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.

Representative 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-shaped PV loop during ventricular systole. This should be distinguished from a spike caused by direct contact of the ventricular muscle on the pressure transducer (Figure 2). Importantly, using atrial pacing, the diastolic function could be clearly distinguished between the control mice and HFpEF mice⁵ (Figure 3 and Figure 4). In the control group, both the Tau and EDPVR improved as the pacing rate increased, whereas, in the HFpEF group, both the Tau and EDPVR worsened as the HR increased with atrial pacing.

Figure 1: Representative pressure-volume relationship at baseline in the absence of pacing, depicted in a screenshot. The results showed that HFpEF mice exhibited higher arterial elastance and ventricular pressure compared to the control mice. Please click here to view a larger version of this figure.

Figure 2: A representative image of a spike-shaped PV loop. This type of PV loop shape is a result of direct compression of the pressure transducer by the ventricular muscle (showed by the orange arrowhead) and should be excluded from the analysis to maintain accuracy in the results. Please click here to view a larger version of this figure.

Figure 3: Representative chart illustrating the differences in hemodynamic parameters in response to atrial pacing between the heart failure with preserved ejection fraction (HFpEF) model mice and the control mice. The chart clearly distinguishes between the two groups, with the HR ranging from 400 beats per minute to 700 beats per minute. Abbreviations: LVP = left ventricular pressure; dP/dt = first derivative of LVP; EDPVR = end-diastolic pressure-volume relationship; LVV = left ventricular volume; Tau = relaxation time constant. Please click here to view a larger version of this figure.

Figure 4: The hemodynamic response of the diastolic parameters from the pressure-volume loop analysis depicted in terms of the heart rate (HR). In the HFpEF model mice, the diastolic function (Tau and EDPVR) deteriorated as the heart rate increased during atrial pacing. The two-way ANOVA analysis showed a significant main effect of HFpEF (F = 28.95, p < 0.001) and HR (F = 3.035, p = 0.08644) on the EDPVR, as well as a significant interaction effect between group and heart rate (F = 3.938, p = 0.02454). For Tau, there was a significant effect of group (F = 25.56, p < 0.001) and HR (F = 0.1088, p = 0.7425), as well as a significant interaction effect between group and heart rate (F = 3.461, p = 0.03759). The data are displayed as the mean ± standard error. n = 6 mice/group. Please click here to view a larger version of this figure.

Figure 5: Representative illustration of the saline calibration procedure. The infusion of hypertonic saline alters the electrical conductivity of the blood, thus enabling the calculation of the signal component attributed to the surrounding cardiac tissue. The blood pressure should remain stable during the injection, with a slight volume increase (shown in the orange arrow). Abbreviations: LVP = left ventricular pressure; LVV = left ventricular volume Please click here to view a larger version of this figure.

Figure 6: Representative illustration of the proper placement of the transesophageal pacing catheter. The proper placement of the transesophageal pacing catheter enables a narrow QRS rhythm. The blue arrows depict a normal sinus rhythm, and the red arrows show the atrial pacing rhythm. Please click here to view a larger version of this figure.

Figure 7: Representative image of an incorrectly adjusted stimulus amplitude in atrial pacing, resulting in a distorted pressure-volume loop. The stimulation intensity induced unwanted motion artifacts in the conductance signal, depicted as the PV loop with a shaking line (indicated by the arrows). Please click here to view a larger version of this figure.

	Control  (n = 10)	HFpEF (n = 10)	p value
CO (μL/min)	12436.8 ± 938.4	10923.5 ± 1032.7	0.2897
SV (μL)	23.6 ± 1.85	20.5 ± 1.88	0.2515
Ved (μl)	37.6 ± 3.45	34.0 ± 1.32	0.4242
Pes (mmHg)	95.2 ± 3.56	109.3 ± 1.74	0.00032*
Ped (mmHg)	6.16 ± 1.53	6.95 ± 1.22	0.6889
HR (beat/min)	532.4 ± 20.8	534.0 ± 13.9	0.9505
EF (%)	66.5 ± 2.95	63.68 ± 2.37	0.4718
Ea (mmHg/μL)	4.02 ± 0.30	5.90 ± 0.72	0.03224*
dP/dt max (mmHg/s)	10812.1 ± 1042.9	9481.1 ± 262.02	0.2444
dP/dt min (mmHg/s)	-9540.7 ± 748.9	-9003.9 ± 320.0	0.5177
Tau (ms)	7.30 ± 0.50	8.02 ± 0.39	0.268
ESPVR (mmHg/μL)	3.41 ± 0.51	4.69 ± 0.41	0.09147
EDPVR (mmHg/μL)	0.096 ± 0.0061	0.103 ± 0.013	0.6103

Table 1: Baseline cardiac parameters in the control and HFpEF mice. The data are displayed as the mean ± standard error; *p < 0.05 versus control by t-test.

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 meticulously executed^{3,4,5,7,8,11,12,13,14}: (1) the animals must be anesthetized carefully, and a consistent body temperature of 37-37.5 °C must be maintained using a heating pad; (2) the animals must be intubated appropriately, and the ventilation must be controlled effectively; (3) the proper placement of the intravenous access must be ensured; (4) the conductance catheter must be properly positioned within the LV; (5) the transesophageal catheter should be placed judiciously, and appropriate pacing should be ensured; 6) the data acquisition system must be connected with care, and the gain and offset values must be adjusted appropriately; (7) the conductance signals should be calibrated using hypertonic saline; (8) the proper measurement of the aortic flow with a flow probe should be verified; (9) the animals' well-being should be continuously monitored throughout the procedure to minimize any stress- or movement-induced artifacts.

Optimizing the dose of anesthesia is crucial in obtaining a reproducible and high-quality PV loop in mice. Typically, a dose of 800 mg/kg of urethane and 5-10 mg/kg of etomidate is administered. However, in cases of pathological heart failure, it is advisable to administer a lower dose of anesthetic. During the procedure, it is essential to maintain a warm body temperature of 37-38 °C by gently placing the anesthetized animal on a heating pad. This is especially important for mice because a drop in body temperature can cause a significant decrease in the HR. In addition, adequate exposure of the heart is crucial to obtain a clear view and facilitate the procedure. In some cases, cutting the 12th to 11th ribs can be helpful in exposing the heart.

The intubation process should be cautiously performed to avoid damage to the carotid arteries and vagus nerves near the trachea. The ventilator setting must be adjusted based on the animal's body weight using the formulas provided³:

Tidal volume (Vt, mL) = 6.2 × W^1.01 (W = body weight, kg)
Respiratory rate (RR, min⁻¹) = 53.5 × W^−0.26
For example, Vt = 149.4 µL, RR = 140/min in a 25 g mouse.

Before canulation, the venous catheter (with a 30 G needle) must be fully primed with 10% albumin and inserted into the vein at a shallow angle to prevent tearing of the fragile vein walls. The proper positioning of the conductance catheter within the left ventricle (LV) is paramount for obtaining accurate results. The catheter should be aligned with the LV longitudinal axis, with all the electrodes positioned between the LV outflow tract and the apical endocardial border. A stable PV loop with no notches should be obtained throughout the entire procedure, including during the intravenous occlusion, hypertonic saline calibration, and transesophageal pacing. In saline calibration, the LV pressures should be stable during the hypertonic salineinjection, and the beats during the initial wash-in phase of risingvolume signals are used (Figure 5). One needs to be careful not to inject volumes of hypertonic saline higher than 20 µL because hypertonic saline could easily depress the heart function by volume overload.The pacing catheter introduced through the esophagus must be confirmed to be in the proper position through atrial capture (Figure 6), and the stimulus amplitude should be appropriately adjusted (usually 3 mA, with a pulse width of 1 ms). Stronger stimulation would affect the conductance catheter and cause a shaking-shaped PV loop (Figure 7).

The accurate calculation of the absolute volume requires the utilization of two types of calibration: saline calibration and aortic flow calibration. The conductance catheter technique necessitates an evaluation of the parallel conductance (Vp) offset to account for the conductance measured not only from the blood pool within the ventricular cavity but also from the surrounding structures. This assessment can be accomplished through the administration of a hypertonic saline bolus infusion. Aortic flow calibration enables the direct measurement of the aortic flow, which, in turn, permits the determination of the absolute stroke volume. However, it should be noted that this calibration provides only the absolute stroke volume and not the absolute ventricular volume. To obtain the absolute ventricular volume, both saline calibration and aortic calibration must be performed.

There are some limitations to this method. First, a transapical approach was employed when introducing the conductance catheter. To access the LV apex, the pericardium needs to be removed. This could affect the diastolic parameters, especially the Pediatric. Second, some blood might be lost during the long procedure time, which could also affect the cardiac functional parameters, but these issues can be avoided by becoming more proficient in the procedures. It is worth noting that the HFpEF model utilized in this protocol does not completely replicate human HFpEF, which is a syndrome with several phenotypes depending on the associated comorbidities, such as obesity, diabetes mellitus, hypertension, atrial fibrillation, or multiple organ failure. There is no available mice model that mimics all of these comorbidities. The double-hit HFpEF mice model, however, is most relevant to human HFpEF with metabolic comorbidities¹⁰. The genetic background of the mice could affect the diastolic function. While C57BL/6J mice have been reported to show differential responses to cardiovascular stress and a potentially milder disease phenotype compared to C57BL/6N mice, this protocol has detected diastolic impairment in the two-hit model even on the C57BL/6J background⁵, which might be difficult with other modalities usually employed in mice.

This manuscriptaims to provide guidance for performing the pacing-associated PV loop procedures effectively, which can be helpful in assessing HR-associated cardiac function and advancing research on heart failure.

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