Name: noninvasive electrocardiography in the perinatal mouse
Uploaded: 2020-06-12T13:30:00.000+00:00
Duration: 4 min 36 s
Description: Watch this Scientific Journal Video about Noninvasive Electrocardiography in the Perinatal Mouse at JoVE.com

The authors acknowledge generous support from the Saving tiny Hearts Society (KLT), the UNE COBRE Program (NIGMS grant number P20GM103643; LAF), and the SURE Fellowship Program at the University of New England (VLB), as well as patient technical support from Ashish More (iWorx, Dover, NH). Figure 3, Figure 4, and Figure S1 were created with Biorender software.

The following protocol follows the standards of the Institutional Animal Care and Use Committee of the University of New England. Close observation of the protocol should deliver satisfactory ECG reads in all examined neonates (n &#62; 70).
1. Device preparations
<ol>
	<li>Plug the device into the USB port of a computer with the ECG software downloaded on it. The measuring device will automatically begin heating up to (37 &#176;C/98.6 &#176;F). The internal heating unit is contained within the measuring unit and heats only the plastic surface. The silver wire electrodes are not heated.</li>
	<li>Allow approximately 15 min for the surface to reach the temperature. Use this time to gather and set up animals. 
	NOTE: The protocol may be paused at this point and the platform can remain plugged in and heating for an extended period of time. In the absence of a self-heating electrode platform, an animal safe heating pad may also be used to keep mother and pups from becoming hypothermic.</li>
</ol>
2. Animal preparations
<ol>
	<li>Collect the mother and pups and keep within the housing cage until ready to collect.</li>
	<li>Once the measuring unit has heated to the temperature, remove the mouse pup from the cage and wipe the thorax with 70% ethanol sprayed on a wipe. Place the pup on the heated surface of the plastic.</li>
	<li>Allow the mouse to acclimate to the surface in the dark for approximately 2-5 minutes.</li>
</ol>
3. Mouse and electrode platform setup (electrode application)
<ol>
	<li>Use a metal spatula, probe, or wooden dowel to collect a small droplet of adhesive, electrical conducting gel (a quick-drying high-conductivity electrode gel commonly used for placement of rodent electrodes). 
	NOTE: Any nonfibrous, solid object can be used to apply the conducting gel, as long as the object will not leave behind synthetic fibers or similar material on the electrodes that could interfere with the quality of the electrical signal.</li>
	<li>Using the spatula/dowel, gently touch the top of each of the four, flattened electrode surfaces by pressing gently down and pulling the conducting gel at an oblique angle away from the center of the electrode construct. Make sure that each individual electrode is completely covered with the gel. 
	CAUTION: This step is extremely important to ensure that the conductive, electrode gel does not adhere to more than a single electrode. Adhesive strands that form between electrodes can conduct charge and potentially interfere with or short out the desired electrical signal. The protocol should not be paused at this time as the gel will begin to solidify and become adherent. Make sure to set up the mouse to the platform within 5-10 minutes of applying conducting gel (or equivalent conductive electrode gel substitution).</li>
	<li>Place the metal spatula or wooden dowel with the remainder of the gel to the side.</li>
	<li>Place the neonatal mouse pup sternum down and prone with the head of the pup facing the outgoing USB edge of the platform. Make sure that a portion of the pup&#8217;s chest is covering each of the four electrodes. Gently restrain the pup&#8217;s forearms by their side while simultaneously holding down for approximately 1 min to allow the conducting gel to set.</li>
	<li>Place rubber silicone bumpers on the right and left sides of the pup. Bumpers should secure the pup on each side and provide stability to prevent excessive movements of the mouse but should NOT prevent all movement of the mouse. Once installed, watch the mouse for a moment and adjust bumper placement as needed. 
	CAUTION: Do not compress the mouse too tightly as this can interfere with respiratory mechanics and respiratory rate.</li>
	<li>Use the dowel that was set aside to apply remaining conducting gel to the grounding tail electrode and place on the rump of the pup. Apply gentle pressure to allow the gel to set before releasing the pup.</li>
	<li>Place the final silicon bumper on top of the rump of the mouse to hold the grounding electrode in place. 
	CAUTION: Do not apply excessive force while placing the final bumper as this could cause discomfort to the pup and/or displace the grounding electrode.</li>
	<li>Grab ahold of the entire platform and gently place inside the Faraday cage. 
	CAUTION: Use caution and ensure the top silicone bumper does not become displaced once the Faraday cage is in place.</li>
	<li>Prior to recording, make sure the mouse pup is not moving excessively and make sure the body and head of the mouse appears secure. 
	CAUTION: Make sure the mouse pup&#8217;s head is able to move somewhat freely within the bumpers and is not completely snout down into the platform. The raised platform is designed to elevate the mouse thorax slightly and prevent suffocation, but this should be closely monitored.</li>
</ol>

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The authors report no conflicts of interest.

The data points collected in perinatal day 1 mouse pups are slightly below the average expected values for adult mice (500-700 beats per minute).8 There is an increase in heart rate as the mouse ages, which falls more in line for the expected values (Table 1). However, it is important to emphasize that neonatal values were on the lower end of this range, supporting the idea that normative values should be documented in an age-specific manner. This method is different than other electrocardiogram protocols in that there is no physical trauma to the mouse. The protocol is totally noninvasive, does not require the use of anesthesia, and is optimal for mice immediately after birth. No other electrocardiogram device allows for pups this young to be analyzed in this manner9,10,11. This protocol aims to establish a reliable reference method to generate normative data specific to the neonatal mouse population, but applicable to human pediatric populations.
When performing an electrocardiogram on such a small animal, it is important to be cautious with all steps. However, there are a few key steps that can change the quality of the results. The first is applying the conductive gel. If there is too much gel, there will be a higher chance for the electrodes to connect and short. If there is not enough gel, there will not be a secure connection. The best method to apply the gel is to approach the electrode from the outside corner and roll the gel over the top of the electrode. It is very important that extreme caution is taken to assure there are no threads between electrodes, which would interfere with the presence and/or quality of the electrical activity. It may be useful to take a thin tool (e.g., forceps), and run it between the electrodes to collect any stray threads which may not be apparently visible. While not formally required as part of the protocol, this extra step could serve as an additional precaution to ensure optimal conduction and minimal noise.
If the presence of noise of static causes the ECG to be unreadable (Figure 2), it may be useful to remove all electronic devices from the immediately (table-top) vicinity. This is especially helpful if any of the electronic devices present nearby are moving, as this movement can be picked up by the ECG recording device12. It is also important not to introduce any outside movements during data collection. Outside movements that could interfere with the quality of the ECG could include setting objects down on the same nearby surface, and must be avoided until after the reading is complete. In addition to outside devices, very active mouse pups may also cause electrical interference associated with excessive body movements. The likelihood of this type of musculoskeletal interference increases as the pups mature, which should be considered when selecting ages for data collection. In the event that the pup shifts from the electrodes in a way that significantly compromises the quality of the ECG reading, repositioning of the pup should be considered. Repositioning the mouse before opting to reapply the electrode gel can provide improved results in most cases and save additional time and reagents. Before repositioning the pup, select the pause button in the software. Pausing the run will stop active recording of the ECG but will continue to track time. Of note, when the recording is resumed, the ECG will appear at a later time than paused at. Slide the device platform out from the Faraday age with the mouse still positioned between the bumpers. Remove the bumpers surrounding the mouse, and gently lift the pup off of the electrodes. Reposition pup on the electrodes following the same protocol of gently holding the mouse in place for 1 min for gel to adhere (step 3.4-3.5). Try to reposition the mouse so that the electrodes are on the thorax between the upper limbs (Figure 3). While designed as an ideal, noninvasive method for collecting ECG in neonatal mice, one limitation associated with this protocol would be the increased mobility associated with data collection on an un-anesthetized mouse, as the mouse may also move and shift on the device which will affect the quality of the reading. While movement may be limited with positioning of silicone bumpers, this cannot be prevented without the use of sedation or anesthesia.
In the situation in which an ECG recording comes with heavy interference (Figure 2) despite having minimized all electrical interference, the next step that should be taken is to reposition the external wiring connecting the recording platform to the Faraday cage. It is very important that the external wiring remains properly connected to the recording platform during data acquisition. If external wiring is repositioned, be sure to reattach this wiring carefully at both ends, until a clearer recording can be obtained. If the use of the Faraday cage provided with the device is not suitable, the device can be used in other Faraday cages.
If the recording is not clear or the mouse has moved from the electrodes, remove the mouse from the device and clean the electrodes by taking forceps and removing all the conductive gel. Because the conducting gel is water-soluble, one can also use warm water to gently remove excess gel from the skin of the pup. Reapply the gel and reposition the pup.
To obtain the best results make sure the device is properly cleaned before and after each use. The gel does dry and can be removed using forceps to pull it from the device, but the gel is water soluble, so a damp cloth can be used to clean the electrodes of the recording platform.
Older mice have been more active in the recording process, so it is important to closely monitor them as they often move from the electrodes and can even move off the device platform. While a clear read may not happen right away, with troubleshooting and repositioning, there has been success in getting usable recordings with this device (Figure 1). Active mice may need to be returned to their mother and reanalyzed after a break. They can also be held in the palm of a hand and gently covered to provide heat and darkness until the pup settles down.
This device is designed to collect ECG data on mouse pups from the age of birth to P10 (Figure 4). Pups older than P10 will likely not be able to fit into the device with the Faraday cage, an essential component to maximizing signal to noise ratios. Even at P10, positioning adjustments will likely need to be made to accommodate a larger body size into the device. Use extreme caution when moving the device into and out of the Faraday cage. Removal of the top bumper will allow the mouse to lay on the electrode platform with the surrounding Faraday cage. Given that the mice at this age are more active, they are more apt to move off the electrodes without the stabilization of the top bumper. The top bumper can also be placed in front of the pup to help discourage the pup moving off the device.
The novelty of this device and corresponding protocol include optimization for use immediately after birth, the ability of the system to accommodate a broader age range (P1-P10) and the need addressed by this method to expand the translational applications of in vivo research methods in the field of cardiovascular physiology and beyond. Although sophisticated devices utilizing echocardiography to quantify cardiac cycles in neonate mice are available13, one great advantage of this protocol is that it allows for a relatively simple and affordable means to address basic electrophysiological parameters, which is very attractive in the current parlous scientific funding environment.

Cardiac function can be measured in different ways, the most common of which includes the use of electrocardiography (ECG) to analyze the conduction of electric current through the heart as well as its overall cardiac cycle and function1. Electrocardiography continues to be a useful diagnostic tool for identifying and characterizing cardiac anomalies in both human and animal models of disease1,2. Irregularities in an electrocardiogram reading can be found in abnormal cardiac development (i.e., congenital heart disease (CHD)), and can include arrhythmias manifesting as changes in heart rate (e.g., bradycardia), and rhythm (e.g., &#8220;heart blocks&#8221;), suggestive of defects in the integrity and/or function of the underlying myocardium. Changes such as these may predispose patients to life-threatening cardiac dysfunction (e.g., congestive heart failure and/or cardiac arrest) and increased mortality3,4. Given the high rates of mortality with severe and untreated CHD, developing a standardized and repeatable method for collecting ECG during this early postnatal period is critical.
Although we are not the first to address this problem, previous methods of collecting ECG on a mouse pups have traditionally included invasive procedures (subcutaneous needle or wire electrodes) and/or the use of anesthetics5,6,7. Advantages of performing noninvasive ECG analysis include minimizing pain and undo stress on the animal. While the experimenter must still be cautious about causing the pup stress, the device is designed to avoid common stressors in order to produce accurate data. In the context of evaluating cardiac function, introducing anesthesia to animals that may have cardiopulmonary abnormalities could potentially mask or even exacerbate underlying conditions. Anesthetics may affect the electrical conduction by altering depolarization and/or repolarization of the cells. Finally, the use of anesthesia can put the newborn pup at an increased risk for hypothermia, which could further confound any inherent pathology. The following protocol does not introduce any anesthetics, invasive procedures, or pronounced discomfort to the pup. Once equipment setup is finalized, device setup and data collection involving the animal can be completed efficiently, after which the pups can be returned to their mother. Additionally, this system allows for repeat and/or serial analyses to be performed, which is ideal for experiments requiring analysis over time, introduction of pharmacological therapies, etc.

<table><tbody><tr><td>LabScribe4</td><td>iWorx</td><td>LabScribe4</td><td>Software used to record ECG</td></tr><tr><td>Neonatal Mouse ECG &amp;amp; Respiration System</td><td>iWorx</td><td>RS-NMECG : Neonatal Mouse ECG</td><td>ECG device</td></tr><tr><td>Tensive Conductive Adhesive Gel</td><td>Parker Laboratories, Inc</td><td>22-60</td><td>Tac-gel used as conductive gel for ECG</td></tr></tbody></table>

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).

An ideal ECG would have a clear, prominent signal that allows all waves to be analyzed in several different time frames (Figure 1). The laboratory initially employed a custom application of an electromyography apparatus to produce ECGs of an unsatisfactory quality, which only allowed us to analyze basic parameters such as heart rate (Figure S1). This inspired work with a company to develop a novel prototype ECG device specifically for the analysis of early postnatal mouse pups.
A poor-quality reading has no discernable beats, shows clear interference, and has waves or inconsistency across the reading (Figure 2). To achieve the highest quality ECG, follow instructions carefully. Use caution with application of conducting gel, as it is moderately adhesive, and may require additional time to allow the mouse to accommodate to the device. By doing this, it lowers the risk of the mouse moving, there being a shorting out of electrodes, and for correct use of the device. Mice should be placed on the device so the head is facing the cords that connect the device to the USB-port and in a prone position (Figure 3). The mouse should be secured by rubber bumpers to hold them securely in place, with two on the side and one on the top (Figure 3). These bumpers should secure the mouse, but should not inhibit the mouse from moving its head. The layout of the mouse is important for the reading, as the leads are stationary. The leads are set up so that the front two electrodes are Lead I (Figure 3). The rear two electrodes are Leads II and III, with the ground electrode being on the rump of the pup (Figure 3). Setting the mouse up in this way will allow for better results.
The program used allows for the analysis of the ECG in the program. This provides analysis of key aspects including heart rate, R-R intervals, QRS complex interval, QT interval, and PR interval. Given this ability, it was possible to establish a data set of normative values for a perinatal mouse (Table 1). These normative results were based off mice who were analyzed within one day after birth. It was found that an average heartbeat was 357.2 beats per minute (bpm). The average R-R, QRS, QT, and PR intervals were 169.1, 16.9, 45.4, and 36.3 milliseconds (ms), respectively (Table 1). Importantly, the setup can be used to analyze ECG patterns from neonatal mice suffering from congenital heart defects (Figure S2).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Pup Age</td>
			<td>Ave/STDEV</td>
			<td>Heart Rate (bpm)</td>
			<td>R-R Interval (ms)</td>
			<td>PR Duration (ms)</td>
			<td>QRS Duration (ms)</td>
			<td>QT Duration (ms)</td>
			<td>ST Duration (ms)</td>
			<td>T Duration (ms)</td>
			<td>P Duration (ms)</td>
		</tr>
		<tr>
			<td rowspan="2">P1</td>
			<td>Averages</td>
			<td>357.2</td>
			<td>169.1</td>
			<td>36.3</td>
			<td>16.9</td>
			<td>45.4</td>
			<td>16.4</td>
			<td>18</td>
			<td>12.8</td>
		</tr>
		<tr>
			<td>Standard Deviation</td>
			<td>36.3</td>
			<td>20</td>
			<td>10.9</td>
			<td>5.8</td>
			<td>16</td>
			<td>7.4</td>
			<td>7.2</td>
			<td>3.1</td>
		</tr>
		<tr>
			<td rowspan="2">P3</td>
			<td>Averages</td>
			<td>412.4</td>
			<td>149.2</td>
			<td>46.4</td>
			<td>14.5</td>
			<td>53</td>
			<td>22.3</td>
			<td>16.2</td>
			<td>14.8</td>
		</tr>
		<tr>
			<td>Standard Deviation</td>
			<td>55.4</td>
			<td>21.4</td>
			<td>6.8</td>
			<td>11</td>
			<td>12.2</td>
			<td>6.9</td>
			<td>4.6</td>
			<td>3.1</td>
		</tr>
		<tr>
			<td rowspan="2">P5</td>
			<td>Averages</td>
			<td>505.5</td>
			<td>119.2</td>
			<td>46.7</td>
			<td>11.7</td>
			<td>51.3</td>
			<td>20.8</td>
			<td>18.8</td>
			<td>14.2</td>
		</tr>
		<tr>
			<td>Standard Deviation</td>
			<td>19.2</td>
			<td>4.6</td>
			<td>13.3</td>
			<td>5.8</td>
			<td>8.1</td>
			<td>11.4</td>
			<td>4.6</td>
			<td>2.3</td>
		</tr>
		<tr>
			<td rowspan="2">P7</td>
			<td>Averages</td>
			<td>555.3</td>
			<td>108.7</td>
			<td>40</td>
			<td>9.5</td>
			<td>43.6</td>
			<td>20.3</td>
			<td>13.7</td>
			<td>14</td>
		</tr>
		<tr>
			<td>Standard Deviation</td>
			<td>34.2</td>
			<td>7</td>
			<td>2.5</td>
			<td>0.6</td>
			<td>6</td>
			<td>7.1</td>
			<td>3.2</td>
			<td>2.7</td>
		</tr>
	</tbody>
</table>
Table 1: Representative results of ECG measurements for the average perinatal mouse pup P1, P3, P5, and P7.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61074/61074fig01.jpg" /> 
Figure 1: Representative electrocardiographic reads from neonatal mice on the first (A, P1.0), third (B, P3.0), and seventh (C, P7.0) postnatal day. 
(A-C) Images represent examples of good quality ECG tracings using the 2-lead, noninvasive device, captured in a 1.5 s frame of the reading. Notable characteristics of good ECG reads include clear, discernable beats, as described collectively by the presence of consistent P-waves followed by a QRS complex and subsequent T-waves, visible in both Leads I-II of each postnatal time point. Examples also include a low signal-to-noise ratio (minimal artifact) and a discernable isoelectric line. Top ECG strip (red): Lead I; bottom ECG strip (green): Lead II. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61074/61074fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61074/61074fig02.jpg" /> 
Figure 2: Representative ECG read with complications. 
This image is representative of a poor-quality ECG reading using the 2-lead, noninvasive device on the first postnatal day (P1.0). The above images were captured in a 1.5 s reading frame. Poor quality ECG tracings are characterized by the absence of discernable beats (and specific cardiac cycle waveforms), along with pronounced artifact (high signal:noise ratio), and notable inconsistencies between Leads I and II from a given mouse pup. To improve this ECG, both the device and the silicone bumpers securing the pup would require repositioning within the Faraday cage. To minimize electromagnetic interference, removal of all moving devices near the apparatus would need to be carried out. The final troubleshooting measure would involve repositioning of the mouse pup on the device electrodes and/or more conductive gel would need to be (re-)applied. Top ECG strip (red): Lead I; bottom ECG strip (green): Lead II. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61074/61074fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61074/61074fig03.jpg" /> 
Figure 3: Placement of the mouse pup and limb lead electrodes for collection of early postnatal ECG. 
(A) Left: Anterior perspective of mouse placement on electrode platform within the Faraday cage (black). Right: Lateral view illustrating proper mouse placement on top of raised electrodes/platform; supportive silicone bumpers (not pictured) are placed to either side and across the top of the mouse pup within the Faraday cage.&#160;(B) Bipolar limb leads and electrode placement on the neonatal mouse. Illustration depicts the point of contact for each raised electrode on the ventral thoracic surface of the mouse pup. (B,C) Electrode placement, chest lead directionality, and (C) corresponding, representative ECG tracings from a neonatal mouse pup at P1.0 (Lead I (red); Lead II (green)). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61074/61074fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61074/61074fig04.jpg" /> 
Figure 4: Representative ECG tracings of neonatal mice at multiple postnatal time points. 
Representative ECG reads (top 2 traces) and illustrated cardiac cycles (bottom row) from neonatal mouse pups on the first (A, P1.0), third (B, P3.0), and seventh (C, P7.0) postnatal day. Each image represents an exemplary ECG tracing using the 2-lead, noninvasive device, captured in a 1.5 second frame of the reading (A-C, Lead I (top/red); Lead II (bottom/green)). While individual waveforms do appear to undergo morphological changes with increasing age, notable and consistent characteristics include clear, discernable beats, as described collectively by the presence of consistent P-waves followed by a QRS complex and subsequent T-waves, visible in both Leads I-II of each postnatal time point. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61074/61074fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Figure S1: Illustration of traditional limb lead electrodes for noninvasive collection of early postnatal ECG. (A, left) Lateral view of mouse and electrode placement within Faraday cage (box). (B) Traditional self-stick skin electrodes are positioned on the dorsal surface of the pup. (A, right) ECG-like signal may be interpreted with the use of traditional electromyography transducer to produce a minimalistic ECG tracing discernable only in Lead II (C, bottom). (B-C) Electrode placement, chest lead directionality, and corresponding, representative ECG tracing from a neonatal mouse pup at P1.0 (Lead II; purple). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61074/61074supfig01.jpg" target="_blank">Please click here to download this figure.</a>
Figure S2: Comparative electrocardiographic reads from littermate control pups and mutant pups with congenital heart disease on the first postnatal day (P1.0). (A,B) Images represent examples of good quality ECG tracings from healthy neonatal pups (A, CONTROL) compared to pups born with CHD (B, CHD MUT) at P1.0. The 2-lead, noninvasive device was used to capture ECG tracings at 10.0 (A,B, top) and 1.5-second intervals (A, B, bottom). Noticeable differences in heart rate are apparent in the CHD MUT (B), as visualized by the decreased number of cardiac cycles (complexes) visible in the given time frame. Comparison also reveals irregularities in the general morphology of QRS waveforms, frequency, and overall regularity of cardiac cycles in the CHD MUT (B) when compared to the control (A). Lead I (red); Lead II (green)). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61074/61074supfig02.jpg" target="_blank">Please click here to download this figure.</a>

Watch this Scientific Journal Video about Noninvasive Electrocardiography in the Perinatal Mouse at JoVE.com

Noninvasive Electrocardiography in the Perinatal Mouse

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 ...

noninvasive-electrocardiography-in-the-perinatal-mouse

Nanyang Technological University

Research

JoVE Journal

Medicine

5.9K Views.  University of Maine, Orono, ME. Here, we present a noninvasive electrocardiography (ECG) protocol, optimized for early postnatal mice, that does not require the use of anesthetics.

Video: Noninvasive Electrocardiography in the Perinatal Mouse

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable around embryonic day (E) 8.51. Systolic left ventricular (LV) pressure is 2 mmHg at E9.5 and 11 mmHg at E14.52. At these mid-embryonic stages, the LV is clearly visible through the chest wall for invasive pressure measurements because the ribs and skin are not fully developed. Between E14.5 and birth (approximately E21) imaging methods must be used to view the LV. After birth, mean arterial pressure increases from 30 - 70 mmHg from postnatal day (P) 2 - 353. Beyond P20, arterial pressure can be measured with solid-state catheters (i.e. Millar or Scisense). Before P20, these catheters are too big for developing mouse arteries and arterial pressure must be measured with custom pulled plastic catheters attached to fluid-filled pressure transducers3 or glass micropipettes attached to servo null pressure transducers4.
Our recent work has shown that the greatest increase in blood pressure occurs during the late embryonic to early postnatal period in mice5-7. This large increase in blood pressure may influence smooth muscle cell (SMC) phenotype in developing arteries and trigger important mechanotransduction events. In human disease, where the mechanical properties of developing arteries are compromised by defects in extracellular matrix proteins (i.e. Marfan's Syndrome8 and Supravalvular Aortic Stenosis9) the rapid changes in blood pressure during this period may contribute to disease phenotype and severity through alterations in mechanotransduction signals. Therefore, it is important to be able to measure blood pressure changes during late embryonic and neonatal periods in mouse models of human disease.
We describe a method for measuring LV pressure in late embryonic (E18) and early postnatal (P1 - 20) mice. A needle attached to a fluid-filled pressure transducer is inserted into the LV under ultrasound guidance. Care is taken to maintain normal cardiac function during the experimental protocol, especially for the embryonic mice. Representative data are presented and limitations of the protocol are discussed.

Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable ...

Bioengineering

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

Murine Fetal Echocardiography

Transgenic mice displaying abnormalities in cardiac development and function represent a powerful tool for the understanding the molecular mechanisms underlying both normal cardiovascular function and the pathophysiological basis of human cardiovascular disease. Fetal and perinatal death is a common feature when studying genetic alterations affecting cardiac development 1-3. In order to study the role of genetic or pharmacologic alterations in the early development of cardiac function, ultrasound imaging of the live fetus has become an important tool for early recognition of abnormalities and longitudinal follow-up. Noninvasive ultrasound imaging is an ideal method for detecting and studying congenital malformations and the impact on cardiac function prior to death 4. It allows early recognition of abnormalities in the living fetus and the progression of disease can be followed in utero with longitudinal studies 5,6. Until recently, imaging of fetal mouse hearts frequently involved invasive methods. The fetus had to be sacrificed to perform magnetic resonance microscopy and electron microscopy or surgically delivered for transillumination microscopy. An application of high-frequency probes with conventional 2-D and pulsed-wave Doppler imaging has been shown to provide measurements of cardiac contraction and heart rates during embryonic development with databases of normal developmental changes now available 6-10. M-mode imaging further provides important functional data, although, the proper imaging planes are often difficult to obtain. High-frequency ultrasound imaging of the fetus has improved 2-D resolution and can provide excellent information on the early development of cardiac structures 11.

Fetal and perinatal death is a common feature when studying genetic alterations affecting cardiac development. High-frequency ultrasound imaging has improved 2-D resolution and can provide excellent information on early cardiac development and is an ideal method to detect the impact on cardiac structure and function prior to death.

Transgenic mice displaying abnormalities in  cardiac development and function represent a powerful tool for the understanding  the molecular mechanisms ...

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound (30/45MHZ) System

Congenital heart defects (CHDs) are the most common cause of childhood morbidity and early mortality. Prenatal detection&#160;of the underlying molecular mechanisms of CHDs is&#160;crucial for inventing new preventive and therapeutic strategies. Mutant mouse models are powerful tools to discover new&#160;mechanisms&#160;and environmental stress modifiers that drive cardiac development and their potential&#160;alteration in CHDs. However, efforts to establish the causality of these putative contributors have been limited to histological&#160;and molecular studies&#160;in non-survival animal experiments, in which&#160;monitoring the key physiological and hemodynamic parameters is&#160;often absent. Live imaging technology has become an essential tool to establish the&#160;etiology of CHDs. In particular,&#160;ultrasound imaging can be used prenatally without surgically exposing the fetuses,&#160;allowing maintaining&#160;their baseline physiology while monitoring the impact of environmental stress&#160;on the hemodynamic and structural aspects of cardiac chamber development. Herein, we use the High-Frequency Ultrasound (30/45)&#160;system to examine the cardiovascular system in fetal mice at E18.5 in utero at the baseline and in response to prenatal hypoxia exposure. We demonstrate the feasibility of the system to measure cardiac chamber size, morphology, ventricular function, fetal heart rate, and umbilical artery flow indices, and their alterations in fetal mice exposed to systemic chronic hypoxia in utero in real time.

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound 30/45MHZ System

High-frequency ultrasound imaging of the fetal mouse has improved imaging resolution and can provide precise non-invasive characterization of cardiac development and structural defects. The protocol outlined herein is designed to perform real-time fetal mice echocardiography in vivo.

Congenital heart defects (CHDs) are the most common cause of childhood morbidity and early mortality. Prenatal detection&#160;of the underlying molecular ...

Developmental Biology

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Congenital heart disease (CHD) is the most frequent noninfectious cause of death at birth. The incidence of CHD ranges from 4 to 50/1,000 births (Disease and injury regional estimates, World Health Organization, 2004). Surgeries that often compromise the quality of life are required to correct heart defects, reminding us of the importance of finding the causes of CHD. Mutant mouse models and live imaging technology have become essential tools to study the etiology of this disease. Although advanced methods allow live imaging of abnormal hearts in embryos, the physiological and hemodynamic states of the latter are often compromised due to surgical and/or lengthy procedures. Noninvasive ultrasound imaging, however, can be used without surgically exposing the embryos, thereby maintaining their physiology. Herein, we use simple M-mode ultrasound to assess heart rates of embryos at E18.5 in utero. The detection of abnormal heart rates is indeed a good indicator of dysfunction of the heart and thus constitutes a first step in the identification of developmental defects that may lead to heart failure.

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Ultra-high frequency ultrasound is a powerful live imaging tool to examine cardiac abnormalities in small animals. Its noninvasiveness allows maintaining the physiologic state of embryos. Herein, we show the use of M-mode ultrasound to measure the heart rates of embryos at E18.5 in utero.

Congenital heart disease (CHD) is the most frequent noninfectious cause of death at birth. The incidence of CHD ranges from 4 to 50/1,000 births (Disease ...

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

Mouse Maternal Autonomic Changes from Pregnancy to Pup-Rearing

Analysis of Electrocardiograms and Behavior in Mice from Pregnancy to Lactation Period

Changes in the mother-offspring relationship are presumably accompanied by dynamic changes in the autonomic nervous system. Although temporal measurements of autonomic activity have been performed in human mothers and infants, the analysis of long-term changes remains unexplored. Mouse mothers can form social bonds with their pups and have a short period of pregnancy and lactation, which makes them useful for the examination of physiological changes from pregnancy to pup-rearing. Therefore, a telemetry system was used for several weeks to measure the changes in the autonomic nervous system and the behavior of mouse mothers. The current results showed that an electrocardiogram (ECG) could be stably recorded regardless of the movements of mothers and parturition. ECG analysis showed that the heart rate gradually decreased from pregnancy to lactation, and sympathetic activity sharply increased as the pups developed. Furthermore, the simultaneous recording of behavior and ECG in the home cage enabled us to understand the behavior-dependent influences on the ECG, thereby revealing the characteristics of autonomic nervous activity during each behavior. Thus, the present experimental method helps to understand how the physiological characteristics of mothers change from pregnancy through pup rearing, supporting the healthy development of pups.

A simultaneous recording of autonomic activity and detailed maternal behavior of mother mice from pregnancy to lactation was achieved using a telemetry system. This method helps to understand the dynamics of the physiological and behavioral characteristics in mothers from pregnancy to weaning.

Changes in the mother-offspring relationship are presumably accompanied by dynamic changes in the autonomic nervous system. Although temporal measurements ...

Behavior

Myocardial Infarction in Neonatal Mice, A Model of Cardiac Regeneration

Myocardial infarction induced by coronary artery ligation has been used in many animal models as a tool to study the mechanisms of cardiac repair and regeneration, and to define new targets for therapeutics. For decades, models of complete heart regeneration existed in amphibians and fish, but a mammalian counterpart was not available. The recent discovery of a postnatal window during which mice possess regenerative capabilities has led to the establishment of a mammalian model of cardiac regeneration. A surgical model of mammalian cardiac regeneration in the neonatal mouse is presented herein. Briefly, postnatal day 1 (P1) mice are anesthetized by isoflurane and placed on an ice pad to induce hypothermia. After the chest is opened, and the left anterior descending coronary artery (LAD) is visualized, a suture is placed around the LAD to inflict myocardial ischemia in the left ventricle. The surgical procedure takes 10-15 min. Visualizing the coronary artery is crucial for accurate suture placement and reproducibility. Myocardial infarction and cardiac dysfunction are confirmed by triphenyl-tetrazolium chloride (TTC) staining and echocardiography, respectively. Complete regeneration 21 days post myocardial infarction is verified by histology. This protocol can be used to as a tool to elucidate mechanisms of mammalian cardiac regeneration after myocardial infarction.

This protocol describes a highly reproducible model of cardiac regeneration by surgical induction of myocardial infarction in the left ventricle of postnatal day 1 mice. The method involves induction of hypothermic anesthesia and ligation of the left anterior descending coronary artery.

Myocardial infarction induced by coronary artery ligation has been used in many animal models as a tool to study the mechanisms of cardiac repair and ...

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 ...

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.

Noninvasive Electrocardiography in the Perinatal Mouse

Summary

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Chapters in this video

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

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

Murine Fetal Echocardiography

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound (30/45MHZ) System

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Ambulatory ECG Recording in Mice

Analysis of Electrocardiograms and Behavior in Mice from Pregnancy to Lactation Period

Myocardial Infarction in Neonatal Mice, A Model of Cardiac Regeneration

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

Electrocardiogram Recordings in Anesthetized Mice using Lead II