Name: a preclinical model of exertional heat stroke in mice
Uploaded: 2021-07-01T14:30:00.000+00:00
Duration: 8 min 22 s
Description: Watch this Scientific Journal Video about A Preclinical Model of Exertional Heat Stroke in Mice at JoVE.com

This work was funded by the Department of Defense W81XWH-15-2-0038 (TLC) and BA180078 (TLC) and the BK and Betty Stevens Endowment (TLC). JMA was supported by financial aid from the Kingdom of Saudi Arabia. Michelle King was with the University of Florida at the time this study was conducted. She is currently employed by the Gatorade Sports Science Institute, a division of PepsiCo R&#38;D.

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
<ol>
	<li>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.</li>
	<li>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&#34; (W) x 11.75&#34; (L) x 5&#34; (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 &#176;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.</li>
	<li>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).</li>
	<li>Use eye lube, such as a vet ointment, to protect the animal&#39;s eyes from damage or injury during surgery.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.&#160;If gas sterilization is not available, immersion in sterilization solutions (following manufacturer&#8217;s recommendation for dilution and immersion time) is accepted to&#160; disinfect and sterilize the telemeters.</li>
	<li>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&#39;s ability to obtain the signal from the emitter.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
2. Familiarization: Voluntary and forced wheel running
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Perform the four training sessions (one/day) in the environmental chamber at room temperature (~25 &#176;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>After a brief rest period (&#60;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.</li>
	<li>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.</li>
</ol>
3. EHS protocol
<ol>
	<li>The night before the EHS protocol, place the mouse in the environmental chamber at room temperature (~25 &#176;C, &#8776;30% relative humidity) to acclimate to the chamber.</li>
	<li>Use a data acquisition system to collect continuous Tc, averaged over 30-s intervals overnight.</li>
	<li>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 &#176;C). This ensures the mouse does not have a fever and&#160;has not experienced undue stress during this period.</li>
	<li>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 &#176;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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#176;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 &#176;C). This endpoint is slightly above symptom limitation in essentially all mice.</li>
	<li>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&#160;and return the incubator set point to room temperature to allow the chamber to cool rapidly. This procedure results in &#62;99% long-term survival.</li>
	<li>To perform a more Severe (S) EHS exposure, keep the animal&#39;s home cage within the 37.5 &#176;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).</li>
	<li>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.</li>
	<li>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.</li>
	<li>For the S protocol, carefully monitor the mouse&#160;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.</li>
	<li>Mice are unlikely to recover if their core temperature reverses direction during the recovery phase, eventually exceeding 40 &#176;C; at this time, terminate the experiment and evaluate the mouse for standard humane endpoints.</li>
</ol>

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The authors have no conflicts of interest to disclose. All work performed and all support for this project were generated at the University of Florida.

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 victims11,19. Furthermore, this model allows for the examination of the mechanism underlying short- and long-term EHS recovery<...

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 dehydration3,4. 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 outcomes6,7. 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 characterized8,9,10. 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 rates12,13,14,15,16 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&#39; knowledge, this is the only preclinical model of EHS with such characteristics, making it possible to pursue relevant EHS research in a hypothesis-driven manner11,17,18.

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a preclinical model of exertional heat stroke in mice

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 &#176;C and 40%-50% relative humidity (RH) until displaying symptom limitation (e.g., loss of consciousness) at Tc of 42.1-42.5 &#176;C, although suitable results can be obtained at chamber temperatures between 34.5-39.5 &#176;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 na&#239;ve 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.

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&#160;the time required to reach T...

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A Preclinical Model of Exertional Heat Stroke in Mice

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.

Heat stroke is the most severe manifestation of heat-related illnesses. Classic heat stroke (CHS), also known as passive heat stroke, occurs at rest, ...

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3.6K Views.  University of Florida. 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.

Video: A Preclinical Model of Exertional Heat Stroke in Mice

Assessment of Murine Exercise Endurance Without the Use of a Shock Grid: An Alternative to Forced Exercise

Using laboratory mouse models, the molecular pathways responsible for the metabolic benefits of endurance exercise are beginning to be defined. The most common method for assessing exercise endurance in mice utilizes forced running on a motorized treadmill equipped with a shock grid. Animals who quit running are pushed by the moving treadmill belt onto a grid that delivers an electric foot shock; to escape the negative stimulus, the mice return to running on the belt. However, avoidance behavior and psychological stress due to use of a shock apparatus can interfere with quantitation of running endurance, as well as confound measurements of postexercise serum hormone and cytokine levels. Here, we demonstrate and validate a refined method to measure running endurance in na&#239;ve C57BL/6 laboratory mice on a motorized treadmill without utilizing a shock grid. When mice are preacclimated to the treadmill, they run voluntarily with gait speeds specific to each mouse. Use of the shock grid is replaced by gentle encouragement by a human operator using a tongue depressor, coupled with sensitivity to the voluntary willingness to run on the part of the mouse. Clear endpoints for quantifying running time-to-exhaustion for each mouse are defined and reflected in behavioral signs of exhaustion such as splayed posture and labored breathing. This method is a humane refinement which also decreases the confounding effects of stress on experimental parameters.

A method to assess exercise endurance in laboratory mice without the use of a shock grid is demonstrated. This method is a humane refinement that can decrease the confounding effects of stress on experimental parameters.

Using laboratory mouse models, the molecular pathways responsible for the metabolic benefits of endurance exercise are beginning to be defined. The most ...

Behavior

Normothermic Cardiac Arrest and Cardiopulmonary Resuscitation: A Mouse Model of Ischemia-Reperfusion Injury

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused by whole-body hypoperfusion.5,6 Successful reproduction of whole-body hypoperfusion in rodent models has been fraught with 
difficulty.7-9,9,10 Models which employ focal ischemia have repeatedly demonstrated results which do not translate to the clinical setting, and larger 
animal models which allow for whole body hypoperfusion lack access to the full toolset of genetic manipulation possible in the mouse.11,12 However, in recent 
years a mouse model of cardiac arrest and cardiopulmonary resuscitation has emerged which can be adapted to model AKI.13 This model reliably reproduces 
physiologic, functional, anatomic, and histologic outcomes seen in clinical AKI, is rapidly repeatable, and offers all of the significant advantages of a murine surgical 
model, including access to genetic manipulative techniques, low cost relative to large animals, and ease of use. Our group has developed extensive experience 
with use of this model to assess a number of organ-specific outcomes in AKI.14,15

A powerful model for perioperative and critical care related acute kidney injury is presented. Using whole body hypoperfusion induced by cardiac arrest it is possible to nearly replicate the histologic and functional changes of clinical AKI.

Acute Kidney Injury (AKI) is a common, highly lethal, complication of critical illness which has a high mortality1-4 and which is most 
frequently caused ...

Protocol for Long Duration Whole Body Hyperthermia in Mice

Hyperthermia is a general term used to define the increase in core body temperature above normal. It is often used to describe the increased core body temperature that is observed during fever. The use of hyperthermia as an adjuvant has emerged as a promising procedure for tumor regression in the field of cancer biology. For this purpose, the most important requirement is to have reliable and uniform heating protocols. We have developed a protocol for hyperthermia (whole body) in mice. In this protocol, animals are exposed to cycles of hyperthermia for 90 min followed by a rest period of 15 min. During this period mice have easy access to food and water. High body temperature spikes in the mice during first few hyperthermia exposure cycles are prevented by immobilizing the animal. Additionally, normal saline is administered in first few cycles to minimize the effects of dehydration. This protocol can simulate fever like conditions in mice up to 12-24 hr. We have used 8-12 weeks old BALB/Cj female mice to demonstrate the protocol.

This paper describes a protocol for whole body hyperthermia in mice that can stimulate fever like conditions up to 12-24 hr.

Hyperthermia is a general term used to define the increase in core body temperature above normal. It is often used to describe the increased core body ...

Supramaximal Intensity Hypoxic Exercise and Vascular Function Assessment in Mice

Exercise training is an important strategy for maintaining health and preventing many chronic diseases. It is the first line of treatment recommended by international guidelines for patients suffering from cardiovascular diseases, more specifically, lower extremity artery diseases, where the patients' walking capacity is considerably altered, affecting their quality of life. 
Traditionally, both low continuous exercise and interval training have been used. Recently, supramaximal training has also been shown to improve athletes' performances via vascular adaptations, amongst other mechanisms. The combination of this type of training with hypoxia could bring an additional and/or synergic effect, which could be of interest for certain pathologies. Here, we describe how to perform supramaximal intensity training sessions in hypoxia on healthy mice at 150% of their maximal speed, using a motorized treadmill and a hypoxic box. We also show how to dissect the mouse in order to retrieve organs of interest, particularly the pulmonary artery, the abdominal aorta, and the iliac artery. Finally, we show how to perform ex vivo vascular function assessment on the retrieved vessels, using isometric tension studies.

High-intensity training in hypoxia is a protocol that has been proven to induce vascular adaptations potentially beneficial in some patients and to improve athletes&#39; repeated sprint ability. Here, we test the feasibility of training mice using that protocol and identify those vascular adaptations using ex vivo vascular function assessment.

Exercise training is an important strategy for maintaining health and preventing many chronic diseases. It is the first line of treatment recommended by ...

Measuring Skeletal Muscle Thermogenesis in Mice and Rats

Skeletal muscle thermogenesis provides a potential avenue for better understanding metabolic homeostasis and the mechanisms underlying energy expenditure. Surprisingly little evidence is available to link the neural, myocellular, and molecular mechanisms of thermogenesis directly to measurable changes in muscle temperature. This paper describes a method in which temperature transponders are utilized to retrieve direct measurements of mouse and rat skeletal muscle temperature.
Remote transponders are surgically implanted within the muscle of mice and rats, and the animals are given time to recover. Mice and rats must then be repeatedly habituated to the testing environment and procedure. Changes in muscle temperature are measured in response to pharmacological or contextual stimuli in the home cage. Muscle temperature can also be measured during prescribed physical activity (i.e., treadmill walking at a constant speed) to factor out changes in activity as contributors to the changes in muscle temperature induced by these stimuli.
This method has been successfully used to elucidate mechanisms underlying muscle thermogenic control at the level of the brain, sympathetic nervous system, and skeletal muscle. Provided are demonstrations of this success using predator odor (PO; ferret odor) as a contextual stimulus and injections of oxytocin (Oxt) as a pharmacological stimulus, where predator odor induces muscle thermogenesis, and Oxt suppresses muscle temperature. Thus, these datasets display the efficacy of this method in detecting rapid changes in muscle temperature.

Mice and rats are surgically implanted with remote temperature transponders and then habituated to the testing environment and procedure. Changes in muscle temperature are measured in response to pharmacological or contextual stimuli in the home cage or during prescribed physical activity (i.e., treadmill walking at a constant speed).

Skeletal muscle thermogenesis provides a potential avenue for better understanding metabolic homeostasis and the mechanisms underlying energy expenditure. ...

A Behavioral Screen for Heat-Induced Seizures in Mouse Models of Epilepsy

Transgenic mouse models have proved to be powerful tools in studying various aspects of human neurological disorders, including epilepsy. The SCN1A-associated genetic epilepsies comprise a wide spectrum of seizure disorders with incomplete penetrance and clinical variability. SCN1A mutations can result in a large variety of seizure phenotype ranging from simple, self-limited fever-associated febrile seizures (FS), moderate-level genetic epilepsy with febrile seizures plus (GEFS+) to more severe Dravet Syndrome (DS). Although FS are commonly seen in children below 6-7 years of age who do not have genetic epilepsy, FS in GEFS+ patients continue to occur into adulthood. Traditionally, experimental FS have been induced in mice by exposing the animal to a stream of dry air or heating lamps, and the rate of change in body temperature is often not well controlled. Here, we describe a custom-built heating chamber, with a plexiglass front, that is fitted with a digital temperature controller and a heater-equipped electric fan, which can send heated forced air into the test arena in a temperature-controlled manner. The body temperature of a mouse placed in the chamber, monitored through a rectal probe, can be increased to 40-42 &#176;C in a reproducible manner by increasing the temperature inside the chamber. Continual visual monitoring of the animals during the heating period demonstrates induction of heat-induced seizures in mice carrying an FS mutation at a body temperature that does not elicit behavioral seizures in wild-type litter mates. Animals can be easily removed from the chamber and placed on a cooling pad to rapidly return body temperature to normal. This method provides for a simple, rapid, and reproducible screening protocol for the occurrence of heat-induced seizures in epilepsy mouse models.

The goal of the method is to screen for hyperthermia or heat-induced seizures in mouse models. The protocol describes the use of a custom-built chamber with continuous monitoring of the body temperature to determine whether elevated body temperature leads to seizures.

Transgenic mouse models have proved to be powerful tools in studying various aspects of human neurological disorders, including epilepsy. The ...

Neuroscience

Establishing A Mouse Model for Respiratory Tract Extreme Low-temperature Air Inhalation

Currently, constructing a mouse model for cold environmental stimulation employs cold-heat plates and wearable cooling devices. These methods can partially fulfill the requirements for studying the responses and regulatory effects of mouse skin or neural circuits to cold stimulation. Numerous clinical studies have substantiated the correlation between exposure to low-temperature environments and the development of various diseases. Recently, there has been a growing emphasis on the continuous exchange of information between organs and tissues, providing a novel perspective on addressing longstanding issues within the human body. However, existing installations are unable to construct a model for mice inhaling cold air. 
Although placing mice in a cold environment seems attractive, it has considerable limitations. While mice inhale cold air, their skin is also being stimulated by the cold environment, making it unclear whether resulting pathological changes are due to lung stimulation through the interaction of distant organs or due to the skin receptors and neural signal transmission. This creates considerable confusion in related research. This scheme presents a new approach for constructing a mouse model for extreme cold air inhalation stimulation. This device allows mice to inhale extremely low-temperature gases while their bodies remain at a normal temperature. It maximizes the simulation of the stimulating effects of extreme ambient temperatures on mice and meets the research needs for studying the relationship between extreme environmental temperatures and related diseases.

Cold environmental stimulation has been implicated in the development of various chronic diseases. Therefore, establishing animal models for preclinical research is crucial. This system addresses this need by offering a device that creates a stimulus model, meeting the requirements for basic research on pathogenic mechanisms.

Currently, constructing a mouse model for cold environmental stimulation employs cold-heat plates and wearable cooling devices. These methods can ...

Mouse Model for Studying Sepsis-Induced Myopathy with Hindlimb Suspension

A Preclinical Model of Sepsis-Induced Myopathy with Disuse in Mice

Sepsis is a major cause of in-hospital deaths. Improvements in treatment result in a greater number of sepsis survivors. Approximately 75% of the survivors develop muscle weakness and atrophy, increasing the incidence of hospital readmissions and mortality. However, the available preclinical models of sepsis do not address skeletal muscle disuse, a key component for the development of sepsis-induced myopathy. Our objective in this protocol is to provide a step-by-step guideline for a mouse model that reproduces the clinical setting experienced by a bedridden septic patient. Male C57Bl/6 mice were used to develop this model. Mice underwent cecal ligation and puncture (CLP) to induce sepsis. Four days post-CLP, mice were subjected to hindlimb suspension (HLS) for seven days. Results were compared with sham-matched surgeries and/or animals with normal ambulation (NA). Muscles were dissected for in vitro muscle mechanics and morphological assessments. The model results in marked muscle atrophy and weakness, a similar phenotype observed in septic patients. The model represents a platform for testing potential therapeutic strategies for the mitigation of sepsis-induced myopathy.

Author Spotlight: Advanced Integrated Model for Sepsis-Induced Myopathy and Single-Cell Metabolic Analysis

This murine model combines a septic insult with hindlimb muscle disuse to recapitulate the bedridden feature of the typical septic patient. The model represents a significant departure from previous models to study muscle dysfunction in sepsis and is a reproducible approach to addressing therapeutic strategies to treat this condition.

Sepsis is a major cause of in-hospital deaths. Improvements in treatment result in a greater number of sepsis survivors. Approximately 75% of the ...

Exertional Heat Stroke Mouse Model: A Protocol to Study Mechanisms Underlying Exertional Heat Stroke

Source: King, M. A. et al. <a href="https://www.jove.com/video/62738" target="_blank">A Preclinical Model of Exertional Heat Stroke in Mice</a>. J. Vis. Exp. (2021).
In this video we describe a step-by-step protocol to study exertional heat stroke (EHS) in a mouse model.

Source: King, M. A. et al. A Preclinical Model of Exertional Heat Stroke in Mice. J. Vis. Exp. (2021).
In this video we describe a step-by-step protocol ...

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Biology

Rodent Models

Adult

Surgical Implantation of Temperature Telemetric Monitoring System in Mouse Model: A Surgical Procedure to Implant Temperature Telemetric Device to Measure Mouse Core Body Temperature

Source: King, M. A. et al.&#160; <a href="https://www.jove.com/video/62738" target="_blank">A Preclinical Model of Exertional Heat Stroke in Mice</a>. J. Vis. Exp. (2021)
In this video, we describe a surgical procedure to implant a temperature telemetric device&#160; in a mouse model to continuously measure its core body temperature.