&#65279;This work was supported by research grants from the Fukuda Foundation for Medical Technology (to E.T. and G. N.) and the JSPS KAKENHI Scientific Research Grant-in-Aid 21K08047 (to E.T.).

This animal protocol was approved by the Institutional Animal Care and Use Committee and followed the regulations for animal experiments and related activities at the University of Tokyo. For the present study, 8-12 week old male C57/Bl6J mice were used. The animals were obtained from a commercial source (see the Table of Materials). A model of HFpEF was established by administering a high-fat diet for 15 weeks in conjunction with NG-nitro-L-arginine methyl ester, as described previously10.
1. Catheter preparations and pressure/volume calibration
<ol>
	<li>Place a conductance catheter in normal saline, and attach it to a module consisting of the PowerLab 8/35 and a pressure-volume unit (MPVS module, see the Table of Materials).</li>
	<li>Electronically calibrate the pressure and volume through the recording of predetermined pressure (0 mmHg and 100 mmHg) and volume parameters (these vary between MPVS modules) on the MPVS module3,11&#160;(see also the manufacturer&#39;s instructions).</li>
</ol>
2. Preparing an animal for catheterization
<ol>
	<li>Anesthesia and ventilation
	<ol>
		<li>Administer an intraperitoneal injection of 5 mg/kg of etomidate and 500 mg/kg of urethane (see the Table of Materials) 5-10 min prior to intubation. 
		NOTE: Urethane, while effective as an anesthetic agent in animal studies, is suspected to be carcinogenic to humans. Therefore, when urethane is necessary for the achievement of experimental objectives and no alternative agents suffice, it must be handled with caution. Appropriate protective measures, such as wearing gloves and masks and utilizing a fume hood during preparation, are mandated. As a possible alternative, ketamine (80 mg/kg, ip) might be employed.</li>
		<li>Place the mouse in an anesthesia chamber previously saturated with 2% isoflurane, and transfer the animal to a pre-warmed heating pad maintained between 38 &#176;C and 40 &#176;C upon the induction of anesthesia.</li>
		<li>Shave the surgical area. Then, disinfect the surgical site with three alternating rounds of betadine and alcohol.</li>
		<li>Make a horizontal incision (1-2 cm) in the neck, excise the tracheal muscle, and expose the trachea. Pass a surgical 2-0 silk suture beneath the trachea, elevate it, and make a small incision (1-2 mm) to open it.</li>
		<li>Insert an endotracheal tube into the trachea, and connect it to a ventilator that delivers a mixture of 100% oxygen and 2% isoflurane (reduced to 0.5% to 1% later).</li>
	</ol>
	</li>
	<li>Central venous (CV) catheter insertion and fluid injection
	<ol>
		<li>Locate the internal jugular vein beneath the sternocleidomastoid muscle3.</li>
		<li>Insert the central venous catheter, consisting of PE-10 silastic tubing (see the Table of Materials) attached to a 30 G needle, into the jugular vein.</li>
		<li>Administer a bolus infusion of 5-6 &#181;L/g of body weight of 10% albumin/NaCl over 3 min, followed by a constant infusion rate of 5-10 &#181;L/min. 
		&#8203;NOTE: This step is crucial for preventing hypotension resulting from the peripheral vasodilation caused by the anesthesia. The internal jugular vein is located between the sternocleidomastoid muscle and the carotid artery, and it appears darker in color than the artery.</li>
	</ol>
	</li>
</ol>
3. Surgical procedure for left ventricular catheterization (open chest approach)
<ol>
	<li>Shave the surgical area of the anesthetized mouse. Then, disinfect the surgical site with three alternating rounds of betadine and alcohol.</li>
	<li>Confirm the depth of anesthesia by performing a toe pinch. Then, make a horizontal incision (2-3 cm) below the xiphoid process, and separate the skin from the chest wall using blunt scissors.</li>
	<li>Cut through the chest wall laterally on both sides using electrical cautery (see the Table of Materials).</li>
	<li>Expose the heart by cutting through the diaphragm, and remove the pericardium gently from the heart using forceps.</li>
	<li>Insert a 27 G needle into the apex of the left ventricle (LV), and retrogradely insert a conductance catheter into the LV via the puncture hole.</li>
	<li>Adjust the catheter position so that a square-shaped pressure-volume loop is obtained.</li>
	<li>Verify that the catheter does not contact the papillary muscle when changes in loading conditions occur by checking the shape of the PV loop during inferior vena cava (IVC) occlusion. 
	&#8203;NOTE: Adequate heart exposure facilitates the procedure and helps to obtain a clear view.</li>
</ol>
4. Recording PV loop data and determining the end-systolic pressure-volume relationship (ESPVR) and the end-diastolic pressure-volume relationship (EDPVR)
NOTE: Reducing the preload by IVC occlusion enables the determination of the ESPVR and EDPVR.
<ol>
	<li>Record and analyze the baseline pressure-volume (PV) loop with LabChart software (see the Table of Materials), PowerLab, and the MPVS module after signal stabilization (5-10 min after canulation).</li>
	<li>Perform IVC occlusion by compressing the IVC with forceps, and record the PV loop for at least 20 cardiac cycles during the IVC occlusion. Determine the ESPVR by fitting a linear regression line through the end-systolic points of the PV loop and the EDPVR by fitting a curvilinear line through the end-diastolic points of the PV loop using LabChart software. 
	&#8203;NOTE: Stop the ventilator during the IVC occlusion to prevent lung motion artifacts. A paralytic agent like pancuronium (0.5-1 mg/kg) may be helpful when lung motion is excessive and should be used only after a stable anesthetic plane is confirmed.</li>
</ol>
5. Transesophageal pacing
<ol>
	<li>Insert a 2-Fr tetrapolar electrode catheter into the esophagus, connect the catheter to a pulse stimulator (see the Table of Materials), and determine the atrial capture threshold (normally, the stimulus amplitude is 3 mA, and the pulse width is 1 ms).</li>
	<li>Slow the HR below 400 beats/min using 20 mg/kg of ivabradine (see the Table of Materials) administered intraperitoneally.</li>
	<li>Following stabilization, acquire 20 continuous cardiac cycles of PV loops at different pacing rates from 400 beats/min to 700 beats/min, with an increment of 100 beats/min; acquire the cycles over 5 min at each pacing rate.</li>
</ol>
6. Saline calibration and aortic flow calibration
<ol>
	<li>Inactivate the ventilator, and administer a 5-10 &#181;L of hypertonic saline solution intravenously through the CV catheter.</li>
	<li>Document the fluctuations in pressure and volume during the saline injection, and calculate the Vp value using PowerLab3,11.</li>
	<li>Repeat the saline calibration to enhance the accuracy and reproducibility.</li>
	<li>Turn the mouse onto its left side in order not to disturb the volume signal.</li>
	<li>Make a lateral thoracotomy between Th3 to Th5 toward the spine, and gently dissect a small part of the descending aorta with forceps.</li>
	<li>Place a vascular flow probe (see the Table of Materials) over the aorta to measure the cardiac output. 
	&#8203;NOTE: The accurate calculation of the absolute volume requires the use of two types of calibration: saline calibration and aortic flow calibration. It is important to recognize the potential risks associated with a hypertonic saline infusion in animal subjects, as excessive salt loading can result in a decline in contractility.</li>
</ol>
7. Euthanasia
<ol>
	<li>After the study, euthanize the mice under an anesthetic overdose via cervical dislocation. 
	NOTE: To ensure the complete cessation of vital function, a secondary method of euthanasia is employed, such as exsanguination under anesthesia with subsequent cardiac tissue harvesting.</li>
</ol>

Exploring HR-Dependent Cardiac Performance Using an Atrial-Pacing Protocol

<ol>
	<li>Backhaus, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exercise+stress+real-time+cardiac+magnetic+resonance+imaging+for+noninvasive+characterization+of+heart+failure+with+preserved+ejection+fraction.">Exercise stress real-time cardiac magnetic resonance imaging for noninvasive characterization of heart failure with preserved ejection fraction.</a> Circulation. 143 (15), 1484-1498 (2021).</li><li>Borlaug, B. A., Nishimura, R. A., Sorajja, P., Lam, C. S. P., Redfield, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exercise+hemodynamics+enhance+diagnosis+of+early+heart+failure+with+preserved+ejection+fraction.+Circulation.">Exercise hemodynamics enhance diagnosis of early heart failure with preserved ejection fraction. Circulation.</a> Heart Failure. 3 (5), 588-595 (2010).</li><li>Pacher, P., Nagayama, T., Mukhopadhyay, P., B&#225;tkai, S., David, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+cardiac+function+using+pressure-volume+conductance+catheter+technique+in+mice+and+rats.">Measurement of cardiac function using pressure-volume conductance catheter technique in mice and rats.</a> Nature Protocols. 3 (9), 1422-1434 (2008).</li><li>Cingolani, O. H., Kass, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pressure-volume+relation+analysis+of+mouse+ventricular+function.">Pressure-volume relation analysis of mouse ventricular function.</a> American Journal of Physiology - Heart and Circulatory Physiology. 301 (6), 2198-2206 (2011).</li><li>Numata, G., 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=A+pacing-controlled+protocol+for+frequency-diastolic+relations+distinguishes+diastolic+dysfunction+specific+to+a+mouse+HFpEF+model.">A pacing-controlled protocol for frequency-diastolic relations distinguishes diastolic dysfunction specific to a mouse HFpEF model.</a> American Journal of Physiology - Heart and Circulatory Physiology. 323 (3), H523-H527 (2022).</li><li>Pi&#241;a, I. L., 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=Exercise+and+heart+failure.">Exercise and heart failure.</a> Circulation. 107 (8), 1210-1225 (2003).</li><li>Georgakopoulos, D., Kass, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Minimal+force-frequency+modulation+of+inotropy+and+relaxation+of+in+situ+murine+heart.">Minimal force-frequency modulation of inotropy and relaxation of in situ murine heart.</a> Journal of Physiology. 534 (2), 535-545 (2001).</li><li>Takimoto, E., 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=Frequency-+and+afterload-dependent+cardiac+modulation+in+vivo+by+troponin+I+with+constitutively+active+protein+kinase+A+phosphorylation+sites.">Frequency- and afterload-dependent cardiac modulation in vivo by troponin I with constitutively active protein kinase A phosphorylation sites.</a> Circulation Research. 94 (4), 496-504 (2004).</li><li>Meyer, M., Lewinter, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Heart+rate+and+heart+failure+with+preserved+ejection+fraction:+Time+to+slow+&#946;-blocker+use?+Circulation.">Heart rate and heart failure with preserved ejection fraction: Time to slow &#946;-blocker use? Circulation.</a> Heart Failure. 12 (8), 006213 (2019).</li><li>Schiattarella, G. G., 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=Nitrosative+stress+drives+heart+failure+with+preserved+ejection+fraction.">Nitrosative stress drives heart failure with preserved ejection fraction.</a> Nature. 568 (7752), 351-356 (2019).</li><li>Abraham, D., Mao, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+pressure-volume+loop+analysis+using+conductance+catheters+in+mice.">Cardiac pressure-volume loop analysis using conductance catheters in mice.</a> Journal of Visualized Experiments. (103), e52942 (2015).</li><li>Zhang, B., Davis, J. P., Ziolo, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+catheterization+in+mice+to+measure+the+pressure+volume+relationship:+Investigating+the+Bowditch+effect.">Cardiac catheterization in mice to measure the pressure volume relationship: Investigating the Bowditch effect.</a> Journal of Visualized Experiments. (100), e52618 (2015).</li><li>Townsend, D. W. <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. (111), e53810 (2016).</li><li>Georgakopoulos, D., Kass, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimation+of+parallel+conductance+by+dual-frequency+conductance+catheter+in+mice.">Estimation of parallel conductance by dual-frequency conductance catheter in mice.</a> American Journal of Physiology - Heart and Circulatory Physiology. 279 (1), H47 (2000).</li></ol>

There are no competing financial interests.

We present a methodology to assess pressure-volume relationships with the application of transesophageal pacing. Exercise intolerance is one of the key characteristics of HFpEF, yet there are no techniques available for the evaluation of cardiac function in mice during exercise. Our pacing protocol offers a valuable tool for detecting diastolic dysfunction, which may not be apparent under resting conditions.
To achieve a PV loop of accurate and consistent quality, the following steps must be m...

Heart failure represents a leading cause of hospitalization and death across the globe, and heart failure with preserved ejection fraction (HFpEF) accounts for around 50% of all heart failure diagnoses. HFpEF is characterized by diastolic dysfunction and impaired exercise tolerance, and the associated hemodynamic abnormalities, such as diastolic dysfunction, can be clearly detected through exercise-stressed hemodynamic testing or MRI scans1,2.
In experimental models, however, available modalities for assessing the physiological abnormalities related to HFpEF are limited3,4. Treadmill exercise testing (TMT) is used to determine running time and distance, which might reflect exercise-stress cardiac hemodynamics; however, this method is susceptible to interference from extraneous variables such as the body weight, skeletal muscle strength, and mental status.
To circumvent these limitations, we have devised an atrial-pacing protocol that detects subtle but crucial changes in diastolic performance based on the heart rate (HR) and have validated its usefulness in a mouse model of HFpEF5. Several physiological factors contribute to exercise-related cardiac function, including the sympathetic nerve and catecholamine response, peripheral vasodilation, the endothelial response, and the heart rate6. Among these, however, the HR-pressure relationship (also called the Bowditch effect) is known as a critical determinant of cardiac physiological features7,8,9.
The protocol involves performing a conventional pressure-volume analysis at baseline to assess the systolic and diastolic function, including parameters such as the rate of pressure development (dp/dt), the end-systolic pressure-volume relationship (ESPVR), and the end-diastolic pressure-volume relationship (EDPVR). However, it should be noted that these parameters are influenced by the HR, which can vary between animals due to differences in their intrinsic heart rate. Additionally, the effects of anesthesia on the HR should also be considered. To address this, the HR was standardized by administering atrial pacing concomitantly with ivabradine, and cardiac parameter measurements were performed at incremental heart rates. Notably, the HR-dependent cardiac response distinguished HFpEF mice from the control group mice, while no significant differences were observed in the baseline PV loop measurements (using the intrinsic heart rate)5.
While this pacing protocol may seem relatively complicated, its success rate exceeds 90% when it is well understood. This protocol would provide a useful way to study the underlying mechanisms of diastolic dysfunction in HFpEF mouse models and help in the development of new treatments for this condition.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-0 silk suture, sterlie</td>
			<td>Alfresa Pharma Corporation, Osaka, Japan</td>
			<td>62-9965-57</td>
			<td>Surgical Supplies</td>
		</tr>
		<tr>
			<td>2-Fr tetrapolar electrode catheter</td>
			<td>Fukuda Denshi, Japan and UNIQUE MEDICAL, Japan</td>
			<td>custom-made</td>
			<td>Surgical Supplies</td>
		</tr>
		<tr>
			<td>Albumin Bovine Serum</td>
			<td>Nacalai Tesque, Inc., Kyoto, Japan</td>
			<td>01859-47</td>
			<td>Miscellaneous</td>
		</tr>
		<tr>
			<td>C57/BI6J mouse</td>
			<td>Jackson Laboratory</td>
			<td></td>
			<td>animals</td>
		</tr>
		<tr>
			<td>Conductance catheter</td>
			<td>Millar Instruments, Houston, TX</td>
			<td>PVR 1035</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical cautery, Electrocautery Knife Kit</td>
			<td>ellman-Japan,Osaka, Japan</td>
			<td>1-1861-21</td>
			<td>Surgical Supplies</td>
		</tr>
		<tr>
			<td>Etomidate</td>
			<td>Tokyo Chemical Industory Co., Ltd., Tokyo Japan</td>
			<td>E0897</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>Grass Instrument S44G Square Pulse Stimulator</td>
			<td>Astro-Med, West Warwick, RI</td>
			<td></td>
			<td>Pacing equipment</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Viatris Inc., Tokyo, Japan</td>
			<td>8803998</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>Ivabradine</td>
			<td>Tokyo Chemical Industory Co., Ltd., Tokyo Japan</td>
			<td>I0847</td>
			<td>Miscellaneous</td>
		</tr>
		<tr>
			<td>LabChart software</td>
			<td>ADInstruments, Sydney, Australia</td>
			<td>LabChart 7</td>
			<td>Hemodynamic equipment</td>
		</tr>
		<tr>
			<td>MPVS Ultra</td>
			<td>Millar Instruments, Houston, TX</td>
			<td>PL3516B49</td>
			<td>Hemodynamic equipment</td>
		</tr>
		<tr>
			<td>Pancronium bromide</td>
			<td>Sigma Aldrich Co., St. Louis, MO</td>
			<td>15500-66-0</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>PE10 polyethylene tube</td>
			<td>Bio Research Center&#160; Co. Ltd., Tokyo, Japan</td>
			<td>62101010</td>
			<td>Surgical Supplies</td>
		</tr>
		<tr>
			<td>PowerLab 8/35</td>
			<td>ADInstruments, Sydney, Australia</td>
			<td>PL3508/P</td>
			<td>Hemodynamic equipment</td>
		</tr>
		<tr>
			<td>PVR 1035</td>
			<td>Millar Instruments, Houston, TX</td>
			<td>842-0002</td>
			<td>Hemodynamic equipment</td>
		</tr>
		<tr>
			<td>Urethane (Ethyl Carbamate)</td>
			<td>Wako Pure Chemical Industries, Ltd., Osaka, Japan</td>
			<td>050-05821</td>
			<td>Anesthetic</td>
		</tr>
		<tr>
			<td>Vascular Flow Probe</td>
			<td>Transonic, Ithaca, NY</td>
			<td>MA1PRB</td>
			<td>Surgical Supplies</td>
		</tr>
	</tbody>
</table>

a pacing-controlled procedure for the assessment of heart rate-dependent diastolic functions in murine heart failure models

Heart failure with preserved ejection fraction (HFpEF) is a condition characterized by diastolic dysfunction and exercise intolerance. While exercise-stressed hemodynamic tests or MRI can be used to detect diastolic dysfunction and diagnose HFpEF in humans, such modalities are limited in basic research using mouse models. A treadmill exercise test is commonly used for this purpose in mice, but its results can be influenced by body weight, skeletal muscle strength, and mental state. Here, we describe an atrial-pacing protocol to detect heart rate (HR)-dependent changes in diastolic performance and validate its usefulness in a mouse model of HFpEF. The method involves anesthetizing, intubating, and performing pressure-volume (PV) loop analysis concomitant with atrial pacing. In this work, a conductance catheter was inserted via a left ventricular apical approach, and an atrial pacing catheter was placed in the esophagus. Baseline PV loops were collected before the HR was slowed with ivabradine. PV loops were collected and analyzed at HR increments ranging from 400 bpm to 700 bpm via atrial pacing. Using this protocol, we clearly demonstrated HR-dependent diastolic impairment in a metabolically induced HFpEF model. Both the relaxation time constant (Tau) and the end-diastolic pressure-volume relationship (EDPVR) worsened as the HR increased compared to control mice. In conclusion, this atrial pacing-controlled protocol is useful for detecting HR-dependent cardiac dysfunctions. It provides a new way to study the underlying mechanisms of diastolic dysfunction in HFpEF mouse models and may help develop new treatments for this condition.

The baseline PV loop data are displayed in Figure 1 and Table 1. At baseline (in the absence of pacing), there were no significant differences in diastolic parameters such as the relaxation time constant (Tau), the minimum rate of pressure change (dP/dt min), and EDPVR between the control and HFpEF mice. However, the HFpEF mice exhibited higher blood pressure and arterial elastance (Ea), as shown in Figure 1, and demonstrated a typical mountain-...

Watch this Scientific Journal Video about Investigating HR-Dependent Cardiac Function in Mouse Models Through a Novel Atrial-Pacing Approach at JoVE.com

Author Spotlight: Investigating HR-Dependent Cardiac Function in Mouse Models Through a Novel Atrial-Pacing Approach 

The present protocol describes obtaining the pressure-volume relationship through transesophageal pacing, which serves as a valuable tool in evaluating diastolic function&#160;in mouse models of heart failure.

A Pacing-Controlled Procedure for the Assessment of Heart Rate-Dependent Diastolic Functions in Murine Heart Failure Models

Heart failure with preserved ejection fraction (HFpEF) is a condition characterized by diastolic dysfunction and exercise intolerance. While ...

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1.2K Views.  The University of Tokyo Hospital. The present protocol describes obtaining the pressure-volume relationship through transesophageal pacing, which serves as a valuable tool in evaluating diastolic function in mouse models of heart failure.

Video: Author Spotlight: Investigating HR-Dependent Cardiac Function in Mouse Models Through a Novel Atrial-Pacing Approach 

Implantation of Engineered Tissue in the Rat Heart

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in common laboratory rats or mice and these quite frequently serve as an intermediate step between bench-top experiments and large animal testing or human trials. Given that rodents provide an established, cost-effective, and physiologically-relevant model system in which to test novel combinations of scaffolding materials and cells, they are particularly well-suited for cardiovascular tissue engineering studies. Presently, we describe an open-heart surgical procedure to implant engineered tissue containing myogenic progenitor cells in the atrioventricular (AV) groove of a rat heart. These implants are intended to create an electrical conduit between the right atrium and right ventricle with the ultimate goal of providing an alternative treatment to conventional pacemaker implantation in pediatric patients with complete heart block[1]. The engineered tissue is implanted in the AV-groove by means of a thoracotomy. For our purposes, Lewis rats are anesthetized and invasively ventilated to maintain positive airway pressure during the sterile surgical procedure. The approach to the heart is performed by a right thoracotomy through an antero-lateral incision at the 5th intercostal space. The tissue construct is fixed in the AV groove using a single 7-0 Prolene suture and positioned between the right ventricle and atrium at the ventral portion of the heart. The epicardium is partially removed to allow direct contact between the recipient myocardial cells and those contained in the engineered tissue. Following implantation, the chest wall is closed in layers, any pneumothorax is evacuated, and the animal is extubated and treated with analgesic.

Here, we describe a cardiac surgical procedure to implant engineered tissue in the atrioventricular (AV)-groove of an adult Lewis rat.

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in ...

Murine Renal Transplantation Procedure

Renal orthotopic transplantation in mice is a technically challenging procedure. Although the first kidney transplants in mice were performed by Russell et al over 30 years ago (1) and refined by Zhang et al years later (2), few people in the world have mastered this procedure. In our laboratory we have successfully performed 1200 orthotopic kidney transplantations with &gt; 90% survival rate. The key points for success include stringent control of reperfusion injury, bleeding and thrombosis, both during the procedure and post-transplantation, and use of 10-0 instead of 11-0 suture for anastomoses.
Post-operative care and treatment of the recipient is extremely important to transplant success and evaluation. All renal graft recipients receive antibiotics in the form of an injection of penicillin immediately post-transplant and sulfatrim in the drinking water continually. Overall animal health is evaluated daily and whole blood creatinine analyses are performed routinely with a portable I-STAT machine to assess graft function.

Renal transplantation in mice is a technically challenging procedure that requires careful post-operative care and treatment for success.

Renal orthotopic transplantation in mice is a technically challenging procedure. Although the first kidney transplants in mice were performed by Russell ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

Visualizing the Beating Heart in Drosophila

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in myogenic hearts. A key step in examining heart function in the fly is finding a way to access the heart in a manner that preserves its myogenic function while still allowing the beating heart organ to be observed and recorded. Two different methods for observing and recording the beating heart in both larva and adult Drosophila are described here. Our semi-intact preparation using adult flies allows clear visualization of the abdominal heart without interference from the pigmented cuticle and overlying fat bodies. To record larval heart beats it is necessary to immobilize the larva, which minimizes body wall movements thereby reducing heart movements that are not associated with myocardial contractions. Our methodologies produce stable adult and larval heart preparations that can beat for hours at rates of 1-3 Hz.

Visualizing the Beating Heart in Drosophila

Technique required for visualizing the beating heart in larval and adult Drosophila are presented. Each life stage requires a different methodology.

The Drosophila heart has recently emerged as a good model system for examining the genetic, cellular, and molecular mechanisms underlying function in ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Quantification of &gamma;H2AX Foci in Response to Ionising Radiation

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA lesions with respect to survival and preservation of genomic integrity. An early response to the induction of DSBs is phosphorylation of the H2A histone variant, H2AX, at the serine-139 residue, in the highly conserved C-terminal SQEY motif, forming &gamma;H2AX1. Following induction of DSBs, H2AX is rapidly phosphorylated by the phosphatidyl-inosito 3-kinase (PIKK) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)2. Typically, only a few base-pairs (bp) are implicated in a DSB, however, there is significant signal amplification, given the importance of chromatin modifications in DNA damage signalling and repair. Phosphorylation of H2AX mediated predominantly by ATM spreads to adjacent areas of chromatin, affecting approximately 0.03% of total cellular H2AX per DSB2,3. This corresponds to phosphorylation of approximately 2000 H2AX molecules spanning ~2 Mbp regions of chromatin surrounding the site of the DSB and results in the formation of discrete &gamma;H2AX foci which can be easily visualized and quantitated by immunofluorescence microscopy2. The loss of &gamma;H2AX at DSB reflects repair, however, there is some controversy as to what defines complete repair of DSBs; it has been proposed that rejoining of both strands of DNA is adequate however, it has also been suggested that re-instatement of the original chromatin state of compaction is necessary4-8. The disappearence of &gamma;H2AX involves at least in part, dephosphorylation by phosphatases, phosphatase 2A and phosphatase 4C5,6. Further, removal of &gamma;H2AX by redistribution involving histone exchange with H2A.Z has been implicated7,8. Importantly, the quantitative analysis of &gamma;H2AX foci has led to a wide range of applications in medical and nuclear research. Here, we demonstrate the most commonly used immunofluorescence method for evaluation of initial DNA damage by detection and quantitation of &gamma;H2AX foci in &gamma;-irradiated adherent human keratinocytes9.

Quantification of &#947;H2AX Foci in Response to Ionising Radiation

Quantification of DNA double-strand streaks using &gamma;H2AX formation as a molecular marker has become an invaluable tool in radiation biology. Here we demonstrate the use of an immunofluorescence assay for quantification of &gamma;H2AX foci after exposure of cells to radiation.

DNA double-strand breaks (DSBs), which are induced by either endogenous metabolic processes or by exogenous sources, are one of the most critical DNA ...

Evaluation of the Spatial Distribution of &gamma;H2AX following Ionizing Radiation

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone H2AX1,2. This phosphorylation of H2AX is mediated by the phosphatidyl-inosito 3-kinase (PI3K) family of proteins, ataxia telangiectasia mutated (ATM), DNA-protein kinase catalytic subunit and ATM and RAD3-related (ATR)3. The phosphorylated form of H2AX, referred to as &gamma;H2AX, spreads to adjacent regions of chromatin from the site of the DSB, forming discrete foci, which are easily visualized by immunofluorecence microscopy3. Analysis and quantitation of &gamma;H2AX foci has been widely used to evaluate DSB formation and repair, particularly in response to ionizing radiation and for evaluating the efficacy of various radiation modifying compounds and cytotoxic compounds4.
Given the exquisite specificity and sensitivity of this de novo marker of DSBs, it has provided new insights into the processes of DNA damage and repair in the context of chromatin. For example, in radiation biology the central paradigm is that the nuclear DNA is the critical target with respect to radiation sensitivity. Indeed, the general consensus in the field has largely been to view chromatin as a homogeneous template for DNA damage and repair. However, with the use of &gamma;H2AX as molecular marker of DSBs, a disparity in &gamma;-irradiation-induced &gamma;H2AX foci formation in euchromatin and heterochromatin has been observed5-7. Recently, we used a panel of antibodies to either mono-, di- or tri- methylated histone H3 at lysine 9 (H3K9me1, H3K9me2, H3K9me3) which are epigenetic imprints of constitutive heterochromatin and transcriptional silencing and lysine 4 (H3K4me1, H3K4me2, H3K4me3), which are tightly correlated actively transcribing euchromatic regions, to investigate the spatial distribution of &gamma;H2AX following ionizing radiation8. In accordance with the prevailing ideas regarding chromatin biology, our findings indicated a close correlation between &gamma;H2AX formation and active transcription9. Here we demonstrate our immunofluorescence method for detection and quantitation of &gamma;H2AX foci in non-adherent cells, with a particular focus on co-localization with other epigenetic markers, image analysis and 3D-modeling.

Evaluation of the Spatial Distribution of &#947;H2AX following Ionizing Radiation

Microscopic analysis of &gamma;H2AX foci, which form following the phosphorylation of H2AX at Ser-139 in response to DNA double-strand breaks, has become an invaluable tool in radiation biology. Here we used an antibody to mono-methylated histone H3 at lysine 4 as an epigenetic marker of actively transcribing euchromatin, to evaluate the spatial distribution of radiation-induced &gamma;H2AX formation within the nucleus.

An early molecular response to DNA double-strand breaks (DSBs) is phosphorylation of the Ser-139 residue within the terminal SQEY motif of the histone ...

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. Thus, protocols to isolate and culture individual organs of interest are essential to provide a method of both visualizing changes in development and allowing novel treatment strategies. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel. This substrate provides enough support to maintain the three-dimensional structure but is flexible enough to allow continued contraction. In brief, hearts are excised from the embryo and placed in a mixture of cold Matrigel diluted 1:1 with growth medium. After the diluted Matrigel solidifies, growth medium is added to the culture dish. Hearts excised as late as embryonic day 16.5 were viable for four days post-dissection. Analysis of the coronary plexus shows that this method does not disrupt coronary vascular development. Thus, we present a novel method for long-term culture of embryonic hearts.

A Novel Ex vivo Culture Method for the Embryonic Mouse Heart

Developmental studies in the mouse are hampered by the inaccessibility of the embryo during gestation. To promote the long-term culture of the embryonic heart at late stages of gestation, we developed a protocol in which the excised heart is cultured in a semi-solid, dilute Matrigel.

Developmental studies in the mouse are hampered  by the inaccessibility of the embryo during gestation. Thus, protocols to  isolate and culture individual ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Isolation of Murine Valve Endothelial Cells

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and surrounded by a monolayer of valve endothelial cells (VECs). VECs play essential roles in establishing the valve structures during embryonic development, and are important for maintaining life-long valve integrity and function. In contrast to a continuous endothelium over the surface of healthy valve leaflets, VEC disruption is commonly observed in malfunctioning valves and is associated with pathological processes that promote valve disease and dysfunction. Despite the clinical relevance, focused studies determining the contribution of VECs to development and disease processes are limited. The isolation of VECs from animal models would allow for cell-specific experimentation. VECs have been isolated from large animal adult models but due to their small population size, fragileness, and lack of specific markers, no reports of VEC isolations in embryos or adult small animal models have been reported. Here we describe a novel method that allows for the direct isolation of VECs from mice at embryonic and adult stages. Utilizing the Tie2-GFP reporter model that labels all endothelial cells with Green Fluorescent Protein (GFP), we have been successful in isolating GFP-positive (and negative) cells from the semilunar and atrioventricular valve regions using fluorescence activated cell sorting (FACS). Isolated GFP-positive VECs are enriched for endothelial markers, including CD31 and von Willebrand Factor (vWF), and retain endothelial cell expression when cultured; while, GFP-negative cells exhibit molecular profiles and cell shapes consistent with VIC phenotypes. The ability to isolate embryonic and adult murine VECs allows for previously unattainable molecular and functional studies to be carried out on a specific valve cell population, which will greatly improve our understanding of valve development and disease mechanisms.

The ability to isolate heart valve endothelial cells (VECs) is critical for understanding mechanisms of valve development, maintenance, and disease. Here we describe the isolation of VECs from embryonic and adult Tie2-GFP mice using FACS that will allow for studies determining the contribution of VECs in developmental and disease processes.

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and ...