Acclimation Prior to an Intraperitoneal Insulin Tolerance Test to Mitigate Stress-Induced Hyperglycemia in Conscious Mice

Summary

Stress-induced elevations in glucose levels can confound interpretation of data derived from a conscious intraperitoneal insulin tolerance test in mice. In this article, we describe a method to acclimate the mice to handling, injections and blood sampling prior to performing the insulin tolerance test in order to limit stress-induced hyperglycemia.

Abstract

The insulin tolerance test is commonly used in metabolic studies to assess whole body insulin sensitivity in rodents. It is a relatively simple test that involves measurement of blood glucose levels over time following a single intraperitoneal injection of insulin. Given that it is performed in the conscious state and blood is often collected via a tail snip, it has the potential to elicit a stress response from animals due to anxiety associated with handling and blood collection. As such, a stress-induced rise in blood glucose can occur, making it difficult to detect and interpret the primary endpoint measure, namely an insulin-mediated reduction in blood glucose. This has been seen in many mouse strains, and is quite common in diabetic db/db mice, where glucose levels can increase, rather than decrease, after insulin administration. Here, we describe a method of acclimating mice to handling, injections and blood sampling prior to performing the insulin tolerance test. We find that this lowers stress-induced hyperglycemia and results in data that more accurately reflects whole body insulin sensitivity.

Introduction

Metabolic tests in rodents are routinely performed to assess various parameters that regulate glucose homeostasis¹. The gold standard for assessing whole-body insulin action in vivo is the hyperinsulinemic-euglycemic clamp². This test involves administration of insulin to raise circulating insulin levels while glucose is infused to maintain euglycemia. The glucose infusion rate required to maintain euglycemia is indicative of insulin action. While it is a powerful tool in metabolic research, the clamp technique in mice is technically challenging and labor intensive, and thus is not well suited as an initial screening tool in characterizing a metabolic phenotype. For these reasons, the simpler intraperitoneal insulin tolerance test (ITT) is often chosen.

The ITT is performed in the conscious state following a fasting period (typically 4-6 hours). A bolus of insulin is administered intraperitoneally, after which blood glucose is monitored over a timeframe that usually lasts 60 min. Blood glucose levels are expected to fall due to the ability of insulin to facilitate glucose uptake into insulin-sensitive tissues; the degree to which this occurs is indicative of whole-body insulin action. In some cases, it has been shown that glucose levels paradoxically increase, rather than decrease, after insulin administration. This phenomenon is likely attributed to a stress response. Handling, injections and blood sampling can all induce stress^3,4,5, resulting in activation of the hypothalamic-pituitary-adrenal axis (HPA) and the autonomic nervous system (ANS)^6,7,8. It is well known that both the HPA and ANS contribute to increases in circulating glucose levels^9,10,11. The presence of stress-induced hyperglycemia at the beginning of an ITT is problematic, as it interferes with the rate and magnitude of the fall in glucose upon insulin administration¹². This can lead to erroneous conclusions regarding the presence of insulin resistance. Thus, to mitigate the confounding impact of stress on glucose levels during an ITT, we have developed a method of acclimating mice to handling, injections and blood sampling prior to performing the ITT.

Protocol

All methods described here have been approved by the VA Puget Sound Health Care System’s Institutional Animal Care and Use Committee.

NOTE: Local requirements for monitoring and/or intervention of animals that experience hypoglycemia may differ from those described here.

1. Fasting (t= -210 min)

1. After the dark cycle has ended, transfer mice to a new cage with non-nutritious bedding such as cellulose or paper bedding (not corn-cob bedding, which will affect metabolic endpoints if consumed by mice¹³). Provide mice with ad libitum access to water throughout the fasting period. Be consistent with the fasting time of day and duration across groups of mice. For the data shown, food was removed between 0700 and 0800 hours, which was 1-2 hours after the dark cycle had ended.
2. Move cage(s) to the location where the ITT will be performed. This should be a quiet space where stressors such as temperature, noise, light or movement are minimized.

2. Acclimation (t= -150 to -60 min)

IMPORTANT: Handle mice as gently as possible. Avoid use of a restraint device if possible.

1. At -150 min, measure body weight. This will be used for calculation of the volume of insulin that will be administered for the ITT.
2. Pick up the mouse gently by the tail and rest on a flat tabletop surface while still gently gripping the tail. Use a 20 G needle (or surgical scissors) to make a small incision in the tip of the tail. A drop of blood should begin to form at the site.
3. Set the mouse on a smooth hard surface and restrain the mouse gently by the tail.
   1. To record blood glucose at this time, place a drop of blood from the tail tip on the test strip of a handheld glucometer. If necessary, very gently massage the tail to obtain a drop of blood.
4. Draw up 100 µL of sterile saline into an insulin syringe. Pick up the mouse using gentle scruffing and inject intraperitoneally. Record the time of saline injection.
   1. If multiple mice are being studied, inject the mice with saline at 1 min intervals to allow time for subsequent steps below.
5. At 15 min after saline injection, pick up the mouse gently by the tail. Use gauze to gently dislodge any blood clot on the tail tip.
   1. Optional: Record blood glucose again at this time as described in step 2.3.
6. At 30 min after the saline injection gently pick up the mouse by the tail. If desired, obtain another blood glucose measurement as described in step 2.3.
7. Return the mouse to the cage, and repeat for other mice as necessary. Leave mice undisturbed until -90 min.
8. At -90 min, repeat steps 2.3 through 2.6, including measurement of blood glucose levels if desired.
9. Return the mouse to the cage, repeat for other mice as necessary. Leave mice undisturbed until the ITT (for which the baseline blood glucose measurement is done at -5 min).

3. Insulin Tolerance Test (t= -5 to +60 min)

1. Prepare a working solution of regular insulin (CAUTION) in sterile saline, such that the desired dose can be injected at 4 µL/g body weight.
   NOTE: Caution should be used when handling insulin as accidental injection can result in hypoglycemia.
2. Prepare a 25% (v/v) dextrose solution in sterile saline to have on hand in case mice develop hypoglycemia requiring intervention.
3. At -5 min, pick up the mouse gently by the tail. Determine the baseline blood glucose level at this time, by placing a drop of blood from the tail tip on the test strip of a handheld glucometer. If necessary, very gently massage the tail to obtain a drop of blood.
4. Draw up insulin working solution into an insulin syringe (4 µL/g body weight, for a typical dose range of 0.5-2.0 U/kg). Pick up the mouse using gentle scruffing and inject intraperitoneally. Record time of insulin injection.
   1. If multiple mice are being studied, inject the mice with insulin at 1 min intervals to allow time for subsequent blood sampling and data recording below.
5. At 15 min after insulin injection, pick up the mouse gently by the tail. Use gauze to gently dislodge any blood clot on the tail tip. Determine the blood glucose again at this time as described in step 3.3.
6. Return the mouse to the cage and monitor for signs of hypoglycemia (e.g., excessive lethargy). If mice develop symptoms of hypoglycemia, measure blood glucose as described in step 5 and administer dextrose if necessary. A mouse undergoing dextrose intervention would be removed from the ITT protocol at this time.
7. Repeat steps 3.5 and 3.6 at 30 min after insulin injection.
8. Repeat steps 3.5 and 3.6 at 45 min after insulin injection.
9. Repeat steps 3.5 and 3.6 at 60 min after insulin injection.
10. Return the mice to their home cage, containing a few food pellets on the floor of the cage to aid in recovery from the ITT. Continue to monitor for 30 min and if mice have not regained normal activity/behavior follow procedure described in step 3.6.
    NOTE: Figure 1 summarizes the above acclimation and ITT procedure.

Results

Figure 2A and Figure 3A are representative data (individual and mean data, respectively) showing a paradoxical rise in blood glucose levels in diabetic db/db mice in the 15 minutes following insulin administration, consistent with stress-induced hyperglycemia. Note, no rise in blood glucose was evident in the control non-diabetic littermates (db/+ or +/+) that underwent the same procedure. To determine whether acclimation to the ITT procedure was effective in mi...

Discussion

The hyperinsulinemic-euglycemic clamp is considered the gold standard for assessing insulin action in vivo. Modifications to the methodology for performing the clamp have resulted in the technique being done in conscious, unrestrained mice² that have been previously catheterized using a two-catheter system¹⁴ to enable blood sampling via the carotid artery and infusions via the jugular vein. This limits the need to handle or restrain mice during the procedure, thereby reduci...

Materials

Name	Company	Catalog Number	Comments
Dextrose-50	Pfizer Injectables	00409-6648-16	For use if mouse experiences hypoglycemia.
Gauze pads	Fisher Scientific	22037907	To dislodge blood clot on the tail tip.
Glucometer	Accu-Chek	M001_us	To measure blood glucose.
Gram scale			To measure body weight.
Insulin (Novolin R)	Novo Nordisk	0169-1833-11	For injection.
Insulin syringes	VWR	BD-329461	For injections.
Minute timer
Sterile 20 G needle	VWR	BD-305175	For tail snip.
Sterile saline	Lifeshield	1261699	For injections.
Surgical scissors	Fine Science Tools	14088-10	For tail snip.
Test strips	Accu-Chek	06908217001_us	To measure blood glucose.