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

Summary

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.

Abstract

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.

Introduction

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 scans^1,2.

In experimental models, however, available modalities for assessing the physiological abnormalities related to HFpEF are limited^3,4. 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 features^7,8,9.

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.

Protocol

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

1. 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).
2. 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 module^3,11 (see also the manufacturer's instructions).

2. Preparing an animal for catheterization

1. Anesthesia and ventilation
   1. 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.
   2. 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 °C and 40 °C upon the induction of anesthesia.
   3. Shave the surgical area. Then, disinfect the surgical site with three alternating rounds of betadine and alcohol.
   4. 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.
   5. 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).
2. Central venous (CV) catheter insertion and fluid injection
   1. Locate the internal jugular vein beneath the sternocleidomastoid muscle³.
   2. 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.
   3. Administer a bolus infusion of 5-6 µL/g of body weight of 10% albumin/NaCl over 3 min, followed by a constant infusion rate of 5-10 µL/min.
      ​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.

3. Surgical procedure for left ventricular catheterization (open chest approach)

1. Shave the surgical area of the anesthetized mouse. Then, disinfect the surgical site with three alternating rounds of betadine and alcohol.
2. 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.
3. Cut through the chest wall laterally on both sides using electrical cautery (see the Table of Materials).
4. Expose the heart by cutting through the diaphragm, and remove the pericardium gently from the heart using forceps.
5. 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.
6. Adjust the catheter position so that a square-shaped pressure-volume loop is obtained.
7. 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.
   ​NOTE: Adequate heart exposure facilitates the procedure and helps to obtain a clear view.

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.

1. 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).
2. 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.
   ​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.

5. Transesophageal pacing

1. 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).
2. Slow the HR below 400 beats/min using 20 mg/kg of ivabradine (see the Table of Materials) administered intraperitoneally.
3. 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.

6. Saline calibration and aortic flow calibration

1. Inactivate the ventilator, and administer a 5-10 µL of hypertonic saline solution intravenously through the CV catheter.
2. Document the fluctuations in pressure and volume during the saline injection, and calculate the Vp value using PowerLab^3,11.
3. Repeat the saline calibration to enhance the accuracy and reproducibility.
4. Turn the mouse onto its left side in order not to disturb the volume signal.
5. Make a lateral thoracotomy between Th3 to Th5 toward the spine, and gently dissect a small part of the descending aorta with forceps.
6. Place a vascular flow probe (see the Table of Materials) over the aorta to measure the cardiac output.
   ​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.

7. Euthanasia

1. 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.

Results

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-...

Discussion

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...

Materials

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