This work was supported by the German Research Foundation (DFG; Clinician Scientist Program in Vascular Medicine (PRIME), MA 2186/14-1 to P. Tomsits), the German Centre for Cardiovascular Research (DZHK; 81X2600255 to S. Clauss), the Corona Foundation (S199/10079/2019 to S. Clauss), and the ERA-NET on Cardiovascular Diseases (ERA-CVD; 01KL1910 to S. Clauss). The funders had no role in manuscript preparation.

The authors have nothing to disclose.

Current guidelines recommend regular physical activity as it has been demonstrated to be an important modifier of cardiovascular risk factors30. There is also a growing body of evidence that moderate physical activity may protect against atrial fibrillation (AF) both in primary and secondary prevention31,32,33. On the contrary, endurance athletes such as marathon runners have a higher risk to develop AF indicating that endurance training may also have negative effects34,35. Such a U-shaped relationship between arrhythmia risk and training intensity has been clearly shown for AF in otherwise healthy athletes9,36,37,38 and in patients with underlying heart disease, however, only little is known regarding training intensity and arrhythmogenesis4,5,6,7.
To overcome this limitation and to improve patient care, further research on exercise-related effects on cardiac electrophysiology is warranted. To investigate fundamental mechanisms and molecular/cellular adaptations in response to training different models in a number of animal species have been developed15. Given the immanent benefits but also limitations of every model/species, researchers have to choose the most suitable one for each individual research question; regarding electrophysiology and arrhythmia research mouse13,14,39,40 and pig models are widely used13,14,41,42,43. Although training protocols using a motorized treadmill have been developed in pigs, there are a number of significant challenges including (i) the pigs&#39; sedentary behavior, which requires a time- and labor-intense conditioning prior to the experiment as well as stimuli to keep pigs compliant during experiment and (ii) the body size and weight, which may prevent training in older pigs or training over long periods of time15,44. In mice, several exercise protocols have been developed including motorized treadmill training, VWR, or swimming17,18. Although VWR mimics the natural running pattern in rodents and is less stressful compared to forced exercise methods such as swimming and treadmill training, it also has certain disadvantages45. The spontaneous nature of VWR does not allow controlling intensity, duration, or frequency of exercise thus preventing well-controlled experiments. In swimming models, the duration and intensity of training can be easily regulated, the necessary equipment is simple and available at low costs, and the method can be established in most research labs46. Despite these advantages, studying electrophysiology in a swimming model is difficult as there is currently no option to monitor the ECG during swimming. The approach described in this protocol combines an implantable telemetry system with a treadmill exercise model and thus, overcomes the limitations of other training models in the context of electrophysiology research47,48. Using a treadmill allows to control various exercise conditions such as intensity (slope inclination and running speed) or duration. In addition, different training protocols can be studied including endurance exercise training, interval training, and acute exercises. Following this protocol, it is now also possible to record and monitor the ECG using implantable telemetry transmitters while the mouse is running on the treadmill.
Given that mice usually run willingly for only a few minutes, stimuli such as tapping their back with little sticks, blowing puffs of compressed air or electrical stimuli are necessary. These stimuli, however, can induce psychological stress, which can affect the quality of experimental data significantly. Hence, we tried to minimize these stress factors by letting the mouse adapt to the treadmill during an acclimatization phase, with a steady increment of speed and using minimum-to-zero shock intensity as previously described15,17,45.
In general, when recording ECGs, motion artefacts are a major issue, especially during physical activity. Following our proposed protocol, researchers will be able to acquire ECG signals in good quality allowing to clearly distinguish and annotate P, Q, R, S, T (Figure 3). Thus, various ECG parameters such as heart rate, heart rate variability, PR interval, QRS duration, or QT duration can reliably be assessed before, during, and after the training using automated software algorithms. Also, arrhythmias such as tachyarrhythmia, bradyarrhythmia, or pauses can be detected. As heart rate variability analyses-usually performed to investigate the effects of the autonomic nervous system on the heart27,28-depend on sufficient R-wave annotation, data quality can be verified by similarly low SDRR values obtained at rest and during training by automated annotation as shown in Figure 6.
As every experimental technique, this method does not come without pitfalls and contains several critical steps. Sterile conditions and a short operation time are requirements for successful transmitter implantation, proper wound healing, and fast animal recovery post-surgery. Sutures must not be too tight, or they will cause skin necrosis. In general, the surgical procedure requires practical experience, and results will improve over time. Lead positioning influences the recorded main vector, best results are obtained with a steep lead two position, as it results in higher P- and R-wave amplitudes, which in turn are critical requirements for later ECG analysis. Training mice can be challenging as not all animals train willingly. A well-designed acclimatization protocol, including introduction to the treadmill environment, slow increments in conveyor belt speed and positive enhancement of good training behavior, e.g., with food pellets, can help to condition the animals to train better and to reduce the need for potentially interfering stimuli during the experiments. It is important to reduce all stimuli to the absolute minimum as they may affect data quality. However, the most critical step is the optimal positioning of the telemetry receiver during the treadmill training as it directly determines the quality of the data obtained. The receiver position must be determined for each pair of animals training at the same time, as it varies depending on the exact position of telemetry device and leads as well as on the individual animals running pattern. The position is found by trial and error, visually judging the signal quality in real time. All ECG traits to be analyzed must be clearly visible before experiments can start. Given the high murine heart rate, many data points accumulate even with short recording periods. This and the overall low signal amplitude, naturally leading to a lower signal-to-noise ratio in rodents than in humans or large animals, make data analysis extremely challenging, as we have previously discussed23. A major limitation of this protocol besides the costly equipment needed to perform telemetry and treadmill training is the high technical demand on the surgical procedure and on data analysis, limiting accessibility to beginners in the field.
In sum, the ECG is a brilliant tool to study cardiac electrophysiology and arrhythmogenesis. In humans, stress tests to record ECGs during exercise are routinely performed and allow to assess training-associated effects on cardiac electrophysiology. Mice are the most commonly used species in research, several exercise protocols have been developed, but monitoring the real-time ECG during training was not possible so far. Our proposed protocol allows to obtain ECG recordings during periods of exercise in mice for the first time. This will enable researchers to study both exercise-related mechanisms leading to beneficial cardiac adaptations and maladaptive, proarrhythmic remodeling and will, thus, eventually result in improved patient care in the future.

Regular physical activity is important for a healthy life. Certain cardiovascular conditions, however, lead to situations where this common-sense agreement is at least questionable. In patients with myocarditis, current data even shows adverse effects of exercise and, thus, it is recommended to pause all exercise for a certain period in these patients1,2,3. In other cardiovascular diseases (CVD) such as inherited arrhythmia syndromes comparatively less evidence on the appropriate level of exercise exists4,5,6,7, making clinical counselling in these cases, mostly for young and physically active patients, very challenging.
Adverse remodeling leading to reduced contractility and heart failure and proarrhythmogenic remodeling leading to arrhythmias and sudden cardiac death have been suggested as hallmarks of exercise-associated harmful effects on the heart8. A large number of studies indicate beneficial effects of moderate exercise over a broad spectrum of different diseases9,10. Extensive training, however, may have detrimental effects on the heart leading to arrhythmias especially in otherwise healthy athletes11. Although structural remodeling processes leading to a vulnerable proarrhythmic substrate production may underlie this paradox situation as demonstrated in marathon runners12, the specific mechanisms of exercise-related adverse remodeling both in healthy people and in patients with cardiovascular diseases remain largely unknown.
In animals, especially in mice, several suitable models have been developed to mimic a broad range of cardiovascular diseases13,14. Also, various exercise models and training protocols have been established in mice15,16,17, including motorized treadmill training, voluntary wheel running (VWR), and swimming17,18. Evaluation of cardiac electrophysiology by ECG monitoring classically depends on a direct conducting connection between the animal and some sort of detection device. Thus, either animals need to be anesthetized, e.g., to obtain ECG recordings using sharp electrodes19, or animals need to be immobilized by a restrainer 20, or data quality is reduced due to motion artifacts, e.g., when using paw-electrodes21 or conductive platforms22 allowing only basic analysis. Thus, none of the above-mentioned approaches are compatible with training protocols and consequently prevent studies on exercise-related mechanisms leading to adverse remodeling in mice. Implantable telemetry devices can overcome these hurdles and are nowadays the most powerful tool and gold standard to evaluate murine electrophysiology in vivo in conscious and moving animals23,24. Current telemetry hardware solutions have been developed to monitor mice in their cages25,26,&#160;and commonly require a receiver to be placed underneath the cage for data acquisition, thus making real-time monitoring outside these circumstances challenging. Here we provide an approach to investigate the effects of exercise on cardiac electrophysiology and arrhythmogenesis by real-time ECG recording during treadmill training in mice using implanted telemetry devices. All parameters obtained were analyzed as previously described by Tomsits et al.23.

<table><tbody><tr><td>14-gauge needle</td><td>Sterican</td><td>584125</td><td></td></tr><tr><td>Any mouse</td><td>e.g. Jackson Laboratories</td><td></td><td></td></tr><tr><td>Bepanthen</td><td>Bayer</td><td>1578675</td><td></td></tr><tr><td>Carprofen 0.005 mg/&amp;micro;L</td><td>Zoetis</td><td>53716-49-7</td><td></td></tr><tr><td>Data Exchange Matrix 2.0 (MX2)</td><td>Data Science International</td><td></td><td>Manages communication between PhysioTel and PhysioTel HD telemetry implants and the acquisition computer.</td></tr><tr><td>Enrofloxacin 25 mg/ml</td><td>Baytril</td><td>400614.00.00</td><td></td></tr><tr><td>Fentanyl 0.5 mg/10 mL</td><td>Braun Melsungen</td><td></td><td></td></tr><tr><td>Fine forceps</td><td>Fine Science Tools</td><td>11295-51</td><td></td></tr><tr><td>Five Lane Treadmill for Mouse</td><td>Panlab - Harvard Apparatus</td><td>76-0896</td><td>Includes treadmill unit, touchscreen control unit, a sponge , and cables</td></tr><tr><td>Iris scissors</td><td>Fine Science Tools</td><td>14084-08</td><td></td></tr><tr><td>Isoflurane 1 mL/mL</td><td>Cp-Pharma</td><td>31303</td><td></td></tr><tr><td>Isoflurane vaporizer system</td><td>Hugo Sachs Elektronik</td><td>34-0458, 34-1030, 73-4911, 34-0415, 73-4910</td><td>Includes an induction chamber, a gas evacuation unit and charcoal filters</td></tr><tr><td>LabChart Pro 8.1.16</td><td>ADInstruments</td><td></td><td></td></tr><tr><td>Magnet</td><td>Data Science International</td><td></td><td></td></tr><tr><td>Modified Bain circuit</td><td>Hugo Sachs Elektronik</td><td>73-4860</td><td>Includes an anesthesia mask for mice</td></tr><tr><td>Modular connectors</td><td>Data Science International</td><td></td><td>Connecting cables between Reciever, Signal Interface and Matrix 2.0 (MX2)</td></tr><tr><td>Novafil s 5-0</td><td>Medtrocin/Covidien</td><td>88864555-23</td><td></td></tr><tr><td>Octal BioAmp</td><td>ADInstruments</td><td>FE238-0239</td><td>Amplifier for recording Surface ECG</td></tr><tr><td>Octenisept</td><td>Sch&amp;uuml;lke</td><td>121418</td><td></td></tr><tr><td>Oxygen 5 L</td><td>Linde</td><td>2020175</td><td>Includes a pressure regulator</td></tr><tr><td>PhysioTel ETA-F10 transmitter</td><td>Data Science International</td><td></td><td></td></tr><tr><td>PhysioTel receiver RPC-1</td><td>Data Science International</td><td></td><td>Signal reciever</td></tr><tr><td>Ponemah 6.42</td><td>Data Science International</td><td></td><td>ECG Analysis Software</td></tr><tr><td>Powerlab</td><td>ADInstruments</td><td>3516-1277</td><td>Suface ECG Acquisition hardware device. Includes ECG electrode leads</td></tr><tr><td>Prism 8.0.1</td><td>Graph Pad</td><td></td><td></td></tr><tr><td>Radio Device (Sony AF/AM)</td><td>Sony</td><td></td><td></td></tr><tr><td>Signal Interface</td><td>Data Science International</td><td></td><td>Acquires and synchronizes digital signals with telemetry data in Ponemah v6.x.</td></tr><tr><td>Spring scissors</td><td>Fine Science Tools</td><td>91500-09</td><td></td></tr><tr><td>Surgical platform</td><td>Kent Scientific</td><td>SURGI-M</td><td></td></tr><tr><td>Tergazyme 1%</td><td>Alconox</td><td>13051.0</td><td>Commercial cleaning solution</td></tr><tr><td>Tweezers</td><td>Kent Scientific</td><td>INS600098-2</td><td></td></tr></tbody></table>

real-time electrocardiogram monitoring during treadmill training in mice

Regular physical exercise is a major contributor to cardiovascular health, influencing various metabolic as well as electrophysiological processes. However, in certain cardiac diseases such as inherited arrhythmia syndromes, e.g., arrhythmogenic cardiomyopathy (ACM) or myocarditis, physical exercise may have negative effects on the heart leading to a proarrhythmogenic substrate production. Currently, the underlying molecular mechanisms of exercise-related proarrhythmogenic remodeling are largely unknown, thus it remains unclear which frequency, duration, and intensity of exercise can be considered safe in the context of disease(s).
The proposed method allows to study proarrhythmic/antiarrhythmic effects of physical exercise by combining treadmill training with real-time monitoring of the ECG. Implantable telemetry devices are used to continuously record the ECG of freely moving mice over a period of up to 3 months both at rest and during treadmill training. Data acquisition software with its analysis modules is used to analyze basic ECG parameters such as heart rate, P wave duration, PR interval, QRS interval, or QT duration at rest, during and after training. Furthermore, heart rate variability (HRV) parameters and occurrence of arrhythmias are evaluated. In brief, this manuscript describes a step-by-step approach to experimentally explore exercise induced effects on cardiac electrophysiology, including potential proarrhythmogenic remodeling in mouse models.

Depending on individual research objectives, subsequent analysis of obtained telemetry data will differ widely. Here, we demonstrate feasibility of the method by obtaining good quality data recorded during training periods and provide exemplary results of ECGs and heart rate variability analyses before, during, and after training. Data are presented as mean ± standard error of mean (SEM), all statistical analyses were conducted with a suitable statistical software (see Table of Materials). Statistical significance was assessed by the student's t test. The QT interval is corrected as previously discussed by Roussel et al. using the formula QTc = QT / (√(RR / 100))29.
Successful telemetric ECG recording during training 
With this protocol, it is possible to obtain ECG data with clear P, Q, R, S, and T waves in animals during training as demonstrated in Figure 3.
All the measurements from one animal were taken from the same day. Baseline measurements were taken at 10 am ± 10 min before training when animals were still in their permanent housing. Measurements during training were taken from the middle of the 60 min training session ± 10 min on day 3 in the third week of training, post training measurements were taken from the 5 min resting period after training and before retransfer to the permanent housing and recovered measurements were taken 1 h after training ± 10 min. Suitable sections of the ECG tracing for analysis were chosen manually from these defined sections with regard to the readout, e.g., 40 consecutive cycles for data presented in Figure 4.
Evaluation of ECG-derived parameters 
Data are used to analyze physiologic changes before, during, and after exercise as shown for one example animal in Figure 4. Heart rate (Figure 4A), PR interval (Figure 4B), QRS duration (Figure 4C), and QTc interval (Figure 4D) are evaluated by averaging 40 consecutive ECG cycles. Heart rate increases to around 800 bpm when the animal is exercising and gradually recovers toward baseline after training. PR interval, QRS duration, and QTc intervals shorten under stress and once stress is over, return to baseline. Exemplary data from one animal are shown.
Detection of tachycardia 
Search definitions were used as described in step 3.2.4 for detection of tachycardia and bradycardia episodes. Figure 5A shows sinus rhythm at baseline. A representative trace of sinus tachycardia during training is shown in Figure 5B. Exemplary data from one animal is shown here.
Data quality assessment by evaluation of heart rate variability parameters 
HRV analysis is done as described in step 3.3. 5 min sections for HRV analysis is presented in Figure 6. Figure 6A shows the heart rate of a single animal over the course of an experiment. Heart rate increases during training and gradually returns to baseline post training, this trend can also be visualized by the median RR interval as shown in Figure 6B. Figure 6C shows comparable standard deviation of RR intervals (SDRR) obtained at baseline and during training by automated RR annotation, demonstrating data quality. Data obtained is from three mice. The SDRR is the standard deviation of all interbeat intervals (IBI) and is calculated automatically by the software as positive square root of the IBI variance around the mean IBI using the formula:
σx = <img alt="Square root of variance formula, √Var(IBI), used in statistical analysis or data variability." src="/files/ftp_upload/63873/63873eq01.jpg" style="margin: auto;" data-altupdatedat="1747911167000" data-alt="Equation 1">
<img alt="Rodent telemetry setup, diagram showing electrodes and transmitter for physiological data collection." class="xfigimg" src="/files/ftp_upload/63873/63873fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Schematic illustration of the telemetry transmitter and lead positioning. Mouse is in supine position; the transmitter is placed intraperitoneally, and leads are fixed subcutaneously in a lead II configuration. Created with Biorender. <a href="https://www.jove.com/files/ftp_upload/63873/63873fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Signal acquisition and visualization system, animal activity monitoring, treadmill setup, experiment diagram." class="xfigimg" src="/files/ftp_upload/63873/63873fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Experimental setup. (A) Setup for ECG recording using implantable telemetry before and after training with the signal receiver being kept underneath the animal cage. (B) Setup for real-time ECG monitoring during treadmill training. For optimal signal quality, the signal receiver is placed on the transparent box. Created with Biorender. <a href="https://www.jove.com/files/ftp_upload/63873/63873fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Electrocardiogram waveform showing PQRST sequence and RR interval; cardiac cycle analysis diagram." class="xfigimg" src="/files/ftp_upload/63873/63873fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Representative ECG during training. Normal sinus rhythm, P-wave, QRS, and T-wave are indicated using capital letters, RR-interval is marked with a bar. <a href="https://www.jove.com/files/ftp_upload/63873/63873fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Cardiac response graphs; heart rate, PR, QRS, QTc intervals; training phases; ECG analysis." class="xfigimg" src="/files/ftp_upload/63873/63873fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Telemetry over time. Trend graphs show representative results for (A) heart rate (BPM). (B) PR interval (ms). (C) QRS duration (ms). (D) QTc Interval (ms) before (baseline), during (training), immediately after training (post training), and after full recovery (recovered). Data is obtained from one animal by averaging 40 consecutive ECG cycles. <a href="https://www.jove.com/files/ftp_upload/63873/63873fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Sinus rhythm vs. sinus tachycardia ECG graph showing baseline, training heart electrical activity." class="xfigimg" src="/files/ftp_upload/63873/63873fig05.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Representative ECGs before and during training. (A) Sinus rhythm before training. (B) Sinus tachycardia during training. Data is from one animal. <a href="https://www.jove.com/files/ftp_upload/63873/63873fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Heart rate trend chart; bar graphs show median RR interval and SDRR post-training recovery data." class="xfigimg" src="/files/ftp_upload/63873/63873fig06.jpg" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
Figure 6: Data quality assessment by HRV analysis. (A) Representative heart rate trend of a single animal before (baseline), during (training), and after (post training) exercise. (B) Median RR interval before (baseline) and during training (training) and after full recovery (recovered), shown as mean ± SEM, unpaired Student's t-test, ***p &lt; 0.001. (C) SDRR before (baseline) and during training (training) and after full recovery (recovered), n = 3, shown as mean ± SEM. <a href="https://www.jove.com/files/ftp_upload/63873/63873fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">5-day acclimatization phase</td>
		</tr>
		<tr>
			<td>Day</td>
			<td>Speed (cm/sec)</td>
			<td>Time(min)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>16.7</td>
			<td>10</td>
		</tr>
		<tr>
			<td>2</td>
			<td>18.3</td>
			<td>20</td>
		</tr>
		<tr>
			<td>3</td>
			<td>20</td>
			<td>30</td>
		</tr>
		<tr>
			<td>4</td>
			<td>21.7</td>
			<td>40</td>
		</tr>
		<tr>
			<td>5</td>
			<td>23.3</td>
			<td>50</td>
		</tr>
		<tr>
			<td colspan="3">Remark: 2 min rest intervals after every 15 min</td>
		</tr>
	</tbody>
</table>
Table 1: Training regime during acclimatization phase.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="3">5-day training phase</td>
		</tr>
		<tr>
			<td>Day</td>
			<td>Speed (cm/sec)</td>
			<td>Time(min)</td>
		</tr>
		<tr>
			<td>1</td>
			<td>25.0</td>
			<td>60</td>
		</tr>
		<tr>
			<td>2</td>
			<td>25.0</td>
			<td>60</td>
		</tr>
		<tr>
			<td>3</td>
			<td>25.0</td>
			<td>60</td>
		</tr>
		<tr>
			<td>4</td>
			<td>25.0</td>
			<td>60</td>
		</tr>
		<tr>
			<td>5</td>
			<td>25.0</td>
			<td>60</td>
		</tr>
		<tr>
			<td colspan="3">Remark: 2 min rest intervals after every 15 min</td>
		</tr>
	</tbody>
</table>
Table 2: Training regime during training phase.

Watch this Scientific Journal Video about Real-Time Electrocardiogram Monitoring During Treadmill Training in Mice at JoVE.com

Real-Time Electrocardiogram Monitoring During Treadmill Training in Mice

Electrocardiogram (ECG) is the key variable to understanding cardiac electrophysiology. Physical exercise has beneficial effects but may also be harmful in the context of cardiovascular diseases. This manuscript provides a method of recording real-time ECG during exercise, which can serve to investigate its effects on cardiac electrophysiology in mice.

Regular physical exercise is a major contributor to cardiovascular health, influencing various metabolic as well as electrophysiological processes. ...

real-time-electrocardiogram-monitoring-during-treadmill-training

Universidad Científica del Sur

Research

JoVE Journal

Medicine

2.3K Views.  Ludwig-Maximilians University Munich (LMU). Electrocardiogram (ECG) is the key variable to understanding cardiac electrophysiology. Physical exercise has beneficial effects but may also be harmful in the context of cardiovascular diseases. This manuscript provides a method of recording real-time ECG during exercise, which can serve to investigate its effects on cardiac electrophysiology in mice.

Video: Real-Time Electrocardiogram Monitoring During Treadmill Training in Mice

Programmed Electrical Stimulation in Mice

Genetically-modified mice have emerged as a preferable animal model to study the molecular mechanisms underlying conduction abnormalities, atrial and ventricular arrhythmias, and sudden cardiac death.1 Intracardiac pacing studies can be performed in mice using a 1.1F octapolar catheter inserted into the jugular vein, and advanced into the right atrium and ventricle. Here, we illustrate the steps involved in performing programmed electrical stimulation in mice. Surface ECG and intracardiac electrograms are recorded simultaneously in the atria, atrioventricular junction, and ventricular myocardium, whereas intracardiac pacing of the atrium is performed using an external stimulator. Thus, programmed electrical stimulation in mice provides unique opportunities to explore molecular mechanisms underlying conduction defects and cardiac arrhythmias.

Programmed electrical stimulation provides the ability to determine conduction properties of the heart, and the possibility to induce and terminate cardiac arrhythmias using various pacing protocols. Using a transvenous catheter, intracardiac electrogram recordings can be obtained in mice following programmed electrical stimulation protocols to identify arrhythmogenic substrates.

Genetically-modified mice have emerged as a preferable animal model to study the molecular mechanisms underlying conduction abnormalities, atrial and ...

Transthoracic Echocardiography in Mice

In recent years, murine models have become the primary avenue for studying the molecular mechanisms of cardiac dysfunction resulting from changes in gene expression. Transgenic and gene targeting methods can be used to generate mice with altered cardiac size and function,1-3 and as a result, in vivo techniques are needed to evaluate their cardiac phenotype. Transthoracic echocardiography, pulse wave Doppler (PWD), and tissue Doppler imaging (TDI) can be used to provide dimensional measurements of the mouse heart and to quantify the degree of cardiac systolic and diastolic performance. Two-dimensional imaging is used to detect abnormal anatomy or movements of the left ventricle, whereas M-mode echo is used for quantification of cardiac dimensions and contractility.4,5 In addition, PWD is used to quantify localized velocity of turbulent flow,6 whereas TDI is used to measure the velocity of myocardial motion.7 Thus, transthoracic echocardiography offers a comprehensive method for the noninvasive evaluation of cardiac function in mice.

Transthoracic echocardiography offers a noninvasive method for the evaluation of cardiac function in mice. A combination of ultrasound and Doppler imaging modalities can be used to obtain dimensional measurements of the heart and intracardiac blood flow, which together provide an assessment of cardiac systolic and diastolic performance.

In recent years, murine models have become the primary avenue for studying the molecular mechanisms of cardiac dysfunction resulting from changes in gene ...

Simultaneous Video-EEG-ECG Monitoring to Identify Neurocardiac Dysfunction in Mouse Models of Epilepsy

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can potentially increase risk of sudden unexpected death in epilepsy (SUDEP). Electroencephalography (EEG) and electrocardiography (ECG) are widely used clinical diagnostic tools to monitor for abnormal brain and cardiac rhythms in patients. Here, a technique to simultaneously record video, EEG, and ECG in mice to measure behavior, brain, and cardiac activities, respectively, is described. The technique described herein utilizes a tethered (i.e., wired) recording configuration in which the implanted electrode on the head of the mouse is hard-wired to the recording equipment. Compared to wireless telemetry recording systems, the tethered arrangement possesses several technical advantages such as a greater possible number of channels for recording EEG or other biopotentials; lower electrode costs; and greater frequency bandwidth (i.e., sampling rate) of recordings. The basics of this technique can also be easily modified to accommodate recording other biosignals, such as electromyography (EMG) or plethysmography for assessment of muscle and respiratory activity, respectively. In addition to describing how to perform the EEG-ECG recordings, we also detail methods to quantify the resulting data for seizures, EEG spectral power, cardiac function, and heart rate variability, which we demonstrate in an example experiment using a mouse with epilepsy due to Kcna1 gene deletion. Video-EEG-ECG monitoring in mouse models of epilepsy or other neurological disease provides a powerful tool to identify dysfunction at the level of the brain, heart, or brain-heart interactions.

Here, we present a protocol to record brain and heart bio signals in mice using simultaneous video, electroencephalography (EEG), and electrocardiography (ECG). We also describe methods to analyze the resulting EEG-ECG recordings for seizures, EEG spectral power, cardiac function, and heart rate variability.

In epilepsy, seizures can evoke cardiac rhythm disturbances such as heart rate changes, conduction blocks, asystoles, and arrhythmias, which can ...

Genetics

Implantation of Radiotelemetry Transmitters Yielding Data on ECG, Heart Rate, Core Body Temperature and Activity in Free-moving Laboratory Mice

The laboratory mouse is the animal species of choice for most biomedical research, in both the academic sphere and the pharmaceutical industry. Mice are a manageable size and relatively easy to house. These factors, together with the availability of a wealth of spontaneous and experimentally induced mutants, make laboratory mice ideally suited to a wide variety of research areas.
In cardiovascular, pharmacological and toxicological research, accurate measurement of parameters relating to the circulatory system of laboratory animals is often required. Determination of heart rate, heart rate variability, and duration of PQ and QT intervals are based on electrocardiogram (ECG) recordings. However, obtaining reliable ECG curves as well as physiological data such as core body temperature in mice can be difficult using conventional measurement techniques, which require connecting sensors and lead wires to a restrained, tethered, or even anaesthetized animal. Data obtained in this fashion must be interpreted with caution, as it is well known that restraining and anesthesia can have a major artifactual influence on physiological parameters1, 2.
Radiotelemetry enables data to be collected from conscious and untethered animals. Measurements can be conducted even in freely moving animals, and without requiring the investigator to be in the proximity of the animal. Thus, known sources of artifacts are avoided, and accurate and reliable measurements are assured. This methodology also reduces interanimal variability, thus reducing the number of animals used, rendering this technology the most humane method of monitoring physiological parameters in laboratory animals3, 4. Constant advancements in data acquisition technology and implant miniaturization mean that it is now possible to record physiological parameters and locomotor activity continuously and in realtime over longer periods such as hours, days or even weeks3, 5.
Here, we describe a surgical technique for implantation of a commercially available telemetry transmitter used for continuous measurements of core body temperature, locomotor activity and biopotential (i.e. onelead ECG), from which heart rate, heart rate variability, and PQ and QT intervals can be established in freeroaming, untethered mice. We also present pre-operative procedures and protocols for post-operative intensive care and pain treatment that improve recovery, well-being and survival rates in implanted mice5, 6.

A surgical technique for implantation of commercially available telemetry transmitters used for continuous measurement of biopotential (one-lead ECG), heart rate, core body temperature and locomotor activity in freely moving mice is shown. Recommendations and protocols for post-operative care and pain relief, improving recovery, well being and survival rate are also presented.

The laboratory mouse is the animal species of choice for most biomedical research, in both the academic sphere and the pharmaceutical industry. Mice are a ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

Biology

Wireless Intramuscular EMG System for Locomotion Studies in Stroke Model Rats

A Real-Time Wearable Electromyography Measurement System for Small Animals

The intramuscular electromyography (EMG) measurement method for experimental animals has been implemented in various ways. Among these methods, tethering cables to external measurement devices can restrict the movement of experimental animals, while implantable devices may cause unwanted side effects due to the constant presence of a device with considerable size and weight. To address these issues, we propose a low-cost, wireless, detachable EMG measurement system and experimental procedure. This article focuses on the surgical installation of intramuscular wire electrodes with small connectors and the development of the wireless system. Notably, in this system, only the wire electrodes are inserted into the animal's body. Using this system, EMG measurements can be easily performed by attaching the circuit system to a connector installed on the animal's back, with real-time monitoring achievable on a laptop. The proposed method is explained in a detailed, step-by-step manner, followed by a demonstration involving the insertion of intramuscular electrodes into the hindlimbs of a rat. A treadmill experiment is conducted for a locomotion study, and the resulting electrophysiological signals are subsequently obtained and analyzed.

A protocol is presented for the installation and preparation of intramuscular electrodes, along with an animal-mountable, miniaturized measurement device for electromyography (EMG) analysis in locomotion studies. This system enables the wireless transmission of real-time EMG data for investigating locomotion recovery in stroke model rats.

The intramuscular electromyography (EMG) measurement method for experimental animals has been implemented in various ways. Among these methods, tethering ...

Engineering

Confirmation of Myocardial Ischemia and Reperfusion Injury in Mice Using Surface Pad Electrocardiography

Many animal models have been established for the study of myocardial remodeling and heart failure due to its status as the number one cause of mortality worldwide. In humans, a pathologic occlusion forms in a coronary artery and reperfusion of that occluded artery is considered essential to maintain viability of the myocardium at risk. Although essential for myocardial recovery, reperfusion of the ischemic myocardium creates its own tissue injury. The physiologic response and healing of an ischemia/reperfusion injury is different from a chronic occlusion injury. Myocardial ischemia/reperfusion injury is gaining recognition as a clinically relevant model for myocardial infarction studies. For this reason, parallel animal models of ischemia/reperfusion are vital in advancing the knowledge base regarding myocardial injury. Typically, ischemia of the mouse heart after left anterior descending (LAD) coronary artery occlusion is confirmed by visible pallor of the myocardium below the occlusion (ligature). However, this offers only a subjective way of confirming correct or consistent ligature placement, as there are multiple major arteries that could cause pallor in different myocardial regions. A method of recording electrocardiographic changes to assess correct ligature placement and resultant ischemia as well as reperfusion, to supplement observed myocardial pallor, would help yield consistent infarct sizes in mouse models. In turn, this would help decrease the number of mice used. Additionally, electrocardiographic changes can continue to be recorded non-invasively in a time-dependent fashion after the surgery. This article will demonstrate a method of electrocardiographically confirming myocardial ischemia and reperfusion in real time.

During murine myocardial ischemia/reperfusion surgery, correct placement of the occluding ligature is typically confirmed by visible observation of myocardial pallor. Herein, a method of electrocardiographically confirming ischemia and reperfusion, to supplement observed myocardial pallor, is demonstrated in male C57Bl/6 mice.

Many animal models have been established for the study of myocardial remodeling and heart failure due to its status as the number one cause of mortality ...

Noninvasive Electrocardiography in the Perinatal Mouse

Electrocardiography (ECG) has long been relied upon as an effective and reliable method of assessing cardiovascular (and cardiopulmonary) function in both human and animal models of disease. Individual heart rate, rhythm, and regularity, combined with quantitative parameters collected from ECG, serve to assess the integrity of the cardiac conduction system as well as the integrated physiology of the cardiac cycle. This article provides a comprehensive description of the methods and techniques used to perform a noninvasive ECG on perinatal and neonatal mouse pups as early as the first postnatal day, without requiring the use of anesthetics. This protocol was designed to directly address a need for a standardized and repeatable method for obtaining ECG in newborn mice. From a translational perspective, this protocol proves to be entirely effective for characterization of congenital cardiopulmonary defects generated using transgenic mouse lines, and particularly for analysis of defects causing lethality at or during the first postnatal days. This protocol also aims to directly address a gap in the scientific literature to characterize and provide normative data associated with maturation of the early postnatal cardiac conduction system. This method is not limited to a specific postnatal timepoint, but rather allows for ECG data collection in neonatal mouse pups from birth to postnatal day 10 (P10), a window that is of critical importance for modeling human diseases in vivo, with particular emphasis on congenital heart disease (CHD).

Here, we present a noninvasive electrocardiography (ECG) protocol, optimized for early postnatal mice, that does not require the use of anesthetics.

Electrocardiography (ECG) has long been relied upon as an effective and reliable method of assessing cardiovascular (and cardiopulmonary) function in both ...

Electrocardiogram Recordings in Anesthetized Mice using Lead II

The electrocardiogram is a valuable tool for evaluating the cardiac conduction system. Animal research has helped generate novel genetic and pharmacological information regarding the electrocardiogram. However, making electrocardiogram measurements in small animals in vivo, such as mice, has been challenging. To this end, we used an electrocardiogram recording method in anesthetized mice with many advantages: it is a technically simple procedure, is inexpensive, has short measuring time, and is affordable, even in young mice. Despite the limitations with using anesthesia, comparisons between control and experimental groups can be performed with enhanced sensitivity. We treated mice with agonists and antagonists of the autonomic nervous system to determine the validity of this protocol and compared our results with previous reports. Our ECG protocol detected increased heart rates and QTc intervals on treatment with atropine, decreased heart rates and QTc intervals after carbachol treatment, and higher heart rates and QTc intervals with isoprenaline but did not note any change in ECG parameters on administration of propranolol. These results are supported by previous reports, confirming the reliability of this ECG protocol. Thus, this method can be used as a screening approach to making ECG measurements that otherwise would not be attempted due to high cost and technical difficulties.

We present an ECG protocol that is technically easy, inexpensive, fast, and affordable in small mice, and can be performed with enhanced sensitivity. We suggest this method as a screening approach for studying pharmacological agents, genetic modifications, and disease models in mice.

The electrocardiogram is a valuable tool for evaluating the cardiac conduction system. Animal research has helped generate novel genetic and ...

Revealing Subtle Changes in Cardiac Function using Transthoracic Dobutamine Stress Echocardiography in Mice

Left ventricular (LV) dysfunction paves the final pathway for a multitude of cardiac disorders. With the non-invasive high-frequency transthoracic dobutamine stress echocardiography in humans, a reductionist investigation approach to unmask subtle changes in cardiac function has become possible. Here, we provide a protocol for using this technique in mice to facilitate expanded analysis of LV architecture and function in physiology and pathology enabling the observation of alterations in models of cardiac disease hidden in unstressed hearts. This investigation can be performed in one and the same animal and allows both, basal and pharmacologically stress-induced measurements. We outline detailed criteria for appropriate anesthesia, imaging-based LV analysis, consideration of intra- and interobserver variability, and obtaining positive inotrope response that can be attained in mice after intraperitoneal injection of dobutamine under near physiological conditions. To recapitulate the characteristics of human physiology and disease in small animal models, we highlight critical pitfalls in evaluation, e.g., a pronounced Bowditch effect in mice. To further meet translational objectives, we compare stress-induced effects in humans and mice. When used in translational studies, attention must be paid to physiological differences between mice and human. Experimental rigor dictates that some parameters assessed in patients can only be used with caution due to restrictions in spatial and temporal resolution in mouse models.

Left ventricular dysfunction constitutes the final common pathway for a host of cardiac disorders. We here provide a detailed protocol of transthoracic dobutamine stress echocardiography approach for comprehensive evaluation of the left ventricular function of mouse models of cardiac disease as well as cardiac phenotyping.