Measuring Pressure Volume Loops in the Mouse

Summary

This manuscript describes a detailed protocol for the collection of pressure-volume data from the mouse.

Abstract

Understanding the causes and progression of heart disease presents a significant challenge to the biomedical community. The genetic flexibility of the mouse provides great potential to explore cardiac function at the molecular level. The mouse's small size does present some challenges in regards to performing detailed cardiac phenotyping. Miniaturization and other advancements in technology have made many methods of cardiac assessment possible in the mouse. Of these, the simultaneous collection of pressure and volume data provides a detailed picture of cardiac function that is not available through any other modality. Here a detailed procedure for the collection of pressure-volume loop data is described. Included is a discussion of the principles underlying the measurements and the potential sources of error. Anesthetic management and surgical approaches are discussed in great detail as they are both critical to obtaining high quality hemodynamic measurements. The principles of hemodynamic protocol development and relevant aspects of data analysis are also addressed.

Introduction

Cardiovascular disease continues to be a significant cause of mortality and morbidity throughout the world ¹. Diseases of the heart present particularly difficult challenges in developing new therapies. Advances in genetics provide for the possibility to identify a multitude of potential genetic contributors to the development of heart disease. The integrative nature of the cardiovascular system requires that these genetic targets be validated in intact animal models. The genetic flexibility and low housing costs of the mouse have brought it to the forefront for the assessment of the physiological role of a given gene. The small size of the mouse presents some unique challenges for the assessment of cardiac function. There are several modalities that can provide information regarding cardiac function, but only the simultaneous measurement of ventricular pressure and volume allows pressure-volume (PV) loop analysis of ventricular function. PV loops allow cardiac function to be analyzed independent of its connection to the vasculature; an important factor in determining the functional role of a particular genetic element.

The assessment of pressure-volume loops has been used both experimentally and clinically for many years and extensive literature exists regarding the analysis of these data sets ^2,3. The adaptation of PV loop technology to the mouse has been an important advancement for the understanding of murine cardiac physiology ^4-6. Catheter based PV loop technologies couple a pressure transducer and the use of conductance to estimate ventricular volume. The ventricular volume is determined by examining changes in an electrical field generated by the catheter. This method models the ventricle as a cylinder, the height of which is defined by the distance between the electrodes on the catheter and the radius is calculated from conduction of an electrical field through the blood in the ventricle ^7-9. The conductance signal measured by the catheter has two components. The first is the conduction through the blood; this varies with the volume of the ventricle and constitutes the primary signal used to determine ventricular volume. The second component results from conduction through and along the wall of the ventricle. This is called parallel conductance and must be removed in order to determine the absolute ventricular volume. There are two commercially available systems for the collection of pressure-volume data in the research laboratory and the method used to calculate and remove the parallel conductance is the primary difference between them ^6,10,11. Conductance catheters require the injection of hypertonic saline for the calculation of parallel conductance. This injection transiently changes the conductivity of the blood in the ventricle, while the conductivity of the wall remains constant. From this data it is possible to determine the component of the conductance signal that originates from the blood and what comes from the ventricular wall. This approach assumes that parallel conductance does not vary during the cardiac cycle. The admittance method relies on phase changes in the electrical field to assess the contribution of the ventricular wall to the overall volume signal. This method relies on a variety of predetermined constants for the conductivity of the blood and myocardium to determine the final volume, but makes continuous measures of parallel conductance during the cardiac cycle. Both of these systems provide good estimates of left ventricular volume and the differences between them are not likely to be physiologically significant. The cylindrical model of the ventricle and other assumptions render these catheter-based approaches not as accurate as other modalities, but this data is provided on a beat-by-beat basis that is essential for the assessment of load independent measures of cardiac function.

The procedure outlined here is used in my laboratory and has provided data for a large number of studies examining the basic pathophysiological mechanisms of dystrophic cardiomyopathy ^12-18. The procedure outlined below is one of two that can be used to obtain PV loop data. While many of the principles are applicable for either approach, this protocol will focus on an open-chest apical approach; a closed chest protocol has been detailed elsewhere ^19,20. While the procedure will be described in detail, the important overarching principles are to expose the heart with minimal damage to either the heart or the lungs. Throughout the protocol it is important to remember that this is a non-survival procedure and that having a good exposure of the heart is critically important for the proper placement of the catheter.

Protocol

Before performing any of the procedures described in this protocol, obtain approval by the local institutional animal care and use committee.

1. Setting Up the Experimental Rig

Note: This procedure is performed on anesthetized animals and the quality of the data is proportional to the quality of the anesthetic support offered to the animal. This first section will detail the equipment and procedures necessary to provide anesthesia to the mouse while performing this protocol.

1. Select an anesthetic protocol. Inhalant anesthetics have many beneficial properties for performing PV-loop analysis, although some injectable protocols have been used as well. See the discussion for more information on choosing an anesthetic regime.
2. Secure compressed oxygen tanks to the surgical table or wall near the surgical site.
3. If using inhalant anesthetics, use a vaporizer to insure proper dosing. Calibrate vaporizers annually to ensure they are providing the appropriate dose of anesthetic gas. Connect vaporizers to a flow meter which allows control of the rate at which gas enters the anesthetic circuit. Set at 0.5 - 1.0 L/min.
4. Use a manifold to allow directed flow of anesthetic gas to 1) the induction chamber, 2) a mask, and 3) the ventilator. Scavenging anesthetic gas is critically important and should be performed by an active system that either vents into a fume hood (or other similar building infrastructure) or through a canister designed to remove anesthetic gases.
   Note: Check with local occupational health officials to ensure compliance with all local regulations.
5. Maintain core body temperature by using heating pads and/or warming lamps. Continuously monitor body temperature with a rectal thermometer. This will allow for proactive increases or decreases in heating to ensure a physiological body temperature (≈ 37 °C) during the collection of hemodynamic data.
6. Provide fluid support to counteract blood loss and insensible volume loss.
   1. Prepare a 10% solution of albumin in 0.9% NaCl by taking 1 ml of 25% albumin and adding 1.5 ml of 0.9% NaCl in a syringe.
   2. Prepare a low residual volume intravascular catheter.
      1. Use pliers to crush the plastic hub of a 0.5 inch 30 gauge needle. Using needle holders, grasp the needle and remove the hub. Scrape the remaining adhesive from the needle using a hemostat. Insert the blunt end of the needle into a length of microbore tubing. Use less than 20 inches of length for the tubing.
   3. Use a syringe pump to allow accurate volumes to be delivered.
7. Ensure proper ventilation for the collection of high quality PV data. There are a variety of mouse ventilators available for purchase. Pressure-controlled ventilators provide the closed environment required for inhalant anesthetic gases and provide better control of the ventilation during the procedure.
   1. Ensure that inspiratory pressures are limited to <15 cm H₂O to prevent barotrauma. Set the ventilator to deliver the inspiratory pulse during 35% of the respiratory cycle. The use of positive end-expiratory pressure (PEEP) at a level of 4 - 5 cm H₂O will greatly improve the ventilation of the mouse, by preventing atelectasis of the lung and supporting gas exchange.
   2. Limit the dead space distal to the Y-joint in the ventilatory circuit. This is critical because the mouse's tidal volume is very small and any dead space subtracts from the delivery of fresh inspired air.
   3. Create a mouse sized endotracheal (ET) tube by cutting the tip off of a 20 gauge indwelling intravascular catheter. This provides a tapered point for easier insertion. Place the cut end into the Y-joint of the anesthetic circuit.
   4. Use flexible tubing in the respiratory circuit. Any structural memory in the tubing will create external forces that have the potential to pull the endotracheal tube out of the mouse's airway.
   5. Let the mouse determine the respiratory rate by using the lowest rate that suppresses the endogenous respiratory drive. Start at a relatively slow respiratory rate of around 60 breaths per minute.
      Note: With appropriate ventilation the mouse should make very little effort to breathe. However, if ventilation is inadequate, the buildup of CO₂ in the blood will initiate respiratory effort by the mouse. If this is observed, increasing the respiratory rate is a straight-forward way to increase alveolar ventilation. It is often necessary to increase respiratory rate in response to elevations in cardiac workload associated with beta-adrenergic receptor stimulation.
   6. Once the respiratory circuit is prepared, pressure test the system by plugging the tip of the endotracheal (ET) tube with a finger. Ensure an airway pressure of ≈10 cmH₂O. This test should be performed prior to each procedure.

2. Surgical Approach

1. Use small forceps and scissors and other equipment from Table 1. All instruments are quite small to allow for easy use in the magnified surgical field. Use a surgical stereomicroscope to provide adequate magnification for several aspects of the surgical procedure.
2. Use a cautery to maximize hemostasis during the procedure.
   Note: There are a couple of broad classes of cautery. Thermocautery heats a thin metal element that will cut through muscle tissue and stop bleeding. These systems are initially relatively inexpensive; however it is important to note that the wire tips are both fragile and relatively costly. Electrocautery systems are more costly to purchase initially, but the tips are very sturdy and will not need to be replaced.
3. Induction and Surgical Preparation
   1. Obtain the body weight of the mouse.
   2. Place the mouse into an induction chamber that is filled with 5% isoflurane.
      Note: The mouse cannot survive more than a few minutes in this environment. Only 45 - 60 sec are required for the mouse to lose its righting reflex (efforts to flip over when placed on its back or side).
   3. Once the righting reflex is lost, reduce the isoflurane concentration to 2% and open the anesthetic gas to the mask.
   4. Quickly transfer the mouse to the operating table and place it in dorsal recumbency with its nose within the mask.
   5. If using electrocautery, use saline soaked gauze to electrically couple the mouse to the grounding pad of the electrocautery system.
   6. Secure the limbs with surgical tape. This tape provides adhesive properties even when wet.
   7. Insert a rectal thermoprobe for monitoring core body temperature. Secure with tape.
   8. Apply a depilatory to the neck and chest of the mouse. Wait 2 - 3 min for the depilatory to work, and then remove the fur from these areas using a cotton-tipped applicator and/or laboratory wipes.
      Note: Draping of the surgical field is not required, as this is a non-survival procedure, but may be desired to limit the surgeon's exposure to the cautery grounding pad, if used.
   9. Once the mouse is prepped for surgery, assess the surgical plane of the mouse by performing a toe-pinch. Once assured of an appropriate anesthetic depth the mouse is ready for the first incision.
4. Obtain control of the airway through oral intubation or tracheotomy. Tracheotomy is a convenient approach that is relatively simple to perform.
   Note: Throughout this procedural description all directions and orientations will be relative to the surgeon.
   1. Make an incision at the level of the sternal notch extending from ≈ 5 mm right of the mid-line to ≈ 5 mm left of the midline.
   2. Make a second incision extending along the right edge of the first incision, extending rostrally to a level ≈ 2 mm caudal to the end of the mandible.
   3. Make a third incision extending from the rostral end of the second incision over to ≈ 5 mm left of mid-line. Retract the resulting skin flap to the left to expose the underlying tissues.
   4. Separate the parotid and submandibular salivary glands at the midline by blunt dissection. This will expose the underlying musculature overlying the trachea.
   5. Bluntly separate the right and left sternohyoideus muscles to expose the trachea.
   6. Pass a ≈10 cm piece of 3-0 silk suture under the trachea, taking care to not include the esophagus.
   7. Identify location for the tracheotomy: just caudal to the larynx there is a gap before the first tracheal ring, this is an ideal location to perform the tracheotomy.
   8. Adjust the manifold to provide anesthetic gas to the ventilator and turn on the ventilator.
   9. Check for leaks by plugging the tip of the endotracheal (ET) tube with a finger. Ensure an airway pressure of ≈10 cmH2O.
   10. Using a 20 gauge needle as a scalpel, incise the trachea. Make the incision relatively wide, as the ET tube will fill much of the tracheal lumen.
   11. Moving quickly, remove the mask and carefully insert the ET tube into the trachea. Do not force it as the tissues are very fragile and breaking through the wall of the trachea can result in a pneumothorax.
       Note: Immediately upon insertion, chest excursions should become evident.
   12. Secure the ventilatory circuit with tape to prevent the ET tube from being pulled out.
   13. Tie a single overhand knot in the 3-0 suture to form a seal around the ET tube.
       Note: At this point chest excursions should be clearly evident. If not it is usually the positioning of the ET tube within the trachea that is the problem. Pull the ET tube back and try to reposition it, focusing on the orientation of the trachea as a guide.
5. Preparation of the Jugular Vein Site
   1. Retract the left salivary glands rostral-laterally exposing the external jugular vein.
   2. Bisect the thin muscle (sternomastoideus) covering the vein with blunt dissection. This will expose the outer surface of the jugular vein.
   3. Carefully clear off any major pieces of tissue, although caution must be used as the walls of the vein are very thin. Once this task is complete, cover the jugular vein with the salivary glands to preserve it for cannulation later.
6. Thoracotomy; entering the pleural space without damaging the heart or lungs
   1. Remove much of the skin covering the chest extending the right edge of the original skin incision down to the level of the xiphoid process, then across the midline to ≈ 1.5 cm left of the midline.
      Note: Care should be taken when cutting through the external mammary vessels, which can be a significant source of bleeding. Cauterizing these vessels before cutting them will largely prevent this bleeding.
   2. Using blunt dissection, retract the skin flap laterally to expose underlying musculature.
   3. Isolate the insertion of the pectoralis major on the right side near the caudal aspect of the sternum using vessel dilating forceps. Cauterize and cut the muscle.
   4. Cut through the pectoralis major along its attachment to the sternum. Cauterize the cut edges to ensure hemostasis.
   5. Next undermine the latissimus dorsi on the same side, which is a large sheet of muscle covering the lateral aspect of the mouse. Cauterize and cut this muscle and then retract the cut end cranially. This may require some blunt dissection.
      Note: The ribs are now clearly evident and the heart may also be visible in some mouse strains. In most mice, entering the chest in the caudal half of the fourth intercostal space will provide good access to the heart. The fourth intercostal space is the second most caudal space.
   6. To enter the chest, use a pair of sharp forceps to carefully dissect down through the intercostal muscle layers.
   7. Once the pleural space has been opened, carefully insert the blunt-tipped vessel dilators. Using the vessel dilators to provide gentle upward force on the chest wall, use the blunt end spring scissors to carefully incise the remainder of the intercostal muscles.
   8. First cut laterally, being careful not to cut the lung lobe underneath. Next extend the incision medially, but stay 3 - 4 mm lateral of the midline to avoid the internal mammary artery.
      Note: The internal mammary vessels run parallel to the sternum and can result in significant blood loss if cut accidentally.
   9. Carefully cauterize the cut edge of the intercostal muscles, using a small cotton tipped applicator to roll the tissue upward to allow cautery contact with cut tissue without coming in contact with the lungs or the heart.
   10. Place a saline soaked small cotton tipped applicator through the incision pointing toward the midline. Provide a gentle upward traction to pull the chest wall away from the underlying structures. Begin cauterizing the chest wall at the medial edge of the incision and ending ≈ 1 cm lateral of the midline on the left side.
   11. Advance the applicator to the left so that it is continuously under the cautery tip.
   12. Once the tissue is thoroughly cauterized, use scissors to carefully cut through the sternum. The apex of the heart should be clearly visible at this point.
   13. Using blunt dissection, disrupt the pericardium and identify the caudal vena cava.
   14. Check for evidence of any bleeding and cauterize it now. Once all bleeding has been addressed, carefully remove any draping, the cautery grounding pad, and the saline soaked gauze.
7. Placing the Jugular Vein Catheter
   1. Connect the catheter made in step 1.6.2 to the syringe containing 10% albumin by carefully slipping the tubing over a 30 gauge needle.
   2. Begin infusing the albumin through the catheter.
   3. Orient the catheter such that the needle lies upon the jugular vein on its own, with the needle bevel up. If necessary, use a needle holder to rotate the needle in the tubing to move the bevel to the proper orientation.
   4. When the catheter is fully flushed stop the infusion.
   5. Grasp the catheter needle with a forceps. With the other hand, use a tissue forceps to retract the salivary gland to allow visualization of the jugular vein. Apply gentle traction to the tissues surrounding the distal jugular vein creating tension on the vessel wall. Using a shallow angle of approach, carefully insert the needle into the vein. Advance the tip of the needle 3 - 4 mm into the vessel.
   6. Before releasing the catheter needle secure the catheter tubing with a piece of tape, this will limit any movement of the needle once released.
   7. Release the needle and gently pull back on the syringe plunger to confirm that the catheter is in the lumen of the vessel, by visualizing blood in the line.
   8. Once positioned properly secure the catheter with surgical glue to attach the needle to the underlying salivary glands.
   9. Calculate the total volume to be infused. If there was not significant blood loss a volume of 5 µl/g body weight will be sufficient. If there was significant blood loss infusing 6 - 6.5 µl/g may be required. Set the flow rate such that the entire infusion will be complete in 10 - 15 min.
8. PV Catheter Placement in the Left Ventricle
   1. During the surgical procedure described above, place the PV catheter in a syringe containing a saline solution to allow it to equilibrate.
   2. Just prior to placement, move the syringe and catheter next to the mouse. With the catheter tip at roughly the same height as the heart, zero the pressure reading.
   3. Using a saline soaked small cotton tipped applicator, maneuver the heart to allow visualization of the apex.
   4. Using a 25 gauge needle, make a stab incision as close to the center of the apex as possible.
   5. Following the removal of the needle, quickly insert the catheter through the incision. It does not take much force to insert the catheter, so show restraint when advancing the catheter into the ventricle. Occasionally it is necessary to perform an additional stab incision. If this is necessary, try to perform the subsequent incision near the initial location to minimize damage to the heart.
      Note: Once the catheter is advanced into the ventricle the final placement is critically important. The ventricular pressure tracing will be evident by a low diastolic pressure (<10 mmHg) and a high systolic pressure (>80 mmHg at this point). Ideally, the catheter will be centered in the ventricle with the outer electrodes just within the ventricle. Fine adjustments in catheter position can be performed by observing the PV-loop data, looking for a box-shaped tracing with ≈ 90° angles between the sides.

3. Procedural Details

Note: Once the catheter is in place a brief stabilization period (10 - 15 min) is necessary to allow the animal to recover from some of the acute surgical stress and to allow time for the infusion of fluids. After this stabilization period the actual protocol can begin.

1. Once the PV catheter is placed and surgical manipulation has ceased, turn down the isoflurane to ≈ 1% as the need for a deep surgical plane of anesthesia is lessened.
   1. During this period, carefully monitor the mouse to ensure that an appropriate level of anesthesia is maintained. Carefully assess any movement; movement of the respiratory muscles suggests a level of hypoventilation and can be addressed by increasing the respiratory rate of the ventilator. Movement of limbs or twitching of whiskers are signs that the mouse is getting too light and requires more anesthetic.
      Note: There are a wide variety of permutations of treatments that can be used in combination with this protocol. Many of these treatments will require infusion of drugs. It is essential to manage the dead-space volume effectively. Solution switches can be accomplished by sliding the catheter tubing from the needle of one syringe pump to the other. Doing this shortly before the end of the earlier infusion allows the catheter tubing to be loaded with the next drug. It is necessary to know the volume within the catheter tubing to determine the timing of the solution switch. The introduction of a small air bubble in the line allows the precise timing of the infusion switch to be determined; this bubble infused into the venous circulation is well tolerated.
2. Alter the loading conditions of the heart by lowering preload and increasing afterload.
   1. Reduce preload by blocking the venous return to the heart. In this preparation, visualize and occlude the caudal vena cava as it passes from the diaphragm to the heart. Perform this occlusion smoothly and relatively quickly, lasting no more than 2 - 3 sec. Increase left ventricular afterload transiently by performing a gentle abdominal compression lasting 1 - 2 sec.
   2. During these changes in loading of the heart, pause respirations to eliminate any artifact introduced by the ventilator.
3. Calibrate the volume signal using the conductance catheters. These procedures are not necessary with catheters using admittance technology.
   1. After the experimental protocol inject 5 - 10 µl of hypertonic saline (20% NaCl) to calculate the parallel conductance.
   2. Collect blood by removing the catheter and drawing blood from the left ventricle into a heparinized syringe. Place this blood into cuvettes of known volume and use the catheter to measure the conductance.
   3. Use the cuvette conductance measures to convert the conductance signal to volume and the parallel conductance is necessary to define the absolute volume measured by the catheter.
4. Once the protocol is complete, euthanize the mouse by removing the heart following transection of the vena cava and aortic attachments.

Results

By convention, volume is plotted on the X-axis and pressure on the Y-axis as in Figure 1. The pressure-volume loops resulting from plotting pressure against volume should resemble a rectangle, the vertical edges representing isovolumic changes in pressure (i.e., when both mitral and aortic valves are closed). The bottom horizontal represents ventricular filling through the mitral valve and the upper horizontal portion represents ventricular emptying through the a...

Discussion

There are three critical steps in this procedure: 1) the placement of the endotracheal tube and appropriate ventilation, 2) placement of the jugular IV catheter, and 3) the proper placement of the PV catheter in the left ventricle. Determining the appropriate respiratory rate is an important part of providing ventilatory support. Conscious mice generally maintain alveolar ventilation with rapid shallow breaths. In general, ventilated mice will have much larger tidal volumes. Thus a slower respiratory rate is required. Th...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Dumont 5/45 (2)	Fine Science Tools	11251-33
Vessel Dilating Forceps	Fine Science Tools	18153-11
Castroviejo Micro Dissecting Spring Scissor	Roboz Instruments	RS-5668
Octogon Forceps - Serrated/Curved	Fine Science Tools	11041-08
Octogon Forceps - Serrated/Straight	Fine Science Tools	11040-08
Dissector Scissors- Heavy Blade	Fine Science Tools	14082-09
Transpore Surgical Tape	3M	1527-1
3-0 Silk Suture	Fine Science Tools	18020-30
TOPO Ventilator	Kent Scientific	TOPO
Martin ME 102 Electrosurgical Unit	Harvard Apparatus	PY2 72-2484
Syringe Pump	Lucca Technologies	GenieTouch
Stereomicroscope with boom stand	Nikon	SMZ-800N
Thermocouple Thermometer	Cole Parmer	EW-91100-40
T/Pump Warm Water Recirculator	Kent Scientific	TP-700
ADVantage Pressure-Volume System	Transonic	ADV500
Data Acquision and Analysis	DSI	Ponemah ACQ-16