Cardiac Response to β-Adrenergic Stimulation Determined by Pressure-Volume Loop Analysis

Summary

Here we describe a cardiac pressure-volume loop analysis under increasing doses of intravenously infused isoproterenol to determine the intrinsic cardiac function and the β-adrenergic reserve in mice. We use a modified open-chest approach for the pressure-volume loop measurements, in which we include ventilation with positive end-expiratory pressure.

Abstract

Determination of the cardiac function is a robust endpoint analysis in animal models of cardiovascular diseases in order to characterize effects of specific treatments on the heart. Due to the feasibility of genetic manipulations the mouse has become the most common mammalian animal model to study cardiac function and to search for new potential therapeutic targets. Here we describe a protocol to determine cardiac function in vivo using pressure-volume loop measurements and analysis during basal conditions and under β-adrenergic stimulation by intravenous infusion of increasing concentrations of isoproterenol. We provide a refined protocol including ventilation support taking into account the positive end-expiratory pressure to ameliorate negative effects during open-chest measurements, and potent analgesia (Buprenorphine) to avoid uncontrollable myocardial stress evoked by pain during the procedure. All together the detailed description of the procedure and discussion about possible pitfalls enables highly standardized and reproducible pressure-volume loop analysis, reducing the exclusion of animals from the experimental cohort by preventing possible methodological bias.

Introduction

Cardiovascular diseases typically affect cardiac function. This issue points out the importance in assessing in vivo detailed cardiac function in animal disease models. Animal experimentation is surrounded by a frame of the three Rs (3Rs) guiding principles (Reduce/Refine/Replace). In case of understanding complex pathologies involving systemic responses (i.e., cardiovascular diseases) at the current developmental level, the main option is to refine the available methods. Refining will also lead to a reduction of the required animal numbers due to less variability, which improves the power of the analysis and conclusions. In addition, combination of cardiac contractility measurements with animal models of heart disease including those induced by neurohumoral stimulation or by pressure overload like aortic banding, which mimics for example altered catecholamine/β-adrenergic levels^1,2,3,4, provides a powerful method for pre-clinical studies. Taking into account that the catheter-based method remains the most widely used approach for in depth assessment of cardiac contractility⁵, we aimed to present here a refined measurement of in vivo cardiac function in mice by pressure-volume loop (PVL) measurements during β-adrenergic stimulation based on previous experience including the evaluation of specific parameters of this approach^6,7.

To determine cardiac hemodynamic parameters approaches that include imaging or catheter-based techniques are available. Both options are accompanied by advantages and disadvantages that carefully need to be considered for the respective scientific question. Imaging approaches include echocardiography and magnetic resonance imaging (MRI); both have been successfully used in mice. Echocardiographic measurements involve high initial costs from a high-speed probe required for the high heart rate of the mice; it is a relatively straightforward non-invasive approach, but it is variable among operators who ideally should be experienced recognizing and visualizing cardiac structures. In addition, no pressure measurements can be performed directly and calculations are obtained from combination of size magnitudes and flow measurements. On the other hand, it has the advantage that several measurements can be performed on the same animal and cardiac function can be monitored for example during disease progression. Regarding the volume measurement, the MRI is the gold standard procedure, but similar to echocardiography, no direct pressure measurements are possible and only preload dependent parameters can be obtained⁸. Limiting factors are also the availability, analysis effort and operating costs. Here catheter-based methods to measure cardiac function are a good alternative that additionally allow for the direct monitoring of intracardiac pressure and the determination of load-independent contractility parameters like preload recruitable stroke work (PRSW)⁹. However, ventricular volumes measured by a pressure-conductance catheter (through conductivity determination) are smaller than those from the MRI but group differences are maintained in the same range¹⁰. In order to determine reliable volume values the corresponding calibration is required, which is a critical step during the PVL measurements. It combines ex vivo measurements of blood conductivity in volume-calibrated cuvettes (conversion of conductance to volume) with the in vivo analysis for the parallel conductance of the myocardium during the bolus injection of the hypertonic saline^11,12. Beyond that, the positioning of the catheter inside the ventricle and the correct orientation of the electrodes along the longitudinal axis of the ventricle are critical for the detection capability of the surrounding electrical field produced by them. Still with the reduced size of the mouse heart it is possible to avoid artifacts produced by changes in the intraventricular orientation of the catheter, even in dilated ventricles^5,10, but artifacts can evolve under β-adrenergic stimulation^6,13. Additional to the conductance methods the development of admittance based method appeared to avoid the calibration steps, but here the volume values are rather overestimated^14,15.

Since the mouse is one of the most important pre-clinical models in cardiovascular research and the β-adrenergic reserve of the heart is of central interest in cardiac physiology and pathology, we here present a refined protocol to determine in vivo cardiac function in mice by PVL measurements during β-adrenergic stimulation.

Protocol

All animal experiments were approved and performed according to the regulations of the Regional Council of Karlsruhe and the University of Heidelberg (AZ 35-9185.82/A-2/15, AZ 35-9185.82/A-18/15, AZ 35-9185.81/G131/15, AZ 35-9185.81/G121/17) conform to the guidelines from Directive 2010/63/EU of the European Parliament on the protection of animals used for scientific purposes. Data shown in this protocol are derived from wild type C57Bl6/N male mice (17 ± 1.4 weeks of age). Mice were maintained under specified pathogen-free conditions at the animal facility (IBF) of the Heidelberg Medical Faculty. Mice were housed in a 12-hour light-dark cycle, with a relative humidity between 56-60%, a 15-times air change per hour and room temperature of 22°C +/- 2°C. They were kept in conventional cages type II or type II long provided with animal bedding and tissue papers as enrichment. Standard autoclaved food and autoclaved water were available to consume ad libitum.

1. Preparation of instruments and drug solutions

1. Central venous catheter: Cut the micro tube (0.6 mm outer diameter) into ~20 cm long catheter tubes. Use forceps to pull one end of the tube onto the tip of a 23-gauge cannula. Cut the other end of the tubing diagonally to create a sharp tip that can pierce the femoral vein.
2. Endotracheal tube: For an intubation tube cut a 20-gauge venipuncture-cannula 3 cm in length to remove the syringe attachment.
   1. If the intubation tube does not fit the ventilator connection perfectly, wrap parafilm over the end of the tube where the ventilation device is connected. The connection must be stable and sealed by the thickening (Figure 1A). Shorten the metal guide pin of the 20-gauge venipuncture-cannula to 2.7 cm and use it as an intubation aid. Refined approaches for intubation including light fibers to facilitate visualization of the trachea are also well described, for example by Das and collaborators¹⁶.
3. Anesthetic mixture used for intubation: Mix 200 µL of heparin (1000 IU/mL) with 50 µL of 0.9% NaCl and 750 µL of 2 mg/mL etomidate from an oil-in-water emulsion based product. Use 7 µL/g body weight (BW) for each mouse (0.1 mg/kg BW Buprenorphine 10 mg/kg BW etomidate).
4. Muscle relaxant: Dissolve 100 mg of Pancuronium-bromide in 100 mL of 0.9% NaCl. Use 1.0 µL/g body weight (1 mg/kg BW) for each mouse.
5. Isoproterenol solutions: Dissolve 100 mg of isoproterenol in 100 mL of 0.9% NaCl (1 µg/µL). Prepare the following dilutions (Table 1) and transfer each in a 1 mL syringe.
   1. To obtain dilution 1, dilute the stock 1:1.8. To obtain dilution 2, dilute the stock 1:6. To obtain dilution 3, dilute dilution 1 into 1:10. Finally, obtain dilution 4 by a 1:10 dilution of dilution 2.
6. 15% Hypertonic NaCl (w/v): Dissolve 1.5 g of 0.9% NaCl in 10 mL of double distilled H₂O. Filter the solution with a 0.45 µm pore syringe filter.
7. Preparation of 12.5% albumin solution (w/v): Dissolve 1.25 g of bovine serum albumin in 10 mL of 0.9% NaCl. Incubate the solution at 37 °C for 30 min. Cool down to room temperature and filter the solution with a 0.45 µm pore syringe filter.
8. Preparation of the setup: First switch on the heating plate and set it to 39-40 °C. Place a syringe filled with saline on the heating pad and transfer the pressure-volume loop (PVL) catheter into the syringe. Pre-incubate the catheter for at least 30 min before use for stabilization. The setup we use consist of a 1.4-F pressure-conductance catheter, a control unit and the corresponding software, and it is graphically described on Figure 1B and provider references are listed in the Table of Materials.

2. Anesthesia

1. Inject buprenorphine (0.1 mg/kg BW intraperitoneally) 30 min before intubation.
2. Place the mouse into an acrylic glass-chamber pre-saturated with 2.5% isoflurane and pre-warmed with a heating pad placed on the base of the chamber.
3. As soon as the mouse sleeps (lack of reflex), inject the anesthetic mixture (7 mL/kg BW) containing 10 mg/kg etomidate and heparin (1,200 IU/kg BW) intraperitoneally.

3. Ventilation

1. Transfer the animal to the intubation platform (Figure 1C) 3-4 minutes after the anesthetic injection. The mouse hangs from the teeth with the dorsal view facing the operator.
2. Gently lift the tongue with forceps. To identify the glottis, lift the mouse's lower jaw slightly with second forceps.
3. Carefully insert the endotracheal tube (Figure 1A) into the trachea and remove the guide rod.
4. Transfer the animal onto the heating plate, place it on the back and connect the intubation tube to the small animal respirator.
5. Adjust respiratory rate to 53.5 x (Body weight in grams)^-0.26 [min^-1], as described by others¹², and tidal volumes to peak inspiratory pressures of 11 ± 1 cmH₂O. Establish a PEEP of 2 cmH₂O.
6. Fix carefully the extremities of the mouse on the heating plate with adhesive strips and apply eye ointment on both eyes to prevent dryness.
7. Insert a rectal temperature probe and maintain core body temperature at 37 ± 0.2 °C.
8. Install a 1-lead ECG and monitor the heart rate on-line as an indicator for anesthesia depth and stability.
9. Upon absence of interdigital reflexes, inject 1 mg/kg BW of the muscle relaxant pancuronium-bromide intraperitoneally. This prevents respiratory artifacts during PVL measurements.

4. Surgery

1. General recommendations
   1. During surgery, ventilate with ~1.5-2% isoflurane vaporized with O₂. The isoflurane concentration can also depend on variables like mouse strain, gender, age and weight of the animals, but it needs to be individually and experimentally determined and the values here are reference for the C57BL6/N mouse strain. Importantly, the ventilator is connected to an extraction system to prevent the operator from inhaling isoflurane.
   2. Use a magnification between 1.5-4x from the stereo microscope for surgical procedures.
      NOTE: Refer to institutional/local guidance on preparation of the animal for non-survival surgeries.
2. Femoral cannulation
   1. Rinse the hindlimb with 70% ethanol, incise the left inguinal region and expose the left femoral vein.
   2. Blast the epigastric artery and vein with a cautery.
   3. Ligate the femoral vein with a suture placed distal to the catheter access.
   4. Pass a suture underneath the femoral vein and prepare a knot cranial of puncture site. Puncture the femoral vein with the prepared micro tube (see step 1.1) attached to a 1 mL syringe.
   5. Tie down the knot to fix the tube inside the vessel.
   6. Counteract fluid loss by the infusion of 0.9% NaCl supplemented with 12.5% albumin at an infusion rate of 15 µL/min with an automatic syringe pump. Additionally, keep exposed tissue humid using pre-warmed 0.9% NaCl.
3. Thoracotomy
   1. Rinse the thorax with 70% ethanol.
   2. Incise the skin just beneath the xyphoid process and bluntly separate the pectoral muscles from the chest wall with forceps or a cautery.
   3. Lift the xyphoid process with forceps, and then cut through the chest wall moving laterally on both sides with a cautery until the diaphragm is fully visible from beneath.
   4. Incise the diaphragm from beneath and expose the cardiac apex. Then carefully remove the pericardium with forceps.
   5. Perform a limited costotomy on the left side as previously described⁶.
   6. Pass a suture beneath the inferior caval vein to perform preload reduction during later stages.
   7. Gently puncture the cardiac apex with a 25-gauge cannula (maximal 4 mm). Remove the cannula and insert the PV catheter until all electrodes are within the ventricle.
   8. Adjust the position of the catheter by gentle movements and turns until rectangular shaped loops are obtained (Figure 2A).
   9. Keep always all exposed tissue humid using pre-warmed 0.9% NaCl.

5. Measurements

1. General recommendations
   1. During measurements, ventilate with ~1.5-2% isoflurane vaporized with 100% O_2.
   2. Perform 2 baseline measurements as well as 2 vena cava occlusions on each step of the dose response protocol.
      NOTE: It is important that after the first and second vena cava occlusion, both pressure and volume values return to steady-state values as before the first occlusion. This observation is necessary in order to recognize a shift in catheter position due to serial reductions in intraventricular volume. If a shift in catheter position would be the case, especially volume values would be shifted.
2. Perform an on-line analysis of parameters (heart rate, stroke volume, dP/dt_max) and wait until steady-state cardiac function is obtained. For the expected parameter range with the here used setting in C57Bl6/N mice please refer to published results⁶.
3. Stop the respirator at end-expiratory position and record baseline parameters. After 3 to 5 seconds reduce cardiac preload by lifting the suture beneath the inferior caval vein with forceps in order to obtain preload independent parameters (Figure 2B). Turn the ventilator on. Wait at least 30 seconds for the second occlusion until hemodynamic parameters are stabilized.
4. After obtaining the measurements under basal conditions proceed to the dose-response of isoproterenol by switching to the prepared syringes. Here the infusion rate stays unchanged in order to avoid modifications of the cardiac preload. Take care not to infuse air bubbles when changing the syringe.
   1. Wait at least 2 minutes until new steady-state cardiac function is obtained than again stop the respirator at end-expiratory position and record baseline parameters. After 3 to 5 seconds reduce cardiac preload by lifting the suture beneath the inferior caval vein in order to obtain preload independent parameters.
   2. Wait at least 30 seconds for the second occlusion. Afterwards switching to the prepared syringe with the next isoproterenol concentration and repeat the recordings of baseline and preload independent parameters.
      NOTE: Artifacts like the end-systolic pressure-spike (ESPS, Figure 2C) can occur during the increase in the dosage of isoproterenol, which results from catheter entrapment. Artefacts that occur before the start of basal parameters can be easily corrected via re-positioning of the catheter.

6. Calibration

NOTE: Calibration procedures may vary depending on the PVL system used.

1. Parallel-conductance calibration
   1. Connect a syringe containing a 15% NaCl solution to the femoral cannula after the last measurement from the isoproterenol dose-response. Carefully infuse 5 µL of the hypertonic solution remaining in the tube until PVL slightly shift to the right during on-line visualization. Then wait until the loops come back to steady-state.
   2. Stop the respirator at end-expiration and inject one bolus of 10 µL of 15% NaCl within 2 to 3 seconds. Check if PVL largely broaden and are shifted to the right during on-line visualization.
2. Conductance-to-volume calibration
   1. Wait 5 min, no less, so that the hypertonic saline bolus is completely diluted. Afterwards remove the catheter and draw at least 600 μL blood from the left ventricle of the beating heart using a 1 mL syringe and a 21-gauge cannula. At this time point the animal is euthanized under deep anesthesia and analgesia by massive bleeding, by stopping the ventilation and removal of the heart.
   2. Transfer the blood into the pre-warmed (in a water bath at 37 °C) calibration cuvette with cylinders of known volume. Place the PV catheter centrally in each cylinder and record the conductance. By calculating a standard curve for each animal, the conductance units can be converted into absolute volume values.

7. Analysis

1. After successful PVL measurements under basal conditions and isoproterenol stimulation, visualize, digitalize, calculate and extract parameters characterizing cardiac function (like PRSW, dP/dt, end-diastolic pressure and volume, end-systolic pressure and volume, relaxation constant Tau, among others) using an appropriate PVL analysis software. Further statistical analysis and graphical representations can be performed with standard analysis software.
2. Analysis of preload independent parameters
   NOTE: For this step it is crucial to standardize the procedure.
   1. Select the first 5-6 PVLs showing decreasing preload throughout all measurements for the analysis of preload independent parameters (Figure 2D). A constant number of PVLs selected for analysis during preload reduction will decrease the variability among measurements of the obtained parameters.
   2. Calculate the mean value of the two measurements on each step of the protocol.

Results

The pressure volume-loop (PVL) measurement is a powerful tool to analyze cardiac pharmacodynamics of drugs and to investigate the cardiac phenotype of genetically modified mouse models under normal and pathological conditions. The protocol allows the assessment of cardiac β-adrenergic reserve in the adult mouse model. Here we describe an open-chest method under isoflurane anesthesia combined with buprenorphine (analgesic) and pancuronium (muscle relaxant), which focuses on the cardiac response to β-adrenergic stimulation...

Discussion

Here, we provide a protocol to analyze the in vivo cardiac function in mice under increasing β-adrenergic stimulation. The procedure can be used to address both, baseline parameters of cardiac function and the adrenergic reserve (e.g., inotropy and chronotropy) in genetically modified mice or upon interventions. The most prominent advantage of pressure-volume loop (PVL) measurements as compared to other means of determining cardiac function is the analysis of intrinsic, load-independent cardiac function. All other m...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
1.4F SPR-839 catheter	Millar Instruments, USA	840-8111
1 ml syringes	Beckton Dickinson, USA	REF303172
Bio Amplifier	ADInstruments, USA	FE231
Bridge-Amplifier	ADInstruments, USA	FE221
Bovine Serum Albumin	Roth, Germany	8076.2
Buprenorphine hydrochloride	Bayer, Germany	4007221026402
Calibration cuvette	Millar, USA	910-1049
Differential pressure transducer MPX	Hugo Sachs Elektronik- Harvard Apparatus, Germany	Type 39912
Dumont Forceps #5/45	Fine Science tools Inc.	11251-35
Dumont Forceps #7B	Fine Science tools Inc.	11270-20
Graefe Forceps	Fine Science tools Inc.	11051-10
GraphPad Prism	GraphPad Software	Ver. 8.3.0
EcoLab-PE-Micotube	Smiths, USA	004/310/168-1
Etomidate Lipuro	Braun, Germany	2064006
Excel	Microsoft
Heparin	Ratiopharm, Germany	R26881
Hot plate and control unit	Labotec, Germany	Hot Plate 062
Isofluran	Baxter, Germany	HDG9623
Isofluran Vaporizer	Abbot	Vapor 19.3
Isoprenalinhydrochloride	Sigma-Aldrich, USA	I5627
Fine Bore Polythene tubing 0.61 mm OD, 0.28 mm ID	Smiths Medical International Ltd, UK	Ref. 800/100/100
MiniVent ventilator for mice	Hugo Sachs Elektronik- Harvard Apparatus, Germany	Type 845
MPVS Ultra PVL System	Millar Instruments, USA
NaCl	AppliChem, Germany	A3597
NaCl 0.9% isotonic	Braun, Germany	2350748
Pancuronium-bromide	Sigma-Aldrich, USA	BCBQ8230V
Perfusor 11 Plus	Harvard Apparatus	Nr. 70-2209
Powerlab 4/35 control unit	ADInstruments, USA	PL3504
Rechargeable cautery-Set	Faromed, Germany	09-605
Scissors	Fine Science tools Inc.	140094-11
Software LabChart 7 Pro	ADInstruments, USA	LabChart 7.3 Pro
Standard mouse food	LASvendi GmbH, Germany	Rod18
Stereo microscope	Zeiss, Germany	Stemi 508
Surgical suture 8/0	Suprama, Germany	Ch.B.03120X
Venipuncture-cannula Venflon Pro Safty 20-gauge	Beckton Dickinson, USA	393224
Vessel Cannulation Forceps	Fine Science tools Inc.	00574-11
Water bath	Thermo Fisher Scientific, USA
Syringe filter (Filtropur S 0.45)	Sarstedt, Germany	Ref. 83.1826