The overall goal of this procedure is to establish a method for the inbivo measuring of gastric emptying in mice using a 13 carbon Octa NOIC acid breath Test. Mice and Other small animals have of course been really useful to help us understand the pathophysiology of disease our laboratory studies gastroparesis, and one of the issues with gastroparesis is that, you know, to study disease we need to follow gastric empty over time. This has been a problem with current techniques because current techniques require you to put a substance in the, in either liquid or solid in the mouse stomach, and then sometime later kill the mouse and estimate what has left the stomach.
What we wanted to do was establish a non-invasive technique of measuring gastric emptying, and what we did is we adapted the known 13 carbon ano acid breath test to enable us to use it on mice. This allows us to do repeated measurements over time and then correlate what we see with both histology as well as physiology. The equipment we use here allows us to automate the procedure, although of course it can also be done manually.
The advantage of automation is not only the fact that we can do many more gastric emptys over time, but that we can actually do it in multiple mice at the same time. What we'll be showing is a method where we can do up to 12 mice at one time. The following animation shows the setup of our gastric emptying experiment for the in vivo measurement of gastric emptying.
The mice are put in transparent plastic tubes with constant airflow. After the mice get habituated to the tubes and after adjustment of the airflow, we administer a test meal consisting of egg yolk mixed with 13 carbon labeled Octo NOIC acid. Because the mice are fastened and trained, they generally eat the test meal.
Within two minutes, the administered OCT NOIC acid will be taken up in the duodenum and will get metabolized in the liver released and exhaled, resulting in an enrichment of 13 carbon dioxide in the surrounding air, which is then measured by our isotope analyzer. Let's have a more detailed look into what happens in the mouse body. After eating the test meal containing 13 carbon labeled OCT NOIC acid, the meal enters the stomach.
The OCT NOIC acid remains intact in the stomach. After passage through the pylorus, it is rapidly absorbed in the duodenum. The labeled compound reaches the liver through the portal vein and is metabolized to 13 carbon labeled carbon dioxide.
The labeled carbon dioxide is released in the venous blood and is exhaled. The rate limiting step in this whole process is gastric emptying. This means that the pulmonary excretion of 13 carbon dioxide directly corresponds with gastric emptying of the labeled meal.
Air samples are collected at certain time intervals and are analyzed by the carbon isotope analyzer. Various isotope analyzers can be used. In this study, we chose to use a low GATOS research LGR 13 carbon isotope analyzer.
Collecting samples for four hours gives reproducible 13 carbon dioxide kinetics, which reflect the rate of gastric emptying. Curve fitting. Using non-linear regression allows calculation of half emptying times.
This procedure is suitable for the follow-up of mice with chronic disease. In this particular study, we use female non-obese diabetic, N-O-D-L-T-J mice, which are a model for human type one diabetes. Prior to analysis, all mice are allowed to habituate in the testing chambers to minimize stress levels.
Since a delay in gastric emptying is a characteristic indicator of stress and mice, Non habituated mice continue to move around for about one hour after transfer to the chamber and defecate and urinate. Frequently the mice are considered successfully habituated when they quickly settle into their new environment. Upon transfer into the chamber, after a brief period of exploration, the habituated mice will rest quietly in the chamber.
In habituated mice gastric emptying test results should show a difference of less than 10%intra mouse variability. Once trained and ready for gastric emptying, the mice must be fasted the day prior to the experiment. The mice are fasted overnight on a metal mesh bottom fasting rack.
To prevent phasia, make sure they have free access to drinking water. Diabetic mice should not be fasted more than 16 hours. Consistency is extremely important during a gastric MCN experiment.
Factors such as handling the mice the same way, fasting the mice at the same time every day, checking glucose levels and administering insulin at the same time each day. As a user wearing the same cologne and the same lab coat every day is extremely important cause all of these factors will change the gastric MTN data. The materials we use for the preparation of the egg meal are an egg OC acid, a glass beaker, a falcon tube, and a bent end spatula.
Start with weighing out five grams of egg yolk in a 50 milliliter falcon tube. Next, add 10 microliters of okin NOIC acid with a concentration of two microliters per gram to the Falcon tube containing the egg and mix vigorously with the spatula in the falcon tube. For one minute, the egg is then transferred to a glass beaker and heated over a buns and burner until it coagulates and its consistency is suitable.
To make small balls, this typically takes about 30 seconds. The balls of egg yolk should weigh 0.2 grams per mouse. It is important to keep the cumulative dose constant in all the mice.
Our range for acceptable cumulative dose recovery is 55 to 100%Start by setting up the gastric emptying chambers. Use clean chambers and covers that have been air dried.Also. Any tubes connecting the chamber to the analyzer or the carbon dioxide air supply should be moisture free.
Moisture can interfere with the signal red by the analyzer. Connect the chambers to the inlet tubes that provide hydrocarbon scrubbed airflow. It is important that this air is always consistent because any variation in the carbon dioxide amount could change the baseline value.
Then connect the outlet tubes from the chambers to the machine, close the tubes and turn on the airflow. Apply a very small amount of Vaseline at the end of the covers so they close and are easily and securely sealed. This tight seal is necessary to collect all the carbon dioxide produced by the mice.
Weigh the mice using a small mouse holder to reduce stress levels. Body weight is determined as a measure of their continuing good health. Then place each mouse in the appropriate chamber.
It is important to have the air flowing into the chambers at this time. Each mouse is implanted with a tracking device. The data on the mouse's weight, blood glucose and treatments are tracked using a software database that uses this number.
We use software that allows us to automatically link the mouse ID number directly to the store data. This, of course, can also be done manually. To start the measurement, allow the mice to acclimate to the chambers before adjusting the air levels.
Once the mice appear calm, which may take a few minutes, adjust the airflow rate for each mouse chamber. This may be different for each mouse. Typically, the airflow is adjusted at the beginning of the experiment to make sure that exhaled carbon dioxide reaches levels detectable by whatever equipment is being used, and to make sure that the level stays low enough to ensure healthy air turnover.
We use initial carbon dioxide levels between 1000 and 1500 parts per million if having difficulty with adjustments, check for air leaks. Next, repeat the process for each of the chambers and watch for another round of measurements to see if adjustments made to airflow have corrected the carbon dioxide level. It is important to obtain a steady baseline reading prior to the feeding of the mice.
When this is achieved, give the egg meal to the first mouse and record the time each mouse receives their food. We run the procedure for four hours to obtain enough values for fitting the 13 carbon dioxide enrichment curve for each mouse. Check on the mice every 30 to 60 minutes to make sure that the carbon dioxide levels are still safe.
For the mice, our machine measures 12 carbon dioxide, 13 carbon dioxide and water concentrations every 25 seconds. Therefore, 12 mice can be simultaneously analyzed with a five minute interval between readings. If manually sampling or using another device, make sure your readings are obtained at a minimum of five to 10 minute intervals.
Prepare new boxes containing food before the end of the test so the mice can start eating immediately after the test is Over. This graph represents the data points of a mouse with normal gastric emptying. It shows a fraction of 13 carbon that is recovered in the exhaled air expressed as a percentage of the administered dose per hour expressed as a function of time.
The data points are fitted by a curve with the following equation. Why is the percentage of the 13 carbon recovered in the breath per hour and a B and CA regression constant? The T half is calculated by putting the parameters of this curve into the inverse gamma function as established in a previous study in our lab, the normal gastric half emptying time for a non-diabetic NOD mouse age nine to 15 weeks ranges from 62 to 1 31 minutes as represented by the gray box.
The blue curve is from a mouse with an accelerated gastric emptying with a T half value of 40 minutes, and the red curve is from a mouse with a delayed gastric emptying with a T half value of 168 minutes. In conclusion, this video demonstrates how to measure solid gastric emptying by means of a non-invasive breath test. In mice, we showed how to prepare the animals for testing the making of the food and the collection of the data.
This method of measuring solid gastric emptying with the administration of 13 carbon OCT oak acid was previously reported for humans, and we adapted the protocol for use in mice. This protocol provides us with a valuable method to repeatedly measure gastric emptying in the same mouse, which allows for the investigation of gastric emptying for the study of long-term diseases. There are several future applications of this method.
For example, the testing of different drugs for the treatment of diabetic gastroparesis.
