of insulin injection were assessed in nonfasted male mice. Similar to the glucose-tolerance test, blood was withdrawn from the tail without anesthesia before administration of human insulin (1 unit/kg, i.p.; Sigma–Aldrich). Samples were collected 15, 30, 60, and 120 min after the insulin challenge. Blood glucose levels were determined by a blood glucose meter (Home Diagnostics) (n = 6 for each group).
Plasma Levels of Leptin and Insulin. Plasma leptin and insulin levels were determined at 6:00 a.m. and 6:00 p.m., at the onset of the light and dark cycle, respectively. Mice were euthanized with isoflorane (5%), decapitated, and bled into EDTA-coated tubes. Blood was centrifuged at 10,000 x g for 10 min at 4°C, and supernatants were taken and stored at –70°C until further analysis. Plasma leptin was measured by using a Mouse Leptin ELISA kit 96-well plate (catalog no. EZML-82K; Millipore, Billerica, MA) used for a nonradioactive quantification of leptin, and values were collected and averaged ± SEM. Mouse insulin was determined by RIA using a Rat Insulin RIA kit (250 tubes; catalog no. RI-13K; Millipore) according to manufacturer's instructions (n = 6 for each group).
Telemetry Device Implant. EP3R–/– and WT littermate male mice were anesthetized with isoflorane (induction, 3–5%; maintenance, 0.9–1.5%) and implanted with radio telemetry devices (TA10TA-F20; Data Sciences, Inc., St. Paul, MN) into the peritoneal cavity for core body temperature measurement. Mice were allowed to recover for 2 weeks and were then submitted for freely moving recording (n = 10 for each group). Mice were maintained in a temperature-controlled room (25°C) on a 12-h light–dark cycle (light on at 6 a.m.). Core body temperature and motor activity sensors were located in the transmitter implant. The cages were positioned onto the receiver plates. Radio signals from the core body temperature and motor activity of each animal (number of horizontal movements) were continuously monitored with a fully automated data-acquisition system (Dataquest A.R.T.; Data Sciences, Inc.).
Data Analysis. Data were grouped and analyzed by using the paired t test or ANOVA with repeated measures followed by post hoc Newman–Keuls test. All results are expressed as means ± SE. Metabolic efficiency was calculated as the energy intake divided by the body weight gain over a certain period. Linear relationships were estimated by using Pearson's moment correlation coefficient.
Acknowledgements
We thank Professor Jerold Chun for insightful suggestions. This work was supported by funds from the Skaggs Institute of Chemical Biology at The Scripps Research Institute.
Footnotes
Abbreviations: PGE2, prostaglandin E2.
*To whom correspondence should be addressed. E-mail: [email protected]
Freely available online through the PNAS open access option.
Author contributions: M.S.-A., I.V.T., C.N.D., B.C., and T.B. designed research; M.S.-A., I.K., S.E.B., I.V.T., C.N.D., and B.C. performed research; I.K., S.E.B., I.V.T., and C.N.D. contributed new reagents/analytic tools; M.S.-A., I.K., I.V.T., C.N.D., B.C., and T.B. analyzed data; and M.S.-A. and T.B. wrote the paper.
The authors declare no conflict of interest.
© 2007 by The National Academy of Sciences of the USA
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