Glucose cii test

The concentrations of undigested starch in feces were determined after hydrolysis with thermostable α‐amylase and amyloglucosidase, followed by an enzymatic colorimetric measurement of the released glucose using a commercial assay kit (Glucose CII‐test, Wako Pure Chemical Industries, Ltd.) (Unno, Hisada, & Takahashi, 2015). Results were converted to a starch basis by multiplying them by 0.9.

Serum biochemical parameters were analyzed by the following methods. SUN measurements were determined by a colorimetric assay using a urease-indophenol method (Wako Pure Chemicals, Osaka, Japan). IGF-1 and glucose measurements were analyzed with an enzyme-linked immunosorbent assay (ELISA; MG100; R&D Systems, Minneapolis, MN, USA) and a colorimetric assay by the mutarotase-glucose oxidase method (Glucose CII-test; Wako), respectively. Insulin and glucagon were measured using ELISA kits (AKAIN-010T; Shibayagi, Gunma, Japan, and Wako, respectively). Cortisol was analyzed by an ELISA kit (KGE008; R&D Systems). AST and ALT were measured by the pyruvate oxidase-N-ethyl-N-(2-hydroxy-3-sulfopro-pyl)-m-toluidine method with a commercial kit (Transaminase CII-test; Wako). To measure renal tissue cAMP content, kidneys were prepared by homogenizing with HCl. Extracted cAMP levels were analyzed by using a cAMP radioimmunoassay kit (Yamasa, Chiba, Japan), and were corrected by protein concentration.
All biochemical parameters were measured according to the manufacturers’ instructions and are expressed as the mean ± standard deviation (SD) (n = 6 rats per treatment group).

Six‐week‐old male Sprague‐Dawley (SD) rats (Japan SLC, Hamamatsu, Japan; n = 16) were maintained in an air‐conditioned environment (23 ± 3°C) under an automatic lighting schedule (08:00–20:00 light) with free access to water and a standard laboratory diet (MF, Oriental Yeast Co., Ltd., Tokyo, Japan). IPGTT was conducted using our previous published process (Kato, Nakanishi, Tani, & Tsuda, 2017; Kato, Nishikawa, et al., 2017; Nagamine et al., 2014). Briefly, rats were deprived of food for 13 hr at 7 weeks of age, before oral administration by direct stomach intubation of vehicle (control; 0.9% NaCl) or BCE 5 mg/kg body weight (1 mg D3R/kg body weight). The dose of BCE was determined based on a preliminary experiment in order to show that the dose level was significant impact on improving glucose tolerance. After 30 min, rats received IP administration of glucose solution (2 g/kg). Blood samples were collected at 0 (before glucose loading), 15, 30, 60, and 120 min after glucose injections, and serum glucose and insulin concentrations were measured with Glucose CII‐Test (Wako) and Ultra Sensitive Rat Insulin ELISA kit (Morinaga Institute of Biological Science, Yokohama, Japan), respectively (Kato, Nakanishi et al., 2017; Kato, Nishikawa, et al., 2017; Nagamine et al., 2014).

We collected the blood of hatchlings from clutches laid at Kochi Beach, excluding that of hatchlings used in the swimming performance and rearing analyses. Hatchlings included in blood collection were maintained in a tank at 28 °C without feeding until blood collection was completed (72 h). We did not use hatchlings whose blood had already been collected. Approximately 500 μl of blood was collected from the dorsal cervical sinus using a 26 G needle (Termo, Japan) and a 1 mL syringe (Termo, Japan) at 0, 4, 24, 48, and 72 h after size measuring (n = 3–4 at each sampling time). Blood was poured into a heparinised tube (Fujifilm, Japan) and centrifuged at 3600 g for 10 min. Plasma was moved into another tube and stored at −80 °C until glucose measurements. Blood glucose concentrations were measured enzymatically using the glucose-oxidase method (Glucose CII-test, Wako Pure Chemical Industries, Japan) according to the manufacturer’s protocol.

Systolic blood pressure was measured by a tail-cuff method using a blood pressure monitor (Model MK-2000, Muromachi Kikai Co., Chuo-ku, Tokyo, Japan) one week before the experimental date of vasodilation studies, as described previously [45 (link)]. The systolic blood pressure values were derived from an average of at least five measurements per animal at each time point.
Before collecting tissues from animals for experiments, body weight, body length and waist length were measured under anesthesia (sodium pentobarbital, 80 mg/kg, i.p.), and then blood was drawn from the abdominal aortas of non-fasting rats. The serum was separated by centrifugation at 1000× g for 10 min at 4 °C. Serum levels of triglyceride, glucose, insulin, and thiobarbituric acid reactive substances (TBARS), as an index of oxidative stress, were determined using commercial kits; the triglyceride E-test (Wako Pure Chemical Ind. Ltd., Osaka, Osaka, Japan), glucose CII-test (Wako Pure Chemical), the rat insulin detection kit (Morinaga Biochemistry Lab., Yokohama, Kanagawa, Japan), and TBARS Assay Kit (Cayman Chemical Co., Ann Arbor, MI, USA).

Non-structural carbon (NSC: starch and soluble sugars) within the sapwood was measured in three plant parts (twigs 6 mm in diameter, stem bases and taproots). The xylem sapwood (without the bark, phloem and pith) from each part was ground to a fine powder and then extracted in 80% ethanol (v/v). The supernatant was extracted via centrifugation and used to quantify the soluble sugar content via the phenol–sulfuric acid method^{48 (link)}. The starch in the remaining pellets was depolymerized to glucose by the addition of KOH, acetic acid and an amyloglucosidase buffer. After quantifying the glucose extracted via the mutarotase–glucose oxidase method (Glucose C-II test; Wako, Tokyo, Japan), the starch content within the xylem sapwood was measured.

The urinary glucose concentration was performed at the last week (sixth week) of treatment, mice were fasted for 12 h and urine was taken during this period. Urinary glucose concentrations were measured by a commercially available kit (Glucose CII-test, Wako, WAKO, Tokyo, Japan).