A Preclinical Model of Exertional Heat Stroke in Mice

Summary

The protocol describes the development of a standardized, repeatable, preclinical model of exertional heat stroke (EHS) in mice free from adverse external stimuli such as electric shock. The model provides a platform for mechanistic, preventative, and therapeutic studies.

Abstract

Heat stroke is the most severe manifestation of heat-related illnesses. Classic heat stroke (CHS), also known as passive heat stroke, occurs at rest, whereas exertional heat stroke (EHS) occurs during physical activity. EHS differs from CHS in etiology, clinical presentation, and sequelae of multi-organ dysfunction. Until recently, only models of CHS have been well established. This protocol aims to provide guidelines for a refined preclinical mouse model of EHS that is free from major limiting factors such as the use of anesthesia, restraint, rectal probes, or electric shock. Male and female C57Bl/6 mice, instrumented with core temperature (Tc) telemetric probes were utilized in this model. For familiarization with the running mode, mice undergo 3 weeks of training using both voluntary and forced running wheels. Thereafter, mice run on a forced wheel inside a climatic chamber set at 37.5 °C and 40%-50% relative humidity (RH) until displaying symptom limitation (e.g., loss of consciousness) at Tc of 42.1-42.5 °C, although suitable results can be obtained at chamber temperatures between 34.5-39.5 °C and humidity between 30%-90%. Depending on the desired severity, mice are removed from the chamber immediately for recovery in ambient temperature or remain in the heated chamber for a longer duration, inducing a more severe exposure and a higher incidence of mortality. Results are compared with sham-matched exercise controls (EXC) and/or naïve controls (NC). The model mirrors many of the pathophysiological outcomes observed in human EHS, including loss of consciousness, severe hyperthermia, multi-organ damage as well as inflammatory cytokine release, and acute phase responses of the immune system. This model is ideal for hypothesis-driven research to test preventative and therapeutic strategies that may delay the onset of EHS or reduce the multi-organ damage that characterizes this manifestation.

Introduction

Heat stroke is characterized by central nervous system dysfunction and subsequent organ damage in hyperthermic subjects¹. There are two manifestations of heatstroke. Classic heat stroke (CHS) affects mostly elderly populations during heat waves or children left in sun-exposed vehicles during hot summer days¹. Exertional heat stroke (EHS) occurs when there is an inability to thermoregulate adequately during physical exertion, typically, but not always, under high ambient temperatures resulting in neurological symptoms, hyperthermia, and subsequent multi-organ dysfunction and damage². EHS occurs in recreational and elite athletes as well as military personnel and in laborers with and without concomitant dehydration^3,4. Indeed, EHS is the third leading cause of mortality in athletes during physical activity⁵. It is extremely challenging to study EHS in humans as the episode can be lethal or lead to long-term negative health outcomes^6,7. Therefore, a reliable preclinical model of EHS could serve as a valuable tool to overcome the limitations of retrospective and associative clinical observations in human EHS victims. Preclinical models of CHS in rodents and pigs have been well characterized^8,9,10. However, preclinical models of CHS do not directly translate into EHS pathophysiology due to the unique effects of physical exercise on the thermoregulatory profile and innate immune response¹¹. In addition, previous attempts to develop preclinical EHS models in rodents posed significant restrictions, including superimposed stress stimuli induced by electric shock, insertion of a rectal probe, and predefined maximum core body temperatures with high mortality rates^{12,13,14,15,16} that do not match current epidemiological data. These represent significant limitations that may confound data interpretation and provide unreliable biomarker indexes. Therefore, the protocol aims to characterize and describe the steps of a standardized, highly repeatable, and translatable preclinical model of EHS in mice that is largely free from the limitations mentioned above. Adjustments to the model that can result in graded physiological outcomes from moderate to fatal heat stroke are described. To the authors' knowledge, this is the only preclinical model of EHS with such characteristics, making it possible to pursue relevant EHS research in a hypothesis-driven manner^11,17,18.

Protocol

All procedures have been reviewed and approved by the University of Florida IACUC. C57BL/6J male or female mice, ~4 months old, weighing within a range of 27-34 g and 20-25 g, respectively, are used for the study.

1. Surgical implantation of the telemetric temperature monitoring system

1. Upon arrival from the vendor, allow the animals to rest in the vivarium for at least 1 week prior to surgery to minimize the stress of transportation.
2. Group house the mice (maximum of 5 per cage under local IACUC guidelines) until the day of surgery for temperature telemetric device implantation. House them in standard 7.25" (W) x 11.75" (L) x 5" (H) cages containing corncob bedding. Maintain the light cycle on a 12 x 12 light cycle (on: 7 AM; off: 7 PM). Maintain the housing temperature at 20-22 °C and relative humidity (RH) at 30%-60%. Provide the standard chow diet and water ad libitum until the EHS protocol.
   NOTE: The rationale for individual housing is to avoid frequent fighting injury in male C57bl/6J mice and to provide ample opportunity for spontaneous wheel running for each mouse.
3. For placement of the telemetry devices, anesthetize the mouse with isoflurane (4%, 0.4-0.6 L/min of O₂ flow) in an induction chamber. Then, place the mouse under continuous anesthesia via a nose cone (1.5%, 0.6 L/min).
4. Use eye lube, such as a vet ointment, to protect the animal's eyes from damage or injury during surgery.
5. To prepare the surgical site, shave the lower abdomen with small animal hair clippers or use a commercially available hair remover. Administer the first dose of subcutaneous buprenorphine (0.1 mg/kg) during this time.
6. Scrub the area with three washes of povidone-iodine (or similar germicidal scrub) followed by 70% isopropyl alcohol rinse (or sterile saline depending on local veterinary requirements). Then, transfer the mouse to the surgical area.
7. Use an adhesive drape to isolate the surgical site on the mouse. Using sterile instruments and aseptic technique, make a ~1 cm incision on the midline along the linea alba, about 0.5 cm from the costal margin. Then, separate the skin from the muscle layer and make a slightly smaller incision on the linea alba, careful not to damage the bowels or internal organs.
8. Once the muscle layer is open, place the sterile telemeter (miniature reusable battery-free radiotelemetry device; 16.5 x 6.5 mm) into the intraperitoneal cavity in front of the caudal arteries and veins and dorsal to the digestive organs to allow it to float freely.
   NOTE: All telemeters are cleaned with soap and water, thoroughly rinsed and gas sterilized with ethylene oxide between usages. If gas sterilization is not available, immersion in sterilization solutions (following manufacturer’s recommendation for dilution and immersion time) is accepted to  disinfect and sterilize the telemeters.
9. Close the abdominal opening with a sterile 5-0 absorbable suture, and close the skin using a simple interrupted stitch with 5-0 proline suture.
   NOTE: Allowing the telemeter to float in the abdominal compartment without tying it to the abdominal wall (a method recommended by the manufacturer) has been demonstrated to be successful and preferred by the authors to eliminate excess tension in the abdominal wall during healing. Further, this has no impact on the receiver's ability to obtain the signal from the emitter.
10. Place the mouse in its clean cage with a portable heating pad under the cage. Monitor the mouse every 15 min during the first hour of recovery from anesthesia, and then return to the animal housing facility.
11. Provide mice with subcutaneous buprenorphine injections every 12 h for 48 h during recovery and continue to monitor for signs of distress. If available, give slow-release buprenorphine subcutaneously every 24 h (1 mg/kg) for 48 h. Allow the mice to recover for ~2 weeks following surgery before introducing a voluntary wheel running.

2. Familiarization: Voluntary and forced wheel running

1. Following recovery from surgery, place the voluntary running wheels in the cage for free access to the wheel. Other running wheel selections may be equally effective, but ensure it fits within the limited cage sizes available.
   NOTE: The running wheels had to be slightly reduced in dimension to fit in a standard cage.
2. Acclimate the mouse to the voluntary wheel in the cage for 2 weeks. Once acclimated, the mouse is ready for training with familiarization procedures for the forced running wheels.
3. Perform the four training sessions (one/day) in the environmental chamber at room temperature (~25 °C, 30% relative humidity).
   NOTE: Although this is ideal, mice were also successfully trained in identical forced running wheels outside the chamber. Several mice can then be trained simultaneously without interfering with the use of the chamber.
4. To begin the first training session, allow the mouse to free wheel in the modified running wheel for 15 min by removing or loosening the motor drive belt to allow the mouse to determine the speed of the wheel and acclimatize to it in a non-stressful manner.
   NOTE: Protocols can be run with software and hardware supplied by the running wheel manufacturer or may be substituted by an external programmable power supply that is wired directly to the wheel motor, which allows for automation of the incremental exercise protocol.
5. Calibrate the system for each running wheel to determine the relationship between the power supply voltage and meters/minute (m/min) of each wheel.
   NOTE: The forced running wheels were also modified to elevate the motor 15 cm, invert and move the pulley driving the wheel down to 5 cm above the telemetry receiver platform. This ensured that the receiver platform obtained accurate telemetry data during the running protocol without interference from the motor.
6. After a brief rest period (<5 min), initiate the forced running wheel protocol. Start the wheel at 2.5 m/min and increase 0.3 m/min every 10 min for a total of 1 h to mimic the first hour of the actual EHS trial, but at room temperature. Return the mouse to its home cage and allow for 24 h recovery. Conduct the subsequent three forced running sessions in the same manner on consecutive days. After day 1, the free-wheeling acclimation portion is unnecessary.
7. Allow the mouse 2-3 days of wash-out or recovery from the stress of the forced running wheel practice, but allow the mouse free access to the home cage voluntary wheel. The mouse is now prepared to undergo the EHS protocol.

3. EHS protocol

1. The night before the EHS protocol, place the mouse in the environmental chamber at room temperature (~25 °C, ≈30% relative humidity) to acclimate to the chamber.
2. Use a data acquisition system to collect continuous Tc, averaged over 30-s intervals overnight.
3. On the morning of the EHS protocol, make sure the mouse is at or below a normal range of diurnal temperature before increasing the chamber temperature (i.e., 36-37.5 °C). This ensures the mouse does not have a fever and has not experienced undue stress during this period.
4. Once the mouse is stable and within a range of normal resting core temperature, remove the food and water and weigh the animal. Shut the chamber door and increase the chamber temperature to a target of 37.5 °C and 40%-50% relative humidity, or the desired environmental temperature and humidity¹⁹. Verify the chamber temperature and humidity with a calibrated temperature and humidity monitor.
5. Surround the chamber with a black-out curtain to keep light and disturbances minimal during the protocol. Monitor the mouse continuously during the protocol via remote IR illuminated cameras. Focus a second camera on the temperature and humidity monitor, placed close to the running wheel. Make any adjustments to the controller for the environmental chamber set-point to ensure accurate temperature readings near the animal.
6. Once the chamber has reached its target temperature as measured by the second camera on the temperature monitor (this can take ~30 min), quickly open the chamber door and place the mouse in the forced running wheel.
7. Initiate the forced running wheel protocol at a speed of 2.5 m/min and increase the speed 0.3 m/min every 10 min until the mouse reaches a Tc of 41 °C. Once the mouse has reached this core temperature, allow the speed to remain constant until symptom limitation, characterized by an apparent loss of consciousness, a backward fall or fainting, and the inability to continue to run or hold on to the wheel. Confirm this time point when the mouse has three backward rotations on the wheel without signs of a physical response. Alternatively, identify a humane endpoint following local IACUC rules to determine when to stop the protocol (e.g., when Tc ~43 °C). This endpoint is slightly above symptom limitation in essentially all mice.
8. To perform the Rapid Cooling protocol (R), once the mouse reaches symptom limitation, stop the wheel, and remove it immediately from the forced running wheel. Weigh the mouse and place it back in its home cage to recover at room temperature. During this time, leave the chamber door open and return the incubator set point to room temperature to allow the chamber to cool rapidly. This procedure results in >99% long-term survival.
9. To perform a more Severe (S) EHS exposure, keep the animal's home cage within the 37.5 °C chamber during the EHS protocol. When the animal reaches symptom limitation, allow them to remain in the running wheel until they return to consciousness as observed by the remote camera (~5-9 min).
10. Then quickly remove the mouse from the running wheel and return it directly to its pre-warmed cage to result in a much slower cooling profile (Figure 1A, red dashed line), essentially eliminating the EHS hypothermic phase. Remove the filter top from the cage during this time to improve equilibration with the chamber.
11. Use a recovery cage precooled to room temperature to perform a less severe alternative procedure to result in a suppressed hypothermic phase but with a 100% survival rate²⁰.
12. For the S protocol, carefully monitor the mouse during recovery and check continuously for humane endpoints. Although it is difficult to remotely test for commonly used humane endpoints (e.g., righting reflex), observe the mice remotely for normal movements during recovery such as grooming, normal breathing, licking, etc. Monitor the Tc during this time.
13. Mice are unlikely to recover if their core temperature reverses direction during the recovery phase, eventually exceeding 40 °C; at this time, terminate the experiment and evaluate the mouse for standard humane endpoints.

Results

The typical thermoregulatory profiles during the entirety of the EHS protocol and early recovery of a mouse is illustrated in Figure 1A. This profile comprises four distinct phases that can be defined as the chamber heating stage, incremental exercise stage, steady-state exercise stage, and a recovery stage by either a rapid cooling (R) or severe (S) method¹⁷. The main thermoregulatory outcomes include maximum Tc achieved (Tc,max) and the time required to reach Tc,max...

Discussion

This technical review aims to provide guidelines for the performance of a preclinical model of EHS in mice. Detailed steps and materials required for the execution of a reproducible EHS episode of variable severities are provided. Importantly, the model largely mimics the signs, symptoms, and multi-organ dysfunction observed in human EHS victims^11,19. Furthermore, this model allows for the examination of the mechanism underlying short- and long-term EHS recovery<...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1080P HD 4 Security Cameras 4CH Home Video Security Camera System w/ 1TB HDD 2MP Night View Cameras CCTV Surveillance Kit	LaView
5-0 Coated Vicryl Violet Braided	Ethicon
5-0 Ethilon Nylon suture Black Monofilament	Ethicon
Adhesive Surgical Drape with Povidone 12x18	Jorgensen Labset al.
BK Precision Multi-Range Programmable DC Power Supplies Model 9201	BK Precision
DR Instruments Medical Student Comprehensive Anatomy Dissection Kit	DR Instruments
Energizer Power Supply	Starr Life Sciences
G2 Emitteret al.	Starr Life Sciences
Layfayette Motorized Wheel Model #80840B	Layfayette
Patterson Veterinary Isoflurane	Patterson Veterinary
Platform receiveret al.	Starr Life Sciences
Scientific Environmental Chamber Model 3911	ThermoForma
Training Wheels	Columbus Inst.