経口ブドウ糖負荷試験 (OGTT) とインスリン負荷試験 (ITT) を使用して高脂肪食飼育マウスの生体内糖代謝の研究

The overall goal of these procedures is to characterize the general metabolic phenotype of mice and to specifically assess alterations in glucose metabolism in vivo. These methods can help answer key questions in the field of metabolism. They are representative and powerful tools to investigate the influence of genetic, pharmacological, or dietary factors on glucose metabolism in vivo.

The main advantage of these techniques is that they are relatively easy to perform. The day before the oral glucose tolerance test, transfer mice into a cage with fresh bedding and drinking water but without food to fast them overnight before testing. The next day, prepare 10 milliliters of 20%glucose solution by diluting 45%D-glucose in distilled water.

Then, prepare a plate for plasma collection by adding five microliters of sodium EDTA per time point for each mouse to the wells of a 96 well plate. During the experiment, store this plate on ice. Next, measure the body weight of all mice and mark their tails with a permanent marker to make the mice easily distinguishable.

To collect blood for baseline sampling, use sharp scissors to carefully cut off one to two millimeters of the tail tip. Wipe off the first drop of blood to avoid hemolysis or contamination with tissue fluid before taking new blood samples for blood glucose determination. Draw a small blood sample for the measurement of the basal blood glucose level with the glucometer.

Then collect a larger blood sample using a fresh capillary tube. Empty the capillary tube using a pipette by putting the pipette tip at the top of the capillary tube end and carefully pushing the collected blood into a well of the 96 well plate while avoiding air bubbles. Prepare 20%D-glucose dissolved in distilled water.

To administer a glucose solution, first restrain the mouse by firmly grasping it by the scruff. Apply enough firmness to the skin around the neck to prevent the mouse from twisting out and to property tilt its head back. Ensure that the mouse can breathe properly.

Then, cautiously direct the feeding needle through the mouth towards the esophagus. Allow the mouse to swallow the needle. After the needle sinks into the lower esophagus of the mouse, inject the glucose solution.

Start the timer immediately after the first gavage, and administer glucose to all other mice in one minute intervals. Once glucose administration is started, good time management is very important. After 15 minutes, measure blood glucose levels with the glucometer as before.

And then take another 30 microliter blood sample from each mouse in the same order as they were injected. Blood collection should be repeated 30, 45, 60, 90, 120, 150, and 180 minutes after glucose administration. Ensure mice have access to drinking water during this time.

When the measurements are finished, return the mice to their home cages equipped with food and water. Then centrifuge the blood samples at 2, 500 times G for 30 minutes at four degrees Celsius. Transfer the plasma supernatant to empty wells of the 96 well plate.

And then store the plate at minus 20 degrees Celsius until analysis. At least one week after the glucose tolerance test, fast the mice for a minimum of two hours prior to insulin injection ensuring that the mice have access to drinking water. Dilute insulin stock solution one to one thousand with 0.9%sodium chloride, and prepare 20%D-glucose dissolved in distilled water to be administered if the mice become hypoglycemic.

After weighing the mice and marking their tails, calculate the volume of insulin required to administer 0.75 units of insulin per kilogram of body weight. Then, cut the tail tip using sharp scissors, and measure basal blood glucose levels as before. To inject insulin intraperitoneally, restrain the mouse by the scruff method, and tilt the mouse head down at a slight angle to expose the ventral side of the animal.

Place the filled syringe at a 30 degree angle in the lower right quadrant of the animals abdomen, and inject the volume with the bevel of the sterile 30 gauge needle turned up. Start the timer immediately after the first mouse is injected. Measure blood glucose levels at selected time points.

After the final time points, place the mice back into their home cages prepared with plenty of food and water. The oral glucose tolerance test was employed to compare the metabolic phenotype of mice fed on a high fat diet for 12 weeks with mice fed on a low fat diet. In the glucose tolerant LFD mice, the peak blood glucose level of approximately 240 milligrams per deciliter was reached approximately 15 minutes after glucose administration and immediately followed by a decrease towards the baseline level indicating proper glucose elimination.

In contrast, HFD mice peaked at approximately 320 milligrams per deciliter glucose and showed nearly no disposal of glucose. Because blood glucose levels between the two groups differ in the fasting state, a calculation of the area under the curve above baseline glucose was performed, and this validated the prior results. Additionally, blood insulin levels were determined using an insulin ELISA.

The mice fed at HFD showed elevated fasting insulin levels compared to the control group as well as an increased insulin response indicating HFD induced compensatory hyperinsulinemia. To measure insulin sensitivity in the HFD fed mice, an insulin tolerance test was performed one week after the oral glucose tolerance test. Compared to the LFD fed control group, the HFD fed mice showed an impaired reduction of blood glucose levels after insulin administration suggesting insulin resistance.

While attempting this procedure, it's especially important to maintain a good time management as well as to reduce the stress levels for the mice to a minimum in order to generate robust, reproducible results. Differences in glucose tolerance, insulin levels, and insulin sensitivity which are all obtained by the percent of methods can help to blend the next complex steps. This can include, for example, hyperglycemic or hyperinsulinemic claims or experiments with isolated pancreatic islets.