EXAMEN NACIONAL PARA LA EDUCACIÓN SUPERIOR (ENES)

3.5.1 Body weight and food consumption

Body weight (g) was recorded daily with digital scales (TE4101, Precision Balance Sartorius AG, Goettingen, Germany). Daily food consumption was identified by weighing feed clearance. The calculated nutritional parameters of the high fat diet and standard chow diet are presented in Table 3.7.

Table 3.7: Calculated nutritional parameters of animal fed diets

Nutritional parameters High fat diet Standard chow diet

Protein (%) 19.0 20.0

Total fat (%) 21.0 5.0

Crude fibre (%) 4.7 5.0

Digestible energy (MJ/kg) 19.4 17.25

Digestible energy from lipids (%) 40.0 10.7

Digestible energy from protein (%) 17.0 19.7

3.5.2 Body composition

Abdominal circumference was assessed at the end of the study when the rodent was anaesthetized with the rat placed in the ventral position. Abdominal circumference was taken on the largest zone of the rat abdomen using a plastic non-extensible measuring tape with an accuracy of 0.1 cm.

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Whole body composition (fat mass, lean mass, free water and total body water) was determined in conscious rats by using the EchoMRI-900 body composition analyser (EchoMedical systems, Houston, Texas, USA) at pre-intervention and post-intervention. Calibration of the machine was completed according to the manufacturer’s instructions using canola oil as the calibration medium. The body scan was completed in duplicate to ensure accuracy of measurements and the mean of the two scans was used as the final reading. Data were exported to Windows XP Professional Edition (Microsoft).

Following the treatment period, rats were anaesthetised and abdominal adipose tissue was dissected and weighed. Visceral fat mass was determined by weighing the left perirenal and epididymal adipose fat pads.

3.5.3 Blood pressure

Systolic blood pressure and diastolic blood pressure values were measured in conscious, pre- warmed, restrained animals by non-invasive tail-cuff plethysmography (CODA Non-invasive blood pressure System for rats, Kent Scientific Corporation, Connecticut, USA) at baseline and at the end of the supplementation period. To increase tail vein circulation, rats were pre- warmed in a heating chamber for 10 minutes at 30 °C. The tail cuff and volume pressure sensor were placed on the tail of each rat and then underwent five acclimatisation cycles followed by 10-20 cycles of cuff inflation (5 seconds between cycle sets). Systolic and diastolic blood pressure readings were analysed using CODA software.

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3.5.4 Urine collection

To monitor possible effects on kidney function, 24-hour urine collection was taken at 3-time points during the experiment. Metabolic cages were used to collect the rodent’s urine over a 24 hour period at baseline and at the end of the supplementation period. One rat was placed in each metabolic cage provided with a wire mesh bottom and a funnel to collect urine. Urine was collected into pre-weighed plastic containers to identify total volume of urine voided. Rats were provided with their group specific feed and tap water ad libitum. After 24 hours of collection, the urine samples were transported to the laboratory on ice immediately after collection. The urine was then centrifuged to remove solid debris such as food, faecal matter or fur at 4000 x g (RCF 3399) at 4 °C for 10 minutes. Urine was then collected into 2 mL Eppendorf tubes and frozen at -80 °C for further analysis.

3.5.5 Urinary sodium excretion

Urinary sodium content was measured at baseline and post-intervention using the flame photometer (Corning Clinical Flame Photometer 410c, Corning 805 Dilutor, Sherwood Scientific Limited, Cambridge, United Kingdom). The flame photometer was calibrated according to the manufacturer’s instructions. Undiluted samples were used to measure sodium content and were expressed as mmol/L.

3.5.6 Insulin sensitivity

Animals were deprived of food for at least 2 hours but were allowed to drink tap water. The animal’s weight was recorded to determine the insulin dosage. Rats were given 0.75 U/kg

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body weight of insulin (Humalog, Eli Lilly, NSW, Australia, Pty Ltd). The insulin solution was prepared with 50 µL of insulin and 20 mL of 0.9 % of saline.

Following the two hour fast, the baseline blood glucose was measured with a blood glucose meter kit (Optium Freestyle blood glucose meter kit, Abbott Laboratories Limited, England) via the tail snip method. The tail tip of each rat was clipped to obtain blood (approximately 25 µL) for blood glucose determination (mmol/L). The insulin solution was administrated when the rat was placed in a towel roll. The insulin was injected intraperitoneally into the abdominal cavity just below the abdominal muscle layers. Blood glucose was monitored at 15, 30, 60, 90 and 120 minutes following administration of insulin. At the completion of the test, all rats were given food and water. The insulin sensitivity test was completed at baseline and at the end of the supplementation period.

3.5.7 Glucose tolerance

Two days following the insulin tolerance test, an intraperitoneal glucose tolerance test (IPGTT) was conducted. The rodents were fasted for 24 hours prior to the commencement of the test. Animals were weighed to determine the volume of glucose to be injected. Before the glucose load, a fasting measure was obtained using the blood glucose meter kit (Optium Freestyle blood glucose meter kit, Abbott Laboratories Limited, England). The glucose load (2 g/kg BW) was administrated via an intraperitoneal injection. Blood was monitored via the tail snip method post IP by reopening the initial incision at 15, 30, 60, 90, 120 minutes and a commercial glucose meter (MediSense, Precision Plus, Abbott Diabetes Care Inc.) was used to determine the blood glucose level. The IPGTT was performed at baseline and at the end of the supplementation period.

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