The authors would like to acknowledge the help and collaboration of Queen&#8217;s University animal facility personal. The authors would like to acknowledge the help of Austin Read, TMED MSc candidate.
This study was supported in part by U.S. National Institutes of Health (NIH) grants NIH 1R01HL113003-01A1 (S.L.A.), NIH 2R01HL071115-06A1 (S.L.A), Canada Foundation for Innovation and the Queen&#8217;s Cardiopulmonary Unit (QCPU) 229252 and 33012 (S.L.A.), Tier 1 Canada Research Chair in Mitochondrial Dynamics and Translational Medicine 950-229252 (S.L.A.), Canadian Institutes of Health Research (CIHR) Foundation Grant CIHR FDN 143261, the William J. Henderson Foundation (S.L.A.), Canadian Vascular Network Scholar Award (F.P.), and the Paroian Family scholarship from the pulmonary hypertension association of Canada (F.P.)

All experiments were performed in accordance with Queen&#8217;s University biosafety and ethical guidelines (ROMEO/TRAQ#6016826). The procedures followed were performed in accordance with institutional guidelines. This is a terminal procedure. Due to the invasiveness of the right and left catheterization, the animals should be euthanized immediately after data acquisition. Euthanasia should be performed according to the institution&#8217;s animal studies guidelines.
1. Experimental preparation and setup
<ol>
	<li>Put the catheter in a 10 mL syringe with saline/heparin, at room temperature 30 min before starting the experiment (Figure 1A).</li>
	<li>After 30 min, calibrate the catheter (e.g., baseline and acquisition system) according to the manufacturer&#39;s recommendations. The acquisition system displays high and low calibration values that are used to calibrate the acquisition system before starting an experiment. Output these values and make sure that they match.
	<ol>
		<li>Use the button &#8220;Pressure Balance Control&#8221;, &#8220;Coarse +/- &#8220;, or &#8220;Fine +/-&#8220; to set up the baseline pressure value at zero.</li>
		<li>Perform a two point calibration for high and low signal.
		<ol>
			<li>On the control console press &#8220;System Setting&#8221; in the &#8220;Catheter Menu&#8221;.</li>
			<li>Press &#8220;Send Calibration Signal&#8221; in the &#8220;System Setting Menu&#8221; to send the low signal. Ensure that the pressure, volume, phase, and magnitude are at 0 mm Hg, 0 &#181;L, 0&#730;, and 0 &#181;s respectively.</li>
			<li>Press &#8220;Enter&#8221; to send the high signal. Ensure that pressure, volume, phase, and magnitude are at 100 mm Hg, 150 &#181;L, 20&#730; and 5,000 &#181;s respectively.</li>
			<li>Press &#8220;Enter&#8221; to return to the &#8220;System Setting Menu&#8221;.</li>
			<li>Press &#8220;6&#8221; to return to the &#8220;Catheter Menu&#8221;. Then press &#8220;Acquire Data&#8221;.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Bend a 30 G needle to approximately 90&#730; (Figure 1B,C). This bent needle will be used to puncture the jugular and carotid vessels.</li>
</ol>
2. Anesthesia and body temperature control&#160;
<ol>
	<li>Place the mouse (28 g, C57BL/6 in this protocol) into an anesthesia chamber containing anesthetic gas (i.e., oxygen 100%, isoflurane 3&#8211;4% for induction).</li>
	<li>When the animal is anesthetized, not responding to paw or tail pinch, place the mouse supine on the heating pad set at 37 &#730;C.</li>
	<li>Connect the mouse to the respirator through a nose cone providing a mixture of 100% oxygen and 2% of isoflurane. To automatically calculate the recommended ventilation settings, enter the animal&#8217;s weight into the ventilator&#8217;s proprietary software using the touch screen. The calculations use the following formula: 
	Tidal volume = 6.2 x animal mass1.01 (kg), 
	Respiration rate = 53.5 x animal mass-0.26 (kg).</li>
	<li>Turn on the anesthetic line from the anesthesia chamber to the nose cone.</li>
	<li>Insert the temperature feedback probe into the rectum, and the pad probe between the pad and the back of the mouse, setting the desired body temperature to 37 &#176;C&#8211;37.5 &#176;C. Control the animal&#8217;s temperature on the monitor screen (Figure 2A,B).</li>
	<li>Tape down the front paws and one distal paw of the mouse to the heating blanket using surgical tape, leaving one hind paw free to monitor the depth of anesthesia.</li>
</ol>
3. Surgical Site Access
<ol>
	<li>Perform a 2 cm H-shape ventral midline cervical incision from the manubrium to the level of the hyoid bone.
	<ol>
		<li>Reflect the skin away from the underlying muscles. If needed, these muscles can be excised for better visualization.&#160;</li>
		<li>Gently move the submandibular gland aside.</li>
		<li>Dissect the cervical soft tissue and expose the sternocleidomastoid and the sternohyoid muscle with forceps using the blunt dissection method.</li>
		<li>Split the fascia in the middle, overlying the paired sternohyoid. Allow the paired sternohyoid to retract laterally to expose the trachea. Be careful to not damage the carotid arteries and the vagus nerves, which run alongside the trachea.</li>
	</ol>
	</li>
	<li>Pass forceps underneath the trachea to elevate it. Then, pass a 4.0 surgical silk suture underneath the trachea and make a potential knot in the middle of the suture, which will later be tightened to secure the endotracheal tube (Figure 3A).</li>
	<li>Using scissors, make a small cut between the cartilage rings of the trachea below the level of the larynx. Insert the endotracheal tube (Figure 3B).</li>
	<li>Connect the tracheostomy tube to the respirator and begin ventilation with 100% oxygen and 2% isoflurane. Tighten the knot around the trachea to secure the endotracheal tube and tape the respirator tubing to the operative table. Ensure that the trachea is not obstructed or collapsed (Figure 3C).</li>
</ol>
4. Right jugular and right carotid isolation
<ol>
	<li>Right carotid isolation
	<ol>
		<li>Using blunt dissection, displace the sternohyoid muscle laterally to expose and isolate the right carotid artery.</li>
		<li>Isolate the carotid artery from the vagus nerve by blunt dissection using forceps.</li>
		<li>Pass three surgical sutures (4.0) underneath the carotid artery, excluding the vagus nerve.</li>
	</ol>
	</li>
	<li>Right jugular vein isolation
	<ol>
		<li>Displace the submandibular and parotid gland laterally to visualize the right jugular vein. Bluntly dissect and expose the right jugular vein using forceps. Carefully dissect the vein and remove the surrounding fascia.</li>
		<li>Pass forceps underneath the jugular vein.</li>
		<li>Pass one surgical suture underneath the jugular vein, then tie it at the cranial side of the vein. Apply gentle traction on this suture in the direction of the head using a hemostatic clamp.</li>
		<li>Pass two additional sutures underneath the jugular vein. Gently pull the most distal suture in a caudal direction using a hemostatic clamp. Make a loose, potential knot in the middle suture.</li>
		<li>Put several drops of warmed, physiological saline on the vessel at the site of anticipated venotomy.</li>
	</ol>
	</li>
</ol>
5. Surgical procedures for right ventricular and left ventricular catheterization
<ol>
	<li>Right ventricular catheterization (Figure 4 A&#8211;D).
	<ol>
		<li>Using the stereomicroscope, identify the jugular vein.</li>
		<li>Gently apply superior traction on the vein. Perform a venotomy by inserting a 30 G curved needle between the cranial suture and the middle suture. Insert the needle at a 140&#730; angle relative to the vein to ensure it enters in a coaxial manner.</li>
		<li>When inserted, dilate the venotomy by moving the needle. Insert the catheter tip in the venotomy, underneath the needle. Then gently tie the middle suture, securing the catheter. 
		NOTE: Take extreme care not to tie the suture too tightly, because excess force can damage the catheter.</li>
		<li>Release the caudal suture, and advance the catheter into the right ventricle, detecting the classical right ventricular pressure waveform on a continuous monitor.</li>
		<li>Stabilize the right ventricular pressure. Ensure the correct positioning of the catheter in the right ventricle to generate an optimal PV loop.
		<ol>
			<li>Stabilize the magnitude, which reflects the blood and muscle, to generate pressure-magnitude loops (i.e., Y axis pressure, X axis magnitude). If required, gently rotate the catheter shaft to achieve optimal placement of the catheter along the axis of the right ventricle. 
			NOTE: The maximal phase value, which reflects the muscle, should be below 7&#176;.</li>
		</ol>
		</li>
		<li>When the pressure-magnitude loop signal is optimal, press &#8220;Enter&#8221; on the console during the acquisition to perform a baseline scan. Ensure that the heart rate reported on the monitor screen in beats per minute (bpm) is in a physiologic range (i.e., 400&#8211;600 bpm).</li>
		<li>Generate the PV loops. Change &#8220;Magnitude&#8221; to &#8220;Volume&#8221; as a parameter for the X axis and keep the pressure as the Y axis. When the PV loops signal is optimal, record for 30 s.</li>
		<li>Stop the recording. Pull back the catheter and gently wipe with gauze. Put the catheter in heparin/sodium chloride solution and tie the caudal suture to stop bleeding from the jugular vein.</li>
	</ol>
	</li>
	<li>Left ventricular catheterization (Figure 5 A&#8211;D).
	<ol>
		<li>Gently elevate the right carotid, which was previously isolated (5A) by sliding curved forceps underneath the artery.</li>
		<li>Tie the previous suture, thereby occluding the artery. Then, gently apply cranially directed traction using a hemostatic clamp.</li>
		<li>Pull the most distal suture in a caudal direction using a hemostatic clamp. Make a loose potential knot on the middle suture.</li>
		<li>Put several drops of warmed, physiological saline on the vessel at the site of the anticipated arteriotomy. Focus on the cranial section, between the caudal and the middle suture, using the stereotaxic microscope.</li>
		<li>Gently apply superior traction on the artery. Perform an arteriotomy, by inserting a 30 G curved needle between the cranial suture and the middle suture. Insert the needle at 140&#730; relative to the artery to ensure it enters in a coaxial manner.</li>
		<li>Insert the catheter tip into the arteriotomy and then tighten the middle suture to secure the catheter. Simultaneously, release the distal suture and advance the catheter into the aorta to start recording. Ensure that the pressure channel shows a typical aorta trace.</li>
		<li>Advance the catheter retrograde across the aortic valve into the left ventricle. Entry into the left ventricle will be evident from the sudden marked drop in diastolic pressure from the aorta.</li>
		<li>Stabilize the left ventricular pressure. Ensure the correct positioning of the catheter in the left ventricle to generate an optimal PV loop.
		<ol>
			<li>Stabilize the magnitude, which reflects the blood and muscle, to generate pressure-magnitude loops (i.e., Y axis pressure, X axis magnitude). If required, gently rotate the catheter shaft to achieve optimal placement of the catheter along the axis of the left ventricle. 
			NOTE: The maximal phase value, which reflects the muscle, should be below 7&#176;.</li>
		</ol>
		</li>
		<li>Stop the recording. Pull back the catheter and put it in heparin/sodium chloride solution. Then tie the caudal suture.</li>
		<li>Clean the catheter with an enzymatic detergent (e.g., endozime). 
		NOTE: After the surgery, euthanize the animal according to the institution&#8217;s animal studies guidelines.&#160;</li>
	</ol>
	</li>
</ol>
6. Data analysis&#160;
<ol>
	<li>Perform the PV loop analysis according to established recommendations.
	<ol>
		<li>Select the optimal pressure-volume trace (ideally an entire, stable 30 s recording). On the software, click &#8220;Advance&#8221;, click &#8220;Loops&#8221;, and then click &#8220;Offline Calculation&#8221;.</li>
		<li>Select volume as a volume channel and pressure as pressure channel.</li>
		<li>For consistent results a minimum of 20 loops is necessary.</li>
	</ol>
	</li>
</ol>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Validation+of+high-resolution+echocardiography+and+magnetic+resonance+imaging+vs.+high-fidelity+catheterization+in+experimental+pulmonary+hypertension.">Validation of high-resolution echocardiography and magnetic resonance imaging vs. high-fidelity catheterization in experimental pulmonary hypertension.</a> American Journal of Physiology-Lung Cellular and Molecular Physiology. 299 (3), 401-412 (2010).</li><li>Ashton, J. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anatomical+and+functional+imaging+of+myocardial+infarction+in+mice+using+micro-CT+and+eXIA+160+contrast+agent.">Anatomical and functional imaging of myocardial infarction in mice using micro-CT and eXIA 160 contrast agent.</a> Contrast Media &amp; Molecular Imaging. 9 (2), 161 (2014).</li><li>Larson, E. R., Feldman, M. D., Valvano, J. W., Pearce, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+Spatial+Sensitivity+of+Conductance/Admittance+Catheter+Ventricular+Volume+Estimation.">Analysis of the Spatial Sensitivity of Conductance/Admittance Catheter Ventricular Volume Estimation.</a> IEEE Transactions on Biomedical Engineering. 60 (8), 2316-2324 (2013).</li><li>Sasayama, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+cardiac+function+by+left+heart+catheterization:+an+analysis+of+left+ventricular+pressure-volume+(length)+loops.">Assessment of cardiac function by left heart catheterization: an analysis of left ventricular pressure-volume (length) loops.</a> Journal of Cardiography. Supplement. (1), 25-34 (1984).</li><li>Lindsey, M. L., Kassiri, Z., Virag, J. A. I., de Castro Br&#225;s, L. E., Scherrer-Crosbie, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Guidelines+for+measuring+cardiac+physiology+in+mice.">Guidelines for measuring cardiac physiology in mice.</a> American Journal of Physiology-Heart and Circulatory Physiology. 314 (4), 733-752 (2018).</li><li>Townsend, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+Pressure+Volume+Loops+in+the+Mouse.">Measuring Pressure Volume Loops in the Mouse.</a> Journal of Visualized Experiments JoVE. (111), e53810 (2016).</li><li>Provencher, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Standards+and+Methodological+Rigor+in+Pulmonary+Arterial+Hypertension+Preclinical+and+Translational+Research.">Standards and Methodological Rigor in Pulmonary Arterial Hypertension Preclinical and Translational Research.</a> Circulation Research. 122 (7), 1021-1032 (2018).</li><li>Lips, D. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Left+Ventricular+Pressure-Volume+Measurements+in+Mice:+Comparison+of+Closed-Chest+Versus+Open-Chest+Approach.">Left Ventricular Pressure-Volume Measurements in Mice: Comparison of Closed-Chest Versus Open-Chest Approach.</a> Basic Res Cardiol. 99 (5), 351-359 (2004).</li></ol>

Assessment of cardiac function is a critical step for preclinical cardiovascular and pulmonary-vascular research. In this work, we proposed a protocol for a closed chest biventricular assessment of cardiac function in mice. Through this approach, one can generate the right ventricle and left ventricle PV loops in the same mouse. This approach provides a robust and complete assessment of cardiac function, allowing measurement of systolic and diastolic function, as well as stroke volume and cardiac output. Unlike the open chest approach classically used for rodent catheterization, this closed chest technique results in more stable physiology and more physiologically relevant data. While technically more challenging and dependent on operator skills to successfully position the catheter in the right and left ventricle, the closed chest approach limits the trauma and hemorrhage associated with open chest surgery and reduces the drastic pressure changes associated with exposing the lungs to atmospheric pressure. The closed chest approach also better emulates the cardiac catheterization procedure performed in patients, which enhances the relevance of using this technique in preclinical research.
The surgical procedure is the critical step of the protocol. Even when using a surgical microscope for catheter insertion in the jugular vein or carotid artery, which is recommended, this procedure requires practice and technical skill. Careful dissection of the vessels free from surrounding fascia by means of gentle, blunt dissection will increase the success of cannulation while minimizing the risk of hemorrhage. To minimize blood loss, it is crucial to cannulate the carotid in sequential steps: 1) introduce the catheter tip in the carotid artery; 2) gently tie the suture around the portion of the artery that contains the catheter; 3) release the secure suture, allowing catheter movement while maintaining gentle upward traction to minimize bleeding; and 4) advance the catheter to the aorta. Positioning the catheter in the ventricle, as determined by real-time waveform monitoring, is the most challenging part of this protocol. All the catheter&#39;s electrodes should be within the ventricular cavity and none should be touching the wall. Any improper positioning of the catheter will result in irregular PV loops and will adversely affect or preclude data acquisition. Recognizing the characteristic pressure-volume waveform that results from having all electrodes within the ventricle allows one to be confident of an appropriate catheter position. It is critical to obtain a stable ventricular pressure waveform and stable pressure-magnitude loops before shifting to the PV mode and volume acquisition. Proper knowledge of cardiac physiology and anatomy is essential for the success of this procedure. Online reading of the PV traces, from the atrium, the tricuspid valve area, and right ventricle, will show the advancing of the catheter and help achieve proper positioning. It is critical to know the normal heart rate (400&#8211;600 bpm), and waveforms and pressures expected (e.g., right ventricular systolic pressure, 18&#8211;25 mm Hg, diastolic pressure &#60;5 mm Hg; left ventricular systolic pressure 60&#8211;120 mm Hg40, diastolic pressure &#60;8 mmHg) in mice to allow the operator to evaluate the veracity of the observed data.
The quality and reproducibility of the data will depend on the speed of the procedure and blood loss or hemorrhage. The procedure from anesthesia to completion of data acquisition takes on average ~30&#8211;40 min/mouse. Right heart catheterization from the insertion of the catheter to data acquisition takes 5&#8211;10 min, left heart catheterization from the insertion of the catheter to the data acquisition takes another 10&#8211;15 min. Publication-quality data is obtained in ~75% of cases. The sequence of steps in the cardiac catheterization should be kept constant between the animals. In this procedure, the mice are intubated first, followed by the right ventricular catheterization, and finally the left ventricular catheterization. The decision to proceed in this order is based on the greater difficulty and bleeding risk of left heart versus right heart catheterization. A nonspecific 50 Hz noise recording artifact can be observed. This noise could be diminished using an FIR filter with a high cutoff at 50 Hz and a low cutoff of 0 on the software. For the volume channel create a new channel/filter/FIR filter. A notch filter of 50 Hz could also be applied during data acquisition to eliminate mains noise and remove any radiofrequency interference.
The faster the catheterization is done, the better the quality of the data. Based on previous experience, it is recommended to acquire the data within 15 min. Increased catheterization time increases the physiologic stress on the animal and increases the risk of arrhythmia due to the presence of the catheter in the cavity. These forces can reduce stroke volume and impair the reproducibility and interpretability of the waveforms. In addition, the tip of the catheter is sharp and can damage or puncture the ventricle. This is particularly important for the right ventricle, which is ~ 1/3rd the thickness of the left ventricle.
Invasive tracheostomy and positive pressure mechanical ventilation result in stable and controlled breathing of the mice and decrease the variability of the PV loops acquisition. However, positive end expiratory pressure (PEEP) is a marked contrast to normal ventilation, which is a negative pressure phenomenon. Together, positive pressure ventilation and PEEP lower cardiac output and reduce right heart pressure. Thus, while required for acquisition of stable data, mechanical ventilation as well as cardiodepressive effects of the anesthesia will affect the PV loops and should be considered as a limitation. Transiently stopping mechanical ventilation during the brief recording of PV loops is used to eliminate this potential source of artifacts. Note that ventilation efficiency can be confirmed by the capnography monitoring of carbon dioxide.
The technical skills required for the closed-chest approach may be a limitation of this technique. Likewise, it is challenging to obtain proper, stable positioning of the catheter in the ventricle. The odds of success increase with operator experience and with the size and weight of the mice. Catheterization of mice below 20 g is extremely challenging. The unique chamber geometry of the right ventricle might affect&#160;volume measurement and should be considered. The anesthetic used, heart rates, temperatures, and animal strain could affect the hemodynamic parameters and should be carefully reported and monitored.
In conclusion, in this protocol both right and left ventricular catheterization are performed in the same mouse. Depending on a scientist&#8217;s specific aims, left or right ventricular catheterization can be performed independently, using the relevant portion of the biventricular procedure. However, the approach presented is optimal for complete assessment of cardiac function.

According to the world health organization (WHO), heart disease is the leading cause of death worldwide for both men and women1,2,3. Many studies focus on diagnosing and improving impaired cardiac function4. For these applications high-quality and reproducible evaluation of cardiac function is critical. High fidelity and reproducible catheter data are required to assess both etiological and therapeutic responses. For example, the assessment of cardiac function is essential to evaluate the efficacy of drugs and other treatments in preclinical models of myocardial infarction5. While many cardiovascular studies focus on left ventricular function, right ventricular function is also a critical determinant of functional capacity and prognosis in patients with pulmonary-vascular disease6,7. In patients with advanced heart failure, persistently elevated right-sided and left-sided filling pressures are predictive of the combined risk of death, cardiovascular hospitalization, and heart transplantation8. In combined aortic and mitral valve disease, preoperative myocardial function (reflected in parameters such as cardiac index and left ventricular ejection fraction) is the main predictor of long-term survival9. Right ventricular function is the major predictor of both morbidity and mortality in pulmonary arterial hypertension10,11. Thus, assessment of the right ventricular function is a necessary component of a comprehensive preclinical study using models of pulmonary arterial hypertension12,13,14.
Left and right ventricular function are often studied independently. However, because the functions of the left and right ventricles are intimately linked, it is ideal to obtain a biventricular assessment of systolic and diastolic function from a single test15. For example, the right ventricle shares oblique fibers in the interventricular septum with the left ventricle, which constitutes one of the mechanical links between the left and right ventricular contractile function16,17. This phenomenon, known as systolic ventricular interaction, allows left ventricular contraction to augment the right ventricular contraction. Ventricular interactions during diastole are also important. During diastole, the volume of one ventricle influences the volume of the opposite ventricle, and thereby alters diastolic compliance and preload18,19. In pathological conditions, decreased function of one ventricle, or impaired volume loading, can directly or indirectly impair the function of the other ventricle20. As a consequence of systolic ventricular interaction, a global decrease in left ventricular function may reduce right ventricular contractile performance15. In patients with heart failure due to left ventricular systolic function and increased end diastolic pressure, pulmonary artery pressure is elevated, indirectly increasing right ventricle afterload21,22. Conversely, increased right ventricular pressure and volume overload in severe pulmonary hypertension exerts a mechanical compression on the left heart. This D-shaped flattening of the left ventricle, caused by a leftward shift in the interventricular septum, reduces left ventricular volumes and impaired systolic and diastolic function23,24,25,26,27. Thus, the assessment of both left and right ventricles is essential to evaluate global cardiac function in preclinical models of human disease.
Cardiac function can also be assessed by noninvasive echocardiography, magnetic resonance imaging (MRI), and invasive catheterization28,29,30. Echocardiography is the most commonly used imaging modality in cardiovascular research because it is relatively inexpensive and accessible31. However, echocardiography has several technical limitations, including indirect measurement of filling pressure and limited ability to quantify diastolic function. In addition, the quality of the data obtained by echocardiography is highly operator dependent. Cardiac MRI is a relatively new addition to preclinical imaging armamentarium that has great potential for quantitative assessment of biventricular function. Quantification with cardiac MRI is accurate, as it does not make geometric assumptions of ventricular shape, unlike echocardiography32. However, the MRI imaging platform is expensive, and is rarely available. Moreover, the processing of MRI data requires skilled support by a physicist or equivalent scientist, which is lacking in many preclinical laboratories33. Similarly, the use of microcomputed tomography (MicroCT) in preclinical studies provides quantitative high-resolution three-dimensional (3D) anatomical data that can be obtained noninvasively, allowing longitudinal studies34. However, MicroCT imaging requires the injection of contrast agents, which are often expensive. The MicroCT imaging platform, like MRI, is also expensive and also requires a skilled technician.
In contrast, catheterization is an invasive technique that consists of the introduction of a catheter in the right and/or left ventricle to measure pressure and/or volume. The tools required to perform cardiac catheterization are not as expensive as echocardiography, CT, or MRI. Substantial technical proficiency for catheterization and small animal anesthesia is required, however. Catheterization allows direct and accurate assessments of cardiac function28. In this protocol, an admittance PV catheter is used to assess cardiac function. This technology, based on the distinct electrical conductance properties of blood and cardiac muscle, allows for the simultaneous recording of pressure and volume within the cardiac cavity and generation of PV loops in real time5,35. Briefly, the catheter is comprised of both excitation electrodes and recording electrodes. The excitation electrodes generate an electrical field inside the right or left ventricle. The inner recording electrode measures voltage change, which is proportional to a change in resistance. Deriving ventricular volume is based on Ohm&#8217;s law (voltage = current x resistance) from which conductance (i.e., the inverse of resistance) is calculated. In this setting, the measured conductance value is a combination of blood conductance and muscle conductance. In the electric field, blood is purely resistive while muscle has both capacitive and resistive properties. The capacitive property of muscle causes a time delay in the measured signal. Tracking this delay, known as the &#8220;phase&#8221; angle, reports heart tissue intrusion into the field as the heart contracts. This measurement is greatest at systole and lowest at diastole. This property allows the separation of the muscle component of the conductance from that of blood and allows a close approximation of absolute systolic and diastolic volumes. Pressure-volume loops provide a range of hemodynamic parameters that are not readily measurable by other methods, such as simple retrograde catheterization using fluid filled catheters to measure cardiac pressures. Pressure-volume loops measure ventricular pressures but also provide data about contractility, elastance, power, energetics, and efficiency. In addition, PV loops provide robust quantitative measurements36. Thus, assessment of cardiac function by PV loops generated by catheterization has emerged as the gold standard in preclinical research37. In addition, preclinical techniques are relevant to human disease where cardiac catheterization, albeit with fluid filled catheters, is common. However, cardiac catheterization in rodents requires impeccable anesthesia and excellent technique to prevent excessive loss of blood, hypoventilation, or changes in body temperature.
In human patients, cardiac catheterization is performed in closed chest configuration and vascular access is achieved via the jugular or subclavian vein for the right ventricle and the radial or femoral artery for the left ventricle. Due to the small size of mice, the closed chest approach is often challenging. Thus, studies conducted in mice commonly adopt an open chest&#160;approach. This technique involves opening the thorax, thereby exposing the heart, and facilitating the insertion of the catheter via puncture of the left and/or right ventricular apex38. While this approach is technically less challenging and fairly reproducible, its major limitations include hemorrhage and other complications of apical insertion of catheters, and a marked drop in intracardiac pressure resulting from opening the thoracic cavity to atmospheric pressure. Opening the thorax in a ventilated rodent induces a 5&#8211;10 mm Hg decrease in left ventricular systolic pressure and 2&#8211;5 mm Hg decrease in right ventricular pressure39. Therefore, a closed chest approach that is less traumatic for the heart and yields more physiologically relevant measurements that are more easily translated to clinical assessment of cardiac function was developed.

<table border="0" cellpadding="0" cellspacing="0" style="width:664px;" width="663">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">ADVantage Pressure-Volume System (ADV500)</td>
			<td style="width:121px;">Transonic</td>
			<td style="width:161px;">FY097B</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Endozime AW triple plus</td>
			<td style="width:121px;">Ruhof</td>
			<td style="width:161px;">34521</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Fiber optic dual Gooseneck</td>
			<td style="width:121px;">Volpi Intralux</td>
			<td style="width:161px;"># 6000-1</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Forceps</td>
			<td style="width:121px;">F.S.T</td>
			<td style="width:161px;">11052-10</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Forceps</td>
			<td style="width:121px;">F.S.T</td>
			<td style="width:161px;">11251-20</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Gauze sponges</td>
			<td style="width:121px;">Dermacea</td>
			<td style="width:161px;">441400</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Hemostatic clamp</td>
			<td style="width:121px;">F.S.T</td>
			<td style="width:161px;">13003-10</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Hemostatic clamp</td>
			<td style="width:121px;">F.S.T</td>
			<td style="width:161px;">13018-14</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Heparin sodium</td>
			<td style="width:121px;">Sandoz</td>
			<td style="width:161px;">023-3086</td>
			<td style="width:167px;">100 U/L</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">High-fidelity admittance catheter</td>
			<td style="width:121px;">Scisence; Transonic</td>
			<td style="width:161px;">FTH-1212B-3518</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Isofluorane</td>
			<td style="width:121px;">Baxter</td>
			<td style="width:161px;">CA2L9108</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">labScribe v4 software</td>
			<td style="width:121px;">iworx</td>
			<td style="width:161px;">LS-30PVL</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Needle (30 gauge)</td>
			<td style="width:121px;">BD</td>
			<td style="width:161px;">305106</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">sodium chloride injection</td>
			<td style="width:121px;">Baxter</td>
			<td style="width:161px;">JB1309M</td>
			<td style="width:167px;">0.9%(wt/vol)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Stereo microscope</td>
			<td style="width:121px;">Cole-Parmer</td>
			<td style="width:161px;">OF-48920-10</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Surgical suture</td>
			<td style="width:121px;">SERAFLEX</td>
			<td style="width:161px;">ID158000</td>
			<td style="width:167px;">black braided silk, 4.0</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Surgical tape</td>
			<td style="width:121px;">3M, Transpore</td>
			<td style="width:161px;">SN770</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Tabletop Single Animal Anesthesia Systems</td>
			<td style="width:121px;">Harvard apparatus</td>
			<td style="width:161px;">72-6468</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Tracheotomy canula 1.45 mm diameter</td>
			<td style="width:121px;">Harvard apparatus</td>
			<td style="width:161px;">72-1410</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:215px;">Ventilator, far infrared warming pad for mice and rats PhysioSuite</td>
			<td style="width:121px;">Kent scientific corporation</td>
			<td style="width:161px;"># PS-02</td>
			<td style="width:167px;"></td>
		</tr>
	</tbody>
</table>

biventricular assessment of cardiac function and pressure-volume loops by closed-chest catheterization in mice

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) generated by recording both pressure and volume during cardiac catheterization are vital when assessing both systolic and diastolic cardiac function. Left and right heart function are closely related, reflected in ventricular interdependence. Thus, recording biventricular function in the same animal is important to get a complete assessment of cardiac function. In this protocol, a closed chest approach to cardiac catheterization consistent with the way catheterization is performed in patients is adopted in mice. While challenging, the closed chest strategy is a more physiological approach, because opening the chest results in major changes in preload and afterload that create artifacts, most notably a fall in systemic blood pressure. While high-resolution echocardiography is used to assess rodents, cardiac catheterization is invaluable, particularly when assessing diastolic pressures in both ventricles.
Described here is a procedure to perform invasive, closed chest, sequential left and right ventricular pressure-volume (PV) loops in the same animal. PV loops are acquired using&#160;admittance technology with a mouse pressure-volume catheter and pressure-volume system acquisition. The procedure is described, beginning with the neck dissection, which is required to access the right jugular vein and the right carotid artery, to the insertion and positioning of the catheter, and finally the data acquisition. Then, the criteria required to ensure the acquisition of high-quality PV loops are discussed. Finally, the analysis of the left and right ventricular PV loops and the different hemodynamic parameters available to quantify systolic and diastolic ventricular function are briefly described.

The catheter was placed in a 10 mL syringe containing a solution of heparinized saline at room temperature 30 min before the catheterization (Figure 1A). A 30 G needle was bent ~90&#730; (Figure 1B, C), and a 1.45 mm diameter tracheotomy canula was prepared (Figure 1C).
Maintenance of physiologic body temperature is critical. The mouse was taped down and connected to the respirator through a nose cone. The feedback probe was placed between the pad and the back of the mouse. A rectal probe was inserted to monitor the animal&#8217;s body temperature (Figure 2A). Body temperature (37.1 &#730;C) and pad (40.7 &#730;C) temperature were monitored (Figure 2B).
Photographs of the critical steps of the intubation procedure are displayed in Figure 3A&#8211;C. Successful and unobstructed intubation resulted in a regular respiratory rate with stable peak pressure (Figure 2B).
Pictures of the critical steps of right heart catheterization, from the isolation of the jugular vein (Figure 4A&#8211;C) to the insertion of the catheter in the jugular vein are shown in Figure 4D. Figure 5 shows the critical steps of left heart catheterization, including right carotid artery isolation (Figure 5 A,B) and catheter insertion (Figure 5 C,D)
The catheter was introduced into the jugular vein and advanced into the right ventricle. Then the right ventricular pressure was stabilized, and the correct positioning verified. All the catheter&#8217;s electrodes (6 mm long axis length) needed to be within the right ventricle chambers and not in contact with the ventricle walls. Optimal positioning of the catheter as schematically represented in Figure 6A generated optimal PV loops (i.e., triangular, regular). Improper positioning as schematically represented in Figure 6B (i.e., contact with the ventricular wall) will result in flawed PV loops (i.e., collapsed and irregular loops).
The catheter was introduced into the carotid, advanced into the aorta, then advanced retrograde across the aortic valve into the left ventricle. The left ventricular pressure was stabilized and right positioning verified. All the catheter&#8217;s electrodes (6 mm long axis length) should be within the left ventricle chambers and not in contact with the ventricle walls. Optimal positioning of the catheter as schematically represented in Figure 6C generated optimal PV loops (i.e., rectangular, regular). Improper positioning as schematically represented in Figure 6D (i.e., contact with the ventricular wall) resulted in flawed PV loops (i.e., collapsed, non-rectangular, and irregular loops).
Representative hemodynamics generated by left and right PV loops showed a heart rate of 410 bpm, cardiac output of 9,107 &#181;L/min, and stroke volume of 24.5 &#181;L. Specific right ventricular parameters showed a right ventricular systolic pressure of 21.9 mm Hg, right ventricular end diastolic pressure 1.049 mm Hg, ejection fraction of 56.1%, dp/dt max of 1,469 mm Hg/s, dp/dt max of -1,504 mm Hg/s, end diastolic volume of 38.4 &#181;L, stroke work of 0.068 mJ, pressure-volume area of 0.089 mJ, pulmonary arterial elastance (Ea) of 0.83 mm Hg/&#181;L, and Tau factor of 12.8 ms. Specific left ventricular parameters showed a left ventricular systolic pressure of 77.1 mm Hg, left ventricular end diastolic pressure of 2.33 mm Hg, ejection fraction of 59.1%, dp/dt max of 4,695 mm Hg/s, dp/dt max of -3,553 mm Hg/s, end diastolic volume of 36.9 &#181;L, stroke work of 0.14 mJ, pressure-volume area of 0.22 mJ, arterial elastance (Ea) of 5.37 mm Hg/&#181;L, and Tau factor of 15.1 ms (Table 1).
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Haemodynamic parameters</td>
			<td></td>
		</tr>
		<tr>
			<td>HR (BPM)</td>
			<td>410.6 &#177; 23.3</td>
		</tr>
		<tr>
			<td>CO (&#956;L/min)</td>
			<td>9107 &#177; 1016</td>
		</tr>
		<tr>
			<td>SV (&#956;L)</td>
			<td>24.5 &#177; 2.3</td>
		</tr>
		<tr>
			<td>RV function</td>
			<td></td>
		</tr>
		<tr>
			<td>RVSP (mmHg)</td>
			<td>21.9 &#177; 2.15</td>
		</tr>
		<tr>
			<td>RVEDP (mmHg)</td>
			<td>1.042 &#177; 0.12</td>
		</tr>
		<tr>
			<td>EF (%)</td>
			<td>56.1 &#177; 4.4</td>
		</tr>
		<tr>
			<td>dP/dt max (mmHg/s)</td>
			<td>1469 &#177; 170</td>
		</tr>
		<tr>
			<td>dP/dt max (- mmHg/s)</td>
			<td>1504 &#177; 215</td>
		</tr>
		<tr>
			<td>EDV (&#956;L)</td>
			<td>38.4 &#177; 3.7</td>
		</tr>
		<tr>
			<td>SW (mJoules)</td>
			<td>0.068 &#177; 0.008</td>
		</tr>
		<tr>
			<td>PVA (mJoules)</td>
			<td>0.084 &#177; 0.009</td>
		</tr>
		<tr>
			<td>Ea (mmHg/&#956;L)</td>
			<td>0.83 &#177; 0.09</td>
		</tr>
		<tr>
			<td>Tau factor (ms)</td>
			<td>12.8 &#177; 0.8</td>
		</tr>
		<tr>
			<td>LV function</td>
			<td></td>
		</tr>
		<tr>
			<td>LVSP (mmHg)</td>
			<td>77.1 &#177; 2.4</td>
		</tr>
		<tr>
			<td>LVEDP (mmHg)</td>
			<td>2.33 &#177; 0.17</td>
		</tr>
		<tr>
			<td>EF (%)</td>
			<td>59.1 &#177; 3.6</td>
		</tr>
		<tr>
			<td>dP/dt max (mmHg/s)</td>
			<td>4695 &#177; 355</td>
		</tr>
		<tr>
			<td>dP/dt max (- mmHg/s)</td>
			<td>3553 &#177; 373</td>
		</tr>
		<tr>
			<td>EDV (&#956;L)</td>
			<td>36.9 &#177; 4.8</td>
		</tr>
		<tr>
			<td>SW (mJoules)</td>
			<td>0.14 &#177; 0.013</td>
		</tr>
		<tr>
			<td>PVA (mJoules)</td>
			<td>0.22 &#177; 0.03</td>
		</tr>
		<tr>
			<td>Ea (mmHg/&#956;L)</td>
			<td>5.37 &#177; 0.9</td>
		</tr>
		<tr>
			<td>Tau factor (ms)</td>
			<td>15.07 &#177; 1.7</td>
		</tr>
		<tr>
			<td colspan="2">CO, cardiac output; Ea, arterial elastance; EDV, end diastolic volume;&#160;HR, heart rate; LVEDP, left ventricular end diastolic volume; LVSP, left&#160;ventricular systolic pressure; PVA, pressure volume area; RVEDP, right&#160;ventricular end diastolic pressure; RVSP, right ventricular systolic&#160;pressure; SV, stroke volume; SW, stroke work; Tau factor, Tau Mirsky.&#160;N= 6 mice. Values are expressed &#177; SEM</td>
		</tr>
	</tbody>
</table>
Table 1: Table of hemodynamic parameters. Left and right ventricular hemodynamic parameter measured in six mice.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61088/61088fig1.jpg" /> 
Figure 1: Experimental preparation and setup. (A) Catheter in a 10 mL syringe of saline/heparin, (B), (C) 30 G needle bent to approximately 90&#176;, (D) tracheotomy canula, 1.45 mm diameter. <a href="https://www.jove.com/files/ftp_upload/61088/61088fig1large.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/61088/61088fig2.jpg" /> 
Figure 2: Anesthesia, body temperature control. (A) Mouse with three paws taped, connected to respirator through a nose cone, with feedback and rectal probes inserted. Note that the warming pad is below the surgical blanket. (B) Temperature monitor control showing body (rectal) and pad (feedback) temperature and the ventilation parameters: respiratory rate (set RR), mean tidal volume (Meas TV), peak pressure (PeakPress), and the minute ventilation (MinVol). <a href="https://www.jove.com/files/ftp_upload/61088/61088fig2large.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/61088/61088fig3.jpg" /> 
Figure 3: Intubation procedure. (A) The skin was pulled away and cut. The submandibular gland was gently moved aside. The sternocleidomastoid and the sternohyoid muscle were pulled apart and then forceps were passed underneath the trachea, using gentle, blunt dissection. (B) Surgical silk (4.0) was passed underneath the trachea and a small cut was made anteriorly between two cartilage rings of the trachea. The tracheostomy was inserted and tied. (C) The tracheostomy tube was connected to the ventilator, and the suture was tied around the tubing. <a href="https://www.jove.com/files/ftp_upload/61088/61088fig3large.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/61088/61088fig4v2.jpg" /> 
Figure 4: Right ventricular catheterization. (A), (B), (C) The right jugular vein was isolated, then one surgical suture was passed underneath and tied at the cranial side of the vein. Gentle traction was applied on this suture in the direction of the head using a hemostatic clamp. Two additional sutures were passed distally, underneath the jugular vein. The most distal suture was pulled gently in a caudal direction using a hemostatic clamp. A loose, potential knot was made in the middle suture. (D) The catheter was inserted in the jugular vein, the middle suture was tied to the catheter. The images in (C) and (D) are magnified through a stereomicroscope. <a href="https://www.jove.com/files/ftp_upload/61088/61088fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61088/61088fig5.jpg" /> 
Figure 5: Left ventricular catheterization. (A), (B) The right carotid was isolated, then one surgical suture was passed underneath the jugular vein and tied at the cranial side of the vein. Gentle traction was applied on this suture in the direction of the head using a hemostatic clamp. Two additional sutures were passed underneath the carotid artery. The most distal suture was gently pulled in a caudal direction using a hemostatic clamp. A loose, potential knot in the middle suture was made. (C) The catheter tip was inserted into the carotid artery, and then the middle suture tied to the catheter to secure it. (D) The catheter was gently advanced retrograde down the carotid toward the aorta. The images in (B), (C), (D) are magnified through a stereomicroscope. <a href="https://www.jove.com/files/ftp_upload/61088/61088fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/61088/61088fig6.jpg" /> 
Figure 6: Schematic representation of catheter positioning and resulting PV loops. (A) Optimal catheter positioning in the right ventricle. The tip of the catheter is in the middle of the ventricle, isolated from the ventricle walls. Representative PV loops resulting from an optimal catheter positioning in the right ventricle (i.e., stable, triangular). (B) Improper catheter positioning in the right ventricle. The tip of the catheter is in contact with the ventricular walls. Representative PV loops noise resulting from a suboptimal catheter positioning in the right ventricle (i.e., collapsed, irregular). (C) Optimal catheter positioning in the left ventricle. The tip of the catheter is in the middle of the ventricle, isolated from the ventricle walls. Representative PV loops resulting from optimal catheter positioning in the left ventricle (i.e., stable, rectangular). (D) Improper catheter positioning in the left ventricle. The tip of the catheter is in contact with the ventricular walls. Representative PV loops resulting from a suboptimal catheter positioning in the left ventricle (i.e., collapsed, irregular). A 50 Hz FIR noise filter was applied to generate the PV loops. <a href="https://www.jove.com/files/ftp_upload/61088/61088fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice at JoVE.com

Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice

Presented here is a protocol to assess biventricular heart function in mice by generating pressure-volume (PV) loops from the right and left ventricle in the same animal using closed chest catheterization. The focus is on the technical aspect of surgery and data acquisition.

Assessment of cardiac function is essential to conduct cardiovascular and pulmonary-vascular preclinical research. Pressure-volume loops (PV loops) ...

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4.0K Views.  Queen's University. Presented here is a protocol to assess biventricular heart function in mice by generating pressure-volume (PV) loops from the right and left ventricle in the same animal using closed chest catheterization. The focus is on the technical aspect of surgery and data acquisition.

Video: Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice

A Murine Closed-chest Model of Myocardial Ischemia and Reperfusion

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ischemia and reperfusion. Immune response includes cytokine expression and release or secretion of endogenous ligands of innate immune receptors. Activation of innate immunity can potentially modulate infarct size. We have modified an existing murine closed-chest model using hanging weights which could be useful for studying myocardial pre- and postconditioning and the role of innate immunity in myocardial ischemia and reperfusion. This model allows animals to recover from surgical trauma before onset of myocardial ischemia.
Volatile anesthetics have been intensely studied and their preconditioning effect for the ischemic heart is well known. However, this protective effect precludes its use in open chest models of coronary artery ligation. Thus, another advantage could be the use of the well controllable volatile anesthetics for instrumentation in a chronic closed-chest model, since their preconditioning effect lasts up to 72 hours. Chronic heart diseases with intermittent ischemia and multiple hit models are other possible applications of this model.
For the chronic closed-chest model, intubated and ventilated mice undergo a lateral blunt thoracotomy via the 4th intercostal space. Following identification of the left anterior descending a ligature is passed underneath the vessel and both suture ends are threaded through an occluder. Then, both suture ends are passed through the chest wall, knotted to form a loop and left in the subcutaneous tissue. After chest closure and recovery for 5 days, mice are anesthetized again, chest skin is reopened and hanging weights are hooked up to the loop under ECG control.
At the end of the ischemia/reperfusion protocol, hearts can be stained with TTC for infarct size assessment or undergo perfusion fixation to allow morphometric studies in addition to histology and immunohistochemistry.

Surgical trauma induces an inflammatory response. Cytokines and endogenous ligands are known to modulate myocardial infarct size following ischemia and reperfusion. We present a modified closed-chest model of murine ischemia and reperfusion using hanging weights to minimize effects of thoracotomy.

Surgical trauma by thoracotomy in open-chest models of coronary ligation induces an immune response which modifies different mechanisms involved in ...

Cardiac Stress Test Induced by Dobutamine and Monitored by Cardiac Catheterization in Mice

Dobutamine is a &beta;-adrenergic agonist with an affinity higher for receptor expressed in the heart (&beta;1) than for receptors expressed in the arteries (&beta;2). When systemically administered, it increases cardiac demand. Thus, dobutamine unmasks abnormal rhythm or ischemic areas potentially at risk of infarction. 
Monitoring of heart function during a cardiac stress test can be performed by either ecocardiography or cardiac catheterization. The latter is an invasive but more accurate and informative technique that the former.
Cardiac stress test induced by dobutamine and monitored by cardiac catheterization accomplished as described here allows, in a single experiment, the measurement of the following hemodynamic parameters: heart rate (HR), systolic pressure, diastolic pressure, end-diastolic pressure, maximal positive pressure development (dP/dtmax) and maximal negative pressure development (dP/dtmin), at baseline conditions and under increasing doses of dobutamine.
As expected, in normal mice we observed a dobutamine dose-related increase in HR, dP/dtmax and dP/dtmin. Moreover, at the highest dose tested (12 ng/g/min) the cardiac decompensation of high fat diet-induced obese mice was unmasked.

We describe the protocol to perform a cardiac stress test induced by dobutamine and monitored by cardiac catheterization in normal mice. Also we show its application to unmask subclinical cardiac disease in high fat diet-induced obese mice.

Dobutamine is a &beta;-adrenergic agonist with an affinity higher for receptor expressed in the heart (&beta;1) than for receptors expressed in the ...

Assessment of Right Ventricular Structure and Function in Mouse Model of Pulmonary Artery Constriction by Transthoracic Echocardiography

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, the RV is significantly affected in pulmonary diseases such as pulmonary artery hypertension (PAH). In addition, the RV is remarkably sensitive to cardiac pathologies, including left ventricular (LV) dysfunction, valvular disease or RV infarction4. To understand the role of RV in the pathogenesis of cardiac diseases, a reliable and noninvasive method to access the RV structurally and functionally is essential.
A noninvasive trans-thoracic echocardiography (TTE) based methodology was established and validated for monitoring dynamic changes in RV structure and function in adult mice. To impose RV stress, we employed a surgical model of pulmonary artery constriction (PAC) and measured the RV response over a 7-day period using a high-frequency ultrasound microimaging system. Sham operated mice were used as controls. Images were acquired in lightly anesthetized mice at baseline (before surgery), day 0 (immediately post-surgery), day 3, and day 7 (post-surgery). Data was analyzed offline using software.
Several acoustic windows (B, M, and Color Doppler modes), which can be consistently obtained in mice, allowed for reliable and reproducible measurement of RV structure (including RV wall thickness, end-diastolic&#160;and end-systolic dimensions), and function (fractional area change, fractional shortening, PA peak velocity, and peak pressure gradient) in normal mice and following PAC.
Using this method, the pressure-gradient resulting from PAC was accurately measured in real-time using Color Doppler mode and was comparable to direct pressure measurements performed with a Millar high-fidelity microtip catheter. Taken together, these data demonstrate that RV measurements obtained from various complimentary views using echocardiography are reliable, reproducible and can provide insights regarding RV structure and function. This method will enable a better understanding of the role of RV cardiac dysfunction.

Right ventricle (RV) dysfunction is critical to the pathogenesis of cardiovascular disease, yet limited methodologies are available for its evaluation. Recent advances in ultrasound imaging provide a noninvasive and accurate option for longitudinal RV study. Herein, we detail a step-by-step echocardiographic method using a murine model of RV pressure overload.

Emerging clinical data support the notion that RV dysfunction is critical to the pathogenesis of cardiovascular disease and heart failure1-3. Moreover, ...

A Rat Model of Ventricular Fibrillation and Resuscitation by Conventional Closed-chest Technique

A rat model of electrically-induced ventricular fibrillation followed by cardiac resuscitation using a closed chest technique that incorporates the basic components of cardiopulmonary resuscitation in humans is herein described. The model was developed in 1988 and has been used in approximately 70 peer-reviewed publications examining a myriad of resuscitation aspects including its physiology and pathophysiology, determinants of resuscitability, pharmacologic interventions, and even the effects of cell therapies. The model featured in this presentation includes: (1) vascular catheterization to measure aortic and right atrial pressures, to measure cardiac output by thermodilution, and to electrically induce ventricular fibrillation; and (2) tracheal intubation for positive pressure ventilation with oxygen enriched gas and assessment of the end-tidal CO2. A typical sequence of intervention entails: (1) electrical induction of ventricular fibrillation, (2) chest compression using a mechanical piston device concomitantly with positive pressure ventilation delivering oxygen-enriched gas, (3) electrical shocks to terminate ventricular fibrillation and reestablish cardiac activity, (4) assessment of post-resuscitation hemodynamic and metabolic function, and (5) assessment of survival and recovery of organ function. A robust inventory of measurements is available that includes &#8211; but is not limited to &#8211; hemodynamic, metabolic, and tissue measurements. The model has been highly effective in developing new resuscitation concepts and examining novel therapeutic interventions before their testing in larger and translationally more relevant animal models of cardiac arrest and resuscitation.

This article describes a rat model of electrically-induced ventricular fibrillation and resuscitation by chest compression, ventilation, and delivery of electrical shocks that simulates an episode of sudden cardiac arrest and conventional cardiopulmonary resuscitation. The model enables gathering insights on the pathophysiology of cardiac arrest and exploration of new resuscitation strategies.

A rat model of electrically-induced ventricular fibrillation followed by cardiac resuscitation using a closed chest technique that incorporates the basic ...

Cardiac Catheterization in Mice to Measure the Pressure Volume Relationship: Investigating the Bowditch Effect

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these strategies is to examine their effects on heart function. There are several techniques to measure in vivo cardiac mechanics (e.g., echocardiography, pressure/volume relations, etc.). Compared to echocardiography, real-time left ventricular (LV) pressure/volume analysis via catheterization is more precise and insightful in assessing LV function. Additionally, LV pressure/volume analysis provides the ability to instantaneously record changes during manipulations of contractility (e.g., &#946;-adrenergic stimulation) and pathological insults (e.g., ischemia/reperfusion injury). In addition to the maximum (+dP/dt) and minimum (-dP/dt) rate of pressure change in the LV, an accurate assessment of LV function via several load-independent indexes (e.g., end systolic pressure volume relationship and preload recruitable stroke work) can be attained. Heart rate has a significant effect on LV contractility such that an increase in the heart rate is the primary mechanism to increase cardiac output (i.e., Bowditch effect). Thus, when comparing hemodynamics between experimental groups, it is necessary to have similar heart rates. Furthermore, a hallmark of many cardiomyopathy models is a decrease in contractile reserve (i.e., decreased Bowditch effect). Consequently, vital information can be obtained by determining the effects of increasing heart rate on contractility. Our and others data has demonstrated that the neuronal nitric oxide synthase (NOS1) knockout mouse has decreased contractility. Here we describe the procedure of measuring LV pressure/volume with increasing heart rates using the NOS1 knockout mouse model.

This article describes the measurement of murine left ventricular function via pressure/volume analysis at different heart rates.

Animal models that mimic human cardiac disorders have been created to test potential therapeutic strategies. A key component to evaluating these ...

Improvement of a Closed Chest Porcine Myocardial Infarction Model by Standardization of Tissue and Blood Sampling Procedures

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved drug to prevent or reduce myocardial I/R injury. The study and understanding of the pathophysiological mechanisms of myocardial I/R injury is essential to develop successful treatments. Large animal experiments are an important step in translational methods. The porcine model of acute myocardial infarction has been established and described by ourselves and others. We aimed to further improve the value of the model by focusing in detail on the sampling techniques for use in future experiments. Furthermore, we emphasize small but important steps that can affect the quality of the final results. To mimic the clinical situation of myocardial I/R injury, a percutaneous coronary intervention (PCI) catheter was inserted into the left anterior descending coronary artery (LAD) of an anesthetized pig. &#176;&#176;&#176;This model mimics acute myocardial infarction and PCI treatment in humans with the possibility of accurately determining the area at risk as well as the necrotic- and viable ischemic tissue. Here the model was used to investigate the effect of a bicyclic peptide inhibitor of FXIIa. The model can also be modified to allow longer reperfusion times to study later effects of myocardial infarction.

Here we demonstrate a protocol to standardize sampling procedures of an established porcine model of acute myocardial infarction in order to increase its translational value in the understanding of the pathophysiology of myocardial ischemia/reperfusion injury and to test novel drug candidates.

Myocardial ischemia reperfusion (I/R) injury contributes to almost half of the necrotic area after myocardial infarction. To date there is no approved ...

A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to perform a partial thoracotomy to visualize the transverse aortic arch. However, the surgical trauma caused by the thoracotomy in open-chest models changes the respiratory physiology as the ribs are dissected and left unattached after chest closure. To prevent this, we established a minimally invasive, closed chest approach via lateral thoracotomy. Herein we approach the aortic arch via the 2nd intercostal space without entering the chest cavities, leaving the mouse with a less traumatic injury to recover from. We perform this operation using standard laboratory settings for open chest TAC procedures with equal survival rates. Apart from maintaining physiological breathing patterns due to the closed chest approach, the mice seem to benefit by showing rapid recovery, as the less invasive technique appears to facilitate a fast healing process and to reduce immune response after trauma.

Here, we present a protocol of Transverse Aortic constriction (TAC) via a lateral thoracotomy. This technique is a minimally invasive, closed chest surgical procedure aiming to simulate pressure overload and heart failure in mice utilizing standard TAC laboratory settings.

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to ...

Real-time Pressure-volume Analysis of Acute Myocardial Infarction in Mice

Acute myocardial infarction can lead to acute heart failure and cardiogenic shock. The evaluation of hemodynamics is critical for the evaluation of any potential therapeutic approach directed against acute left ventricular (LV) dysfunction. Current imaging modalities (e.g., echocardiography and magnetic resonance imaging) have several limitations since data on LV pressure cannot directly be measured. LV catheterization in mice undergoing coronary artery occlusion could serve as a novel method for a real-time evaluation of LV function.
At the beginning of the procedure, mice were anesthetized followed by endotracheal intubation. For LV catheterization, the right carotid artery was exposed via middle-neck incision. The catheter was introduced and placed into the LV cavity. Left thoracotomy was conducted and the left main coronary artery (LCA) was ligated. To induce reperfusion, the suture was released after 45 min. Pressure-volume data was recorded at all times.
Ligation of the LCA caused a decrease in LV systolic function as evidenced by a 30% reduction in stroke volume, LV ejection fraction (EF), and cardiac output. Maximum dP/dt as a parameter for LV contractility was also significantly reduced and diastolic function was severely impaired (minimum dP/dt -40%). Reperfusion over a period of 20 min did not lead to a complete recovery of LV function.
Real-time pressure-volume analysis served as a valid procedure for monitoring cardiac function during acute myocardial infarction in mice. Maintaining stable anesthesia and a standardized surgical approach was crucial to ensure valid results. As the early phase of acute myocardial infarction is critical for morbidity and mortality, the delineated method could be beneficial for preclinical evaluation of new strategies for cardioprotection.

Acute myocardial infarction in mice induces acute but incompletely characterized changes in left ventricular (LV) function. LV catheterization in mice undergoing coronary artery occlusion serves as a novel method for a real-time evaluation of LV function.

Acute myocardial infarction can lead to acute heart failure and cardiogenic shock. The evaluation of hemodynamics is critical for the evaluation of any ...

Assessment of Respiratory Function in Conscious Mice by Double-chamber Plethysmography

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to non-invasively assess some features of the respiratory function in humans as well as in laboratory animals. The present article focuses on the application of the double-chamber plethysmography (DCP) in small animals. It provides background information on the methodology as well as a detailed step-by-step procedure to successfully assess respiratory function in conscious, spontaneously breathing animals in a non-invasive manner. The DCP can be used to monitor the respiratory function of multiple animals in parallel, as well as to identify changes induced by aerosolized substances over a chosen time period and in a repeated manner. Experiments on control and allergic mice are used herein to demonstrate the utility of the technique, explain the associated outcome parameters, as well as to discuss the related advantages and shortcomings. Overall, the DCP provides valid and theoretically sound readouts that can be trusted to evaluate the respiratory function of conscious small animals both at baseline and after challenges with aerosolized substances.

The objective of the present article is to provide a detailed description of the recommended procedures to evaluate respiratory function in conscious mice by double-chamber plethysmography.

Air volume changes created by a conscious subject breathing spontaneously within a body box are at the basis of plethysmography, a technique used to ...

Echocardiographic Assessment of Cardiac Anatomy and Function in Adult Rats

The use of experimental animal models has become crucial in cardiovascular science. Most studies using rodent models are focused on two-dimensional imaging to study the cardiac anatomy of the left ventricle and M-mode echo to assess its dimensions. However, this could limit a comprehensive study. Herein, we describe a protocol that allows an assessment of the heart chamber size, left ventricular function (systolic and diastolic) and valvular function. A conventional medical ultrasound machine was used in this protocol and different echo views were obtained through left parasternal, apical and suprasternal windows. In the left parasternal window, the long and short axis were acquired to analyze left chamber dimensions, right ventricle and pulmonary artery dimensions, and mitral, pulmonary and aortic valve function. The apical window allows the measurement of heart chamber dimensions and evaluation of systolic and diastolic parameters. It also allows Doppler assessment with detection and quantification of heart valve disturbances (regurgitation or stenosis). Different segments and walls of the left ventricle are visualized throughout all views. Finally, the ascending aorta, aortic arch, and descending aorta can be imaged through the suprasternal window. A combination of ultrasound imaging, Doppler flow and tissue Doppler assessment have been obtained to study cardiac morphology and function. This represents an important contribution to improve the assessment of cardiac function in adult rats with impact for research using these animal models.

A non-invasive protocol for transthoracic echocardiography assessment of cardiac anatomy and function for adult rats is presented in the current study. The heart valves, all four cardiac chambers and the ascending aorta, aortic arch and descending aorta are studied in detail.

The use of experimental animal models has become crucial in cardiovascular science. Most studies using rodent models are focused on two-dimensional ...

Methods Article

Biventricular Assessment of Cardiac Function and Pressure-Volume Loops by Closed-Chest Catheterization in Mice